Handbook of Chitin and Chitosan: Volume 1: Preparation and Properties [1, 1 ed.] 0128179708, 9780128179703

Table of contents :
Cover
Handbook of Chitin and Chitosan
Copyright
Contents
List of Contributors
1 Chitin and chitosan: origin, properties, and applications
1.1 Introduction
1.2 Chitin and chitosan
1.2.1 Sources of chitin
1.3 Extraction of chitin
1.3.1 Chemical extraction
1.3.1.1 Demineralization process
1.3.1.2 Deproteinization process
1.3.1.3 Decolorization
1.3.2 Biological extraction
1.3.2.1 Enzymatic demineralization
1.3.2.2 Enzymatic deproteinization
1.3.2.3 Fermentation
1.4 Chitosan preparation methods
1.4.1 Chemical and biological deacetylation of chitin
1.5 Physicochemical properties
1.5.1 Molecular weight
1.5.2 Viscosity
1.5.3 Solubility
1.5.4 Water-binding capacity and fat-binding capacity
1.6 Characterization of chitin and chitosan
1.6.1 Fourier-transform infrared spectroscopy (FT-IR) analysis
1.6.2 X-Ray diffraction analysis
1.6.3 13C Nuclear magnetic resonance analysis
1.6.4 Thermogravimetric analysis for chitin and chitosan
1.6.5 Scanning electron microscope analysis
1.7 Application of chitin and chitosan
1.7.1 Biomedical application
1.7.2 Wastewater treatment
1.7.3 Fuel cell
1.7.4 Packaging of food
1.7.5 Textile industries
1.7.6 Bioplastics
1.7.7 Nanocomposite
References
2 Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications
2.1 Introduction
2.2 Chitin and chitosan: chemistry and solubility
2.2.1 Chitin and chitosan chemistry
2.2.2 Chitin and chitosan extraction
2.2.3 Chitin and chitosan solubility
2.3 Chitin and chitosan: fiber formation
2.3.1 Electrospinning process
2.3.2 Factors affecting the electrospinning process
2.3.2.1 Electrospinning parameters
2.3.2.2 Solution parameters
2.3.2.3 Ambient parameters
2.3.3 Characterization of chitin and chitosan fibers
2.3.4 Applications of chitin and chitosan fibers
2.3.4.1 Chitin and chitosan fibers for biomedical applications
2.3.4.2 Chitin and chitosan fibers for dye removal and wastewaters treatment
2.3.4.3 Chitin and chitosan fibers for cosmetic applications
2.3.4.4 Other applications for chitin and chitosan fibers, in different fields
2.4 Conclusions
References
3 PEGylated chitin and chitosan derivatives
3.1 Introduction
3.2 Chitin and chitosan
3.3 PEGylation and PEGylated chitin/chitosan derivatives
3.4 Fabrication of PEGylated chitosan derivatives
3.4.1 Amino group substitution
3.4.2 O-substitution
3.4.3 Miscellaneous approaches
3.4.4 Solubilization of chitosan prior to derivatization
3.4.5 PEGylated cross-linked chitosan
3.5 Characterization of PEGylated chitosan and chitin derivatives
3.5.1 Structural analysis
3.5.1.1 Fourier-transform infrared spectroscopy analysis
3.5.1.2 Nuclear magnetic resonance analysis
3.5.1.3 Gel permeation chromatography or size exclusion chromatography
3.5.1.4 Thermogravimetric analysis
3.5.1.5 Differential scanning calorimetry
3.5.1.6 Wide-angle X-ray diffraction analysis
3.5.2 Determination of the degree of substitution
3.5.3 Ductility
3.5.4 Solubility
3.5.5 Cytocompatibility
3.5.6 Aggregation
3.6 Applications of PEGylated derivatives of chitosan
3.6.1 Medical applications
3.6.2 Thermoresponsive PEGylated chitosan hydrogels
3.6.3 Gene delivery of chitosan derivatives
3.6.4 Formation of nanofibres
3.7 Conclusions
References
4 Solubility, chain characterization, and derivatives of chitin
4.1 Solubility of chitin
4.1.1 NaOH/urea aqueous solution
4.1.2 CaCl2/methanol solvent
4.1.3 N,N-dimethylacetamide/lithium chloride solvent
4.1.4 Ionic liquid
4.2 Chain characterization of chitin
4.2.1 The configuration of chitin
4.2.1.1 Fourier-transform infrared spectroscopy
4.2.1.2 X-ray powder diffraction spectroscopy
4.2.1.3 Solid 13C nuclear magnetic resonance spectroscopy
4.2.2 The length of chitin chain
4.3 Derivatives of chitin
4.3.1 The etherification of chitin
4.3.1.1 The carboxymethyl chitin
4.3.1.2 The quaternization of chitin
4.3.2 The graft of chitin
4.3.2.1 The graft of chitin powder
4.3.2.2 The graft of chitin nanofiber film
4.3.3 The O-acylation of chitin
4.3.3.1 Preparation of O-acylated chitin in heterogeneous system
4.3.3.1.1 Acid catalyst method
4.3.3.1.2 Activation of chitin
4.3.3.2 Preparation of O-acylated chitin in homogeneous system
4.3.3.2.1 Methanesulfonic acid system
4.3.3.2.2 Trifluoroacetic anhydride system
4.3.3.2.3 The lithium chloride/dimethylacetamide system
4.3.3.2.4 The ionic liquid system
Acknowledgment
References
5 Solubility, degree of acetylation, and distribution of acetyl groups in chitosan
5.1 Introduction
5.2 Chemistry and structure of chitosan
5.3 Acetylation of chitosan
5.3.1 Methods of deacetylation of chitin to chitosan
5.3.1.1 Chemical deacetylation
5.3.1.2 Enzymatic deacetylation
5.3.2 Degree of acetylation of chitosan
5.3.2.1 Measurement of degree of acetylation of chitosan
5.3.2.1.1 Elemental analysis
5.3.2.1.2 Titration methods
5.3.2.1.3 Acid hydrolysis/ high-performance liquid chromatography
5.3.2.1.4 Pyrolysis GC-MS
5.3.2.1.5 Infrared
5.3.2.1.6 Nuclear magnetic resonance spectroscopy
5.3.2.2 Distribution of acetyl groups
5.3.2.3 Relationship between acetylation and properties
5.4 Solubility of chitosan
5.4.1 Chitosan solubility: applications and requirements
5.4.2 Solubility of chitosan in solution
5.4.3 Modifications of chitosan for solubility enhancement
5.5 Conclusion
References
6 Chitin nanomaterials: preparation and surface modifications
6.1 Introduction
6.2 Structure and properties of chitin
6.3 Chitin-based nanomaterials
6.3.1 Chitin nanofiber
6.3.1.1 Pure chitin nanofiber
6.3.1.2 Chitin-based blended nanofiber
6.3.1.3 Modified chitin nanofiber
6.3.1.4 Nanofiber from chitin derivatives
6.3.2 Chitin nanowhisker
6.3.3 Chitin nanocomposite
6.3.4 Polymer/chitin bionanocomposite
6.3.5 Chitin nanogel
6.3.6 Crab chitin-based two-dimensional soft nanomaterials
6.4 Preparation of chitin-based nanomaterials
6.4.1 Electrospinning of chitin
6.4.2 Aqueous counter collision method
6.4.3 Self-assembly
6.4.4 Microcontact printing
6.4.5 Mechanical treatment
6.4.6 Ultrasonication
6.4.7 TEMPO-mediated oxidation
6.4.8 Extraction of chitin nanowhisker
6.4.9 Gelation method
6.4.10 Casting and evaporating technique
6.5 Surface modification of chitin
6.5.1 Chemical modification of chitin surface
6.5.1.1 Acetylation of chitin
6.5.1.2 Surface modification of chitin nanowhiskers
6.5.1.3 Oxidative modification
6.5.2 Hydrophobization of chitin surface
6.5.3 Hydrophilization of chitin surface
6.5.4 Physical modification of chitin surface
6.5.5 Ultrasound-assisted surface modification of chitin
6.5.6 Plasma treatment
6.6 Conclusions
References
7 Importance of electrospun chitosan-based nanoscale materials for seafood products safety
7.1 Optimization
7.1.1 Definition of chitosan molecular weight
7.1.2 Determination of concentrations
7.1.3 Solvent system
7.1.4 Preparation of electrospinning dope solutions
7.2 Determination of electrospinning parameters
7.2.1 Applied voltage
7.2.2 Adjustment proper distance
7.2.3 Flow rate
7.2.4 Environmental conditions
7.2.5 Selection of collector
7.3 Characterization of fabricated nanoscale material(s)
7.3.1 Definition of morphological characteristics of chitosan-based nanomaterial
7.3.2 Encapsulation efficiency
7.3.3 Controlled release property of chitosan-based nanomaterial
7.3.4 Thermal decomposition of chitosan-based nanomaterial
7.3.5 Zeta potential and size of chitosan-based nanomaterial
7.4 Use of electrospun nanomaterials for seafood products safety
7.4.1 Chitosan-based nanofiber coating
7.4.2 Bio/active material-loaded chitosan-based nanofiber coating
7.4.3 Chitosan nanoparticles
7.5 Conclusion
References
8 Alternative methods for chitin and chitosan preparation, characterization, and application
8.1 Introduction
8.2 Chitin production
8.2.1 Synthesis of chitin and enzymes involved in the main commercial sources
8.2.2 Raw materials: fungi and insects
8.2.2.1 Fungi
8.2.2.2 Insects
8.3 Current chitosan production
8.3.1 Alternative methods of production
8.3.1.1 Fermentations
8.3.1.2 Enzymic methods
8.3.1.3 Combined biological methods
8.3.1.4 Other methods
8.3.2 Quality control of chitin production
8.4 Conclusions
References
9 Current research on the blends of chitosan as new biomaterials
9.1 Introduction
9.2 Chitosan biomaterial
9.2.1 Preparation of chitosan
9.2.2 Properties of chitosan
9.2.2.1 Physical properties
9.2.2.2 Chemical properties
9.2.2.3 Biological properties
9.2.3 Applications of chitosan
9.2.3.1 In wastewater treatment
9.2.3.2 In food industry
9.2.3.3 In agriculture
9.3 Modification of chitosan
9.3.1 Polymer blending technique
9.3.2 Chitosan blends
9.4 Natural polymers blends with chitosan
9.4.1 Chitosan/starch
9.4.2 Chitosan/collagen
9.4.3 Chitosan/proteins
9.4.4 Chitosan/natural rubber latex
9.5 Chitosan blends with synthetic polymers
9.5.1 Chitosan/hydrophilic polymers
9.5.2 Chitosan/nylon
9.5.3 Chitosan/polyacrylamide
9.5.4 Chitosan/poly(lactic acid)
9.6 Chitosan-based hydrogels
9.7 Conclusions
Acknowledgment
References
10 Chitin and chitosan-based aerogels
10.1 Introduction
10.1.1 Classification of aerogels
10.1.2 Chitin and chitosan: sources and properties
10.2 Chitin and chitosan-based aerogels: preparation process
10.3 Characterization of chitin and chitosan-based aerogels
10.3.1 Morphological analysis/microscopic analysis
10.3.1.1 Scanning electron microscope
10.3.1.2 Transmission electron microscopy
10.3.2 Porosity
10.3.3 Thermal properties
10.3.3.1 Thermogravimetric analysis
10.3.3.2 Differential scanning calorimetry
10.3.4 Raman spectra
10.3.5 Tensile property of aerogels/mechanical property
10.3.6 Fourier transform infrared spectroscopy
10.3.7 X-ray diffraction
10.3.8 X-ray photoelectron spectroscopy
10.3.9 Nuclear magnetic resonance spectroscopy
10.3.10 Water contact angle
10.4 Future aspects of aerogel
10.5 Conclusions
Acknowledgments
References
11 Chitin, chitosan, marine to market
11.1 Introduction
11.2 Origin and sources of chitin and chitosan
11.3 Synthesis of chitin and chitosan
11.3.1 Synthesis of chitin
11.3.1.1 Synthesis of chitin by chemical method
11.3.1.1.1 Deproteinization
11.3.1.1.2 Demineralization
11.3.1.1.3 Decolorization
11.3.1.2 Synthesis of chitin by biological method
11.3.2 Synthesis of chitosan
11.3.3 Synthesis of derivatives of chitin and chitosan
11.4 Properties of chitin and chitosan
11.4.1 Physiochemical properties of chitin and chitosan
11.4.2 Biological properties of chitosan
11.5 Potential applications of chitin and chitosan
11.5.1 Biomedical application of chitosan
11.5.1.1 Wound dressing/wound healing
11.5.1.2 Burn treatment/artificial skin graft
11.5.1.3 Tissue engineering
11.5.1.4 Drug delivery
11.5.1.5 Ophthalmology
11.5.2 Industrial applications of chitosan
11.5.2.1 Water engineering
11.5.2.2 Agriculture
11.5.2.3 Food processing
11.5.2.4 Cosmetics industries
11.5.2.5 Photography
11.5.2.6 Chitosan gel for light-emitting device
11.5.2.7 Textile industry
11.5.2.8 Paper industry
11.6 Economic potential of chitin and chitosan
11.7 Conclusions
References
Further reading
12 Miscibility, properties, and biodegradability of chitin and chitosan
12.1 Introduction
12.2 Physicochemical properties of chitin and chitosan
12.2.1 Miscibility/solubility of chitin and chitosan
12.2.1.1 Dissolution by inorganic reagents
12.2.1.2 Dissolution by polar solvents and strong acids
12.2.1.3 Solubility in ionic liquids
12.2.1.4 Enhanced solubility of chitin and chitosan on their modification
12.2.2 Dissolution mechanism
12.2.2.1 Acid-catalyzed hydrolysis
12.2.2.2 Base-catalyzed hydrolysis
12.2.2.3 Cleavage/dissolution by ionic liquids
12.3 Biological properties of chitin and chitosan
12.3.1 Antioxidant property
12.3.2 Anticancer/antitumor property
12.3.3 Antimicrobial property
12.3.4 Antiinflammatory property
12.3.5 Neuroprotective property
12.4 Biodegradability of chitin and chitosan
12.4.1 factors affecting the biodegradability
12.5 Concluding remarks
References
13 Chitin and chitosan: current status and future opportunities
13.1 Introduction
13.2 Properties of chitin and chitosan
13.2.1 Structural properties
13.2.2 Biological properties
13.3 Chitin, chitosan, and their derivatives
13.3.1 Biosynthesis of chitin
13.3.2 Chitin and chitosan production
13.3.2.1 Chemical process
13.3.2.2 Biological process
13.4 Applications of chitin and chitosan
13.4.1 Industrial applications
13.4.1.1 Cosmetics
13.4.1.2 Water engineering
13.4.1.3 Paper and textile industry
13.4.1.4 Food industry
13.4.1.5 Other industrial applications
13.4.2 Biomedical applications of chitosan
13.4.2.1 Tissue engineering
13.4.2.2 Wound healing/wound dressing
13.4.2.3 Drug delivery
13.5 Conclusion and future perspectives
References
14 Fungal chitosan: prospects and challenges
14.1 Introduction
14.1.1 Chitin and chitosan
14.1.1.1 Chitin
14.1.1.2 Chitosan
14.1.1.3 Applications of chitosan
14.2 Current commercial production and its disadvantages
14.3 Green synthesis of chitosan
14.4 Fungal chitosan
14.4.1 Significance of fungal sources of chitosan
14.4.2 Production of fungal chitosan
14.4.2.1 Mucor rouxii
14.4.2.2 Rhizopus oryzae
14.4.2.3 Gongronella butleri
14.4.2.4 Other fungal strains
14.4.2.5 Use of industrial residues as culture medium for fungal chitosan production
14.4.2.6 Avenues and challenges of commercial production of fungal chitosan
14.5 Future prospects
14.6 Conclusion
14.7 Acknowledgments
References
Online resource
15 Preparation, properties, and application of low-molecular-weight chitosan
15.1 Introduction
15.2 Preparation of low-molecular-weight chitosan
15.2.1 Chemical methods
15.2.2 Physical methods
15.2.3 Biological methods
15.3 Properties of low-molecular-weight chitosan
15.3.1 Physicochemical properties
15.3.2 Biological activities
15.4 Applications of low-molecular-weight chitosan
15.5 Agriculture
15.5.1 Aquaculture
15.5.2 Food technology
15.6 Conclusions
References
Index
Back Cover

Citation preview

HANDBOOK OF CHITIN AND CHITOSAN

HANDBOOK OF CHITIN AND CHITOSAN PREPARATION AND PROPERTIES VOLUME 1 Edited by

SREERAG GOPI Center for Innovations and Technologies (CIT), ADSO Naturals Private Limited, Bangalore, India

SABU THOMAS Mahatma Gandhi University, Kottayam, India

ANITHA PIUS The Gandhigram Rural Institute (Deemed University), Dindigul, India

Elsevier Radarweg 29, PO Box 211, 1000 AE Amsterdam, Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States Copyright © 2020 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress ISBN: 978-0-12-817970-3 For Information on all Elsevier publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Susan Dennis Editorial Project Manager: Kelsey Connors Production Project Manager: Sujatha Thirugnana Sambandam Cover Designer: Christian J. Bilbow Typeset by MPS Limited, Chennai, India

Contents List of contributors

ix

1. Chitin and chitosan: origin, properties, and applications

1

SUNEETA KUMARI AND RUPAK KISHOR

1.1 Introduction 1.2 Chitin and chitosan 1.3 Extraction of chitin 1.4 Chitosan preparation methods 1.5 Physicochemical properties 1.6 Characterization of chitin and chitosan 1.7 Application of chitin and chitosan References

2 3 5 8 8 13 22 28

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

35

HAKIMA EL KNIDRI, ALI LAAJEB AND AHMED LAHSINI

2.1 Introduction 2.2 Chitin and chitosan: chemistry and solubility 2.3 Chitin and chitosan: fiber formation 2.4 Conclusions References

36 36 41 50 50

3. PEGylated chitin and chitosan derivatives

59

ADIB H. CHISTY, RIFAT A. MASUD, M. MEHEDI HASAN, M. NURUZZAMAN KHAN, ABUL K. MALLIK AND MOHAMMED MIZANUR RAHMAN

3.1 Introduction 3.2 Chitin and chitosan 3.3 PEGylation and PEGylated chitin/chitosan derivatives 3.4 Fabrication of PEGylated chitosan derivatives 3.5 Characterization of PEGylated chitosan and chitin derivatives 3.6 Applications of PEGylated derivatives of chitosan 3.7 Conclusions References

v

60 61 64 65 71 86 93 93

vi

Contents

4. Solubility, chain characterization, and derivatives of chitin

101

MI FENG, XINGMEI LU, DANFENG HOU AND SUOJIANG ZHANG

4.1 Solubility of chitin 4.2 Chain characterization of chitin 4.3 Derivatives of chitin Acknowledgment References

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

102 107 112 125 125

131

E.I. AKPAN, O.P. GBENEBOR, S.O. ADEOSUN AND ODILI CLETUS

5.1 Introduction 5.2 Chemistry and structure of chitosan 5.3 Acetylation of chitosan 5.4 Solubility of chitosan 5.5 Conclusion References

6. Chitin nanomaterials: preparation and surface modifications

132 132 134 145 153 154

165

ABUL K. MALLIK, MD. NURUS SAKIB, MD. SHAHARUZZAMAN, PAPIA HAQUE AND MOHAMMED MIZANUR RAHMAN

6.1 Introduction 6.2 Structure and properties of chitin 6.3 Chitin-based nanomaterials 6.4 Preparation of chitin-based nanomaterials 6.5 Surface modification of chitin 6.6 Conclusions References

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety ¨ ZOGUL AND MUSTAFA TAHSIN YILMAZ ZAFER CEYLAN, RACIYE MERAL, FATIH O

7.1 Optimization 7.2 Determination of electrospinning parameters 7.3 Characterization of fabricated nanoscale material(s) 7.4 Use of electrospun nanomaterials for seafood products safety 7.5 Conclusion References

166 168 169 178 185 189 189

195 196 202 207 212 216 217

Contents

8. Alternative methods for chitin and chitosan preparation, characterization, and application

vii 225

GEORGE M. HALL, CLAUDIA H. BARRERA AND KEIKO SHIRAI

8.1 Introduction 8.2 Chitin production 8.3 Current chitosan production 8.4 Conclusions References

9. Current research on the blends of chitosan as new biomaterials

226 226 231 240 241

247

A. RAJESWARI, SREERAG GOPI, E. JACKCINA STOBEL CHRISTY, K. JAYARAJ AND ANITHA PIUS

9.1 Introduction 9.2 Chitosan biomaterial 9.3 Modification of chitosan 9.4 Natural polymers blends with chitosan 9.5 Chitosan blends with synthetic polymers 9.6 Chitosan-based hydrogels 9.7 Conclusions Acknowledgment References

10. Chitin and chitosan-based aerogels

248 249 256 258 264 274 275 275 275

285

E. JACKCINA STOBEL CHRISTY, A. RAJESWARI, SREERAG GOPI AND ANITHA PIUS

10.1 Introduction 10.2 Chitin and chitosan-based aerogels: preparation process 10.3 Characterization of chitin and chitosan-based aerogels 10.4 Future aspects of aerogel 10.5 Conclusions Acknowledgments References

11. Chitin, chitosan, marine to market

286 292 296 324 327 328 328

335

G.M. OYATOGUN, T.A. ESAN, E.I. AKPAN, S.O. ADEOSUN, A.P.I. POPOOLA, B.I. IMASOGIE, W.O. SOBOYEJO, A.A. AFONJA, S.A. IBITOYE, V.D. ABERE, A.O. OYATOGUN, K.M. OLUWASEGUN, I.E. AKINWOLE AND K.J. AKINLUWADE

11.1 Introduction 11.2 Origin and sources of chitin and chitosan 11.3 Synthesis of chitin and chitosan 11.4 Properties of chitin and chitosan 11.5 Potential applications of chitin and chitosan 11.6 Economic potential of chitin and chitosan 11.7 Conclusions References Further reading

336 337 339 348 354 361 364 364 376

viii

Contents

12. Miscibility, properties, and biodegradability of chitin and chitosan

377

MUHAMMAD ARSHAD, MUHAMMAD ZUBAIR AND AMAN ULLAH

12.1 Introduction 12.2 Physicochemical properties of chitin and chitosan 12.3 Biological properties of chitin and chitosan 12.4 Biodegradability of chitin and chitosan 12.5 Concluding remarks References

13. Chitin and chitosan: current status and future opportunities

378 378 385 389 392 392

401

RUCHI MUTREJA, ABHIJEET THAKUR AND ARUN GOYAL

13.1 Introduction 13.2 Properties of chitin and chitosan 13.3 Chitin, chitosan, and their derivatives 13.4 Applications of chitin and chitosan 13.5 Conclusion and future perspectives References

14. Fungal chitosan: prospects and challenges

402 403 405 407 412 413

419

JOSEPH SEBASTIAN, TAREK ROUISSI AND SATINDER KAUR BRAR

14.1 Introduction 14.2 Current commercial production and its disadvantages 14.3 Green synthesis of chitosan 14.4 Fungal chitosan 14.5 Future prospects 14.6 Conclusion 14.7 Acknowledgments References Online resource

15. Preparation, properties, and application of low-molecular-weight chitosan

420 426 427 429 446 447 448 448 452

453

NGUYEN CONG MINH, NGUYEN VAN HOA AND TRANG SI TRUNG

15.1 Introduction 15.2 Preparation of low-molecular-weight chitosan 15.3 Properties of low-molecular-weight chitosan 15.4 Applications of low-molecular-weight chitosan 15.5 Agriculture 15.6 Conclusions References

Index

454 454 460 461 463 464 464

473

List of Contributors V.D. Abere Department of Mineral Processing, National Metallurgical Development Centre, Jos, Nigeria S.O. Adeosun Department of Metallurgical and Materials Engineering, University of Lagos, Lagos, Nigeria A.A. Afonja Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria K.J. Akinluwade Department of Research and Development, Prototype Engineering Development Institute (National Agency for Science and Engineering Infrastructure, NASENI), Ilesa, Nigeria I.E. Akinwole Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria E.I. Akpan Institute for Kaiserslautern, Germany

Composite

Materials,

Technical

University,

Muhammad Arshad Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, AB, Canada Claudia H. Barrera Biotechnology Department, Laboratory of Biopolymers and Pilot Plant of Bioprocessing of Agro-Industrial and Food By-Products, Autonomous Metropolitan University, Mexico City, Mexico Satinder Kaur Brar INRS-ETE, Universite´ du Que´bec, Que´bec, QC, Canada; Department of Civil Engineering, Lassonde School of Engineering, York University, Toronto, ON, Canada Zafer Ceylan Faculty of Fisheries, Department of Seafood Processing Technology, Van Yuzuncu Yil University, Van, Turkey Adib H. Chisty Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh Odili Cletus Materials and Metallurgical Engineering, University of Lagos, Lagos, Nigeria Hakima El Knidri Catalysis, Materials and Environment Laboratory, Higher School of Technology, Sidi Mohamed Ben Abdellah University, Fez, Morocco T.A. Esan Department of Restorative Dentistry, Obafemi Awolowo University, Ile-Ife, Nigeria Mi Feng CAS Key Laboratory of Green Process and Engineering, State Key Laboratory of Multiphase Complex Systems, Beijing Key Laboratory of Ionic Liquids Clean Process, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, P.R. China; School of Chemical and Engineering, University of Chinese Academy of Sciences, Beijing, P.R. China

ix

x

List of Contributors

O.P. Gbenebor Materials and Metallurgical Engineering, University of Lagos, Lagos, Nigeria Sreerag Gopi Department of Chemistry, The Gandhigram Rural Institute— Deemed to be University, Dindigul, India Arun Goyal Carbohydrate Enzyme Biotechnology Laboratory, Department of Biosciences and Bioengineering, Indian Institute of Technology Guwahati, Guwahati, India George M. Hall Centre for Sustainable Development, University of Central Lancashire, Preston, United Kingdom Papia Haque Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh M. Mehedi Hasan Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering, Bangabandhu Sheikh Mujibur Rahman Science and Technology University, Gopalganj, Bangladesh Danfeng Hou CAS Key Laboratory of Green Process and Engineering, State Key Laboratory of Multiphase Complex Systems, Beijing Key Laboratory of Ionic Liquids Clean Process, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, P.R. China S.A. Ibitoye Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria B.I. Imasogie Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria E. Jackcina Stobel Christy Department of Chemistry, The Gandhigram Rural Institute—Deemed to be University, Dindigul, India K. Jayaraj Department of Chemistry, The Gandhigram Rural Institute— Deemed to be University, Dindigul, India M. Nuruzzaman Khan Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh Rupak Kishor Department of Chemical Engineering, MANIT, Bhopal, India Suneeta Kumari Department of Chemical Engineering, B.I.T. Sindri, Dhanbad, India Ali Laajeb Catalysis, Materials and Environment Laboratory, Higher School of Technology, Sidi Mohamed Ben Abdellah University, Fez, Morocco Ahmed Lahsini Catalysis, Materials and Environment Laboratory, Higher School of Technology, Sidi Mohamed Ben Abdellah University, Fez, Morocco Xingmei Lu CAS Key Laboratory of Green Process and Engineering, State Key Laboratory of Multiphase Complex Systems, Beijing Key Laboratory of Ionic Liquids Clean Process, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, P.R. China; School of Chemical and Engineering, University of Chinese Academy of Sciences, Beijing, P.R. China

List of Contributors

xi

Abul K. Mallik Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh Rifat A. Masud Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering, Bangabandhu Sheikh Mujibur Rahman Science and Technology University, Gopalganj, Bangladesh Raciye Meral Faculty of Engineering, Department of Food Engineering, Van Yuzuncu Yil University, Van, Turkey Nguyen Cong Minh Institute of Biotechnology and Environment, Nha Trang University, Nha Trang, Vietnam Ruchi Mutreja Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India K.M. Oluwasegun Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria A.O. Oyatogun Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria G.M. Oyatogun Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria ¨ zogul Faculty of Fisheries, Department of Seafood Processing Fatih O Technology, C ¸ ukurova University, Adana, Turkey Anitha Pius Department of Chemistry, The Gandhigram Rural Institute— Deemed to be University, Dindigul, India A.P.I. Popoola Deparment of Chemical, Metallurgical and Materials Engineering, Tshwane University of Technology, Pretoria, South Africa Mohammed Mizanur Rahman Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh A. Rajeswari Department of Chemistry, The Gandhigram Rural Institute— Deemed to be University, Dindigul, India Tarek Rouissi

INRS-ETE, Universite´ du Que´bec, Que´bec, QC, Canada

Md. Nurus Sakib Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh Joseph Sebastian

INRS-ETE, Universite´ du Que´bec, Que´bec, QC, Canada

Md. Shaharuzzaman Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh Keiko Shirai Biotechnology Department, Laboratory of Biopolymers and Pilot Plant of Bioprocessing of Agro-Industrial and Food By-Products, Autonomous Metropolitan University, Mexico City, Mexico W.O. Soboyejo Faculty of Engineering, Wisconsin Polytechnic Institute, Menomonie, WI, United States

xii

List of Contributors

Abhijeet Thakur Carbohydrate Enzyme Biotechnology Laboratory, Department of Biosciences and Bioengineering, Indian Institute of Technology Guwahati, Guwahati, India Trang Si Trung Faculty of Food Technology, Nha Trang University, Nha Trang, Vietnam Aman Ullah Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, AB, Canada Nguyen Van Hoa Trang, Vietnam

Faculty of Food Technology, Nha Trang University, Nha

Mustafa Tahsin Yilmaz Faculty of Engineering, Department of Industrial Engineering, King Abdulaziz University, Jeddah, Saudi Arabia; Chemical and Metallurgical Engineering Faculty, Department of Food Engineering, Yıldız Technical University, Istanbul, Turkey Suojiang Zhang CAS Key Laboratory of Green Process and Engineering, State Key Laboratory of Multiphase Complex Systems, Beijing Key Laboratory of Ionic Liquids Clean Process, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, P.R. China; School of Chemical and Engineering, University of Chinese Academy of Sciences, Beijing, P.R. China Muhammad Zubair Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, AB, Canada

C H A P T E R

1 Chitin and chitosan: origin, properties, and applications Suneeta Kumari1 and Rupak Kishor2 1

Department of Chemical Engineering, B.I.T. Sindri, Dhanbad, India, 2 Department of Chemical Engineering, MANIT, Bhopal, India O U T L I N E

1.1 Introduction

2

1.2 Chitin and chitosan 1.2.1 Sources of chitin

3 4

1.3 Extraction of chitin 1.3.1 Chemical extraction 1.3.2 Biological extraction

5 5 6

1.4 Chitosan preparation methods 1.4.1 Chemical and biological deacetylation of chitin

8 8

1.5 Physicochemical properties 1.5.1 Molecular weight 1.5.2 Viscosity 1.5.3 Solubility 1.5.4 Water-binding capacity and fat-binding capacity

8 9 10 12 13

1.6 Characterization of chitin and chitosan 1.6.1 Fourier-transform infrared spectroscopy (FT-IR) analysis 1.6.2 X-Ray diffraction analysis 1.6.3 13C Nuclear magnetic resonance analysis 1.6.4 Thermogravimetric analysis for chitin and chitosan 1.6.5 Scanning electron microscope analysis

13 14 16 17 19 20

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00001-8

1

© 2020 Elsevier Inc. All rights reserved.

2

1. Chitin and chitosan: origin, properties, and applications

1.7 Application of chitin and chitosan 1.7.1 Biomedical application 1.7.2 Wastewater treatment 1.7.3 Fuel cell 1.7.4 Packaging of food 1.7.5 Textile industries 1.7.6 Bioplastics 1.7.7 Nanocomposite

22 22 24 25 25 26 27 28

References

28

1.1 Introduction Chitin and chitosan are naturally abundant and renewable polymers. They have excellent properties such as biodegradability, biocompatibility, and nontoxicity [1]. Chitin is a copolymer of N-acetyl-D-glucosamine and D-glucosamine units linked with β-(1-4) glycosidic bonds, as shown in Fig. 1.1 [2], where N-acetyl-D-glucosamine units are predominant in the polymeric chain [3]. The deacetylated form of chitin refers to chitosan (Fig. 1.1). Chitin and chitosan can be found as supporting materials in many aquatic, terrestrial, and some microorganisms [4], as shown in Fig. 1.1A. Almost as much chitin is estimated to be produced annually as cellulose. It has become of great interest not only as an underutilized resource but also as a new functional biomaterial with high potential in various fields [5]. Chitin is a white, hard, inelastic, nitrogenous polysaccharide found in the exoskeleton and in the internal structure of invertebrates. The production of chitosan from crustacean shells obtained as a food industry waste is economically feasible, especially if it includes the recovery of carotenoids. There are many applications in wastewater treatment, such as the removal of metal ions [6,7] and dyes [8], as a membrane in purification processes [9], in the food industry (anticholesterol and fat binding), as a packaging material, as a preservative and food additive [10], in agriculture (seed and fertilizer coating) [11], for controlled agrochemical release [12], in the pulp and paper industry (surface treatment adhesive paper) [2], in cosmetics (body creams and Inmaculada Aranaz, lotions, etc.), in tissue engineering [13], in wound healing [14], and as excipients for drug delivery [15] and gene delivery [16]. Additionally, It can be easily processed into gels [17], membranes [18], nanofibers [19], beads [10], microparticles, nanoparticles, scaffolds [15], and sponges [20], as shown in Fig. 1.1C.

Handbook of Chitin and Chitosan

1.2 Chitin and chitosan

3

FIGURE 1.1 (A) Sources of chitin, (B) deacetylation of chitin (synthesis of chitosan), and (C) application of chitosan.

The main objective of this chapter to give a brief introduction about chitin and chitosan and its synthesis. The physical and chemical properties of chitosan determined by using different analytical techniques are also discussed. Finally, the most recent applications of chitin and chitosan are discussed.

1.2 Chitin and chitosan Chitin (C8H13O5N)n, is derived from the Greek word “chiton,” meaning a coat of mail. It is a natural polysaccharide of β-(1-4)-N
acetyl-D-glucosamine monomers, first identified by the chemist Henri Braconnot in 1811 [21]. Its structure is similar to cellulose but with 2-acetamido-2-deoxy-β-Dglucose (NAG) monomer units (Fig. 1.1B). Chitin has limited applications because of its acetyl groups, but through the deacetylation process chitin is converted into chitosan. During the deacetylation process, the acetyl group present in chitin is converted into hydroxyl (
OH) and amino (
NH2) groups in the chitosan. The modification of the reactive functional groups present in chitosan opens the possibility of broad application in many fields. Chitosan’s structural modification is possible by chemical methods

Handbook of Chitin and Chitosan

4

1. Chitin and chitosan: origin, properties, and applications

and most of the new applications focus on the properties and modification of its composites [22].

1.2.1 Sources of chitin Chitin is usually isolated from the exoskeletons of arthropods’ chitin-based tissue (30%
40% protein, 30%
50% calcium carbonate, and 20%
30% chitin), such as crustaceans, mollusks, insects, and certain fungi [2,23
26]. It is a biological nanocomposite material strictly hierarchically organized which reveals various structural levels. At the molecular level is the polysaccharide chitin itself (Fig. 1.2). The next structural level is the arrangement of c. 18
25 of such molecules in the form of narrow and long crystalline units, which are wrapped by proteins, forming nanofibrils of about 2
5 nm diameter and about 300 nm length. The next step in the scale consists of the clustering of some of these nanofibrils into long chitin
protein fibers of about 50
300 nm diameter (Fig. 1.2). Chitin is mainly occurs in three different polymeric α-, β-, and γ-forms. The chains are arranged in stacks or sheets in α-chitin and adjacent sheets along the c-axis have the same direction in a parallel arrangement. The α-chitin occurs in the exoskeletons of crustaceans (e.g., crabs, lobsters, and prawns). In the case of β-chitin, the adjacent sheets along

FIGURE 1.2 Hierarchy of the main structural levels and microstructure elements of the exoskeleton material [25].

Handbook of Chitin and Chitosan

1.3 Extraction of chitin

5

the c-axis present in opposite directions in an antiparallel arrangement and it can be found in squid pen, certain diatoms, and vestimentiferans (a class of deep-sea animal) [27]. However every third sheet is in the opposite direction to the preceding sheets in γ-chitin. It mainly exists in fungi and yeast [28].

1.3 Extraction of chitin Cuticles of various crustaceans like crabs, shrimps, and lobster and fish scales are the major sources of chitin. Crustaceans carry an exoskeleton composed of proteins, chitin, and calcium carbonate which bind together to form an external shell [26]. Protein and chitosan bind together and a small part of the protein is available in the polymer complex. Hence, chitin’s separation from the shell requires the elimination of two major constituents, protein and minerals. Protein is removed by a deproteinization process and minerals are removed by a demineralization process. In some cases an additional process of decolorization is carried out to remove pigments. A variety of methods have been adopted to produce chitin. Among all methods chemical and biological processes are primary for the production of chitin. Moreover, among the two method biological methods the sequence of the process offers some advantages specifically in time consumption and quality of chitin produced. Therefore extraction always begins by choosing crustacean shells. For any particular separation, shells of identical size and species are preferred. In the case of shrimp shell, the separation is easier as the shell wall is thinner but the yield is less in comparison with crab and lobster. However, crab and lobster allow a better quality of chitin to be recovered. Thus the availability of crustaceans remains a key factor for the extraction of chitin [29].

1.3.1 Chemical extraction 1.3.1.1 Demineralization process Demineralization of shells is based on acidic treatment to remove minerals like calcium carbonate and calcium phosphate. The most common reagents are HCl, HNO3, H2SO4, CH3COOH, and HCOOH. Then the demineralized shell is filtered under vacuum and washed with distilled water for 30 min until the pH become neutral. Then the demineralized shells are dried in an oven at around 60 C for 24 h. ð1:1Þ This reaction uses hydrochloric acid to decompose calcium carbonate into calcium chloride with the release of water and carbon dioxide [30]. Handbook of Chitin and Chitosan

6

1. Chitin and chitosan: origin, properties, and applications

Similarly minerals also react with the acid and produce soluble salts. The salts are removed by filtration. Chitin is recovered by washing with distilled water and drying [31]. The demineralization process differs with different types of shells, time of extraction, temperature, shell size, concentration of acid, and solute/solvent ratio. Hence, the solute/solvent ratio is based on stoichiometry (2 mol of acid for decomposed 1 mol of calcium carbonate). The demineralization process generally occurs at high temperature as this allows the solvent to diffuse into the chitin matrix more easily and diffusion depends on the particle size. Moreover, a high concentration of acid, high temperature, and prolonged process time adversely affect the properties of chitin [32]. 1.3.1.2 Deproteinization process Deproteinization of demineralized shells is carried out by using an alkali treatment with common reagents such as NaOH, Na2CO3, NaHCO3, KOH, K2CO3, Ca(OH)2, Na2SO3, NaHSO3, CaHSO3, Na3PO4, and Na2S. The protein is removed from demineralized shells using KOH/NaOH with constant stirring for 2 h at around 90 C and filtered under vacuum with distilled water until pH neutral. Then the deproteinized shell is dried in the oven at around 60 C for 24 h. During the deproteinization process cleavage of chemical bonds occurs between protein and chitin. KOH/NaOH has the tendency to depolymerize and degrade the chitin. The deproteinization process reaction condition is dependent on the source of the crustaceans. Hence, alkali treatment leads to the partial deacetylation of chitin with low molecular weight (Mw) [33]. 1.3.1.3 Decolorization Decolorization of chitin is the final stage of chitin preparation. Acetone or another organic solvent mixture is used to remove the pigments. Decolorizing is done with acetone for 10
20 min and then drying for 2 h at ambient temperature.

1.3.2 Biological extraction The chemical treatment process of chitin has many drawbacks. Chemical chitin purification is extremely hazardous, energy consuming, and threatening to the environment, due to the high concentration of minerals and caustic employed. Biological extraction is an alternative method to extract chitin from crustaceans’ shells. It overcomes the environmental problems associated with acidic and alkali treatment. The advantages of biological methods

Handbook of Chitin and Chitosan

1.3 Extraction of chitin

7

include the production of chitin with higher reproducibility. Moreover, the solubility of chitin is limited and the biological approach is limited. 1.3.2.1 Enzymatic demineralization Mineral and protein present in the crustacean shells are dissolved by organic acid with microorganisms such as lactic acid-producing bacteria. This enzymatic demineralization reaction involves organic acid and microorganism reacting with calcium carbonate present in the raw shells and calcium salts precipitate the organic acid. The precipitated salts are removed by the culture medium with special care. The precipitated organic salts are also removed by washing and are used as preservative and antiicing agents [34]. 1.3.2.2 Enzymatic deproteinization In the enzymatic deproteinization process, proteases (alcalase, pepsin, papain, pancreatin, devolvase, and trypsin) from bacteria can eliminate proteins. Commonly, proteolytic enzymes are obtained from plants, microbes, and animal sources. They remove proteins and reduce the steps in preliminary processes. Alcalase is generally preferred for the production of chitin, protein hydrolyzate, and astaxanthin recovery. Its hydrophobic amino group controls hydrolysis. Alcalase is a serine endopeptidase obtained from Bacillus licheniformis. It is selected due to its specificity for terminal hydrophobic amino acids. It generally leads to the production of nonbitter hydrolyzate and allows an easy control of the degree of hydrolysis. Raw shells are demineralized after deproteinization using HCl treatment and the residual protein content is higher in the chitin isolated with the enzymatic deproteinization than that obtained with alkali treatment [35]. 1.3.2.3 Fermentation The enzyme process has a high cost due to the enzymes. The cost of using enzymes can be decreased by performing deproteinization using a fermentation process [36]. Fermentation methods are separated into two major categories: lactic acid fermentation and nonlactic acid fermentation. Fermentation of crustacean waste results in a solid fraction containing crude chitin and the production of liquor rich in natural protein, mineral, and pigments. Moreover, the action of the lactic acid-producing bacteria is twofold [37]. They produce a spectrum of proteases that detach protein from the solid chitin
CaCO3 complex by partial hydrolysis. Thus extraction by biological activity is gaining in importance over chemical treatment. It is an eco-friendly process and the by-products can be recovered and recused [38].

Handbook of Chitin and Chitosan

8

1. Chitin and chitosan: origin, properties, and applications

1.4 Chitosan preparation methods Chitosan is prepared by a degree of acetylation. Acetyl groups are removed during the deacetylation process and Mw changes due to the depolymerization reaction. There are two processes, that is, the enzymatic process and chemical process, and chitosan is produced by chemical process. It is preferable for large-scale production.

1.4.1 Chemical and biological deacetylation of chitin Glycosidic bonds are attracted toward acids and alkalis. Chitin is processed homogeneously or heterogeneously. In the homogeneous method, chitin is diffused in concentrated alkali at 25 C for 3 h and allowed to disperse in compressed ice at around 0 C [39]. In the heterogeneous process the chitin is treated with hot high-concentration alkali and then washed with distilled water until the pH is neutral. It is difficult to produce higher deacetylated chitosan. The addition of thiophenol as a catalyst during the process would minimize the degradation by trapping oxygen and enhance the effective deacetylation. The effective deacetylation process of chitin achieves the preparation of chitosan if the alkali concentration is four times greater than the total amino group in the polysaccharide at a temperature around 100 C for the duration of 1 h. It is recommended to use low concentration alkali and a short contact time between alkali and polymer [40]. Chemical deacetylation has many disadvantages like high energy consumption and environmental pollution problems [41]. An alternative method of enzyme deacetylation has been developed to overcome these drawbacks. Chitin deacetylation enzyme acts as a catalysis to hydrolyze Nacetamide bonds [42]. This enzyme is extracted from the fungi Mucor rouxii, Absidia coerulea, Aspergillus hidulans, and two strains of Celletotrichum lindemuthianum. This enzyme is thermally stable and has a binding affinity toward β-(1, 4)-linked N
acetyl-D-glucosomine polymers [43]. Most of the time the enzyme process is carried out in both batch and continuous culture. In the batch process the Mw of chitosan is lower with respect to time. Moreover, chitosan of higher Mw is obtained in a specific culture even though the yield is comparatively low [44].

1.5 Physicochemical properties Chitin is a colorless, crystalline or amorphous powder that is insoluble in water, organic solvents, dilute acids, and alkalis. It dissolves in concentrated mineral acids with simultaneous degradation of the polymer [45]. Although chitosan is insoluble in water, it does dissolve in aqueous

Handbook of Chitin and Chitosan

1.5 Physicochemical properties

9

organic acids, for example, acetic and formic acids, as well as inorganic acids. Chitin and chitosan preparation can vary due to the source, with compositional differences. Similarly, the physicochemical characteristics of chitin and chitosan differ between crustacean species and preparation methods [46]. Several studies have clearly demonstrated the specific characteristics of these products. The Mw and degree of deacetylation (DD) vary with process conditions [46
48]. The physicochemical characteristics of chitin and chitosan influence their functional properties [24,49]. Mw and DD vary with process conditions or different extraction methods [24]. Chitosan’s application depends on physical, biological, and chemical properties and chitosan depends on two parameters, such as DD and Mw [50].

1.5.1 Molecular weight Chitosan and its derivatives have been used in a wide variety of applications but the effectiveness of these materials has been found to be dependent upon their DD, crystallinity, and Mw [51]. DD and Mw of chitosan are greatly affected by reaction conditions like temperature, reagents concentrations, repetition of alkaline steps, time, and atmospheric conditions of the deacetylation [52]. Chitosan is prepared by chemical and enzymatic processes [1]. The chemical process has several drawbacks such as low product yields and poor structural order (glucose ring). Additionally, acidic and alkali treatments during the chemical process could be a source of environmental problems [24]. The enzymatic process is an alternative way to synthesize chitosan which is more environment-friendly [53]. During the enzymatic process lactic acid-producing bacteria are used for the demineralization of crustacean shells instead of acidic treatment. The obtained lactic acid reacts with calcium carbonate yielding calcium lactate, which can be precipitated and removed. During the deproteination of crustacean shells, proteases from bacteria (Pseudomonas aeruginosa K-187, Serratia marcescens, FS-3, and Bacillus subtilis) are used. Serratia sp. and Bacillus sp. are bacteria that also produce chitin deacetylase and can be used to generate chitosan [54]. The effective deacetylation is attained by intermittently washing the intermediate product with water during the alkali treatment [55]. The average Mw of chitosan is B500 kDa with 100% of DDA. The DDA increases rapidly to about 68% during the first hour of alkali treatment (50% NaOH) at 100 C and further slowly increases with time [48]. There are several methods used for the calculation of Mw, such as light scattering, gel permeation chromatography (GPC), and capillary viscometry. Capillary viscometry is the simplest and most widely used method to determine the Mw of chitosan. During analysis, an Ubbelohde-type

Handbook of Chitin and Chitosan

10

1. Chitin and chitosan: origin, properties, and applications

capillary viscometer is used to measure the passage time of solutions flowing through the capillary at 25 C. Different viscosity solutions of chitosan are used at various concentrations ranging from 0.00125% to 0.15% in 0.1 M acetic acid
0.2 M NaCl solutions. The capillary viscometer is filled with the sample and is equilibrated in a water bath at 25 C. The solution and solvent flow times are measured to calculate relative viscosity. The Mark
Houwink
Sakurada equation given below provides the relation between intrinsic viscosity (η) and Mw [51]: ½η 5 K 3 ½Mwa 5 3:04 3 10
5 ½Mw1:26

(1.2)

Based on Mw, chitosan is classified into three different types, namely low-molecular-weight chitosan (LMWC; ,50 kDa), medium-molecularweight chitosan (MMWC; 50
250 kDa), and high-molecular-weight chitosan (HMWC) ( . 250 kDa). Several authors have reported that LMWC has enhanced properties, such as antibacterial and antifungal [56], antitumor [57], lipid metabolism, intestinal disaccharidase [58], and mucoadhesive properties [59]. Apart from this the Mw also plays an important role in the biopolymer’s rheological properties. It directly impacts the development of chitosan-based biomaterials [60]. LMWC is used as an anticreasing agent to produce a finishing agent and then applied in the anticreasing treatment of cotton fabrics. HMWC is used in quaternized chitosan films with properties of water solubility and free radical scavenging, and HMWC is also used for high-performance cells and its effects on polymer aggregation and phase separation.

1.5.2 Viscosity Chitosan is the best known deacetylated derivative of chitin, as one of its unique properties is its polycationic nature when dissolved in acidic solution (the value of pKa 5 6.0). Hence this biopolymer is favorable for a broad variety of industrial and biomedical applications. When chitosan is dissolved in acidic solution, it gives a viscous solution. The viscosity of the solution is related to the Mw, DD, concentration, pH, and temperature of chitosan solution. The viscosity of chitosan solution, at the molecular level, is a direct measure of the volume of the polymer molecules, which in turn is governed by the molecular size or chain length. The viscosity of the polymer solution is widely measured by a capillary viscometer and its value is used to determine the average Mw of polymer by using the Mark
Houwink equation (Eq. (1.1)):

Handbook of Chitin and Chitosan

11

1.5 Physicochemical properties

Where Mv is the average Mw of the polymer and α and k are constants (α 5 0.83 and k 5 1.4 3 1024 for 0.25 M acetic acid and 0.25 M sodium acetate solvent system). The intrinsic viscosity [η] can be determined from the following equation (Eq. (1.2)): ½ η 5

ðη 2 ηsÞ ðηsCÞ

(1.3)

where η (dL/g) is the solution viscosity and ηs is the reference solvent viscosity and C is the solution concentration. As indicated in (1.2), when (η 2 ηs)/ηsC, that is, reduced viscosity (ηred) is plotted against concentration (C), the intercept corresponds to [η]. The [η] value is directly used in determining the Mw of different grades of chitosan. Some of the typical chitosan viscosities with different Mws are summarized in Table 1.1 [61]. Flow activation energy (Ef) of viscous chitosan sample is analyzed by the Arrhenius equation (Eq. 1.3): ½η 5 AeEf=RT

(1.4)

where A and R are the preexponential factor and universal gas constant absolute temperature (in Kelvin). The value of Ef of two different grade of chitosan 91% DOA and 70% DOA was 25 and 15 kJ/mol, respectively, as reported by Wei Wang [62]. This means that the entanglement of chains increases with the increasing DD of chitosan because of the nature of the polymer. The viscosity and flow properties of the concentrated solutions of chitosan with different degrees of deacetylation are different. The viscosities and the non-Newtonian flow properties of the solutions increase with the increasing DD of chitosan. On the other hand, additional salt decreases the viscosities and the non-Newtonian flow properties of the solutions of chitosan.

TABLE 1.1 Intrinsic viscosity and viscosity average molecular weight of different grades of chitosan. Chitosan

Intrinsic viscosity [η], dL/g

Molecular weight, MV

CHT-1

9.40

654,127

CHT-2

4.72

285,231

CHT-3

2.55

135,839

CHT-4

1.50

71,676

CHT-5

0.535

20,698

Handbook of Chitin and Chitosan

12

1. Chitin and chitosan: origin, properties, and applications

1.5.3 Solubility The three-dimensional chitin crystalline matrix consists of a combination of strong hydrogen bonds and cohesive forces of acetyl (OQC), hydroxyl (2OH) and amine (2NH) functional groups. These function groups link with sNH?OQC and sOH?OQC by hydrogen bonds in the threedimensional matrix. Pure chitin contains around 90% N-acetyl groups in its backbone and some deacetylation reactions take place due to the extraction process of chitin from the natural sources. The first study on the solubility of chitin was performed by Austin, who tested the solubility of chitin in different organic solvents [63]. It was a well-organized evaluation of chitin’s solubility in different types of solvents, such as dichloroacetic (DCA) and trichloroacetic (TCA) acids in the presence or absence of alcohol, etc. Later on, many studies were conducted with the same intentions by many other researchers and chitin’s solubility was verified in many solvents such as dimethylacetamide (DMA)/LiCl mixture, CaBr2 H2O saturated methanol, hexafluoroisopropyl alcohol and hexafluoraceton, lithium thiocyanate, phosphoric acid and N-methyl2-pyrrolidone, etc. [64]. Although the dissolution of chitin is possible by these solvents, many of them are toxic, scarcely degradable, corrosive, or mutagenic. Therefore the choice of an appropriate solvent for chitin and chitosan solubilization is important and a primary issue for lab-scale research and scaling up for industrial practices [65]. There are two monomer units present in the chitin backbone in different fractions namely 2-acetamino-2-deoxy-D-glucopyronase (N-acetyl-D-glucosamine) and 2-amino-2-deoxy-D-glucopyronase (N-amino-D-glucosamine) [66]. The first one group, 2-acetamino-2-deoxy-D-glucopyronase, displays insolubility due to the strong hydrogen bonds between the acetyl groups of the same or adjacent chitin chains. In the other unit N-amino-D-glucosamine is hydrophilic in nature and positively charged in acidic solution. The domination of the hydrophilic character with a high amount of N-amino-D-glucosamine units in the chitin backbone make it soluble in the specific acidic solution. The percentage of N-amino-D-glucosamine unit in the chitin backbone can be determined by DDA. It is the ration of N-amino-D-glucosamine to N-acetyl-D-glucosamine, while the degree of acetylation represents the detection from 100 (100-DDA). When DDA is between 60% and 90%, a new chemical entity “chitosan” is formed and it is soluble in organic acids such as acetic acid. Chitosan can be dissolved in aqueous diluted acids as a polycation at B50% DDA or more due to the presence of N-amino-D-glucosamine units [64]. The other parameters, such as temperature, time of deacetylation, alkali contraction, and prior treatment, applied to chitin isolation and DD also affect the solubility of chitosan. Therefore the fraction of N-amino-D-glucosamine units has a large influence on the solubility and the solution’s properties.



Handbook of Chitin and Chitosan

1.6 Characterization of chitin and chitosan

13

1.5.4 Water-binding capacity and fat-binding capacity Water-binding capacity (WBC) and fat-binding capacity (FBC) of chitin and chitosan is measured by a modified method of Wang and Kinsella [67]. In a typical process, water or fat absorption is initially carried out by weighing a centrifuge tube containing 0.5 g of sample and adding 10 mL of water or soybean oil and mixing with a vortex mixer for 1 min to disperse the sample. The contents are left at ambient temperature for 30 min with shaking for 5 s every 10 min, before being centrifuged at B3200 rpm for 25 min. After the supernatant is decanted, the tube is weighed again. WBC and FBC are calculated as follows [24]. WBC ð%Þ 5

Water bound ðgÞ Initial sample weight ðgÞ

(1.5)

FBC ð%Þ 5

Fat bound ðgÞ Initial sample weight ðgÞ

(1.6)

WBC differed with products ranging from 381% to 673% for chitins and from 458% to 805% for chitosans [24]. Chitosan has a higher WBC than chitin. The WBC values are different for each source (chitin) and chitosan. The differences in WBC between chitin and chitosan are possibly due to dissimilarities in crystallinity, the number of salt-forming groups, and the residual protein content of the products [68]. FBC for products of chitins were in the range of 316%
320%, whereas chitosan shows dissimilar binding capacities ranging from 314%
535%. This apparently suggests that chitin can or cannot have higher FBC than chitosan depending on the products [24].

1.6 Characterization of chitin and chitosan After several decades of intense research concerning the synthesis, characterization, and application of chitin and chitosan, a complete and detailed description of the polymer and its characteristics cannot be deduced yet; even from those studies performed by using advance experimental techniques such as X-ray diffraction or high-resolution scanning electron microscopy (HRSEM). As a consequence, suitable atomistic models have been developed to try to describe their atomic structure in order to understand their different properties and to help in tailoring them for specific purposes. A detailed description of chitin and chitosan must include (1) characteristics of the raw chitin and the produced chitosan, by means of surface functional group; (2) thermal analyses; (3) structural analysis; (4) Mw; (5) viscosity; and (6) the morphology of the product.

Handbook of Chitin and Chitosan

14

1. Chitin and chitosan: origin, properties, and applications

A brief description of different characterization techniques is discussed below.

1.6.1 Fourier-transform infrared spectroscopy (FT-IR) analysis FT-IR analytical technique is used to understand the functional groups present in the chitin and chitosan. The chitins (α-, β-, and γ-chitin) can be distinguished by FT-IR spectra and these are shown in Fig. 1.3. In the case of α-chitin, two separate peaks are observed at B1662 and B1630 cm21. They represent the amide-I band present in the α-chitin. It is associated with the occurrence of the intermolecular hydrogen band
CO. . .HN and
CO. . .. . .HOCH2 [71]. In the case of β-chitin, a single band is observed at 1656 cm21 due to the hydrogen bond present between the amide group (
CQO) of the neighboring intrasheet chain [27]. The
NH stretching band at 3264 and 3107 cm21 is clearly observed in both α- and β-chitin. The bands at 703 and 750 cm21 represent the bending vibration of 2 OH groups and NH
groups present in the α-chitin, respectively. While in the case of β-chitin, they are shifted to 682 and 710 cm21 [71]. However, β-chitin shows a single band at approximately 1656 cm21 which is associated with the intermolecular hydrogen bond of CO. . .HN [69]. The β-chitin have the weaker inter- and intramolecular hydrogen bonding and amore loosely ordered structure compared with α-chitin. γ-Chitin shows the two sharp peaks at 1660 and 1620 cm21 for the amide-I band, which is also available in the α-chitin. Apart from the expected decrease in the band intensity at 1665 and 1550 cm21 for chitin (amide-I), they appeared at nearly 1604, 1598, and 1592 cm21 for chitosan. This demonstrates the effective deacetylation of chitin. Moreover, the bands assigned to the stretching vibrations of the glycosidic bond of chitosan polysaccharide structure at 1151, 1098, and 1021 cm21 weakened distinctly after the deacetylation process. The bands nearly disappeared at 3260 and 3107 cm21, originating from N 2 H stretching after deacetylation indicating the disturbed the regular hydrogen bond of N 2 H in the chitin. In the α-chitin spectra: two separate peaks are observed at 1662 and 1630 cm21. They are associated with the occurrence of the intermolecular hydrogen band
CO. . .HN and
CO. . ...HOCH2. Moreover, a single band is observed at 1656 cm21 in case of the β-chitin. It is commonly related to the stretching of the
CO group hydrogen bond to amide group of the neighboring intrasheet chain [27]. The strong band at 1430 cm21 is noted for β-chitin, while a sharp band at 1416 cm21 present in the case of α-chitin [28]. The NH
stretching band at 3264 and 3107 cm21 is clearly observed in the α-chitin but in the case of β-chitin, it is not easily seen (Fig. 1.3A). The band at 703 cm21 due to the 2 OH out of plane and 750 cm21 due to NH
out of plane is shows in the α-chitin. While in the case of β-chitin, it is shifted to Handbook of Chitin and Chitosan

1.6 Characterization of chitin and chitosan

15

FIGURE 1.3 FT-IR spectra of: (A) different types of chitin (α-, β-, and γ-) [69] and (B) chitosan [70].

682 and 710 cm21 [71]. The major difference between α- and β-chitin is low crystalline and loosely ordered structure showing weaker inter and intramolecular hydrogen banding in the β-chitin compared to the α-chitin. In case of γ-chitin is showed an amide -I band. It is split into two sharp sub peaks at 1660 and 1620 cm21. Similar pattern of α-chitin due to the properties of both α- and β-chitin [69]. In the case of chitosan, the band at 1662 cm21

Handbook of Chitin and Chitosan

16

1. Chitin and chitosan: origin, properties, and applications

corresponded to the stretching of
CQO and CO
NH in amide band decrease. The band 1310 cm21 due to the
CO
NH bending vibration is also reducing. Its indicate the effective deacetylation. Moreover, the bands assigned to the stretching vibrations of glycosidic bond of chitosan polysaccharide structure at 1151, 1098 and 1021 cm21 weakened distinctly after the deacetylation process. Which is confirmed the depolymerization of chitosan. The bands nearly disappeared at 3260 and 3107 cm21 originating from N 2 H stretching after deacetylation indicated. Its indicate the disturbed the regular hydrogen bond of N 2 H in the unreacted chitin (Fig. 1.3B).

1.6.2 X-Ray diffraction analysis XRD analysis is applied to understand the order/disorder degree of the structure of the crystalline isolated chitin and chitosan. The α-, β-, and γ-chitin show two major crystalline reflection peaks along the 2θ angle. These XRD patterns suggest that all the biopolymers are semicrystalline materials. The α-chitin shows the major crystalline reflection peaks at 9.6 degrees (020 plane) and 19.6 degrees (110 plane), β-chitin at 9.1 degrees (020 plane) and 20.3 degrees (110 plane), and γ-chitin at 9.6 degrees (020 plane) and 19.80 degrees (110 plane), as shown in Fig. 1.4. In α- and γ-chitin, the second reflection peak is observed at 12.74 degrees, while for β-chitin the same peak is recorded with a slightly lower intensity at B12.29 degrees. The crystallinity index (CrI) value is higher for α-chitin (greater than 90%), and lower for the β-chitin (greater than 75%
89 %). The CrI value of the γ-chitin is B68.6%, which is lower than both α- and β-chitin. The crystalline peaks for β-chitin extracted from squid pen are at 9.8 and 19.3 degrees, while α-chitin (shrimp, lobster, prawn, and king crab) are at B9.28 6 0.1, 19.36 6 0.3, and 26.20 6 0.1. Chitosan is a twofold helix or a zigzag structure with a fiber repeating unit of O3
O5 at 10.34A . This is due to the incorporation of bound water molecules into the crystal lattice. Chitin and chitosan show broad peaks with a crystalline lattice with interplane distances of d(110) 5 0.34, 0.45, 0.50, and 1.09 nm with a shoulder at 0.71 nm. This is related to the β-(1-4)linked polyglucose polymannose and polyglucosamine. Chitosan chains are packed in an antiparallel fashion in the orthorhombic unit cell (Fig. 1.2). Its chains on the c-axis are up chains and on the unit cell they are down chains. The down chain is bound by hydrogen bonds and form a sheet structure. These sheets are stacked along the a-axis. Water molecules are present between these sheets and stabilize this crystal structure. Its polymorph is the most abundant in chitosan. Its adjacent parallel chains are connected by the O6—N2 hydrogen bond, which forms a sheet structure along the b-axis. There is no hydrogen bond between these sheets along the a-axis. Handbook of Chitin and Chitosan

1.6 Characterization of chitin and chitosan

17

FIGURE 1.4 XRD spectra: (A) α-, β-, and γ-chitin [69]; (B) chitin and chitosan [23].

1.6.3

13

C Nuclear magnetic resonance analysis

The structure of the chemical bridges or bond present in the chitin and chitosan has been extensively examined by 13C Nuclear magnetic resonance (13C-NMR). The conformation of chitin, (1-3)
β-D and (1-4)
α-D-glucans is analyzed by high-resolution solid-state 13C-NMR spectroscopy. 13C solid-state NMR appears to be the most reliable option for the evaluation of the acetyl content and the average degree of acetylation of chitins [67,72]. Chitin consists of eight well-defined resonances, as shown in Fig. 1.5. These resonances for carbon atoms of the N-acetylglucosamine repetitive unit appear at the following chemical shifts (ppm): δ 5 109.91 (C1), 62.16 (C2), 65.58 (C6), 79.89 (C3, C5), 87.89 (C4), carbonyl group (CQO) signal belongs to the acetamido moiety showing little intensity at 173.24 ppm, and methyl group in acetamide moiety at around 28
22 ppm (C7) signals (Table 1.2).

Handbook of Chitin and Chitosan

18

1. Chitin and chitosan: origin, properties, and applications

13

FIGURE 1.5 TABLE 1.2

C-NMR for chitin and chitosan [67].

13

C-NMR peak position for chitosan [72].

Types of carbon

Position (&, ppm)

C1

102.7
105.7

C2

55.2
57.6

C3

73.1
75.7

C4

80.9
85.7

C5

73.1
75.7

C6

59.6
60.8


CH3 (C7)

22.8
23.3

N
CQO (C8)

173.6
173.8

The conformation-dependent 13C chemical shifts of the C1 and CX carbon atoms of polysaccharides have been demonstrated. They are related to glucoside linkages, and the torsion angle at the glycosidic linkages vary appreciably (up to 8 ppm) with the torsion angles at the glycosidic linkages, Cl
Ogly (ϕ) and Ogly
CX (ψ) [67]. The single molecular chain in these polymorphs is always an extended twofold helical structure similar to chitin or common cellulose. 13C chemical shifts of chitosan go from one polymorph to the other. The basis of the peak assignments in the β-anomer of 2-amino-

Handbook of Chitin and Chitosan

1.6 Characterization of chitin and chitosan

19

2
deoxy-D-glucopyranose and the magnitude of displacement of peaks are due to the formation of glycosidic linkages. Therefore it is concluded that the crab shell chitosan assumes the structure of the tendon polymorph. As the most notable feature, the C4 13C-NMR signal of this polymorph appears as a doublet. The separation of which is 3.4
4.4 ppm, but the C1 13C-NMR signal is a singlet (Fig. 1.5). The C1 13C signal of shrimp chitosan is split into a doublet with a peak separation of 1.4 ppm but the C4 signal in this case appears as a plateau whose edges (Fig. 1.5; C4A and C4B) correspond to the doublet peak of the crab chitosan. This finding suggests that the shrimp chitosan assumes a different conformation from that of the “tendon chitosan” (Fig. 1.5). The observation of the latter spectral feature is in good agreement with that of chitin and native cellulose (Cellulose-I), which are known to adopt the extended helical conformation. The lattice of tendon chitosan showed the same symmetry as the unit cell of α-chitin. If so, it is rather difficult to account for the double patterns of the C1 and C4 peaks in the tendon chitosan and L-2 polymorphs. In this connection, it is worthwhile to recall that the C1 and C4 peaks of regenerated cellulose (Cellulose-II) gave peak splitting (of equal peak intensive) as large as 2 and 1 ppm, respectively. The existence of two peaks for the C1 and C4 is first interpreted in terms of a nonequivalence of alternate glycoside linkages (two sets of torsion angles) along the molecular chain. This interpretation requires that dimeric cellobiose, rather than glucose, is considered as the basic repeating unit of cellulose-II.

1.6.4 Thermogravimetric analysis for chitin and chitosan Thermal properties depend on the Mw of the polymer. Thermogravimetric analysis (TGA) and derivative thermogravimetric (DTG) curves are constructed for α-, β-, and γ-chitin, as shown in Fig. 1.6 to evaluate and compare their thermogravimetric profiles with respect to thermal stability and degree of degradation of α-, β-, and γ-chitin in two different stages [28]. The first weight loss is due to the evaporation of water from hydrophilic groups in chitin chains. For α-, β-, and γ-chitin, the first weight loss is recorded at a temperature of B205 C (5.20%), B205 C (7.93%), and B130 C (4.95%), respectively. The first degradation stage for all three of the respective chitin samples shows no notable differences. The second weight loss corresponds to the degradation of polymeric structure, that is, saccharide ring dehydration and deterioration of acetylated chitin units [73]. The DTGmax values recorded for all three chitins are 391.09 C (74.37%) for α-chitin, 348.77 C (72.56%), for β-chitin and 381.99 C (69.87%) for γ-chitin. The DTGmax for α-chitin is slightly higher than for γ-chitin. Moreover, it is much higher than β-chitin. The γ-chitin reveals a low degradation % at its DTGmax temperature compared to α-chitin and β-chitin. It can be

Handbook of Chitin and Chitosan

20

1. Chitin and chitosan: origin, properties, and applications

FIGURE 1.6 (A) TGA thermograms for α-chitin from male blue swimming crab and β-chitin from cuttle fish chitin including DTG thermogram for α- and β-chitin. This suggests that α-chitin exists as a more stable structure with regard to thermal decomposition than β-chitin [69]. TGA/DTG spectra: (α-, β-, and γ-chitin). (B) Chitin and (C) chitosan [73].

attributed to the structural organization of γ-chitin and it comprises thick rod-shaped microfibers. The DTGmax value for α-chitin is recorded as 350 C 2 400 C. Second stage decomposition is at around 350 C
400 C (Fig. 1.6) due to the degradation of the saccharide structure of the molecule, with the dehydration of saccharide rings and the polymerization and decomposition of the acetylation and deacetylation units of chitin. The percentage of residual mass after heating at around 1000 C is 36% due to the presence of minerals [74].

1.6.5 Scanning electron microscope analysis Surface morphology of chitin and chitosan is analyzed by SEM analysis, as shown in Fig. 1.7. The morphology of the exoskeletons of arthropods, including each reaction step for chitosan preparation and its morphology, is shown in Fig. 1.7(a
c). Raw shrimp shell shows a heterogeneous

Handbook of Chitin and Chitosan

1.6 Characterization of chitin and chitosan

21

FIGURE 1.7 (A) SEM images: (a) Raw shrimp shell; (b) demineralization; (c) after deproteinization; and (d) after deacetylation [75]. (B) SEM images: α-chitin (a and b); β-chitin (e and f); and γ-chitin (i and j) [69].

Handbook of Chitin and Chitosan

22

1. Chitin and chitosan: origin, properties, and applications

morphology with a compact structure, well-defined shape, and white spots (Fig. 1.7(a
c)) [75]. White spots indicate that CaCO3 is present in the raw shrimp shell. After the demineralization process, CaCO3 is removed by acid treatment and white spots appear as a rounded hole, as shown in Fig. 1.7(d
f). However, after acidification treatment, the surface becomes a microfibrillar crystalline structure (hierarchical layer patterns) [76]. Furthermore, after the deproteinization and demineralization process, shells produce chitin with a fibrous structure (Fig. 1.7(g
i)). Shrimp chitin also shows a composition of microfibrils. It is also entrapped together with large surface pores. The porous structure of chitin enables good reactivity and a large surface is available for the reaction in the initial stage. The α-, β-, and γ-chitin surface morphologies from different sources are shown in Fig. 1.7B. The α-, β-, and γ-chitins are microfiber structures on the surface and do not have any nanofibers (Fig. 1.7B(i, j)). The α-chitin has some pores between the fibers [28]. β-chitin has slightly visible fibers, but γ-chitin shows cocoon microfibers [69]. After deacetylation, chitosan’s surface is rough and scarred with irregular fibrils [75]. Pores and fibers combine with chitin and chitosan from crustaceans, like pink shrimp, krill, and Gammarus argae, and other sources of chitin as well as other insects. In a similar manner, the external surface of the prawn shell shows homogeneous spots and pore canals with a nanofibrous structure. It is very similar for shrimp, crab, and fungi sources [77]. N-acetyl groups with amino groups form weaker connections including a less porous structure [78].

1.7 Application of chitin and chitosan Chitosan has functional groups like hydroxyl and amino groups, which might be modified by controlled chemical reactions. They are also used to chemically alter its physical and solution properties. Due to its unique properties, there are many other applications in biomedicine, fuel cells, waste water treatment, packaging of food, agriculture, textile industries, bioplastics, and nanocomposites that will be discussed in this section [79].

1.7.1 Biomedical application Most living tissues have a negative charge, but chitosan is positively charged and attracted to tissues, skin, bone, and hair. The external surface of most microbes are also negative charged. Hence, the binding capacity of chitosan is an important property for all types of living cells as well as biomedical applications. Biomedical application of chitosan also depend on its biological properties as shown in Fig. 1.8. Chitosan beads can be easily formed in a variety of porosities and sizes. Its beads are suitable for drug delivery, and enzyme and cell immobilization [80]. Handbook of Chitin and Chitosan

1.7 Application of chitin and chitosan

23

FIGURE 1.8 Chitosan’s biological properties.

Chitosan has exceptional biocompatible, tissue regeneration, antibacterial, antiinflammatory, and hemostatic properties, as shown in Fig. 1.8. Biomedical applications of chitosan are related with its most important properties: biocompatibility and biodegradability. Biocompatibility is directly related to the cytotoxicity of the material. The cell culture method is the most simple and widely used way to study both the toxicity and chitosan
cell interactions. The agglutination of blood protein and platelet activation is related to encouraging fibrin clot formation [81]. Moreover, chitosan’s derivatives are prepared by grafting hydrophobic groups to improve hemostatic property greatly. N-alkylated chitosan (NACS) converts the highest desirable materials for the fabrication of perfect hemostatic dressings activity, and also shows excellent plugging ability to hemorrhage wound surfaces. NACS may transform whole liquid blood into a gel quickly. Thus it can stop bleeding rapidly from both minor and serious injuries. Hence, NACS can accelerate blood clotting significantly. There are many NACS-based hemostatic materials, such as powder, sponge, foam, and gauze, that are currently available. Nanofiber membrane (NM) has a larger surface area and nanoscale size, therefore, NACS-NM might have the potential to accelerate platelet aggregation. It enhances platelet adhesion and aggregation, which are

Handbook of Chitin and Chitosan

24

1. Chitin and chitosan: origin, properties, and applications

the very first steps involved in the early wound-healing process. On the other hand, some sulfate derivatives exhibit noticeable anticoagulant activity. Sulfate chitosan showed an anticoagulant potency similar to heparin [82]. Generally, polymers are usually applied as a film or ointment. A chitosan derivative forming a hydrogel when exposed to ultraviolet light has been recently proposed as a biological adhesive for soft tissues. Most of the time, hydrogel covers a wound effectively, by firmly adhering two pieces of skin together to accelerate closure and healing. The developed bilayer chitosan wound dressing contained silver sulfadiazine for the control of wound infection. Chitosan films can contain basic fibroblast growth factor to accelerate wound healing.

1.7.2 Wastewater treatment Industrial effluents are discharges from various industries, and various organic pollutants have been found in different water resources. They belong to various classes such as pesticides, fertilizers, hydrocarbons, phenols, plasticizers, biphenyls, detergents, oils, greases, pharmaceuticals, etc. [83]. Hence, industrial wastewater creates a serious environmental problem. It poses a danger to water quality when discharged into rivers and lakes. Moreover, organic and inorganic contaminants are effectively removed to meet increasingly stringent environmental quality standards. It is becoming increasingly documented that the nontoxic and biodegradable biopolymers such as chitin and chitosan could be used in wastewater treatment [84]. Their polycationic properties allow them the ability to conglomerate and precipitate in neutral or alkaline pH. Moreover, the long polymer chain may facilitate the contact between the polymer and the contaminated medium [85]. The reactive amino (
NH2) and hydroxyl (
OH) groups in its backbone make chitosan able to be used as an effective adsorbent material for the removal of wastewater pollutants. Hence, a major advantage of chitosan over other polysaccharides (cellulose or starch) is that its chemical structure allows specific modifications to design polymers for selected applications. The reactive groups are able to develop composites with different compounds. Chitosan has a better capacity to adsorb wastewater pollutants and to resist an acidic environment, such as bentonite, kaolinite, oil palm ash, montmorillonite, polyurethane, zeolites, magnetite, etc. Alternatively, the cationic charge (chitosan is a single cationic biopolymer) is able to neutralize and successfully flocculate the anionic suspended colloidal particles and reduce the levels of chemical oxygen demand, chlorides, and turbidity in wastewaters [86]. Chitosan has regeneration capacity and its eco-friendly nature allows its use in adsorption processes [87]. Chitosan is used as coagulating/flocculating agents for polluted wastewaters, in heavy metal or

Handbook of Chitin and Chitosan

1.7 Application of chitin and chitosan

25

metalloid adsorption (Cu(II), Cd(II), Pb(II), Fe(III), Zn(II), Cr(III), etc.), and for the removal of dyes from industrial wastewater, as well as for the removal of other organic pollutants, such as organochloride pesticides, organic oxidized, or fatty and oil impurities.

1.7.3 Fuel cell Fuel cells are eco-friendly devices for energy conversion, power generation, and highly promising as zero-emission power sources [88]. Fuel cells are electrochemical devices. They convert the chemical energy from a redox reaction directly into electrical energy [89]. Mostly this cell contains electrolyte material and it is packed between two thin electrodes (porous anode and cathode). Fuel is passed over the anode and oxygen passes over the cathode. It is dissociated catalytically into ions and electrons and electrons pass through an external electrical circuit to provide power while the ions move through the electrolyte toward the oppositely charged electrode provide power. There are several types of fuel cells. Most of them use solid polymer-based electrolyte membranes that offer advantages such as high efficiency and high energy density. Hence, polymer electrolyte membranes are highly expensive components of a polymer electrolyte-based fuel cell. Moreover, cost-effective and eco-friendly polymer electrolytes from renewable sources are become promising substitutes for synthetic polymers in fuel cells. Natural polymers, like polysaccharides, are among the best candidates due to their abundance in the environment. Chitosan and its derivatives are low-cost biopolymers. They have attracted attention in various scientific and engineering processes with attractive properties, such as biocompatibility, nontoxicity, chemical and thermal stability. Chitosan has a positive charge arising due to the highly protonated amino functionalities that enables chitosan to form polyelectrolyte complexes spontaneously with a wide variety of negatively charged polyanions, for example, lipids, collagen, glycosaminoglycans, lignosulfonate, and alginate, as well as charged synthetic polymers and DNA through electrostatic interactions [90,91].

1.7.4 Packaging of food Chitosan has cationic groups on its backbone. It shows antimicrobial properties against bacteria, yeasts, molds, and fungi [92]. Chitosan performs as an antimicrobial compound. The polycationic nature (positive charge) of chitosan interferes with the bacterial metabolism by electrostatic stacking (negative charge) at the cell surface [93]. Chitosan with low Mw enters the cells’ nucleus and blocks the transcription of RNA

Handbook of Chitin and Chitosan

26

1. Chitin and chitosan: origin, properties, and applications

from DNA due to adsorption to DNA molecules. Chitosan also operates as a chelating agent of some essential minerals [94]. Moreover, chitosan treatment offer protection against contamination and microbial spoilage. It has an excellent film formation property with different coating material or membranes that are semipermeable to gases. It is known that these films possess low oxygen permeability [95]. Hence, chitosan and its derivative have unique properties like biodegradability, biocompatibility, nontoxicity, and renewability [96] as well as being inexpensive and commercially available. Cost-effectiveness is needed as the contribution of packaging material and total product costs is highly significant. Chitosan has certain solubility in acetic acid and hydrochloric acid, which can promote the film-forming ability. Chitosan-based films can be fabricated by different methods, such as casting, coating, and layer-by-layer assembly. The characteristics can be modified, that is, thermal stability, antimicrobial activity, mechanical property, barrier property, antioxidant activity, and optical property. However, sometimes other functional materials are added into chitosan to fabricate composite films to increase the combinational advantages of the obtained films. This film is applied to different foods, such as meat, fruit, and vegetables with exceptional preservative effects, displaying the potential as an alternative food packaging [95].

1.7.5 Textile industries The textile industry is one of the largest producers globally. Textile fibers might be divided into synthetic fibers sourced from petroleum and regenerated fibers derived from natural polymers [97]. Moreover, synthetic fibers are restricted by their inherent nonbiodegradability, serious pollution concerns, and the nonrenewable status of petroleum. Almost 8 million tons of plastic enter the ocean every year and create serious marine pollution. Hence, it is necessary to replace fossil raw materials with renewable alternatives that could be biodegraded in soil and ocean [98,99]. Natural polymers like cellulose, chitin, and chitosan derivatives have attracted incredible research interest due to their renewability and environment-friendliness [100]. Antimicrobial treatment is increasingly becoming a standard finish for some textile products as well as for medical, institutional, and hygienic uses. It has become popular in sportswear, women’s wear, and aesthetic clothing to impart antiodor or biostatic properties [101]. Cellulose and protein fibers are more susceptible to microbial attack than man-made fibers in light due to the hydrophilic porous structure and moisture transport characteristics. Hence, the use of antibacterial agents to avoid the growth of bacteria is becoming a standard finishing for textile goods. Moreover, many antibacterial agents are toxic chemicals and lack

Handbook of Chitin and Chitosan

1.7 Application of chitin and chitosan

27

efficiency and durability [102]. Chitosan is a nontoxic, biodegradable, and biocompatible natural polymer with antimicrobial activity. It is a very suitable material to avoid the growth of bacteria and is becoming a standard finishing for textile goods. For example, cotton fabric treated with water-soluble carboxymethyl chitosan shows better antimicrobial activity against E. coli and S. aureus at 0.1% concentration as well as improved wrinkle recovery [103]. Furthermore, chitosan-based core
shell particles with chitosan [poly(n-butyl acrylate) (PBA)] as the core have been designed as novel antibacterial coating for textiles [102]. Cotton treated with PBA-chitosan particles is used for its excellent antibacterial activity against S. aureus with bacterial reductions more than 99% [104].

1.7.6 Bioplastics Waste plastic is become a global problem. New biodegradable plastics are proposed due to with the same strength as ordinary plastics. They must decompose in soil after use and not cause environmental pollution. The period of decomposition can be controlled with low cost. Natural polymers, such as cellulose, are decomposed by microorganisms in soil and decomposed materials do not pollute the environment. Novel biodegradable plastics can be produced by making natural polymers into composites. These materials are cellulose and chitosan (deacetylation of chitin). Moreover chitosan has a structure like cellulose. The chitin molecule is a linear natural polymer that corresponds to cellulose in which C2 is replaced by an acetylamino group. It has various functions, such as biological compatibility, antibiotic activity, and film-forming capability, and has been used as a material in the food industry [105]. Now, new biodegradable plastics have been derived from cellulose and chitosan. Although chitosan is insoluble in water, it is soluble in acetic acid, and is water soluble with a cationic nature. It can be fabricated into different edible films. This chitosan film preserves various foods, such as banana, pomegranate, carrot, mango, tomato, fish, papaya, and wolfberry, with improved storage stability. Furthermore, pure chitosan prevents the growth of microbes as well as shows antioxidant activity, thus prolonging shelf life. Hence, chitosan-based film with dispersions with a diameter of 600 nm demonstrated better preservation of food compared with submicron chitosan dispersions of 1000 nm. Chitosan is added to plasticizers such as glycerol and sorbitol to improve the strength, extendibility, flexibility, and chain mobility. Some vacuum packaging often uses pure chitosan film to preserve seafood, such as swordfish, hake, sea bass, and rainbow trout [10]. Moreover, chitosan and cellulose both have excellent film-forming capacities, which contribute to the formation of the composite film. This film exhibits

Handbook of Chitin and Chitosan

28

1. Chitin and chitosan: origin, properties, and applications

reduced bacterial adhesive for packaging, great antioxidant activity, and an improved water vapor barrier property. Chitosan is also promising for active packaging film.

1.7.7 Nanocomposite Polymers are considered as a good host material for metal and semiconductor nanoparticles, and, on the other hand, exhibit exceptional optical and electrical properties. The synthetic polymers are frequently used in various applications of nanocomposites. Moreover, they have become a major source of waste after use and show poor biodegradability. Hence, biopolymers, such as biodegradable materials like polysaccharides, proteins, and nucleic acids, are used in various nanocomposite applications. Moreover, these materials have some limitations due to their poor mechanical properties. They can be used to improve the properties of biopolymers as a matrix by means of reinforcement techniques. Chitosan is a polysaccharide biopolymer extensively used as a matrix in nanobiocomposites because of its high biocompatibility and biodegradability [106]. Amongst the many linear polymers, chitosan has the most impressive advantages of low cost, nontoxicity, abundant metal-coordination sites (2NH2 and 2 OH), and can be easily cross-linked [107]. Its functional groups support direct binding of enzymes for immobilization. Covalent attachment of functional groups modifies the polymer properties related to hydrophobicity. Its useful forms include gels, scaffolds, beads, fibers, and films. Chitosan is nonconductive, and thus due to this property, it can also be used in biosensors or biofuel cells. Moreover, chitosan
carbon nanotube composite films are used as a conductive matrix supporting direct electron transfer in biosensors. Chitosan can undergo an amidation reaction with the carboxyl groups of graphene to form a homogenous and well dispersed graphene composite. Graphene/chitosan-based composite with improved physicochemical properties is used for sorptionbased applications [108]. Some magnetic nanocomposites are also used for Pb21, Cu21, and Cd21 removal from water [109].

References [1] K. Kurita, Chitin and chitosan: functional biopolymers from Marine crustaceans, Mar. Biotechnol. 8 (3) (2006) 203. [2] J.L. Shamshina, P. Berton, R.D. Rogers, Advances in functional chitin materials: a review, ACS Sustain. Chem. Eng. 7 (7) (2019) 6444
6457. [3] A. Zamani, et al., Determination of glucosamine and N-acetyl glucosamine in fungal cell walls, J. Agric. Food Chem. 56 (18) (2008) 8314
8318. [4] R.A.A. Muzzarelli, et al., Current views on fungal chitin/chitosan, human chitinases, food preservation, glucans, pectins and inulin: a tribute to Henri Braconnot,

Handbook of Chitin and Chitosan

References

precursor [5] [6]

[7]

[8] [9]

[10] [11] [12]

[13]

[14] [15]

[16] [17]

[18] [19]

[20] [21] [22] [23] [24]

29

of the carbohydrate polymers science, on the chitin bicentennial, Carbohydr. Polym. 87 (2) (2012) 995
1012. P.K. Dutta, J. Dutta, V. Tripathi, Chitin and chitosan: Chemistry, properties and applications, J. Scientific & Industrial Res. 63, 2004, 20
31. V. Zargar, M. Asghari, A. Dashti, A review on chitin and chitosan polymers: structure, chemistry, solubility, derivatives, and applications, ChemBioEng Rev. 2 (3) (2015) 204
226. S. Perumal, et al., Spherical chitosan/gelatin hydrogel particles for removal of multiple heavy metal ions from wastewater, Ind. Eng. Chem. Res. 58 (23) (2019) 9900
9907. H.Q. Le, et al., CO2-activated adsorption: a new approach to dye removal by chitosan hydrogel, ACS Omega 3 (10) (2018) 14103
14110. S. Qiu, et al., Preparation and pervaporation property of chitosan membrane with functionalized multiwalled carbon nanotubes, Ind. Eng. Chem. Res. 49 (22) (2010) 11667
11675. H. Wang, J. Qian, F. Ding, Emerging chitosan-based films for food packaging applications, J. Agric. Food Chem. 66 (2) (2018) 395
413. K. Mikula, et al., Controlled release micronutrient fertilizers for precision agriculture
a review, Sci. Total Environ. 712 (2020) 136365. S. Song, et al., Carboxymethyl chitosan modified carbon nanoparticle for controlled emamectin benzoate delivery: improved solubility, pH-responsive release, and sustainable pest control, ACS Appl. Mater. Interfaces 11 (37) (2019) 34258
34267. S. Kim, et al., Chitosan
lysozyme conjugates for enzyme-triggered hydrogel degradation in tissue engineering applications, ACS Appl. Mater. Interfaces 10 (48) (2018) 41138
41145. C. Chen, et al., Multifunctional chitosan inverse opal particles for wound healing, ACS Nano 12 (10) (2018) 10493
10500. A.K. Mahanta, et al., Nanoparticle-induced controlled drug delivery using chitosanbased hydrogel and scaffold: application to bone regeneration, Mol. Pharm. 16 (1) (2018) 327
338. G. Borchard, Chitosans for gene delivery, Adv. Drug. Deliv. Rev. 52 (2) (2001) 145
150. H. Nagahama, et al., Preparation of biodegradable chitin/gelatin membranes with GlcNAc for tissue engineering applications, Carbohydr. Polym. 73 (3) (2008) 456
463. A. Jafari Sanjari, M. Asghari, A review on chitosan utilization in membrane synthesis, ChemBioEng Rev. 3 (3) (2016) 134
158. J. Kegere, et al., Fabrication of poly (vinyl alcohol)/chitosan/Bidens pilosa composite electrospun nanofibers with enhanced antibacterial activities, ACS Omega 4 (5) (2019) 8778
8785. J. Wu, et al., Green and facile preparation of chitosan sponges as potential wound dressings, ACS Sustain. Chem. Eng. 6 (7) (2018) 9145
9152. D. Elieh-Ali-Komi, M.R. Hamblin, Chitin and chitosan: production and application of versatile biomedical nanomaterials, Int. J. Adv. Res. 4 (3) (2016) 411. M.A. Mohammed, et al., An overview of chitosan nanoparticles and its application in non-parenteral drug delivery, Pharmaceutics 9 (4) (2017) 53. J. Kumirska, et al., Application of spectroscopic methods for structural analysis of chitin and chitosan, Mar. Drugs 8 (5) (2010) 1567
1636. Y.I. Cho, H.K. No, S.P. Meyers, Physicochemical characteristics and functional properties of various commercial chitin and chitosan products, J. Agric. Food Chem. 46 (9) (1998) 3839
3843.

Handbook of Chitin and Chitosan

30

1. Chitin and chitosan: origin, properties, and applications

[25] D. Raabe, C. Sachs, P. Romano, The crustacean exoskeleton as an example of a structurally and mechanically graded biological nanocomposite material, Acta Mater. 53 (15) (2005) 4281
4292. [26] M.S. Islam, S. Khan, M. Tanaka, Waste loading in shrimp and fish processing effluents: potential source of hazards to the coastal and nearshore environments, Mar. Pollut. Bull. 49 (1
2) (2004) 103
110. [27] S. Hajji, et al., Structural differences between chitin and chitosan extracted from three different marine sources, Int. J. Biol. Macromol. 65 (2014) 298
306. [28] F. Al Sagheer, et al., Extraction and characterization of chitin and chitosan from marine sources in Arabian Gulf, Carbohydr. Polym. 77 (2) (2009) 410
419. [29] M.H. Mohammed, P.A. Williams, O. Tverezovskaya, Extraction of chitin from prawn shells and conversion to low molecular mass chitosan, Food Hydrocoll. 31 (2) (2013) 166
171. [30] A. Khanafari, R. Marandi, S. Sanati, Recovery of chitin and chitosan from shrimp waste by chemical and microbial methods, J. Environ. Sci. Eng. 5 (1), 2008, 19
24. [31] W. Arbia, et al., Chitin extraction from crustacean shells using biological methods
a review, Food Technol. Biotechnol. 51 (1) (2013) 12
25. [32] S. Kaur, et al., Mucins in pancreatic cancer and its microenvironment, Nat. Rev. Gastroenterol. Hepatol. 10 (10) (2013) 607. [33] J.W. Wong, R.D. Tyagi, A. Pandey, Current Developments in Biotechnology and Bioengineering: Solid Waste Management, Elsevier, 2016. [34] N. Giyose, N. Mazomba, L. Mabinya, Evaluation of proteases produced by Erwinia chrysanthemi for the deproteinization of crustacean waste in a chitin production process, Afr. J. Biotechnol. 9 (5) (2010) 707. [35] G. Jo, et al., Screening of protease-producing Serratia marcescens FS-3 and its application to deproteinization of crab shell waste for chitin extraction, Carbohydr. Polym. 74 (3) (2008) 504
508. [36] Y. Zhu, et al., The role of abscisic acid in early the development, Plant. Mol. Biol. 72 (2010) 1
2. [37] N. Thirunavukkarasu, K. Dhinamala, R.M. Inbaraj, Production of chitin from two marine stomatopods Oratosquilla spp.(Crustacea), J. Chem. Pharm. Res. 3 (1) (2011) 353
359. [38] P. Beaney, J. Lizardi-Mendoza, M. Healy, Comparison of chitins produced by chemical and bioprocessing methods, J. Chem. Technol. Biotechnol. Int. Res. Process, Environ. Clean. Technol. 80 (2) (2005) 145
150. [39] A. Zamani, M. Taherzadeh, Production of low molecular weight chitosan by hot dilute sulfuric acid, BioResources 5 (3) (2010) 1554
1564. [40] D. Thomas, S. Thomas, Chemical modification of chitosan and its biomedical application, Biopolym. Nanocomp. Process. Prop. Appl. (2013) 33e51. [41] R. Raval, K. Raval, B. Moerschbacher, Enzymatic modification of chitosan using chitin deacetylase isolated from Bacillus cereus, Open Access. Sci. Rep. 2 (1) (2013) 1
4. [42] M.G.B. Pagnoncelli, et al., Chitosanase production by Paenibacillus ehimensis and its application for chitosan hydrolysis, Braz. Arch. Biol. Technol. 53 (6) (2010) 1461
1468. [43] S. Sinha, S. Chand, P. Tripathi, Production, purification and characterization of a new chitosanase enzyme and improvement of chitosan pentamer and hexamer yield in an enzyme membrane reactor, Biocatal. Biotransform. 32 (4) (2014) 208
213. [44] W. Xia, P. Liu, J. Liu, Advance in chitosan hydrolysis by non-specific cellulases, Biores. Technol. 99 (15) (2008) 6751
6762. [45] N.K. Mathur, C.K. Narang, Chitin and chitosan, versatile polysaccharides from marine animals, J. Chem. Edu. 67 (11) (1990) 938.

Handbook of Chitin and Chitosan

References

31

[46] C.J. Brine, P.R. Austin, Chitin variability with species and method of preparation, Comp. Biochem. Physiol. B: Comp. Biochem. 69 (2) (1981) 283
286. [47] P. Sahariah, M. Masson, Antimicrobial chitosan and chitosan derivatives: a review of the structure
activity relationship, Biomacromolecules 18 (11) (2017) 3846
3868. [48] W. Bough, et al., Influence of manufacturing variables on the characteristics and effectiveness of chitosan products. II. Coagulation of activated sludge suspensions, Biotechnol. Bioeng. 20 (12) (1978) 1945
1955. [49] K.Y. Lee, W.S. Ha, W.H. Park, Blood compatibility and biodegradability of partially N-acylated chitosan derivatives, Biomaterials 16 (16) (1995) 1211
1216. [50] P. Morganti, et al., From waste material a new anti aging compound: a chitin nanofi¨ FW J. 138 (7) (2012). ber complex, SO [51] K.T. Hwang, et al., Controlling molecular weight and degree of deacetylation of chitosan by response surface methodology, J. Agric. Food Chem. 50 (7) (2002) 1876
1882. [52] A. Tolaimate, et al., Contribution to the preparation of chitins and chitosans with controlled physico-chemical properties, Polymer 44 (26) (2003) 7939
7952. [53] A. Ilyina, et al., Enzymic preparation of acid-free-water-soluble chitosan, Process. Biochem. 35 (6) (2000) 563
568. ¨ zogul, J.M. Regenstein, Industrial applications of crustacean by[54] I. Hamed, F. O products (chitin, chitosan, and chitooligosaccharides): a review, Trends Food Sci. Technol. 48 (2016) 40
50. [55] S. Mima, et al., Highly deacetylated chitosan and its properties, J. Appl. Polym. Sci. 28 (6) (1983) 1909
1917. [56] I. Bano, et al., Chitosan: a potential biopolymer for wound management, Int. J. Biol. Macromol. 102 (2017) 380
383. [57] M.J. Song, et al., Drug-eluting bead loaded with doxorubicin versus conventional lipiodol-based transarterial chemoembolization in the treatment of hepatocellular carcinoma: a case
control study of Asian patients, Eur. J. Gastroenterol. Hepatol. 23 (6) (2011) 521
527. [58] C.-Y. Chiu, et al., Functional comparison for lipid metabolism and intestinal and fecal microflora enzyme activities between low molecular weight chitosan and chitosan oligosaccharide in high-fat-diet-fed rats, Mar. Drugs 15 (7) (2017) 234. [59] K. Luo, et al., Mucoadhesive and elastic films based on blends of chitosan and hydroxyethylcellulose, Macromol. Biosci. 8 (2) (2008) 184
192. [60] V.C. Souza, G.L. Dotto, Application of chitosan based materials as edible films and coatings, Frontiers in Biomaterials: Chitosan Based Materials and Its Applications, vol. 3, 2017, pp. 249
271. [61] D. Chattopadhyay, M.S. Inamdar, Aqueous behaviour of chitosan, Int. J. Polym. Sci. 2010 (2010). [62] W. Wang, D. Xu, Viscosity and flow properties of concentrated solutions of chitosan with different degrees of deacetylation, Int. J. Biol. Macromol. 16 (3) (1994) 149
152. [63] S. Lu, et al., Preparation of water-soluble chitosan, J. Appl. Polym. Sci. 91 (6) (2004) 3497
3503. [64] Y.-W. Cho, et al., Preparation and solubility in acid and water of partially deacetylated chitins, Biomacromolecules 1 (4) (2000) 609
614. [65] J.C. Roy, et al., Solubility of chitin: solvents, solution behaviors and their related mechanisms, in: Z. Xu (Ed.), Solubility of Polysaccharides, InTech, Vienna, 2017, pp. 109
127. [66] C. Pillai, W. Paul, C.P. Sharma, Chitin and chitosan polymers: chemistry, solubility and fiber formation, Prog. Polym. Sci. 34 (7) (2009) 641
678. [67] H. Saitoˆ, et al., A high-resolution solid-state 13C NMR study of the secondary structure of linear (1-3)-β-D-glucans: a conformational elucidation of noncrystalline and

Handbook of Chitin and Chitosan

32

[68] [69] [70] [71] [72]

[73] [74] [75]

[76]

[77] [78]

[79] [80] [81] [82] [83] [84] [85] [86]

[87]

[88] [89]

1. Chitin and chitosan: origin, properties, and applications

crystalline forms by means of conformation-dependent 13C chemical shifts, Bull. Chem. Soc. Jpn. 60 (12) (1987) 4259
4566. D. Knorr, Functional properties of chitin and chitosan, J. Food Sci. 47 (2) (1982) 593
595. M. Kaya, et al., On chemistry of γ-chitin, Carbohydr. Polym. 176 (2017) 177
186. M. Fernandes Queiroz, et al., Does the use of chitosan contribute to oxalate kidney stone formation? Mar. Drugs 13 (1) (2015) 141
158. R.L. Lavall, O.B. Assis, S.P. Campana-Filho, β-Chitin from the pens of Loligo sp.: extraction and characterization, Biores. Technol. 98 (13) (2007) 2465
2472. D. Silva, F. Zuluaga, H. Carlos, Evaluation of biocompatibility of chitosan films from the mycelium of Aspergillus niger in connective tissue of Rattus norvegicus, J. Mol. Genet. Med. 9 (3) (2015) 1
8. A.T. Paulino, et al., Characterization of chitosan and chitin produced from silkworm crysalides, Carbohydr. Polym. 64 (1) (2006) 98
103. E.S. Abdou, K.S. Nagy, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from local sources, Bioresour. Technol. 99 (5) (2008) 1359
1367. R.S.C. De Queiroz Antonino, et al., Preparation and characterization of chitosan obtained from shells of shrimp (Litopenaeus vannamei Boone), Mar. Drugs 15 (5) (2017) 141. Y. Gao, Q. Yue, B. Gao, High surface area and oxygen-enriched activated carbon synthesized from animal cellulose and evaluated in electric double-layer capacitors, RSC Adv. 5 (40) (2015) 31375
31383. R. Devi, R. Dhamodharan, Pretreatment in hot glycerol for facile and green separation of chitin from prawn shell waste, ACS Sustain. Chem. Eng. 6 (1) (2018) 846
853. B. Bradi´c, et al., A reaction
diffusion kinetic model for the heterogeneous N-deacetylation step in chitin material conversion to chitosan in catalytic alkaline solutions, React. Chem. Eng. 3 (6) (2018) 920
929. P.A. Sandford, A. Steinnes, Biomedical Applications of High-Purity Chitosan: Physical, Chemical, and Bioactive Properties, ACS Publications, 1991. M. Rinaudo, A. Domard, Solution properties of chitosan, Chitin and Chitosan, Elsevier, London, 1989, pp. 71
86. X. Wang, et al., Exploration of blood coagulation of N-alkyl chitosan nanofiber membrane in vitro, Biomacromolecules 19 (3) (2018) 731
739. E. Ruel-Garie´py, J.-C. Leroux, Chitosan: A Natural Polycation with Multiple Applications, ACS Publications, 2006. I. Ali, New generation adsorbents for water treatment, Chem. Rev. 112 (10) (2012) 5073
5091. H.K. No, S.P. Meyers, Application of chitosan for treatment of wastewaters, Reviews of Environmental Contamination and Toxicology, Springer, 2000, pp. 1
27. R. Vidal, J. Moraes, Removal of organic pollutants from wastewater using chitosan: a literature review, Int. J. Environ. Sci. Technol. 16 (3) (2019) 1741
1754. L.A. Picos-Corrales, et al., Environment-friendly approach toward the treatment of raw agricultural wastewater and river water via flocculation using chitosan and bean straw flour as bioflocculants, ACS Omega 5 (8) (2020) 3943
3951. Z.M. Ali, et al., Extraction and characterization of chitosan from Indian prawn (Fenneropenaeus indicus) and its applications on waste water treatment of local ghee industry, Extraction 3 (10) (2013). X. Li, Principles of Fuel Cells, Taylor & Francis Group, New York, 2006. J. Ma, Y. Sahai, Chitosan biopolymer for fuel cell applications, Carbohydr. Polym. 92 (2) (2013) 955
975.

Handbook of Chitin and Chitosan

References

33

[90] B.-K. Chen, et al., Improving the conductivity of sulfonated polyimides as proton exchange membranes by doping of a protic ionic liquid, Polymers 6 (11) (2014) 2720
2736. [91] M. Dash, et al., Chitosan—a versatile semi-synthetic polymer in biomedical applications, Prog. Polym. Sci. 36 (8) (2011) 981
1014. [92] M. Friedman, V.K. Juneja, Review of antimicrobial and antioxidative activities of chitosans in food, J. Food Prot. 73 (9) (2010) 1737
1761. [93] M. Benhabiles, et al., Antibacterial activity of chitin, chitosan and its oligomers prepared from shrimp shell waste, Food Hydrocoll. 29 (1) (2012) 48
56. [94] R.C. Goy, Dd Britto, O.B. Assis, A review of the antimicrobial activity of chitosan, Polı´meros 19 (3) (2009) 241
247. [95] M. Aider, Chitosan application for active bio-based films production and potential in the food industry, LWT-Food Sci. Technol. 43 (6) (2010) 837
842. [96] M. Barikani, et al., Preparation and application of chitin and its derivatives: a review, Iran. Polym. J. 23 (4) (2014) 307
326. [97] K. Zhu, et al., Cellulose/chitosan composite multifilament fibers with two-switch shape memory performance, ACS Sustain. Chem. Eng. 7 (7) (2019) 6981
6990. [98] S. Chandra, N. Karak, Environmentally friendly polyurethane dispersion derived from dimer acid and citric acid, ACS Sustain. Chem. Eng. 6 (12) (2018) 16412
16423. [99] Y. Zhu, C. Romain, C.K. Williams, Sustainable polymers from renewable resources, Nature 540 (7633) (2016) 354
362. [100] B. Duan, et al., Recent advances in chitin based materials constructed via physical methods, Prog. Polym. Sci. 82 (2018) 1
33. [101] E.-R. Kenawy, S. Worley, R. Broughton, The chemistry and applications of antimicrobial polymers: a state-of-the-art review, Biomacromolecules 8 (5) (2007) 1359
1384. [102] W. Ye, et al., Novel core-shell particles with poly (n-butyl acrylate) cores and chitosan shells as an antibacterial coating for textiles, Polymer 46 (23) (2005) 10538
10543. [103] D. Gupta, A. Haile, Multifunctional properties of cotton fabric treated with chitosan and carboxymethyl chitosan, Carbohydr. Polym. 69 (1) (2007) 164
171. [104] A. Itthagarun, N.M. King, Y.-M. Cheung, The effect of nano-hydroxyapatite toothpaste on artificial enamel carious lesion progression: an in-vitro pH-cycling study, Hong Kong Dent. J. 7 (2) (2010) 61
66. [105] M. Nishiyama, et al., Biodegradable Plastics Derived from Cellulose Fiber and Chitosan, ACS Publications, 1996. [106] Z. Moridi, V. Mottaghitalab, A. Haghi, A detailed review of recent progress in carbon nanotube/chitosan nanocomposites, Cellulose Chem. Technol. 45 (9) (2011) 549. [107] K. Li, et al., Selective adsorption of Gd3 1 on a magnetically retrievable imprinted chitosan/carbon nanotube composite with high capacity, ACS Appl. Mater. Interfaces 7 (38) (2015) 21047
21055. [108] M. Sabzevari, D.E. Cree, L.D. Wilson, Graphene oxide
chitosan composite material for treatment of a model dye effluent, ACS Omega 3 (10) (2018) 13045
13054. [109] X. Liu, et al., Magnetic chitosan nanocomposites: a useful recyclable tool for heavy metal ion removal, Langmuir 25 (1) (2009) 3
8.

Handbook of Chitin and Chitosan

C H A P T E R

2 Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications Hakima El Knidri, Ali Laajeb and Ahmed Lahsini Catalysis, Materials and Environment Laboratory, Higher School of Technology, Sidi Mohamed Ben Abdellah University, Fez, Morocco O U T L I N E 2.1 Introduction

36

2.2 Chitin and chitosan: chemistry and solubility 2.2.1 Chitin and chitosan chemistry 2.2.2 Chitin and chitosan extraction 2.2.3 Chitin and chitosan solubility

36 36 38 39

2.3 Chitin and chitosan: fiber formation 2.3.1 Electrospinning process 2.3.2 Factors affecting the electrospinning process 2.3.3 Characterization of chitin and chitosan fibers 2.3.4 Applications of chitin and chitosan fibers

41 42 42 45 46

2.4 Conclusions

50

References

50

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00002-X

35

© 2020 Elsevier Inc. All rights reserved.

36

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

2.1 Introduction Chitin was discovered in 1811 by the chemist Henri Braconnot. It was isolated for the first time from a certain type of mushroom and was named “fungine.” Braconnot observed that this substance is insoluble in aqueous acid solutions. In 1823 Auguste Odier isolated the same fibrous substance from insect exoskeletons and named it chitin, a Greek term meaning tunic or envelope [1 5]. In 1859 Charles Rouget subjected the chitin to an alkaline treatment by using concentrated potash at high temperature; the resulting product is soluble in aqueous acid solutions and this property has allowed differentiating between this new substance and chitin polymer. In 1894 this molecule was named chitosan by the German chemist Fe´lix HoppeSeyler [1,2,6]. From the 1930s chitin and chitosan aroused great interest, mainly in the medical field and in water purification, since they are renewable resources that can be found in abundance in nature. Since 1970 these two biopolymers have begun to be of real interest because of the large quantities of waste produced by the marine processing industries and the crustaceans canneries. Currently industrial production, marketing, and use of these two biopolymers are constantly increasing.

2.2 Chitin and chitosan: chemistry and solubility 2.2.1 Chitin and chitosan chemistry Chitin is the second most abundant biological macromolecule in nature after cellulose; it is a linear amino polysaccharide consisting mainly of β-(1-4)-linked 2-acetamido-2-deoxy-β-D-glucopyranose units and partially of β-(1-4)-linked 2-amino-2-deoxy-β-D-glucopyranose units, this biopolymer is estimated to be produced annually in an amount almost as large as cellulose [7 10]. Chitin is mainly found in the exoskeleton of crustaceans, insect cuticles, and other arthropod shells, however, it is also present in the vast majority of fungi, some mushroom envelopes, green algae, fish scales, cell walls, and yeasts. The main commercial sources of chitin are crab and shrimp shells, which contain a high percentage of chitin [8,11 13]. In general, crustacean shells consist of 30% 50% calcium carbonate and phosphate, 30% 40% proteins, and 20% 30% chitin, as reported by Kumirska et al. [10,14], but these percentages vary depending on the source, or even the species, from which chitin is extracted, for example,

Handbook of Chitin and Chitosan

2.2 Chitin and chitosan: chemistry and solubility

37

Crangon crangon shrimp waste consists of 10% 38% proteins, 31% 44% minerals, and 24% 46% chitin, according to Bajaj and coworkers [15]. Depending on its source, chitin exists as three polymorphic forms in nature, namely, α-chitin, β-chitin, and γ-chitin. The α and β forms are arranged according to monoclinic and orthorhombic cells, respectively, and the third allomorph, γ-chitin, appears to be a combination of α and β forms rather than a different allomorph [10,16 18]. The α-chitin, β-chitin, and γ-chitin correspond to antiparallel, parallel, and alternated arrangements of polymer chains, respectively. The α-chitin is the most abundant and stable polymorph and the β-chitin can be extracted from squid pens and easily converted to α-form by alkaline treatment [10,16,19]. The α-chitin is usually isolated from the exoskeleton of crustaceans, yeast cell walls, and arthropod cuticles, in general [20,21]. For commercial applications, this polymer is mainly extracted from crustacean shells, that is, shrimp and crab, by acid treatment followed by alkaline treatment to remove the calcium carbonates and proteins, respectively. In addition, a decolorization and purification step are often added to eliminate pigments and obtain a high-quality product without impurities [3,22]. Chitosan, the main derivative of chitin, is also a natural polymer obtained by N-deacetylation of chitin under alkaline conditions. This biopolymer is a linear polysaccharide consisting of 2-amino-2-deoxy(1-4)-β-D-glucopyranose residues (D-glucosamine units). A nomenclature border has been defined between chitin and chitosan based on the degree of N-deacetylation. When the acetylation degree is higher than 50%, the polymer is named chitin, and when the acetylation degree is less than 50%, the polymer is named chitosan [6,13]. The structures of chitin and chitosan are presented in Fig. 2.1. Chitosan has attracted, as a renewable biopolymer, considerable attention due to its remarkable properties such as biodegradability, biocompatibility, nontoxicity, ease of chemical modifications, and the excellent chelating ability. This polymer has attracted significant interest in various fields such as chemistry, biochemistry, biology, biotechnology, medicine, pharmacology, food science, marine science, agriculture, and many other related areas [23 26].

FIGURE 2.1 Chemical structure of chitin (A) and chitosan (B).

Handbook of Chitin and Chitosan

38

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

Chitosan has wide applications in medical and pharmaceutical fields, such as wound dressing, drug delivery systems, and as a blood anticoagulant, in addition to other fields such as water purification, wastewater treatment, food additives, wound-healing materials, tissue engineering, cosmetic preparations, textile industry, paper production, and film technologies [23,24,27 29].

2.2.2 Chitin and chitosan extraction The chitin extraction process and its subsequent conversion to chitosan can be carried out using two types of methods: chemical methods and biological methods such as microbial fermentation and enzymatic reactions [30 32]. In addition, many researchers have studied a wide variety of extraction techniques, such as electrochemical, sonochemical, enzymatic, and microwave synthesis methods, to find an alternative method of chitin and chitosan extraction that is efficient, speedy, and environment friendly [33 35]. It has been shown that microwave technology can be very useful to extract chitin and chitosan. Compared with conventional extraction, microwave synthesis allowed a dramatic reduction in reaction time, increasing the yield and the purity of product by reducing unwanted side reactions [36,37]. The extraction process of chitosan involves the following steps: demineralization, deproteinization, and deacetylation. A decolorization step can be added to remove pigment, mainly β-carotene and astaxanthin, using various organic and inorganic solvents, such as acetone, sodium hypochlorite, and hydrogen peroxide [6,10,14,25,38,39]. The extraction process of chitin and chitosan is shown in Fig. 2.2. Demineralization: the demineralization step consists of removing the mineral constituents, that is, calcium carbonate and calcium phosphate, which constitute the main inorganic compounds of the exoskeleton of crustaceans. This step is performed in dilute hydrochloric acid solution. During the digestion reaction, the emission of CO2 gas is more or less an important indicator of the content of mineral constituents [1].

FIGURE 2.2 Extraction process of chitin and chitosan.

Handbook of Chitin and Chitosan

2.2 Chitin and chitosan: chemistry and solubility

39

FIGURE 2.3 Chemical deacetylation of chitin.

Deproteinization: the deproteinization step consists of eliminating proteins with alkaline treatment using dilute sodium hydroxide (NaOH) solution. The product obtained is designated as purified chitin. A decolorization step can be added if a colorless chitin is wanted. Acetone or organic solvent mixtures are used to remove the residual pigments such as carotenoids. It was found that proteins extracted from shrimp shell wastes can be used as an excellent source of animal feed [1,40]. Deacetylation: the deacetylation step consists of transforming chitin into chitosan by the removal of the acetyl group. The deacetylation of chitin is generally achieved by alkaline treatment using concentrated sodium or potassium hydroxide solution at high temperature. The deacetylation reaction is shown in Fig. 2.3. Chitosan polymer is mainly characterized by two parameters: the deacetylation degree (DD) and the molecular weight (Mw). The DD of chitosan and the Mw are the most important parameters as they considerably affect the physicochemical properties. The utilization of chitosan is related to the DD and Mw of the biopolymer [32,41]. During the extraction process, several factors, such as the temperature of treatment, the concentration of alkali, the reaction time, and the previous treatment of the chitin, may influence the Mw of chitosan.

2.2.3 Chitin and chitosan solubility Solubility is one of the most important parameters that affect the processing and the applications of polymers. In general, the water solubility of crystalline polysaccharides such as chitosan and cellulose is poor. Several strategies such as degradation and modification have been adopted to improve the solubility of these polymers. The semicrystalline structure of chitin and the presence of a strong intra- and intermolecular hydrogen bonding network provides the insoluble property of this polymer, in common organic and inorganic solvents [16,17]. However, the insolubility of this polymer is regarded as a hindrance for its potential applications. Chitin is insoluble in common solvents such as water, mild acidic or alkali solution, and organic solvents. However, it is soluble in highly concentrated inorganic acids such as hydrochloric acid, sulfuric acid,

Handbook of Chitin and Chitosan

40

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

and phosphoric acid, and its β-polymorph form dissolves in concentrated formic acid. In addition, chitin solubility has been verified in many other solvents such as N,N-dimethylacetamide DMA containing 5% 8% of LiCl, dichloro- and trichloroacetic acids, lithium thiocyanate, CaBr2 H2O saturated methanol, hexafluoroisopropyl alcohol, hexafluoracetone, and N-methyl-2-pyrrolidone. Another solvent system, methanol saturated with calcium chloride dihydrate, also has been used to dissolve chitin polymer. Although dissolution of chitin is possible by these solvents, many of them are toxic, corrosive, harmful, or scarcely degradable [4,22,42 46]. The most important derivative of chitin is chitosan, composed of N-acetyl-D-glucosamine and D-glucosamine units [26,47]. Each D-glucosamine unit contains a free amino group, these groups can take on a positive charge, leading to protonating the polymer and subsequently providing solubility of chitosan in diluted acidic solutions [28,48,49]. Chitosan is insoluble in either organic solvents or water, however, it is soluble in most aqueous acid solutions, such as acetic, citric, formic, oxalic, and lactic acids, below its pKa (pH 5 6.5), and in some other solvents such as dimethyl sulfoxide, p-toluene sulfonic acid, and 10-camphorsulfonic acid [46,50,51]. Some mineral acids, such as hydrochloric and nitric acids, are also used for chitosan dissolution, but phosphoric and sulfuric acids are not suitable [4]. Many factors affecting chitin and chitosan solubility are described in the literature, mainly, the DD, Mw, pH of solution, ionic strength, and temperature. The relationship between solubility, Mw, and DD has been established by several groups. The solubility of chitin does not depend on its Mw, but is related to the acetylation degree described by the number of N-acetyl amino groups [50]. In addition, the presence of reactive groups, such as amine groups and hydroxyls groups, in chitin and chitosan, allow them to undergo many chemical modifications. This chemical modification has been used as a means of imparting solubility to chitin and chitosan by using appropriate chemical entities that enhance solubility [52]. The chemical modifications of molecular structure of these polymers can enhance their solubility in water and other solvents. For example, monomethyl-modified chitosan is soluble in water, chitin-graftpolystyrene is soluble in dimethyl sulfoxide DMSO, O-alkylated chitosan is soluble in ethanol, chloroform, water, and acetic acid [50,53]. Quaternization, phosphorylation, and carboxymethylation of chitosan are chemical modifications that significantly improve the solubility of this amino polysaccharide in different solvents at ambient conditions. The N-acylation method can also be used to enhance the solubility of chitin and chitosan. Sashiwa and coworkers showed that simple acylation enhanced chitosan solubility [54].




Handbook of Chitin and Chitosan

2.3 Chitin and chitosan: fiber formation

41

As reported in the literature, N-methylene phosphonic chitosan is soluble in water and acidic solutions such as acetic and hydrochloric acid. Chitosan dendrimer hybrid, N,N-dicarboxymethyl chitosan, N, O-carboxymethyl chitosan, and N-[(2-hydroxy-3trimethylammonium) propyl] chitosan chloride (HTACC) are also soluble in water [50]. In addition many preparations of water-soluble chitosan are reported in the literature. Xia and coworkers prepared water-soluble chitosan by hydrolysis using hydrogen peroxide; Minh et al. prepared water-soluble hydrochloric chitosan from low-molecular-weight chitosan in the solid state; Fu et al. used a facile physical approach to make chitosan soluble in acid-free water, it is a physical dissolution precipitation process; Silva and coworkers prepared water-soluble chitosan derivatives by selective C-6 oxidation mediated by TEMPO-laccase redox system [55 58].

2.3 Chitin and chitosan: fiber formation Fiber formation is generally made via three typical processes: melt spinning, dry spinning, and wet spinning. These conventional fiber spinning techniques can produce polymer fibers with diameters down to the micrometer range. All three methods require a homogeneous phase for spinning [59 61]. The melt spinning method requires a polymer melt, which is impossible in the case of chitosan as it has a melting point higher than its degradation temperature. The strong interchain forces, as derived from the amino and the hydroxyl groups, raise the melting point of chitin and chitosan to well above their thermal decomposition temperatures, and thus fiber formation by the melt spinning technique is not possible in the case of chitin and chitosan [60]. Additionally, in the dry spinning method, a polymer solution must be prepared, with the added restriction that the solvent be sufficiently volatile to be removed during spinning. As chitin and chitosan polymers can only be dissolved in polar solvents with high boiling temperatures, because of the strong polar groups in these two polymers, the fiber formation by dry spinning, which produces fibers upon the evaporation of the solvent during extrusion, seems likely to be difficult [60]. Chitin and chitosan fibers formation using the spinning technique can be undertaken successfully only with the wet spinning method. This typically involves polymer dissolution by using the spinning solution, followed by extrusion of the purified dope into a coagulation bath with subsequent washing, drawing, and drying [60,62].

Handbook of Chitin and Chitosan

42

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

2.3.1 Electrospinning process Electrospinning is a simple, low-cost, and versatile technique that can be used to generate nanoscale fibers from a wide variety of synthetic and natural polymers. This technique has emerged as the most popular method of chitosan nanofibers preparation due to the relative ease and flexibility it offers. More recently, electrospinning has replaced the conventional spinning methods, that is, melt, wet, and dry spinning. This technique has been developed as a novel method to produce polymeric fibers of nanometric size, creating mats with a distinctly high surface area to mass ratio. In the electrospinning method, contrary to conventional fiber spinning technologies, an electrical force is used to elongate a polymer jet into nanometer-size fibers. As illustrated in Fig. 2.4, the fibers or nanofibers prepared by this method are formed from a liquid polymer solution that is fed via a capillary tube into a region of an electric field generated by a high-voltage power source to the capillary tube [63 68]. Electrospinning of chitosan for the preparation of nanofibers is considered to be a promising technique that has attracted much interest in recent years. Besides their biocompatibility and antimicrobial activity, the electrospun chitosan nanofibers exhibit important properties including a small diameter, a high surface area, and a high porosity, as reported by Arkoun [69].

2.3.2 Factors affecting the electrospinning process The electrospinning technique is governed by a number of parameters that can play important roles in the fiber formation; they can greatly affect the desired nanofiber size and the final microstructure. These parameters are generally classified as electrospinning parameters, solution parameters, and ambient parameters [70 73].

FIGURE 2.4 Schematic illustration of the electrospinning process.

Handbook of Chitin and Chitosan

2.3 Chitin and chitosan: fiber formation

43

2.3.2.1 Electrospinning parameters Processing parameters that can affect electrospinning during the process of fiber formation include voltage applied between the two electrodes, flow rate at which polymer solution is ejected, distance between the needle and the collector, and needle diameter. During the electrospinning process, the applied electric field is one of the most important processing parameters that can greatly affect fiber formation and structure. Current flow from a high-voltage power supply into a polymer solution via a capillary will cause a spherical droplet to deform into a Taylor cone and form ultrafine nanofibers at a critical voltage, as shown in Fig. 2.5. The fiber formation occurs only after achievement of the critical value of applied voltage, this induces the necessary charges in the solution along with the electric field and initiates the electrospinning process. Suboptimal field strength could lead to bead defects in the spun fibers or even failure in jet formation. An increase in the applied voltage beyond the critical value will result in the formation of beads or beaded nanofibers [70,72,74]. The flow rate of the polymer has also an impact on fiber size and can influence fiber porosity, as well as the morphological structure of the fiber. At the capillary tip, the cone shape cannot be maintained if the flow rate of the polymer solution is insufficient to replace the solution ejected as the fiber jet. A lower flow rate is more preferable as the solvent will get enough time for evaporation. On the other hand, it has been shown that the fiber size and the pore diameter increase with an increase in the polymer flow rate in the case of polystyrene fibers [72,74 76]. Additionally, the distance between the capillary tip and the collector can influence fiber diameter and morphology. It has been found that the fiber diameter decreases with increasing distance from the capillary tip, and a minimum distance is required to give fibers a long enough time to dry before arriving at the collector [72,74,77].

FIGURE 2.5 Three principal stages of deformation of polymer solution droplet under the influence of increasing electric field.

Handbook of Chitin and Chitosan

44

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

2.3.2.2 Solution parameters A number of solution parameters can strongly affect the electrospinning process and present an important role in fiber formation and microstructure; these mainly include the parameters that can be controlled during the formation of the polymer solution, such as the choice of solvent with appropriate volatility, the Mw, the viscosity, the surface tension, the conductivity, and the polymer concentration. Depending on several solution parameters, very different results can be obtained using the same polymer and electrospinning process [70,72]. The polymer concentration determines the spinnability of chitin and chitosan solution. However, the solution cannot be either too concentrated or too dilute. A critical concentration is required in order to obtain fibers from electrospinning. Below this concentration, chain entanglements are insufficient to stabilize the repulsion within the ejected jet, leading to the formation of sprayed droplets. The polymer concentration influences both the viscosity of the solution and the surface tension. These two parameters play important roles in determining the range of concentration from which continuous fibers can be obtained in the electrospinning process [72,74,78]. Similar to other polymers, the viscosity of chitosan solutions change depending on the Mw of chitosan, the concentration of the polymer in solution, and finally the type of solvent [78]. In addition, the Mw of the polymer is an important solution parameter that affects the morphology of electrospun fiber. This parameter has a significant effect on rheological and electrical properties such as conductivity, viscosity, surface tension, and dielectric strength [72]. The choice of the solvent is one of the important factors that affect the fibers formation during the electrospinning process. The formation of beadless electrospun nanofiber requires the use of solvent with certain volatility. In general, volatile solvents are preferred as their high evaporation rates encourage the easy evaporation of the solvent from the fibers during their travel from the needle tip to the collector [72,74]. 2.3.2.3 Ambient parameters Beside the electrospinning and solution parameters, many researchers have evaluated the influence of the ambient parameters on the electrospinning process and the fiber size and microstructure. These studies reveal that ambient parameters, especially humidity and temperature, can affect the electrospinning process and subsequently the fiber formation [72,79]. According to Kim and coworkers, the increase of the temperature to the boiling point of the solvent used, leads to the introduction of pores to the nanofibers, owing to the evaporation of the solvent molecules

Handbook of Chitin and Chitosan

2.3 Chitin and chitosan: fiber formation

45

present on the surface of the fibers. They reported that a further increase in the temperature resulted in an increase in the number and the size of the pores on the fibers due to an accelerated evaporation of the solvent [80]. Mit-uppatham et al. mentioned that there is an inverse relationship between temperature and viscosity; they found that when temperature increased, the fiber diameter decreased and they attributed this decline in diameter to the decrease in the viscosity of the polymer solutions at increased temperatures [81]. Studies into the effects of humidity on the electrospinning process show that an increase in humidity leads to the appearance of small pores on the surface of the fibers, as reported by Casper et al. and Bhardwaj et al. [72,82]. High humidity will lead to thick fibers or nanofibers of enlarged diameter. Temperature and humidity are interrelated parameters embroiled in the technique of fiber formation. An increase in temperature will decrease the humidity and evaporate the solvent faster [79].

2.3.3 Characterization of chitin and chitosan fibers Chitin and chitosan fibers can be characterized by using various types of instruments and analysis techniques, such as infrared spectroscopy, X-ray diffraction, nuclear magnetic resonance, energy-dispersive X-ray spectroscopy, and X-ray photoelectron spectroscopy. [61,83]. Fourier-transform infrared spectroscopy is one of the most widely used characterization techniques. It can be used in quantitative analysis and structure determination of chitin and chitosan fibers. This technique is useful for analyzing specific functional groups present in the polymer structure. In addition, infrared spectroscopy allows the estimation of the DD of chitosan. Raman spectroscopy is an analytical technique complementary to the infrared spectroscopy; it can be used to observe vibrational, rotational, and other low-frequency modes in a structure. This spectroscopic analysis is used to provide a fingerprint by which molecules can be identified. X-ray diffraction is a tool for the investigation of the fine structure of electrospun fibers, in which the crystalline atoms cause a beam of incident X-rays to diffract into many specific directions. It can be used for the determination of the crystal structure of the sample. Scanning electron microscopy is a characterization technique used to study the morphology of the sample and the determination of elemental composition when coupled with energy-dispersive spectroscopy. In addition, X-ray photoelectron spectroscopy is a surface-sensitive quantitative spectroscopic tool that measures elemental composition in a chosen area.

Handbook of Chitin and Chitosan

46

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

Nonetheless, other analytical techniques, including nuclear magnetic resonance spectroscopy, size exclusion chromatography, thermal analysis, fluorescence, and UV, are simple, rapid, convenient, and have gained increased popularity for the characterization of chitin and chitosan fibers [61,84].

2.3.4 Applications of chitin and chitosan fibers Chitin and chitosan are natural biopolymers that have various properties such as antimicrobial activity, hemostatic agent, metal-chelating, molecular affinity, and wound-healing agent. For a wide range of uses of these polymers, different modifications have been studied by different researchers. Electrospun fibers of chitin and chitosan could therefore be of great interest in various application fields such as tissue engineering, biomedical, cosmetics, and many industrial applications, such as wastewater treatment [41,85 88] 2.3.4.1 Chitin and chitosan fibers for biomedical applications Chitin and chitosan fibers have been successfully used by several researchers in various pharmaceutical and biomedical applications. Those fibers have been used in developing drug delivery systems, preparing biodegradable bandages, as inert diluents for drugs, as wound dressing and scaffolds for tissue engineering, as well as many other potential uses [89 91]. Ignatova et al. reported that chitosan nanofibers are promising for wound-healing applications. Quaternized chitosan-containing nanofibers have been successfully prepared by Ignatova and coworkers using electrospinning of mixed aqueous solutions of quaternized chitosan and polyvinyl alcohol PVA. A microbiological screening demonstrates the antibacterial activity of the photocross-linked electrospun mats against Escherichia coli and Staphylococcus aureus [89]. The results obtained by Cremar et al. indicate that the chitosan-based nanofibers produced can serve as potential wound dressing materials given their antimicrobial activity and the similarity to the extracellular matrix, which promotes cell adhesion and growth. Homogeneous chitosan fibers have been successfully produced using the centrifugal spinning technology and the antibacterial effect against S. aureus and the cytotoxicity of the developed composite fiber mats have been evaluated [65]. Zhou and coworkers indicated that quaternized chitosan fibers can be used as wound dressing for skin regeneration. The quaternizationfunctionalized chitosan fibers have been successfully prepared by using 2,3-epoxypropyl trimethyl ammonium chloride as a quaternized reagent reacted with chitosan fiber. The results of antibacterial tests showed that

Handbook of Chitin and Chitosan

2.3 Chitin and chitosan: fiber formation

47

quaternized chitosan fiber had good antibacterial activity toward S. aureus [92]. Abdelgawad and coworkers prepared antibacterial wound dressing nanofiber mats from multicomponent systems: chitosan, silver nanoparticles Ag-NPs, and polyvinyl alcohol PVA. The nanofiber mats were obtained by the electrospinning method and cross-linked with glutaraldehyde. The antibacterial experiment indicated that the nanofiber mats prepared had a good bactericidal activity against the Gram-negative bacteria E. coli [91]. Guibal et al. synthesized very efficient antibacterial supports by using chitosan associated with cellulose fibers; they can be used for elaborating foam structures for the binding of silver ions according to Guibal and coworkers. The composite material had very promising antibacterial properties versus P. aeruginosa, E. coli, Staphylococcus hominis, and S. aureus [93]. Many other researchers have used chitin and chitosan fibers in the biomedical engineering field, such as Haider et al., Abdelgawad et al., and Cremar et al. [65,70,91]. 2.3.4.2 Chitin and chitosan fibers for dye removal and wastewaters treatment Chitin and chitosan have attracted considerable attention in environmental applications and have been extensively investigated in the last decades. The presence of functional groups, that is, free amine and hydroxyl groups, on chitosan endows it with a good capability for the sorption of pollutant. This polymer has a low specific surface area in the form of flakes or powders, which limits its use as an adsorbent, but preparing chitosan in the form of fibers or nanofibers is expected to greatly increase chitosan’s specific surface area and hence improve its efficiency and adsorption capacity. For the efficient removal of heavy metals and other pollutants, more attention should be paid to the use of chitin and chitosan fibers as biosorbents in comparison with the others forms of modified chitosan, such as bead or granule type. Furthermore, the chitosan fibers can be used in many different ways, such as in a filter bed or a packed column. Important works, especially for metal adsorption on chitosan fibers and nanofibers form, have been published by many researchers. Some chitosan fibers, hollow chitosan fibers, chitosan/cellulose fibers, chitosan/graphene oxide fibers, chitosan/carbon fibers, etc., were tested for the removal of dyes and heavy metal ions [19,94 99]. Li et al. prepared a composite material, graphene oxide/chitosan fiber, by a wet spinning method. The chitosan fibers have been successfully used to remove the fuchsin acid dye by adsorption. According to Li and coworkers, the graphene oxide/chitosan fibers had excellent

Handbook of Chitin and Chitosan

48

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

mechanical properties and can serve as a promising adsorbent for dyes removal from aqueous solutions [98]. Mirmohseni and coworkers prepared chitosan hollow fibers, with high mechanical strength, as an effective biosorbent of dye. Chitosan fibers have been produced via a dry wet spinning technique with good mechanical properties. The prepared fibers were used to remove the reactive blue 19 as a model anionic dye. The results obtained showed that the adsorption capacity of chitosan fibers were higher than other adsorbents from earlier reports [96]. Li et al. prepared homogenous electrospun chitosan nanofibers under optimized electrospinning conditions using 5% chitosan in acetic acid as the spinning solution. The chitosan fibers were then cross-linked with glutaraldehyde to remove chromium from water via static adsorption. The results obtained indicated that electrospun chitosan nanofibers can be used as promising sorbents for chromium removal [97]. Haider and Park successfully synthesized electrospun chitosan nanofibers with good properties and examined the metal adsorbability of the prepared chitosan fibers in aqueous solution. The chitosan nanofiber mats were found to be very effective in adsorbing Cu(II) and Pb(II) ions with a high mechanical strength in the swollen state, with adsorption capacities of 485.44 and 263.15 mg/g, respectively. According to Haider and Park, this high adsorption capacity suggests that the chitosan electrospun nanofiber mats can be applied to filter out toxic metal ions without losing their original chitosan properties, such as biocompatibility and nontoxicity [100]. 2.3.4.3 Chitin and chitosan fibers for cosmetic applications Electrospun polymer fibers or nanofibers have been also used in cosmetic industries for the treatment of skin healing and for other medical and therapeutic properties. The electrospinning technique can be used as a choice method because of its potential application in the fabrication of cosmetic masks, which are used for skin healing and skin cleansing [72,101]. Moreover, various factors that are essential for skin health and renewal can be impregnated into the nanofiber masks, which can assist in the skin treatment and skin-revitalizing [102]. The electrospun skin mask has the advantage of a high surface area, which facilitates the flow of additives and speeds up the transfer rate from and to the skin, allowing for a better and more efficient utilization. The electrospun nanofibrous cosmetic skin mask can easily be applied and removed from the skin without inducing pain. Chitosan-based face mask products are gaining popularity in the cosmetic industries. As mentioned by Qin et al., chitosan fibers are ideal materials for the production of face masks, since because of their highly

Handbook of Chitin and Chitosan

2.3 Chitin and chitosan: fiber formation

49

hydrophilic and biocompatible properties, they can be used to carry various bioactive ingredients to achieve sustained release to the skin [103]. 2.3.4.4 Other applications for chitin and chitosan fibers, in different fields In addition to the cosmetic, medical, and environmental applications, chitin and chitosan fibers have been widely used in many other applications. These fibers are usable as ecological and environment-friendly materials in agriculture, tissue engineering, drug delivery, air-cleaning, functional textiles, etc. In the textile industries, chitin and chitosan fibers have been widely used due to several of their important functions, such as antibacterial, nonallergenic, deodorizing, and moisture-controlling activity. These fibers have been commercialized as a functional textile material, especially for underwear production. Chitosan transition metal ion complex fibers are usable as shielding materials for electromagnetic waves and radiation, as reported by Hirano [62]. Chitin and chitosan fibers are also used as water, air, and aerosol filters [62,104 106]. Desai et al. prepared chitosan/polyethylene oxide blend fiber mats by an electrospinning technique by optimizing the effect of Mw, the weight of polyethylene oxide in the blend, and spinning solution temperature, with the goal of forming nanoscale fibers with good properties for filtration applications. According to Desai and coworkers, the results showed potential applications for chitosan blend nanofiber mats as filtration materials for air and water filtration [107]. Liu et al. employed an electrospinning method to prepare polylactic acid/carbon nanotubes/chitosan composite fibers containing different chitosan contents and examined the ability of these fibers for strawberry preservation. The results showed that the composite fiber exhibited high antimicrobial activity against E. coli, S. aureus, Rhizopus, and Botrytis cinerea. Thus these fibers can delay the physiological changes in strawberries and extend their shelf life and therefore have important potential applications for fruit and vegetable preservation in the future, according to Liu and coworkers [108]. In addition, chitin and chitosan fibers have been used in various tissue engineering applications. Turo et al. prepared by electrospinning noncovalently cross-linked chitosan nanofiber mats for use as substrates for soft tissue regeneration. Finally, chitosan fibers have been used as a reinforcement method to improve scaffold mechanical properties, as reported by Albanna and coworkers [78,88,109,110]. Fibers and nanofibers of chitin and chitosan have amazing characteristics and properties such as high porosity with very small pore size and high surface area. Therefore chitin and chitosan fibers can be very promising materials for many biomedical applications, such as medical

Handbook of Chitin and Chitosan

50

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

prostheses, wound dressing, artificial organs, bone tissue engineering, drug delivery, filter formation, and sensing applications [65,73,74,78,88,90].

2.4 Conclusions Chitin and chitosan are amazing amino polysaccharides that have so many applications in several fields. They are found in abundance in nature as a renewable resource, especially in the exoskeleton of crustaceans, principally crabs and shrimps. Chitin and chitosan are biopolymers with interesting chemical and biological properties, such as biodegradability, biocompatibility, nontoxicity, and antibacterial and antimicrobial activities. They have been widely used in various fields, namely, biomedical, pharmaceutical, agricultural, and many industrial applications, such as wastewater treatment, due to the fact that they can adsorb and chelate many metal cations. The presence of reactive functional groups in chitin and chitosan offers these polymers the advantage of being easily processed into films, membranes, fibers or nanofibers, beads, and other physical forms. In recent years, fibers and nanofibers have attracted much interest due to their high surface area and porosity. Chitin and chitosan fibers have been found to be useful as promising biomaterials for potential applications, especially in biomedical fields. One of the most important techniques for the production of chitin and chitosan nanofibers is the electrospinning process. This method is governed by a number of parameters that can play important roles in fiber formation. In the last decade, studies on the electrospinning of chitin and chitosan have increased, but the main challenge to overcome in order to prepare high-quality chitin and chitosan nanofibers is the need to find a suitable nontoxic and environment-friendly solvent.

References [1] I. Hamed, F. Ozogul, J.M. Regenstein, Industrial applications of crustacean byproducts (chitin, chitosan and chitooligosaccharides): a review, Trends Food Sci. Technol. 48 (2016) 40 50. Available from: https://doi.org/10.1016/j.tifs.2015.11.007. [2] K.P. Rahate, An overview on various modifications of chitosan and it’s applications, Int. J. Pharm. Sci. Res. (2013) 4175 4193. Available from: https://doi.org/10.13040/ IJPSR.0975-8232.4(11).4175-93. [3] S.P.M. Castro, E.G.L. Paulı´n, Is chitosan a new panacea? Areas of application, Complex. World Polysacch. (2012) 3 46. [4] K. Kurita, Chitin and chitosan: functional biopolymers from marine crustaceans, Mar. Biotechnol. 8 (2006) 203 226. Available from: https://doi.org/10.1007/s10126-0050097-5.

Handbook of Chitin and Chitosan

References

51

[5] K.K. Gadgey, A. Bahekar, Studies on extraction methods of chitin from crab shell and investigation of its mechanical properties, Int. J. Mech. Eng. Technol. 8 (2017) 220 231. [6] P. Gonil, W. Sajomsang, Applications of magnetic resonance spectroscopy to chitin from insect cuticles, Int. J. Biol. Macromol. 51 (2012) 514 522. Available from: https://doi.org/10.1016/j.ijbiomac.2012.06.025. [7] K. Mohan, S. Ravichandran, T. Muralisankar, V. Uthayakumar, R. Chandirasekar, C. Rajeevgandhi, et al., Extraction and characterization of chitin from sea snail Conus inscriptus (Reeve, 1843), Int. J. Biol. Macromol. 126 (2019) 555 560. Available from: https://doi.org/10.1016/j.ijbiomac.2018.12.241. [8] A. Hassainia, H. Satha, S. Boufi, Chitin from Agaricus bisporus: extraction and characterization, Int. J. Biol. Macromol. 117 (2018) 1334 1342. Available from: https://doi.org/10.1016/j.ijbiomac.2017.11.172. [9] J.-Y. Je, S.-K. Kim, Chitosan as Potential Marine Nutraceutical, first ed., Elsevier Inc, 2012. Available from: https://doi.org/10.1016/B978-0-12-416003-3.00007-X. ´ [10] J. Kumirska, M. Czerwicka, Z. Kaczynski, A. Bychowska, K. Brzozowski, J. Tho¨ming, et al., Application of spectroscopic methods for structural analysis of chitin and chitosan, Mar. Drugs. 8 (2010) 1567 1636. Available from: https://doi.org/10.3390/ md8051567. [11] R. Berger, T. Christina, A. De Miranda, P. Pessoa, M.A. Barbosa, D. Lima, et al., Chitosan produced from Mucorales fungi using agroindustrial by-products and its efficacy to inhibit Colletotrichum species, Int. J. Biol. Macromol. (2017). Available from: https://doi.org/10.1016/j.ijbiomac.2017.11.178. [12] M. Kannan, M. Nesakumari, K. Rajarathinam, Production and characterization of mushroom chitosan under solid-state fermentation conditions, Adv. Biol. Res. (Rennes) 4 (2010) 10 13. [13] F. Croisier, C. Je´roˆme, Chitosan-based biomaterials for tissue engineering, Eur. Polym. J. 49 (2013) 780 792. Available from: https://doi.org/10.1016/j. eurpolymj.2012.12.009. [14] S. Kumari, P. Rath, A.S. Hari, T.N. Tiwari, Extraction and characterization of chitin and chitosan from fishery waste by chemical method, Environ. Technol. Innov. 3 (2015) 77 85. Available from: https://doi.org/10.1016/j.eti.2015.01.002. [15] M. Bajaj, J. Winter, C. Gallert, Effect of deproteination and deacetylation conditions on viscosity of chitin and chitosan extracted from Crangon crangon shrimp waste, Biochem. Eng. J. 56 (2011) 51 62. Available from: https://doi.org/10.1016/j.bej.2011.05.006. [16] X. Yan, P. Gong, L. Wang, S. Wu, B. Liu, J. Feng, et al., Inverse solubility of chitin/ chitosan in aqueous alkali solvents at low temperature, Carbohydr. Polym. 206 (2018) 487 492. Available from: https://doi.org/10.1016/j.carbpol.2018.11.016. [17] W.G. Birolli, J.A.D.M. Delezuk, S.P. Campana-Filho, Ultrasound-assisted conversion of alpha-chitin into chitosan, Appl. Acoust. 103 (2016) 239 242. Available from: https://doi.org/10.1016/j.apacoust.2015.10.002. [18] E.S. Abdou, K.S. a Nagy, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from local sources, Bioresour. Technol. 99 (2008) 1359 1367. Available from: https://doi.org/10.1016/j.biortech.2007.01.051. [19] J. Wang, C. Chen, Chitosan-based biosorbents: modification and application for biosorption of heavy metals and radionuclides, Bioresour. Technol. 160 (2014) 129 141. Available from: https://doi.org/10.1016/j.biortech.2013.12.110. [20] N. Mati-Baouche, P.H. Elchinger, H. De Baynast, G. Pierre, C. Delattre, P. Michaud, Chitosan as an adhesive, Eur. Polym. J. 60 (2014) 198 213. Available from: https:// doi.org/10.1016/j.eurpolymj.2014.09.008.

Handbook of Chitin and Chitosan

52

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

[21] R.A.A. Muzzarelli, Nanochitins and Nanochitosans, Paving the Way to Eco-Friendly and Energy-Saving Exploitation of Marine Resources, Elsevier B.V, 2012. Available from: https://doi.org/10.1016/B978-0-444-53349-4.00257-0. [22] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (2006) 603 632. Available from: https://doi.org/10.1016/j.progpolymsci.2006.06.001. [23] T.D. Rathke, S.M. Hudson, Review of chitin and chitosan as fiber and film formers, J. Macromol. Sci. Part C (2006) 37 41. [24] K.V.H. Prashanth, R.N. Tharanathan, Chitin/chitosan: modifications and their unlimited application potential d an overview, Food Sci. Technol. 18 (2007) 117 131. Available from: https://doi.org/10.1016/j.tifs.2006.10.022. [25] H. Srinivasan, K. Velayutham, R. Ravichandran, Chitin and chitosan preparation from shrimp shells Penaeus monodon and its human ovarian cancer cell line, PA-1, Int. J. Biol. Macromol. (2017). Available from: https://doi.org/10.1016/j.ijbiomac. 2017.09.035. [26] H. EL Knidri, R. Belaabed, R. El khalfaouy, A. Laajeb, A. Addaou, A. Lahsini, Physicochemical characterization of chitin and chitosan producted from Parapenaeus Longirostris shrimp shell wastes, J. Mater. Environ. Sci. 8 (2017) 3648 3653. [27] Z. Shariatinia, Pharmaceutical applications of chitosan, Adv. Colloid Interface Sci. 263 (2019) 131 194. Available from: https://doi.org/10.1016/j.cis.2018.11.008. [28] A.J. Al-manhel, A.R.S. Al-hilphy, A.K. Niamah, Extraction of chitosan, characterisation and its use for water purification, J. Saudi Soc. Agric. Sci. (2016). Available from: https://doi.org/10.1016/j.jssas.2016.04.001. [29] M.N.V. Ravikumar, Chitin and chitosan fibres: a review, Bull. Mater. Sci. 22 (1999) 905 915. [30] M.H. Mohammed, P.A. Williams, O. Tverezovskaya, Extraction of chitin from prawn shells and conversion to low molecular mass chitosan, Food Hydrocoll. 31 (2013) 166 171. Available from: https://doi.org/10.1016/j.foodhyd.2012.10.021. [31] Chitin extraction from shrimp shell using enzymatic treatment. Antitumor, antioxidant and antimicrobial activities of chitosan, Int. J. Biol. Macromol. 69 (2014) 489 498. [32] C.M. De Moura, J.M. De Moura, N.M. Soares, L.A.D.A. Pinto, Evaluation of molar weight and deacetylation degree of chitosan during chitin deacetylation reaction: used to produce biofilm, Chem. Eng. Process. Process Intensif. 50 (2011) 351 355. Available from: https://doi.org/10.1016/j.cep.2011.03.003. [33] M.A. Surati, S. Jauhari, K.R. Desai, A brief review: microwave assisted organic reaction, Arch. Appl. Sci. Res. 4 (2012) 645 661. [34] T. Razzaq, C.O. Kappe, On the energy efficiency of microwave-assisted organic reactions, ChemSusChem. 1 (2008) 123 132. Available from: https://doi.org/ 10.1002/cssc.200700036. [35] A.C. Boye, Microwaves in organic chemistry: is it just a bunch of hot air? Report, (2005) 1 9. https://chemistry.illinois.edu/system/files/inline-files/7_Boye_Abstract_SP05. pdf. [36] H. EL Knidri, R. El Khalfaouy, A. Laajeb, A. Addaou, A. Lahsini, Eco-friendly extraction and characterization of chitin and chitosan from the shrimp shell waste via microwave irradiation, Process. Saf. Environ. Prot. 104 (2016) 395 405. Available from: https://doi.org/10.1016/j.psep.2016.09.020. [37] H. EL Knidri, R. Belaabed, A. Addaou, A. Laajeb, A. Lahsini, Extraction, chemical modification and characterization of chitin and chitosan, Int. J. Biol. Macromol. 120 (2018) 1181 1189. Available from: https://doi.org/10.1016/j. ijbiomac.2018.08.139. [38] M.Y. Arancibia, A. Alema´n, M.M. Calvo, M.E. Lo´pez-Caballero, P. Montero, M.C. Go´mez-Guille´n, Antimicrobial and antioxidant chitosan solutions enriched with

Handbook of Chitin and Chitosan

References

[39]

[40]

[41]

[42]

[43]

[44]

[45]

[46]

[47]

[48]

[49]

[50]

[51] [52]

[53]

53

active shrimp (Litopenaeus vannamei) waste materials, Food Hydrocoll. 35 (2014) 710 717. Available from: https://doi.org/10.1016/j.foodhyd.2013.08.026. A. Abdulkarim, M. Tijani, S. Abdulsalam, A.J. Muhammad, A.O. Ameh, Extraction and characterisation of chitin and chitosan from Mussel Shell, Civ. Environ. Res. 3 (2013) 108 115. M.S. Benhabiles, N. Abdi, N. Drouiche, H. Lounici, A. Pauss, M.F.A. Goosen, et al., Protein recovery by ultrafiltration during isolation of chitin from shrimp shells Parapenaeus longirostris, Food Hydrocoll. 32 (2013) 28 34. Available from: https:// doi.org/10.1016/j.foodhyd.2012.11.035. B.S. de Farias, T.R. Sant’Anna Cadaval Junior, L.A. de Almeida Pinto, Chitosanfunctionalized nanofibers: a comprehensive review on challenges and prospects for food applications, Int. J. Biol. Macromol. 123 (2019) 210 220. Available from: https://doi.org/10.1016/j.ijbiomac.2018.11.042. G. Ma, Y. Liu, J.F. Kennedy, J. Nie, Synthesize and properties of photosensitive organic solvent soluble acylated chitosan derivatives (2), Carbohydr. Polym. 84 (2011) 681 685. Available from: https://doi.org/10.1016/j.carbpol.2010.11.013. O.E. Philippova, E.V. Korchagina, E.V. Volkov, V.A. Smirnov, A.R. Khokhlov, M. Rinaudo, Aggregation of some water-soluble derivatives of chitin in aqueous solutions: role of the degree of acetylation and effect of hydrogen bond breaker, Carbohydr. Polym. 87 (2012) 687 694. Available from: https://doi.org/10.1016/j. carbpol.2011.08.043. S. Popa-Nita, P. Alcouffe, C. Rochas, L. David, A. Domard, Continuum of structural organization from chitosan solutions to derived physical forms, Biomacromolecules 11 (2010) 6 12. Available from: https://doi.org/10.1021/bm9012138. Y. Wu, T. Sasaki, S. Irie, K. Sakurai, A novel biomass-ionic liquid platform for the utilization of native chitin, Polym. (Guildf.) 49 (2008) 2321 2327. Available from: https://doi.org/10.1016/j.polymer.2008.03.027. V.K. Mourya, N.N. Inamdar, Chitosan-modifications and applications: opportunities galore, React. Funct. Polym. 68 (2008) 1013 1051. Available from: https://doi.org/ 10.1016/j.reactfunctpolym.2008.03.002. N.H. Daraghmeh, B.Z. Chowdhry, S.A. Leharne, M.M. Al Omari, A.A. Badwan, Chitin, first ed., Elsevier Inc, 2011. Available from: https://doi.org/10.1016/B978-012-387667-6.00002-6. A. Shajahan, S. Shankar, A. Sathiyaseelan, K.S. Narayan, V. Narayanan, V. Kaviyarasan, et al., Comparative studies of chitosan and its nanoparticles for the adsorption efficiency of various dyes, Int. J. Biol. Macromol. 104 (2017) 1449 1458. Available from: https://doi.org/10.1016/j.ijbiomac.2017.05.128. J. Bonilla, E. Fortunati, L. Atares, A. Chiralt, J.M. Kenny, Physical, structural and antimicrobial properties of poly vinyl alcohol-chitosan biodegradable films, Food Hydrocoll. 35 (2014) 463 470. Available from: https://doi.org/10.1016/j. foodhyd.2013.07.002. J.C. Roy, F. Salau¨n, S. Giraud, A. Ferri, Solubility of chitin: solvents, solution behaviors and their related mechanisms, in: Solubility of Polysaccharides, 2017, pp. 109 127. https://doi.org/10.5772/intechopen.71385. H.M. Struszczyk, Chitin and chitosan, Polymer 47 (2002) 316 325. Y. Wang, E. Wang, Z. Wu, H. Li, Z. Zhu, X. Zhu, et al., Synthesis of chitosan molecularly imprinted polymers for solid-phase extraction of methandrostenolone, Carbohydr. Polym. 101 (2014) 517 523. Available from: https://doi.org/10.1016/j. carbpol.2013.09.078. P.K. Dutta, J. Dutta, V.S. Tripathi, Chitin and chitosan: chemistry, properties and applications, J. Sci. Ind. Res. 63 (2004) 20 31.

Handbook of Chitin and Chitosan

54

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

[54] H. Sashiwa, N. Kawasaki, A. Nakayama, E. Muraki, N. Yamamoto, S. Aiba, Chemical modification of chitosan. 14:1 synthesis of water-soluble chitosan derivatives by simple acetylation, Biomacromolecules 3 (2002) 1126 1128. [55] Z. Xia, S. Wu, J. Chen, Preparation of water soluble chitosan by hydrolysis using hydrogen peroxide, Int. J. Biol. Macromol. 59 (2013) 242 245. Available from: https://doi.org/10.1016/j.ijbiomac.2013.04.034. [56] N.C. Minh, V.H. Nguyen, S. Schwarz, W.F. Stevens, T.S. Trung, Preparation of water soluble hydrochloric chitosan from low molecular weight chitosan in the solid state, Int. J. Biol. Macromol. 121 (2019) 718 726. Available from: https://doi.org/10.1016/j. ijbiomac.2018.10.130. [57] Y. Fu, C. Xiao, A facile physical approach to make chitosan soluble in acid-free water, Int. J. Biol. Macromol. 103 (2017) 575 580. Available from: https://doi.org/10.1016/j. ijbiomac.2017.05.066. [58] S. Botelho da Silva, M. Krolicka, L.A.M. van den Broek, A.E. Frissen, C.G. Boeriu, Water-soluble chitosan derivatives and pH-responsive hydrogels by selective C-6 oxidation mediated by TEMPO-laccase redox system, Carbohydr. Polym. 186 (2018) 299 309. Available from: https://doi.org/10.1016/j.carbpol.2018.01.050. [59] S. Ojha, Structure property relationship of electrospun fibers, Electrospun Nanofibers, Elsevier Ltd, 2017, pp. 239 254. Available from: https://doi.org/ 10.1016/B978-0-08-100907-9.00010-6. [60] O.C. Agboh, Y. Qin, Chitin and chitosan fibers, Polym. Adv. Technol. 8 (1997) 355 365. Available from: https://doi.org/10.1002/(SICI)1099-1581(199706)8:6 , 355:: AID-PAT651 . 3.0.CO;2-T. [61] J. Zhu, S. Jasper, X. Zhang, Chemical characterization of electrospun nanofibers, Electrospun Nanofibers, Elsevier Ltd, 2017, pp. 181 206. Available from: https:// doi.org/10.1016/B978-0-08-100907-9.00008-8. [62] S. Hirano, Wet-spinning and applications of functional fibers based on chitin and chitosan, Macromol. Symp. 168 (2001) 21 30. [63] B.K. Gu, S.J. Park, M.S. Kim, C.M. Kang, J. Kim, C. Kim, Fabrication of sonicated chitosan nanofiber mat with enlarged porosity for use as hemostatic materials, Carbohydr. Polym. (2013). Available from: https://doi.org/10.1016/j. carbpol.2013.04.060. [64] M.A. Kiechel, C.L. Schauer, Non-covalent crosslinkers for electrospun chitosan fibers, Carbohydr. Polym. 95 (2013) 123 133. Available from: https://doi.org/10.1016/j. carbpol.2013.02.034. [65] L. Cremar, J. Gutierrez, J. Martinez, L.A. Materon, R. Gilkerson, F. Xu, et al., Development of antimicrobial chitosan based nanofiber dressings for wound healing applications, Nanomed. J. 5 (2018) 6 14. Available from: https://doi.org/10.22038/ nmj.2018.05.002. [66] R.N. Wijesena, N. Tissera, Y.Y. Kannangara, Y. Lin, G.A.J. Amaratunga, K.M.N. De Silva, A method for top down preparation of chitosan nanoparticles and nanofibers, Carbohydr. Polym. 117 (2015) 731 738. Available from: https://doi.org/10.1016/j. carbpol.2014.10.055. [67] M.P. Lonbani, Production of chitosan-based non-woven membranes using the electrospinning process, Thesis, 2012. https://publications.polymtl.ca/882/. [68] V. Sencadas, D.M. Correia, A. Areias, G. Botelho, A.M. Fonseca, I.C. Neves, et al., Determination of the parameters affecting electrospun chitosan fiber size distribution and morphology, Carbohydr. Polym. 87 (2012) 1295 1301. Available from: https:// doi.org/10.1016/j.carbpol.2011.09.017. [69] M. Arkoun, Elaboration de nouveaux biomate´riaux antibacte´riens a` base de chitosane par le proce´de´ d’e´lectrofilage, Montre´al, 2017.

Handbook of Chitin and Chitosan

References

55

[70] A. Haider, S. Haider, I.-K. Kang, A comprehensive review summarizing the effect of electrospinning parameters and potential applications of nanofibers in biomedical and biotechnology, Arab. J. Chem. 11 (2018) 1165 1188. Available from: https://doi. org/10.1016/j.arabjc.2015.11.015. [71] A. Khalf, S.V. Madihally, Recent advances in multiaxial electrospinning for drug delivery, Eur. J. Pharm. Biopharm. 112 (2017) 1 17. Available from: https://doi.org/ 10.1016/j.ejpb.2016.11.010. [72] N. Bhardwaj, S.C. Kundu, Electrospinning: a fascinating fiber fabrication technique, Biotechnol. Adv. 28 (2010) 325 347. Available from: https://doi.org/10.1016/j. biotechadv.2010.01.004. [73] M.Z. Elsabee, H.F. Naguib, R.E. Morsi, Chitosan based nanofibers, review, Mater. Sci. Eng. C. 32 (2012) 1711 1726. Available from: https://doi.org/10.1016/j. msec.2012.05.009. [74] T.J. Sill, H.A. von Recum, Electrospinning: applications in drug delivery and tissue engineering, Biomaterials. 29 (2008) 1989 2006. Available from: https://doi.org/ 10.1016/j.biomaterials.2008.01.011. [75] X. Yuan, Y. Zhang, C. Dong, J. Sheng, Morphology of ultrafine polysulfone fibers prepared by electrospinning, Polym. Int. 53 (2004) 1704 1710. Available from: https:// doi.org/10.1002/pi.1538. [76] G. Taylor, Electrically driven jets, Proc. Roy. Soc. Lond. A. 313 (1969) 453 475. [77] J. Doshi, D.H. Reneker, Electrospinning process and applications of electrospun fibers, J. Electrostat. 35 (1995) 151 160. [78] C. Salas, Z. Thompson, N. Bhattarai, Electrospun chitosan fibers, Electrospun Nanofibers, Elsevier Ltd, 2017, pp. 371 398. Available from: https://doi.org/ 10.1016/B978-0-08-100907-9.00015-5. [79] S. Thenmozhi, N. Dharmaraj, K. Kadirvelu, H. Yong, Electrospun nanofibers: new generation materials for advanced applications, Mater. Sci. Eng. B. 217 (2017) 36 48. Available from: https://doi.org/10.1016/j.mseb.2017.01.001. [80] J. Kong, Y. Lee, N. Chen, S. Peng, L. Li, L. Tian, Polymer-based composites by electrospinning: preparation & functionalization with nanocarbons, Prog. Polym. Sci. 86 (2018) 40 84. Available from: https://doi.org/10.1016/j.progpolymsci.2018.07.002. [81] C. Mit-uppatham, M. Nithitanakul, P. Supaphol, Ultrafine electrospun polyamide-6 fibers: effect of solution conditions on morphology and average fiber diameter, Macromol. Chem. Phys. 6 (2004) 2327 2338. Available from: https://doi.org/ 10.1002/macp.200400225. [82] C.L. Casper, J.S. Stephens, N.G. Tassi, D.B. Chase, J.F. Rabolt, Controlling surface morphology of electrospun polystyrene fibers: effect of humidity and molecular weight in the electrospinning process, Macromolecules 37 (2004) 573 578. [83] P. Su, C. Wang, X. Yang, X. Chen, C. Gao, X. Feng, et al., Electrospinning of chitosan nanofibers: the favorable effect of metal ions, Carbohydr. Polym. 84 (2011) 239 246. Available from: https://doi.org/10.1016/j.carbpol.2010.11.031. [84] M. Pakravan, M. Heuzey, A. Ajji, A fundamental study of chitosan/PEO electrospinning, Polym. (Guildf.) 52 (2011) 4813 4824. Available from: https://doi.org/10.1016/ j.polymer.2011.08.034. [85] K. Kalantari, A.M. Afifi, H. Jahangirian, T.J. Webster, Biomedical applications of chitosan electrospun nanofibers as a green polymer—review, Carbohydr. Polym. 207 (2019) 588 600. Available from: https://doi.org/10.1016/j.carbpol.2018.12.011. [86] W.Y. Cheah, P.L. Show, I.S. Ng, G.Y. Lin, C.Y. Chiu, Y.K. Chang, Antibacterial activity of quaternized chitosan modified nanofiber membrane, Int. J. Biol. Macromol. 126 (2019) 569 577. Available from: https://doi.org/10.1016/j. ijbiomac.2018.12.193.

Handbook of Chitin and Chitosan

56

2. Chitin and chitosan: chemistry, solubility, fiber formation, and their potential applications

[87] I.R. Sweeney, M. Miraftab, G. Collyer, Absorbent alginate fibres modified with hydrolysed chitosan for wound care dressings—II. Pilot scale development, Carbohydr. Polym. 102 (2014) 920 927. Available from: https://doi.org/10.1016/j. carbpol.2013.10.053. [88] M.Z. Albanna, T.H. Bou-akl, O. Blowytsky, H.L. Walters, H.W.T. Matthew, Chitosan fibers with improved biological and mechanical properties for tissue engineering applications, J. Mech. Behav. Biomed. Mater. 20 (2013) 217 226. Available from: https://doi.org/10.1016/j.jmbbm.2012.09.012. [89] M. Ignatova, K. Starbova, N. Markova, N. Manolova, I. Rashkov, Electrospun nanofibre mats with antibacterial properties from quaternised chitosan and poly (vinyl alcohol), Carbohydr. Res. 341 (2006) 2098 2107. Available from: https://doi.org/ 10.1016/j.carres.2006.05.006. [90] B. Sibaja, E. Culbertson, P. Marshall, R. Boy, R.M. Broughton, A. Aguilar, et al., Preparation of alginate—chitosan fibers with potential biomedical applications, Carbohydr. Polym. 134 (2015) 598 608. Available from: https://doi.org/10.1016/j. carbpol.2015.07.076. [91] A.M. Abdelgawad, S.M. Hudson, O.J. Rojas, Antimicrobial wound dressing nanofiber mats from multicomponent (chitosan/silver-NPs/polyvinyl alcohol) systems, Carbohydr. Polym. (2013). Available from: https://doi.org/10.1016/j. carbpol.2012.12.043. [92] Y. Zhou, H. Yang, X. Liu, J. Mao, S. Gu, W. Xu, Potential of quaternizationfunctionalized chitosan fiber for wound dressing, Int. J. Biol. Macromol. 52 (2013) 327 332. Available from: https://doi.org/10.1016/j.ijbiomac.2012.10.012. [93] E. Guibal, S. Cambe, S. Bayle, J. Taulemesse, T. Vincent, Silver/chitosan/cellulose fibers foam composites: from synthesis to antibacterial properties, J. Colloid Interface Sci. 393 (2013) 411 420. Available from: https://doi.org/10.1016/j. jcis.2012.10.057. [94] Y. Liao, C. Loh, M. Tian, R. Wang, A.G. Fane, Progress in electrospun polymeric nanofibrous membranes for water treatment: fabrication, modification and applications, Prog. Polym. Sci. 77 (2018) 69 94. Available from: https://doi.org/10.1016/j. progpolymsci.2017.10.003. [95] F. Tasselli, A. Mirmohseni, M.S.S. Dorraji, A. Figoli, Mechanical, swelling and adsorptive properties of dry—wet spun chitosan hollow fibers crosslinked with glutaraldehyde, React. Funct. Polym. 73 (2013) 218 223. Available from: https:// doi.org/10.1016/j.reactfunctpolym.2012.08.007. [96] A. Mirmohseni, M.S.S. Dorraji, A. Figoli, F. Tasselli, Chitosan hollow fibers as effective biosorbent toward dye: preparation and modeling, Bioresour. Technol. 121 (2012) 212 220. Available from: https://doi.org/10.1016/j.biortech.2012.06.067. [97] L. Li, Y. Li, L. Cao, C. Yang, Enhanced chromium (VI) adsorption using nanosized chitosan fibers tailored by electrospinning, Carbohydr. Polym. 125 (2015) 206 213. Available from: https://doi.org/10.1016/j.carbpol.2015.02.037. [98] Y. Li, J. Sun, Q. Du, L. Zhang, X. Yang, S. Wu, et al., Mechanical and dye adsorption properties of graphene oxide/chitosan composite fibers prepared by wet spinning, Carbohydr. Polym. 102 (2014) 755 761. Available from: https://doi.org/10.1016/j. carbpol.2013.10.094. [99] E. Guibal, Interactions of metal ions with chitosan-based sorbents: a review, Sep. Purif. Technol. 38 (2004) 43 74. Available from: https://doi.org/10.1016/j. seppur.2003.10.004. [100] S. Haider, S. Park, Preparation of the electrospun chitosan nanofibers and their applications to the adsorption of Cu (II) and Pb (II) ions from an aqueous solution, J. Memb. Sci. 328 (2009) 90 96. Available from: https://doi.org/10.1016/j. memsci.2008.11.046.

Handbook of Chitin and Chitosan

References

57

[101] Z. Huang, Y. Zhang, M. Kotaki, S. Ramakrishna, A review on polymer nanofibers by electrospinning and their applications in nanocomposites, Compos. Sci. Technol. 63 (2003) 2223 2253. Available from: https://doi.org/10.1016/S0266-3538(03)00178-7. [102] S. Ramakrishna, K. Fujihara, W. Teo, T. Yong, R. Ramaseshan, Electrospun nanofibers: solving global issues, Materialstoday 9 (2006) 40 50. [103] Y. Qin, Y. Deng, Y. Hao, N. Zhang, X. Shang, Marine bioactive fibers: alginate and chitosan fibers—a critical review, J. Text. Eng. Fash. Technol. 1 (2017) 228 231. Available from: https://doi.org/10.15406/jteft.2017.01.00037. [104] X. Qin, S. Subianto, Electrospun nanofibers for filtration applications, Electrospun Nanofibers, Elsevier Ltd, 2017, pp. 449 466. Available from: https://doi.org/ 10.1016/B978-0-08-100907-9.00017-9. [105] S. Sundarrajan, K. Luck, S. Huat, S. Ramakrishna, Electrospun nanofibers for air filtration applications, Procedia Engineering, Elsevier B.V, 2014, pp. 159 163. Available from: https://doi.org/10.1016/j.proeng.2013.11.034. [106] F.E. Ahmed, B.S. Lalia, R. Hashaikeh, A review on electrospinning for membrane fabrication: challenges and applications, Desalination 356 (2015) 15 30. Available from: https://doi.org/10.1016/j.desal.2014.09.033. [107] K. Desai, K. Kit, J. Li, S. Zivanovic, Morphological and surface properties of electrospun chitosan nanofibers, Biomacromolecules 9 (2008) 1000 1006. [108] Y. Liu, S. Wang, W. Lan, W. Qin, Fabrication of polylactic acid/carbon nanotubes/ chitosan composite fibers by electrospinning for strawberry preservation, Int. J. Biol. Macromol. 121 (2019) 1329 1336. Available from: https://doi.org/10.1016/j. ijbiomac.2018.09.042. [109] C.T. Turo, F. Ruini, M. Ramella, P.G. Francesca Boccafoschi, E. Gioffredi, G.F.D. Labate, et al., Non-covalently crosslinked chitosan nanofibrous mats prepared by electrospinning as substrates for soft tissue regeneration, Carbohydr. Polym. (2017). Available from: https://doi.org/10.1016/j.carbpol.2017.01.050. [110] N. Soraya, M.G. Kim, S. Bin, The formation of web-like connection among electrospun chitosan/PVA fiber network by the reinforcement of ellipsoidal calcium carbonate, Mater. Sci. Eng. C 60 (2016) 518 525. Available from: https://doi.org/ 10.1016/j.msec.2015.11.079.

Handbook of Chitin and Chitosan

C H A P T E R

3 PEGylated chitin and chitosan derivatives Adib H. Chisty1, Rifat A. Masud2, M. Mehedi Hasan2, M. Nuruzzaman Khan1, Abul K. Mallik1 and Mohammed Mizanur Rahman1 1

Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh, 2 Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering, Bangabandhu Sheikh Mujibur Rahman Science and Technology University, Gopalganj, Bangladesh O U T L I N E 3.1 Introduction

60

3.2 Chitin and chitosan

61

3.3 PEGylation and PEGylated chitin/chitosan derivatives

64

3.4 Fabrication of PEGylated chitosan derivatives 3.4.1 Amino group substitution 3.4.2 O-substitution 3.4.3 Miscellaneous approaches 3.4.4 Solubilization of chitosan prior to derivatization 3.4.5 PEGylated cross-linked chitosan

65 66 67 69 69 70

3.5 Characterization of PEGylated chitosan and chitin derivatives 3.5.1 Structural analysis 3.5.2 Determination of the degree of substitution 3.5.3 Ductility 3.5.4 Solubility

71 71 76 81 82

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00003-1

59

© 2020 Elsevier Inc. All rights reserved.

60

3. PEGylated chitin and chitosan derivatives

3.5.5 Cytocompatibility 3.5.6 Aggregation

83 85

3.6 Applications of PEGylated derivatives of chitosan 3.6.1 Medical applications 3.6.2 Thermoresponsive PEGylated chitosan hydrogels 3.6.3 Gene delivery of chitosan derivatives 3.6.4 Formation of nanofibres

86 86 88 90 92

3.7 Conclusions

93

References

93

3.1 Introduction Chitin and chitosan are the most naturally abundant polymers after cellulose. Several types of products can be obtained from chitin and chitosan since they possess some exciting properties, such as biodegradability, biocompatibility, nontoxicity, nonimmunogenicity, and noncarcinogenicity [1
4]. These biological and physicochemical properties, along with their abundance, make chitin and chitosan strong candidates for ubiquitous application in pharmaceutical and medical applications, paper production, textile wastewater treatment, biotechnology, cosmetics, food processing, and agriculture [5]. But unfortunately the processing of chitin or chitosan to a newer material is facing some difficulties due to their low solubility in dilute acid solutions, which restricts their use in various applications [6
11]. To improve the solubility of chitosan in a broader range of solvents, chemical modification through reactions of the amine and/or hydroxyl groups could be promising options, as demonstrated by cellulose derivatives. Polyethylene glycol (PEG) is an ideal graft-forming polymer because it is soluble in water and organic solvents and it has low toxicity, good biocompatibility, and biodegradability. PEG is a polyether compound. It is a flexible polymer with low toxicity that can be used in many medicinal, biological, commercial, and industrial applications. PEG is soluble in water, methanol, ethanol, acetonitrile, benzene, and dichloromethane, and is insoluble in diethyl ether and hexane [12,13]. To eliminate the problem regarding the solubility of chitosan, PEG is being used as an ideal graft-forming polymer. PEGylation appears to be a very proficient method to obtain higher solubility of chitosan. There are several reports on enhancing the water solubility of chitosan along with its thermal stability by grafting PEG

Handbook of Chitin and Chitosan

3.2 Chitin and chitosan

61

molecules onto the chitosan backbone [14
16]. In this chapter, we will discuss the developments in the synthesis of PEGylated chitin and chitosan derivatives. We will also focus on the characterization of these derivatives and their potential applications in various fields.

3.2 Chitin and chitosan Chitin is a white, hard, inelastic, and nitrogenous compound. It is considered to be a renewable polymer as it can be obtained from the shells of crustaceans such as prawn, shrimp, crab, and lobster. It is also found in the exoskeletons of mollusks and insects, as well as in the cell walls of some fungi. In terms of abundance chitin is the second most naturally available polymer [17
19]. Industrially chitin is extracted from crustaceans by alkali treatment to solubilize the protein, and then acid treatment to dissolve the various minerals present in it. Then chitosan is extracted from the chitin by highly concentrated alkali treatment. A decolorization step is often included to eliminate leftover pigments and a colorless product is obtained [20]. Chitin mainly comprises β-(1-4)-linked 2-acetamido-2-deoxy-β-D-glucose (N-acetylglucosamine) (Fig. 3.1). Chitin is structurally identical to cellulose except for having acetamide groups (-NHCOCH3) at the C-2 position. Thus chitin is frequently regarded as a cellulose derivative, even though it does not exist in organisms generating cellulose. Chitosan is a copolymer of N-acetylglucosamine and glucosamine, mainly consisting of α-(1-4)-linked-2amino-2-deoxy-β-D-glucopyranose (Fig. 3.2). Chitosan is derived from chitin by deacetylation to different levels so that various degrees of deacetylation of chitosan can be obtained [21].

FIGURE 3.1 Chemical structure of chitin.

Handbook of Chitin and Chitosan

62

3. PEGylated chitin and chitosan derivatives

FIGURE 3.2 Chemical structure of chitosan.

Chitin is not completely deacetylated to chitosan, thus chitosan is called partially deacetylated chitin containing some acetamide groups. About 5%
8% nitrogen is present in chitin in the form of acetylated amine groups and in chitosan in the form of primary aliphatic amine groups, respectively, allowing chitin and chitosan performing the reactions of amines [22]. The existence of two hydroxyl groups (primary and secondary) on individual repeating unit and also the presence of amine groups on each partially deacetylated repeat unit make chitosan chemically more active than chitin. Thus these reactive groups of chitosan can undergo more versatile reactions, bringing change in numerous physical, chemical, and mechanical properties through chemical modification. Therefore the amine group of chitosan is very significant. The higher the degree of deacetylation, the more amine groups are present in chitosan. Chitin and chitosan with amine groups can perform as chelating agents [23]. Synthetic polymeric materials are not biocompatible or biodegradable, whereas natural polysaccharides, like chitin and chitosan, are nontoxic, biocompatible, and biodegradable. But there are some limitations in the processing of a natural polymer to the desired product. Consequently, functional derivatives of chitin and chitosan are brought about by chemical modification to enhance their processability for several purposes. Chitin shows three types of allomorphs based on the source from where they are extracted. These are α-chitin, β-chitin, and γ-chitin. Among these three allomorphs γ-chitin is a variant of α-chitin [24
26]. α-Chitin and β-chitin are not soluble in common solvents. Chitin is basic in nature so it is generally soluble in slightly acidic solution. The insolvability of chitin in the usual solvents is the key challenge for the processing and utilization of chitin [27]. The frequently used solvents for chitin are dimethylacetamide/lithium chloride mixtures, hexafluoroisopropyl alcohol, and hexafluoracetone sesquihydrate [28,29]. Chitin can also be dissolved in concentrated phosphoric acid at room temperature,

Handbook of Chitin and Chitosan

3.2 Chitin and chitosan

63

in fresh saturated solution of lithium thiocyanate, and in sodium hydroxide at low temperature [30
34]. Chitin can be dissolved in water under some conditions but chitosan with a lower degree of deacetylation and simple chitin are not soluble. Under alkaline conditions the molecular weight of chitin decreases and some deacetylation takes place, thus the phenomenon of the dissolution of chitin occurs and the degree of deacetylation must be greater than 50%. It is now proven that the trouble regarding the solubilization of chitin is a consequence chiefly of the highly extended hydrogen-bonded semicrystalline structure of chitin [27,35
37]. Chitosan with a degree of deacetylation greater than 50% is semicrystalline in nature. If low-molecular-weight chitin is completely deacetylated crystalline chitosan can be obtained [38]. The solubility of chitosan is mainly consistent with the degree of deacetylation of chitosan, along with the molecular weight and distribution of acetyl groups on the backbone of the chitosan chain [39,40]. Being basic in nature chitosan will be protonated while dissolving in an acidic medium. The degree of ionization varies with pH and pKa values of strong or weak acid, as studied by dissolving chitosan in hydrochloric acid and acetic acid, respectively [41,42]. Chitosan with a lower percentage of deacetylation is soluble in the pH range 4.5
5 for HCl solution. It can be inferred that chitosan is soluble in an acid solution with pH less than 6. One percent acetic acid is frequently used to dissolve chitosan but the volume of acid required to dissolve chitosan fluctuates with the amount of chitosan [42,43]. As chitosan is protonated during dissolving, the amount of proton present in the acid must be equivalent to the concentration of amine groups participating in protonation. Controlling the solubility of chitosan is very challenging as it is associated with a degree of deacetylation, the ionic concentration of strong or weak acid, the pH, the nature of the acid used for protonation, and the distribution of acetyl groups along the backbone. The hydroxyl groups also tend to form intrachain hydrogen bonds, which is another reason for the insolubility of chitosan. Chitosan can be dissolved in a neutral pH solution containing glycerol-2-phosphate [44
47]. At pH 7
7.1 and room temperature chitosan gives a stable solution with glycerol-2-phosphate, but a sol
gel transition occurs on heating to about 40 C. It can be clearly seen that the solubility of chitosan in water and at a neutral pH range is difficult. In order to generate water-soluble derivatives of chitin and chitosan extensive research has been carried out on chemical modification techniques. Nevertheless, the chemical modification can alter the structural change in the fundamental skeleton of chitin and chitosan. The reformed chitin and chitosan will have newer physicochemical and biochemical properties compared with the original chitin and chitosan [48].

Handbook of Chitin and Chitosan

64

3. PEGylated chitin and chitosan derivatives

3.3 PEGylation and PEGylated chitin/chitosan derivatives PEGylation can be defined as a process of both covalent and noncovalent addition or amalgamation of PEG polymer chains to another entity (polymer, molecules, and macrostructures), such as a drug, therapeutic protein, or vesicle, which is then termed as PEGylated [49
51]. PEG is a linear polyether diol which is well-known for its stealth behavior. PEG has been manufactured with a range of molecular weight from 0.3 to 10,000 kDa and can be modified with amino-, carboxyl-, sulfhydryl-, etc. terminal end groups. PEGs with different molecular weight are highly soluble in water and other organic common solvents, such as methanol and dichloromethane. The US Food and Drug Administration (FDA) has approved some polymers for biomedical application, PEG is one of those polymers. PEG is not only soluble in water and organic solvents but is also nontoxic, nonimmunogenic, nonantigenic, and biocompatible. The most valuable benefit of using PEG for the derivatization of polymers or drugs is that it brings improvements in pharmacokinetics and pharmacodynamics properties, which are associated with extended body residence time, better water solubility, abridged renal clearance, and partial toxicity [52
54]. Since PEG is a biocompatible but not biodegradable polymer its use must be monitored so that it can’t create any toxicity with prolonged use in the body [55]. As discussed in the previous section, chitin and chitosan face difficulties in dissolving in water and most organic solvents. In order to use chitosan in tissue engineering, drug delivery, gene delivery, and other biomedical applications more successfully it is necessary to make chitosan solvable either in water or organic solvents. Such challenges can be overcome by the PEGylation of chitosan, that is, grafting chitosan with PEG through chemical modifications by a copolymerization reaction [7]. If chitin or chitosan is chemically modified with a hydrophilic polymer the resulting structure will be hydrophilic in nature, whilst keeping the main skeleton undamaged. Since PEG is a hydrophilic polymer, graft copolymerization of PEG onto chitin and chitosan will produce a copolymerized hydrophilic structure. Some approaches for the graft copolymerization of hydrophilic polymer onto chitin and chitosan were reported as an efficient method for dissolving chitin and chitosan in water and organic solvents [56]. Sugimoto et al. prepared a chitosan
PEG hybrid by modifying chitosan with PEG in various reaction conditions and acetylated it to obtain a chitin
PEG hybrid [7]. Chitosan contains two major functional groups, aOH and aNH2, where the chemical modification could take place. The original physicochemical and biochemical properties of chitosan will be gone if the

Handbook of Chitin and Chitosan

3.4 Fabrication of PEGylated chitosan derivatives

65

amine group of chitosan undergoes chemical modification. So importance has been given to the chemical modification of hydroxyl groups of chitosan, as this change does not alter any fundamental (structural and functional) properties of chitosan. Gorochovceva and Makuska worked with the PEGylation of chitosan through hydroxyl groups [57]. PEG is utilized mainly as a cross-linker and produces interconnected networks to permit drug release [58,59]. Nishimura et al. successfully achieved chemoselective conjugation of PEG at the hydroxyl group of chitosan [60]. Malhotra et al. developed a new method for grafting PEG onto chitosan with a moderate degree of substitution. PEGylated chitosan nanoparticles were prepared using an ionic gelation method and their gene delivery potential was explored [61]. Gorochovceva et al. reported trichlorotriazine to be a coupling reagent, PEG monomethyl ether covalently attached to chitosan. The novel chitosan-O-mPEG graft copolymers are anticipated to merge the valuable properties of chitosan and PEG and can be applicable in several fields including biotechnology, pharmaceuticals, and water treatments [14].

3.4 Fabrication of PEGylated chitosan derivatives The physicochemical properties of chitosan regarding their solubility, together with drug and gene delivery, can be developed through various chemical modifications of amino and hydroxyl groups in each glucosamine unit of chitosan. As the reactivity of the amino group at C-2 is higher than that of hydroxyl groups at C-6 and C-3 positions, maximum modifications are found at the amino group rather than hydroxyl groups [22]. In the synthesis of PEGylated chitosan, simply PEG or PEG derivatives are grafted onto chitosan with different molecular weights, ranging from higher molecular weight to oligochitosans, and with different degrees of substitutions [15,16,57,62
64]. To synthesize PEGylated chitosan, functionalization of PEG with a suitable end group is of prime importance. In this process, PEG is first modified to a useful intermediate which undergoes grafting onto chitosan to produce the final product. Several studies have been found regarding the PEGylation of chitosan with different PEG derivatives, such as PEG-aldehyde [7,65,66], PEG-carboxylic acid [67
71], PEG-carbonate [64,72], PEG-iodide [15,73], PEG-epoxide [74,75], PEG-acrylate [76], PEG-NHS ester [63,77
80], and PEG-sulfonate [81]. In order to overcome the cross-linking between polymers, most publications have described the use of methoxy-poly (ethylene glycol) (mPEG) instead of PEG for PEGylation. Hu et al. [15], working on the derivatization of chitosan, utilized methoxy poly

Handbook of Chitin and Chitosan

66

3. PEGylated chitin and chitosan derivatives

FIGURE 3.3 PEG N-hydroxy succinimidyl ester derivatization method.

(ethylene glycol) iodide for N-substitution of triphenylmethyl chitosan in an organic medium while Jeong et al. [69] successfully employed methoxy poly(ethylene glycol) during PEGylation using 4dicyclohexylcarbodimide (DCC) and N-hydroxysuccinimide (NHS). Though most of the published work emphasizes the PEGylation of chitosan to the amino group of glucosamine unit, one must consider the potential consequence of reduced absorption capacity of the PEGylated chitosan with a high degree of substitution with PEG. Previous studies by Hu et al. [15], Gorochovceva et al. [57], and Makuska et al. [82] found derivatization of chitosan at OH groups more efficient in overcoming such limitations, retaining a free amino group in its structure. Fig. 3.3 shows the chemical structure of a PEGylated chitosan indicating substitution at the amino group by a linker, designated by X, depending on the type of activated PEG.

3.4.1 Amino group substitution The process involves the copolymerization of chitosan with PEG at an amino group. A publication by Harris et al. [65] synthesized PEG grafted on chitosan using PEG-aldehyde whereas the chemical modification of chitosan was done with PEG-aldehyde to imine (Schiff base), which was further converted into PEG-g-chitosan through reduction with sodium cyanoborohydride (NaCNBH3). However, the method used for the preparation of PEG-aldehyde was found to be inconvenient in a study, with a low degree of conversion due to the instability of PEG-aldehyde. A study by Bentley et al. [83] reported air oxidation resulting in a polymerization of the activated PEG. Overcoming such problems, Deng et al. [84] reported a new approach employing oxidation of mPEG to mPEG-aldehyde in anhydrous DMSO/acetic anhydride, grafting it onto chitosan through a subsequent reduction with NaCNBH3, while Bentley et al. [83] and Dal Pozzo et al. [85] synthesized PEG-dialdehyde diethyl acetals which were in situ converted to PEG acetaldehyde hydrates for cross-linking of chitosan, thus avoiding the problem associated with the reactive aldehyde in the former method. Other studies on the formation of a hydrogel based on PEGylated chitosan in drug delivery applications by Kiuchi et al. [75] and Tanuma et al. [74] found that the use of

Handbook of Chitin and Chitosan

3.4 Fabrication of PEGylated chitosan derivatives

67

diepoxide-terminated PEG as a cross-linker reacted easily with the primary amino group of chitosan forming hydrogel films. In these cases, alkylating agents were suggested that conjugate with chitosan in a way that allows the retention of net charge. On the other hand, nonprotonable amide or carbamate linkers with PEG-NHS and PEG-p-nitrophenyl alkylating agents resulted in the loss of the protonable center. During the formation of PEG-NHS derivatives, the most widely used approach in the process of PEGylation of chitosan, the first step is the activation of an end group of PEG through carboxylation, afforded through the use of an anhydride, such as succinic or glutaric. In the second step, the carboxylic group of the activated PEG is converted to a Nhydroxy succinimidyl ester which is finally grafted onto the primary amino group of the chitosan using the 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) coupling agent, in aqueous condition. A procedure proposed by Saito et al. [64] grafted mPEG (Mw 5 kDa) on the chitosan (Mw 70 kDa) backbone starting with mPEG-p-nitrophenyl carbonate with a DS (degree of substitution) of 78.5 wt.%, together with a resulting solubility up to a pH value of 7.4, whereas Jang et al. [72] utilized mPEG-p-nitrophenyl carbonate (Mw 5 kDa) and cholesteryl chloroformate to graft low Mw chitosan (19 kDa), forming nanocarriers (30
150 nm and 1.1
11.5 mV). The same activated intermediate of mPEG was exploited by Jiang et al. [86], who utilized the mPEG-p-nitrophenyl carbonate with a Mw of 5 kDa to graft a chitosan having Mw of 47 kDa with a DS of 94%, where conjugation was carried out in an aqueous/DMSO mixture, leading to a PEGylated chitosan having DS of 3.6% and 9.6% (Fig. 3.4).

3.4.2 O-substitution Although derivatization of chitosan by PEGylation on the amino group is the most ascribed viable method in literature, some authors have explored the derivatization of the polysaccharide on the hydroxyl group, while other studies have described PEGylation at both aOH and aNH2 groups [87,88]. Lately, an investigation by Gorochovceva et al. [57] reported the synthesis of novel PEGylated chitosan derivatives which are soluble in water with different degrees of solubility (DS)

FIGURE 3.4 PEG-p-nitrophenyl carbonate derivatization method.

Handbook of Chitin and Chitosan

68

3. PEGylated chitin and chitosan derivatives

through the aOH functional groups by an etherification reaction between N-phthaloyl chitosan and PEG monomethyl ether iodide while using Ag2O as catalyst, with some limitations regarding the removal of the trace amount of Ag2O dispersed in the final product and achieving the desired solubility of the product in water or common organic solvents, unless the copolymerization is strictly controlled with a high DS [15]. The solubility of such PEGylated chitosans in water and aqueous medium with 2 kDa PEG and a DS from 5% to 197% was found in a range of pH from 4.0 to 10.0. In the normal case, compared to PEGylation at the amino group, mPEG-o-chitosan shows solubility in an aqueous medium, providing less effect on the backbone of chitosan retaining its free amino group when DS is controlled at approximately 15% [89]. A promising approach by Makuska et al. [82] described three different ways of grafting mPEG on the hydroxyl group of chitosan (Mw 400 kDa) at C-6 using 6-oxo-2-N-phthaloylchitosan, 6-O-dichlorotriazine-2-N-phthaloulchitosan, and 3-O-acetyl-2-N-phthaloylchitosan intermediates with a high DS (about 90%) in chitosan with a high density of mPEG (2 kDa). Another study by Yoksan et al. [70] utilized modified mPEG into mPEG-COOH which was further conjugated to chitosan via ester linkage, whereas Huang et al. [90] reported PEGylation of carboxyl derivatives of chitosan at the carboxylic (hydroxyl) group through an esterification reaction between PEG (Mw 1.5 kDa) and 6-O-succinate-Nphthaloyl-chitosan. On the other hand, Ouchi and coworkers [62,91] reported the phenomenon of grafting mPEG-COOH (5 kDa) on 6-triphenylmethyl chitosan (150 kDa) and investigated the aggregation process along with their use as carriers (nanoparticles) for insulin. In their study, they found a continuous aggregation behavior of PEG-g-chitosan in aqueous solution due to intermolecular hydrogen bonds between moieties, while the water-soluble PEG chains acted as a hydrophilic shell stabilizing the nanoparticles. Similar to the previous study, Yang et al. [92] also confirmed the self-aggregation behavior of mPEG-g-chitosan describing the mechanism of weak intermolecular hydrogen bonding between the aNHaCOaand aOH groups in the copolymer. The hydrophobic interaction among the hydrophobic moieties in mPEG-g-chitosan (e.g., aCH2CH2, acetyl groups, and glucosidic rings) along with the ionic strength were also proposed to affect self-aggregation. In recent times, Cai et al. [93] also found the same behavior in PEGylated hexanoly chitosan derivatives. Such micellization behavior of PEGylated chitosan was characterized by a fluorescence probe method using pyrene and a strong pH medium dependency was found (the size of aggregated nanoparticles decreased with decreasing the medium of pH). The critical aggregation concentration (CAC) was 6.61 3 1026 g/mL with a PEGylated DS of 6.5%.

Handbook of Chitin and Chitosan

3.4 Fabrication of PEGylated chitosan derivatives

69

3.4.3 Miscellaneous approaches Together with amino group substitution and O-substitution, researchers working on the derivatization of chitosan have used many unconventional approaches to PEGylate chitosan. For example, a study by Shantha et al. [94] synthesized graft microspheres of chitosan (PEGylated derivatives) following a polymer dispersion technique using PEG-diacrylate by free radical initiation in the presence of ceric ammonium nitrate ((NH4)2Ce(NO3)6), though grafting PEG-diacrylate resulted in increased hydrophilicity of the copolymer, thus enhancing the formation of aggregates. However, Kong et al. [95] first ascribed the synthesis procedure of PEGylated chitosan derivatives (mPEG 2 kDa and medium Mw chitosan) utilizing an alternative approach based on a free radical polymerization in the presence of potassium persulfate at the initiation step. mPEG was first activated under stirring conditions in the presence of triethylamine and acryloyl chloride in dichloromethane and refluxed for 4 h at 40 C. Furthermore, the final product was obtained in a 1% acetic acid aqueous solution, stirred at 50 C for 4 h. Maksuka and coworkers, on the other hand described the utilization of “click chemistry” in the synthesis of PEGylated chitosan. In this process, N-azidation of chitosan was done using trifluoromethane sulfonyl azide or imidazole1-sulfonyl azide hydrochloride with a degree of azidation up to 40% and 65%, respectively. Moreover, mPEG-g-chitosan copolymers comprising triazolyl moiety were obtained by a coupling process via 1,3dipolar cycloaddition between azide and end alkyl groups of chitosan and acetylene-terminated mPEG. Herein, clicking of mPEG alkyne onto soluble azidated chitosan was carried out in a water/methylene chloride mixture (Fig. 3.5).

3.4.4 Solubilization of chitosan prior to derivatization Preparation of an intermediate of chitosan, which is soluble in organic solvents and can be obtained under mild conditions, is an important route as part of PEGylation process. In order to increase the solubility and link different moieties onto the primary aOH groups in the final product, many research groups first synthesized organosoluble N-phthaloyl intermediates protecting the amino moiety on the backbone of chitosan, although the process has an adverse impact on the

FIGURE 3.5 PEGylation of chitosan applying click chemistry.

Handbook of Chitin and Chitosan

70

3. PEGylated chitin and chitosan derivatives

backbone of chitosan. A study by Lebouc et al. [80] reported that the deprotection of the phthaloyl intermediate with hydrazine remains incomplete, retaining 20% of the phthaloyl group in the final product. A significant approach enhancing the solubility of chitosan in water and in organic solvent is its complexation in the presence of negatively charged surfactant. In fact, the strategy can be significantly implemented for all the polycations in common organic solvents. A study proposed by Cai et al. [79] applied sodium dodecylsulfate to sodium dodecylsulfate/chitosan complex solubilizing chitosan in organic solvents. Additionally, the process specified the regions so that organosoluble species could be conjugated selectively onto the hydroxyl groups of chitosan. The solubility of such copolymers was found in solvent DMF and DMSO even when DS value exceeded 24%. On the other hand, Fangkangwanwong and coworkers [88] reported the utilization of hydroxybenzotriazole (HOBt) resulting in complexation to bring chitosan into solution followed by PEGylation (using mPEG-COOH Mw 2 kDa; in a one-pot synthesis) of both the amino and hydroxyl groups in water. During the process, sparingly soluble chitosan attained a good solubility in its derivatized product (ratio 2:1, pH 4.4).

3.4.5 PEGylated cross-linked chitosan Upon cross-linking the resultant insoluble matrix of chitosan shows poor or nonswelling activity at an intestinal pH value of 7.4 in the lower small intestine. Such phenomena also limit the release of active agents from the cross-linked chitosan network. Kulkarni et al. described a PEG cross-linked chitosan together with a prominent swelling behavior at pH of 1.1 and 7.4, found in the stomach and the lower intestinal environment, respectively, in their study. In this scenario, PEG exhibited both the role of a cross-linker and a swelling agent. The cross-linking of chitosan was performed by PEG in the presence of formaldehyde. While treating with formaldehyde, the free amino groups of chitosan formed an intermediate, which further reacted with the hydroxyl groups of PEG to form a cross-linked product with a good swelling ratio in both simulated stomach and intestinal solutions, and which could be beneficial in an oral sustained drug transportation system. Another approach by Tanuma and coworkers [74] reported the formation, and characterization (thermal, mechanical, and swelling properties), of PEG cross-linked chitosan hydrogel employing PEGs of different molecular weights (2 and 4 kDa). In their study, PEG was converted to diepoxy PEGs which were further treated with chitosan synthesizing a three-dimensional network. The content of PEG in such a cross-linked product was found to be between 17% and 44%,

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

71

influencing the behavior of the hydrogel films, and thus potentially applicable in a drug delivery system.

3.5 Characterization of PEGylated chitosan and chitin derivatives It is important to fully characterize the PEGylated chitosan copolymers after the copolymerization, in order to determine the structural conformation of the newly synthesized products and to establish their physicochemical properties. This can be done by characterizing the copolymer with respect to its molecular weight, degree of substitution, and confirming the formation of new bonds on the amino or hydroxyl groups. The analysis of solubility at a wide pH range, mechanical properties such as ductility, aggregation of copolymer, and cytotoxic profile evaluation against different types of healthy cells are also important for complete characterization.

3.5.1 Structural analysis Structural analysis refers to the experiments by which the chemical and physical properties can be evaluated of synthesized PEG-g-chitosan copolymer. PEGylated chitosan derivatives are usually characterized by FT-IR, 1H, and 13CNMR, in some instances with 2D experiments such as heteronuclear multiple quantum coherence (HMQC) a CaH single bond correlation, in order to identify the newly formed bonds. Change in molecular weight is an important criterion to confirm the grafting of PEG on a chitosan backbone, which can be determined with gelpermeation chromatography (GPC). Several techniques, described below, including thermogravimetric analysis (TGA), differential scanning calorimetry (DSC), and wide-angle X-ray diffraction (XRD), have been commonly used to measure the physical properties of PEGylated chitosan. 3.5.1.1 Fourier-transform infrared spectroscopy analysis Structural changes and new bond formation or breaking can be examined by means of FTIR analysis. Pure chitosan shows distinct amide I and amide II peaks at 1650 and 1580 cm21. Because of the presence of three distinct vibrational modes of CaOaC, CaOH, and CaC ring, vibrations peaks are found between 1000 and 1150 cm21. NaH stretching and OaH stretching vibrations can be characterized by the broad peak in the region of 3200
3500 cm21. To increase the yield of PEGylation or grafting-modified chitosan, such as

Handbook of Chitin and Chitosan

72

3. PEGylated chitin and chitosan derivatives

6-O-triphenylmethylchitosan [15], N-phthaloylchitosan [96] has been extensively used in many studies. The presence of these additional groups shows characteristic peaks in the FTIR spectrum. For example, in the case of 6-O-triphenylmethylchitosan, additional peaks at 3080
3027 cm21 for CaH stretching and 763, 747, and 703 cm21 for the aromatic ring of 6-O-triphenylmethyl appear and N-phthaloylchitosan shows characteristic peaks of its keto group at 1776 and 1714 cm21 and aromatic ring stretching at 721 cm21 before the grafting of PEG. Pure PEG shows distinct peaks for OaH, NaH, CaO, and CaOaC stretching at 3472, 2875, 1736, and 1105 cm21, respectively [96]. Peaks at 1280, 947, and 843 cm21 of PEG FTIR spectra represent the crystalline region of PEG [97]. Grafting of PEG on the chitosan backbone causes the shifting, intensifying of existing peaks, or appearance and disappearance of peaks in the spectra. Grafting of mPEG on the 6-O-triphenylmethylchitosan causes the disappearance of the peaks related to the 6-O-triphenylmethyl moieties which suggest the introduction of PEG on chitosan [15]. The FT-IR spectra of mPEG-g-chitosan showed absorption bands associated with the mPEG backbone at 842, 960, and 2915 cm21, in addition to two bands related to ester and amide functionalities at around 1732 and 1651 cm21, respectively, corresponding to the ester and amide groups present in the linker between mPEG and chitosan [67], and an increase in the absorbance of the 1580 cm21 peak compared to that in the pure chitosan spectrum, suggesting an attractive intermolecular interaction between chitosan and PEG [98]. 3.5.1.2 Nuclear magnetic resonance analysis Nuclear magnetic resonance (NMR) is a very important and precise spectroscopic method which can be used for the confirmation of new bond formation. Many nuclei can be studied by NMR techniques, but 1 H and 13C are most readily used. NMR provides information about the number of magnetically distinct atoms of the type being studied. One can determine the number of each of the distinct types of either 1H or 13 C nuclei as well as obtain information regarding the nature of the immediate environment of each type. Fig. 3.6 shows the 1H spectra of mPEG, mPEG-NHS, low-molecular-weight water-soluble chitosan (LMWSC) 10K, and PEG-g-chitosan copolymer [63]. Specific peaks of mPEG are present between 3.5 and 3.7 ppm with methyl group peak at 3.3 ppm. In the activated form, mPEG-NHS, the peaks of the carboxylic group and NHS group are present between 2.5 and 3.0 ppm (peak numbers 8
11). The peaks specific to chitosan (LMWSC 10K) appear between 1.8 and 5.0 ppm. Peaks related to mPEG-g-chitosan copolymer highlight appropriate chemical shifts such as the methyl group of mPEG shifted to about 3.7 from 3.3 ppm.

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

FIGURE 3.6

73

1

H spectra of mPEG, mPEG-NHS, LMWSC 10K, and PEG-g-chitosan copolymer in D2O [63]. Source: Copyright Elsevier Publishers, 2008.

Fig. 3.7 shows the 13C NMR spectrum of the PEG-g-chitosan copolymer. The mPEG peaks appeared between 55 and 80 ppm and those for the carboxylic group and NHS groups were present at 25
30 and 168
172 ppm, respectively. The peaks for chitosan (LMWSC 10K) appeared at 100 ppm (C1), 57 ppm (C2), 72
78 ppm (C3), and 61
62 ppm (C6). In the ChitoPEG copolymer, peaks associated with LMWSC 10K and mPEG both appeared. The 1H
13C HSQC map of LMWSC 10K and the chitoPEG copolymer shows shifts of LMWSC 10K. In the chitoPEG copolymer, C1 (100 ppm) and C2 (57 ppm) appeared clearly, but C3
C6 of LMWSC 10K overlapped with the methyl group of mPEG. The acetyl group appeared at 23 ppm. In comparison with chitosan, peaks corresponding to aNHaCH2CH2Oaappeared at 2.8 and 47.1 ppm on the 1H NMR and 13C NMR spectra of PEG-g-chitosan, respectively [80]. 3.5.1.3 Gel permeation chromatography or size exclusion chromatography Gel permeation chromatography (GPC) and size exclusion chromatography (SEC) are extensively used for the determination of the molecular weight, polydispersity, and conjugation in copolymers. In this method, the polymer of interest is dissolved in a suitable solvent and is

Handbook of Chitin and Chitosan

74

3. PEGylated chitin and chitosan derivatives

13 C NMR spectra of mPEG, mPEG-NHS, LMWSC 10K, and chitoPEG copolymer in D2O [63]. Source: Copyright Elsevier Publishers, 2008.

FIGURE 3.7

eluted through a column made of various inorganic or organic material grafted onto an inorganic surface, such as silica. With the help of a detector the eluate can be examined for the identification and quantification of the polymer. Polymer chains with smaller molecular weight have higher attraction toward column materials compared with larger polymer chains. Because of this, polymer chains with higher molecular weight have a lower retention time than smaller ones. Applying these methods, it is possible to confirm the grafting of PEG on chitosan. Different types of solvent (such as combination of 0.3 M acetic acid and 0.2 M sodium acetate or 0.5 M ammonium acetate buffer) for elution [86] and detectors (such as multiangle light scattering (MALLS), refractive index, and a viscometer) for identification can be used for the precise determination of Mn and Mw of the copolymer. An increase in molecular weight results in a lower retention time for the grafted PEG/ chitosan polymer compared with the pure chitosan and PEG. By comparing the retention time of the grafted polymer with the different feed ratio of PEG, it is possible to find the optimum conditions for producing grafted polymer with the highest yield [99]. For example, retention time for low-molecular-weight water-soluble chitosan and methoxy-PEG was found to be 34 and 30 min, while for the grafted copolymer it was less

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

75

than 30 min; it has been found to be even lower in the case of higher grafted copolymer. These methods can also be used to verify the process efficiency. The copolymer of chitosan and PEG with unimodal molecular weight distribution in the GPC eluograms represents the high efficiency of the grafting process [100]. 3.5.1.4 Thermogravimetric analysis TGA has been utilized to investigate the thermal decomposition temperatures (Td) of the PEGylated chitosan copolymers [88,101]. It provides information about the decomposition pattern of a substance with respect to the change in temperature. This technique allows the determination of changes in weight in relation to the change in temperature and it is commonly used for characterizing polymers. Analysis of unmodified, modified, or grafted chitosan thermograms gives details about the changes in chemical and physical properties. Unmodified chitosan exhibits two sigmoids, one in the 35 C
130 C range due to the loss of absorbed water and the second one from 210 C to 350 C resulting from the degradation of chitosan. On the other hand, PEG-g-chitosan displays a different TGA pattern, with three observed weight loss intervals. The first interval ranges between 40 C and 80 C, corresponding to the evaporation of water, the second from 200 C to 240 C, possibly resulting from decomposition of the chitosan backbone, and the third stage of weight loss is observed from 320 C to
380 C, which could be ascribed to the decomposition of PEG units [67]. In the above studies, the authors described PEG-g-chitosan as being more thermally stable than the original chitosan, a phenomenon which the authors attributed to the destruction of part of the hydrogen bonds between the chitosan chains and part of the chitosan domains at the new mPEG chains [84]. 3.5.1.5 Differential scanning calorimetry DSC is another useful technique for the characterization of PEGylated chitosan [15,84,102]. It is a thermoanalytical technique which enables measurement of the difference in the amount of heat required to increase the temperature of a sample versus a reference, measured as a function of the temperature. This technique can be used to observe fusion and crystallization events as well as glass transition temperatures (Tg). Deng et al. [84] found that the DSC curve of chitosan showed an endothermic peak before 100 C, attributable to the elimination of bound water, together with a small broad transition peak from 210 C to 220 C. In case of the mPEG-g-chitosan copolymer, an endothermic peak was observed at around 185 C, which is attributed to a liquid
liquid transition where PEG side chains might act as internal plasticizers.

Handbook of Chitin and Chitosan

76

3. PEGylated chitin and chitosan derivatives

3.5.1.6 Wide-angle X-ray diffraction analysis XRD is a useful technique that allows the characterization of crystalline materials, determination of their unit cell dimensions, and sample purity. The diffractograms of both pure chitosan and PEG show sharp peaks attributed to their crystalline nature [97]. The crystalline region of pure chitosan shows a strong reflection at 19.7 degrees and a relatively weak reflection centering at 10 degrees, which have been used to estimate crystallinity. In the case of PEG, reflections at 18.74 and 22.86 degrees and weak reflections at 26.77, 30.5, 35.9, and 40 degrees are observed due to its crystalline nature [102]. But the diffractogram of PEGylated chitosan shows a broader peak in the 15
25 degrees regions compared with the native chitosan and PEG. It proves the grafting of PEG on chitosan backbone suppressed the crystallization of both polymers at the molecular level. Three new sharp peaks at 12.8, 13.6, and 27.5 degrees have been reported for PEG-g-chitosan copolymer above 45% DS [16]. These newly formed peaks are not attributed to either chitosan or PEG and are suggested to be due to the possibility of the formation of a new probable tropism arrangement of the PEG side chains.

3.5.2 Determination of the degree of substitution Degree of substitution means the amount of PEG has been grafted onto the polysaccharide backbone of the chitosan by means of different methods as described earlier [86]. DS is responsible for making a copolymer either soluble or insoluble in a specific solvent. It has been reported that there is a lowest limit of DS that must be necessary to allow the copolymer dissolve in water or organic solvents [62]. DS also influences the aggregation, ductility, and cytotoxicity properties of the PEG-g-chitosan copolymers, as described in the following sections. The amount of DS can be controlled by varying the feed ratio of chitosan to PEG [103]. It is not possible to get DS equal to the feed ratio of chitosan and PEG. So, determination of DS is necessary. DS can be expressed in many ways which leads to the confusion and misunderstanding in comparing the properties in different studies. Usually weight/weight (w/ w) or moles/moles (mol/mol) is used to express the DS of PEGylated chitosans, where the former refers to the weight percentage of PEG in the total weight of the copolymer, and the latter usually indicates the percentage of chitosan glucosamine repeating units conjugated with PEG [67]. Several different techniques such as gel permeation [63], 1H NMR, weight ratio%, colorimetric method, colloidal titration, potentiometric titration, elemental analysis, and gravimetric analysis have been applied to calculate the DS of the PEGylated chitosan.

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

77

In GPCthe DS of the PEG-g-chitosan copolymer is determined by comparing the molecular weight of the final PEG-grafted copolymer and the molecular weight of the starting materials (i.e., PEG and chitosan) [67,104]. The degree of chitosan substitution can be calculated using average molecular weight values determined by GPC analysis, as follows [67]: DS% 5

MnðconjÞ 2 MnðcsmÞ MnðmPEGÞ

3

1 3 100% DPðcsmÞ

where Mn(conj) is the number average molecular weight of the mPEG-gchitosan conjugate, Mn(csm) is the number average molecular weight of the chitosan starting material, Mn(mPEG) is the number average molecular weight of the mPEG starting material, and DP(csm) is a number of chitosan glucosamine repeating units. 1 H NMR has been particularly suited for calculating DS (mol/mol) by comparing the integral ratio between OCH3 singlet at approximately 3.2 ppm of the grafted mPEG and specific peaks of the chitosan D-glucosamine repeating units [7,100]. DS can also be calculated by using elementary analysis [84,96,105]; DS of mPEG-g-chitosans was measured by an element analyzer and was calculated using the following equation. DS 5

C=N 3 100% 77:42

where C and N are the contents (wt.%) of carbon and nitrogen element in PEG-g-chitosans, respectively, and 77.42 is the C/N value at DS of 100 mol%, at which all the amine groups in chitosan are substituted by mPEG. However, denominator 77.42 changes with the type of chitosan used to make the PEGylated chitosan. For example, in an another study, where elemental analysis was used to determine DS, N-phthaloylchitosan was used as the starting material [96]. In this case the C/N ratio of the reference for N-phthaloylchitosan-grafted mPEG was 105.23 for a complete substitution of all amine groups. Colloidal titration method has been used also for DS measurement in numerous studies [62,79]. In this method a negative colloid solution of PEG-g-chitosan is titrated with 1/400 N polyanionic solution of potassium poly(vinyl sulfate) by the conventional toluidine blue indicator method. The degrees of amino-substitution for PEG-g-chitosan-SA (SA, stearic acid) were assayed using 2,4,6-trinitrobenzenesulfonic acid solution (TNBS) reaction [66,106]. In this process, a solution of grafted copolymer is added to NaHCO3 at pH 8.5 and an aqueous TNBS solution. The reaction was carried out at 37 C for 2 h and HCl was added prior to the

Handbook of Chitin and Chitosan

78

3. PEGylated chitin and chitosan derivatives

absorbance study conducted by a UV spectrophotometer at 344 nm. The degree of substitution, termed as the substitute number by PEG per one hundred amino groups in chitosan molecules, was then calculated from the UV absorbance of PEG-g-chitosan-SA and chitosan solution with the same content. Another modified colorimetric method is based on the partitioning of a chromophore (present in ammonium ferrothiocyanate reagent) from the aqueous phase to an organic phase (chloroform) in the presence of mPEG [107]. The characteristics (e.g., DS%, method used for DS determination, starting chitosan, and PEG materials) of a range of PEGylated chitosans derivatives are given in Tables 3.1
3.3. TABLE 3.1 Degree of substitution (DS) of N-PEGylated chitosan copolymers [89]. DS%

Method used for DS determination

Chitosan Mw (kDa)

PEG Mw (kDa)

5
37

1

H NMR

28

0.55/5

3.2
27.9

1

H NMR

255

3.4
5

58.5/66.1

1

H NMR

50
100

0.55/5

0.46
1.19

1

H NMR

87

5

5
74

1

H NMR

91

0.55
2

5
8

1

H NMR

10

5

3.57

1

H NMR

7**

5

20***

1

H NMR

19

5

34

1

H NMR




5

14.2

1

H NMR

100

5

1.1/45

1

H NMR

65

1.1/5

0.56

1

H NMR

,150

5

3/90

1

H NMR

5/750*

2

6.6
7.2

1

H NMR

50

3

6
23

1

H NMR

100**

2

19/42

1

H NMR

200*

2

25/91.4

1

H NMR

70
300

2

7/35

1

H NMR

22
79

0.55
2**







1000

2

0.5

1

350**

5**

H NMR

(Continued)

Handbook of Chitin and Chitosan

79

3.5 Characterization of PEGylated chitosan and chitin derivatives

(Continued)

TABLE 3.1 DS%

Method used for DS determination

Chitosan Mw (kDa)

PEG Mw (kDa)

1***




70**

2

15/42

1

H NMR

210*

2

0.5/4.5

1

H NMR

Medium

3.8
5**

1.8

1

H NMR

200

2**

1.8/48.7

1

H NMR

Medium

2

13/40

1

H NMR

305

2

3.6
9.6

1

H NMR

47

5

6.1/19.4

1

H NMR

10

2

2.1/6.8

1

H NMR

28.9
82

1.9
5

5.8/14.3}

Weight ratio%

64.9

5

2.5/11.5}

Weight ratio%

12.2

2

43.7/86.6}

Weight ratio%

400

0.55/5**

3.5
20

Colloidal titration

330*

0.12
2

2
55

Colloidal titration

150

5

0.12/0.48

Potentiometric titration

405*

0.55/5**

78.5
92.7}

Gravimetrically

70

5

21.7
48.7}

Weight ratio%

200

2/3.4

1/52}

Weight ratio%

600

5

22.6/54.8}

Weight ratio%

5
56**

2**

23
89

Colorimetric method

400

2

90

Colorimetric method

48

2

0.1/0.42

Elemental analysis

65*

1.1/5

5
6

Elemental analysis




6.6**

6
22

Elemental analysis

56**

2

9.8/245

Elemental analysis

6
20

3.5
7.5

11.8/19.8

TNBS

450

2

9.4
9.9

TNBS

10

2

4.6
5.3

TNBS

40

2







50

2


means “and”; / means “from. . . to. . .”; *, average molecular viscosity 5 Mv; **, number average molecular weight 5 Mn; ***, as estimated by authors; }, weight% (means the ratio of PEG in PEGylated chitosan calculated by different methods).

Handbook of Chitin and Chitosan

80

3. PEGylated chitin and chitosan derivatives

TABLE 3.2 Degree of substitution (DS) of O-PEGylated chitosan copolymers [89]. DS%

Method used for DS determination

Chitosan Mw (kDa)

PEG Mw (kDa)

16.5

1

H NMR

578

2

4.3/21.6

1

H NMR

300*

2**

9.3
31.8

1

H NMR

Medium

5**

21.67

1

H NMR

48

5

3.3/7

1

H NMR

40/50

3.5
7.5

20/64

1

H NMR

Medium

2

13/42.7

1

H NMR

757*

1.5

6.3
35.1}

Weight ratio%

600*

5**

12.8

1

4

5







480**

2**

7.3/59.8

1

337/870*

2
5

31/96

Colorimetric

400

2

177.69

Weight ratio%

Low

5

B20/379

Weight ratio%

30 mPa s

1.5**

7.83

Elemental analysis

170*

2
5

4.5
6.68

Weight ratio%

72**




26.2

Thermal analysis

260

5







170*

0.55
5**

H NMR

H NMR


means “and”; / means “from. . . to. . .”; *, average molecular viscosity 5 Mv; **, number average molecular weight 5 Mn; }, weight% (means the ratio of PEG in PEGylated chitosan calculated by different methods).

TABLE 3.3 Degree of substitution (DS) of N,O-PEGylated chitosan copolymers and miscellanea [89]. DS%

Method used for DS determination

Chitosan Mw (kDa)

PEG Mw (kDa)

10/42

1

H NMR

560

2

0.68
1.5

1

H NMR

405*

0.55
2

36/320

Weight ratio%

75 cps

0.4







Medium

2**


means “and”; / means “from. . . to. . .”; *, average molecular viscosity 5 Mv; **, number average molecular weight 5 Mn.

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

81

3.5.3 Ductility One of the key concerns about chitosan’s use is the fragile nature of chitosan films. It has a very high modulus of elasticity along with low strain to break owing to the high Tg and crystallinity. By either blending or copolymerizing with other polymers, the morphology and ductility of chitosan can be improved. Several blends of synthetic polymers with chitosan have been evaluated, but PEG is of particular interest due to the hydrophilic, biocompatible, and biodegradable nature of PEG. Evmenko and coworkers suggested the significant improvement in mechanical properties of chitosan upon the addition of PEG [108,109]. A study has been conducted to understand how the ductility of chitosan can be improved by blending and copolymerizing with PEG [97]. The component interactions responsible for the change in properties of chitosan when blended and copolymerized with PEG were investigated using FTIR and X-ray diffraction, and characterized by comparing the properties of chitosan/PEG blends at different compositions and comparing them with a copolymer. Ductility was improved in all compositions of blend, as demonstrated by a decrease in modulus and an increase in strain at break. A 50/50 blend of chitosan and PEG showed the lowest elastic modulus and a 60/40 blend showed the highest strain at break among the compositions of blend tested. The decrease in elastic modulus was 56%, while increase in strain at break was 125% for the 50/50 blend. But for chitosan-g-PEG (31% PEG), the decrease in elastic modulus was 28%, while the increase in strain at break was 88%. So, it can be said that for a comparable PEG composition, the properties of the solution-cast blend were better than those of the grafted copolymer. It suggests that blending is a more efficient way to improve ductility of chitosan. It appears from the FTIR and X-ray diffraction studies that in the case of the graft copolymer the improvement in the properties comes from suppression of the crystallinity of each component and not from component interactions. While in case of the blend, the improvement appears to come predominantly from the well-dispersed, kinetically trapped phase morphology and from the intermolecular interactions. Annealing doesn’t change the properties of the grafted copolymer because it is a one-component system but has significant effect on twocomponent system blends of chitosan and PEG. There was no significant difference in the X-ray diffraction pattern of the blends after annealing, which suggests that crystallinity has no important role in the deterioration in ductility. Decreased intermolecular interactions, phase coarsening, are the driving force behind the deterioration in mechanical properties.

Handbook of Chitin and Chitosan

82

3. PEGylated chitin and chitosan derivatives

3.5.4 Solubility One of the most significant hindrances to chitosan being a perfect material in biomedical application, is its water-insoluble nature. Chitosan is insoluble in most of the commonly used solvents, whether it is water or any other organic solvents like DMF, DMSO, acetone, etc. Chitosan has shown solubility in acidic environments with pH ,3. Derivatization of chitosan with PEG causes the improvement in the solubility of chitosan even in basic pH which is not possible to achieve with normal nonderivativized chitosan. Solubility of the PEG-g-chitosan derivatives depends mainly on two factors: (1) the degree of substitution of PEG on polysaccharide backbone of chitosan and (2) the length (or molecular weight) of the PEG chain chemically linked with chitosan [7,63]. The PEG-g-chitosan can be soluble not only in water, but also in organic solvents such as DMSO, DMF, acetone, and ethanol. An increase in DS of PEG-g-chitosan increases the solubility of the copolymer in water. This is expected as the increase in DS of PEGylated chitosan is attributed to the decrease of intermolecular interactions, such as van der Waals forces and hydrogen bonding [110]. To determine the relationship of DS with the solubility Chan et al. [110] operated an experiment using a new term called pH50. This pH50 is defined as the pH when the transmittance of a polymer solution at 600 nm has reached 50% of the original value. They observed the copolymers with DS between 3 and 28 mol% of PEG (Mw 3, 4, and 5 kDa). It was found that pH50 increased with increasing DS of PEGylation; samples with DS values below 6% were poorly soluble. The solubility of the PEG-g-chitosan in a specific solvent can be controlled by controlling the DS of PEG [62]. For example, Ouchi et al. [62] prepared various PEG-g-chitosans copolymers with various DS values (2, 7, 18, 22, 25, 30, 38, 47, or 55 mol%/sugar unit) by varying the feed ratio of MeO-PEG acid to 6-O-triphenylmethyl-chitosan by the coupling reaction. All of the prepared PEGylated chitosan copolymers were soluble in DMF and DMSO, while only PEG-g-chitosans having a DS value over 22 mol%/sugar unit (determined by colloidal titration) were soluble in chloroform. PEG-g-chitosans with DS values over 10 mol%/sugar unit were soluble in water after sonication. The effect of polymer chain length or molecular weight can be easily understood by comparing the experiments conducted by Ouchi et al. [62] and Hu et al. [15]. Ouchi et al. synthesized PEG-g-chitosan by using MeO-PEG with molecular weight of 5000 Da, while Hu et al. used mPEG with 2000 Da molecular weight. Due to the variation in molecular weight or chain length of the chemically grafted PEG on the chitosan backbone, the solubility of the copolymer varies significantly. While Ouchi et al. found their copolymer to be soluble with all DS values (2, 7, 18, 22, 25, 30, 38, 47, or 55 mol%/sugar unit) in DMSO and DMF, Hu

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

83

et al. showed that PEGylated chitosan dissolved in organic solvents such as DMF or DMSO only if DS is greater than 24%. The solubility of PEG-g-chitosan in water largely depends on the chain length of the attached PEG with the chitosan. Sugimoto et al. [7] reported that PEG-g-chitosans (chitosan Mw 28 kDa) with long PEG side chains (e.g., Mw 2 or 5 kDa) (DS ranging from 0.5 to 37 mol%/sugar unit) were soluble in water, whereas PEG-g-chitosans with shorter PEG side chains (e.g., Mw 550) (DS ranging from 0.3 to 23 mol%/sugar unit) did not exhibit water solubility. Yang et al. [111] suggested that the substitutes on the amino groups can deform the rigid crystalline structure of chitosan, thus disturbing the intra- and intermolecular hydrogen bonding, leading to the enhancement of hydrophilicity. Numerous studies have been done on the solubility of the grafted copolymers in different buffers at different pH values, such as acetate buffer (pH 4.0), phosphate buffer (PBS, pH 7.4), carbonate buffer (pH 10.0) and in different organic solvents, including CH3Cl, DMF, and DMSO [7,15,89]. Jeong et al. [63] found that PEGylated chitosan with a DS ranging from 5% to 20% and PEG molecular weight of 2 kDa showed an aqueous solubility up to pH 11, in contrast to that of a normal chitosan (IS-chitosan), which showed a limit of solubility beyond pH 6 (Fig. 3.8).

3.5.5 Cytocompatibility Chitosan is considered as a safe biomaterial to be used in biomedical application, although it has not yet been certified with GRAS status. It has been extensively used as a nontoxic, safe, and good biomaterial for

FIGURE 3.8 Solubility of chitoPEG copolymer in aqueous solution as a function of pH changes in the buffer solution [63]. Source: Copyright Elsevier Publishers, 2008.

Handbook of Chitin and Chitosan

84

3. PEGylated chitin and chitosan derivatives

topical and oral administration of pharmaceuticals but more research is required for it to be used as a safe material for the parenteral routes [89]. Potential new pharmaceutical and biomedical applications can further be increased by decreasing the present cytotoxicity and increasing the biocompatibility. In general, it has been found that cytotoxicity is reduced with the introduction of a PEG chain to the backbone of chitosan. PEG-gchitosan has shown no toxicity against cancer cells in a study conducted by Bae et al. [104]. They had evaluated the toxicity profile of mPEG-gchitosan (CS Mw 4 kDa, mPEG Mw 5 kDa and DS 12.8% by 1H NMR) by using mouse melanoma B16F10 cell line (a model of a primary tumor) using a CCK-8 cell viability assay, which depends on the mitochondrial dehydrogenase activity. But after making a polyelectrolyte complex of micelles composed of PEG-g-chitosan/heparin, the copolymer exhibited a high cytotoxicity against B16F10 cell line. It was possible to achieve a 42.4% 6 4.5% reduction in cell viability at a dosing concentration of 100 g/mL. Another study evaluating the cytotoxic effect of PEG-g-chitosan on healthy cells like erythrocytes, leukocytes, and platelets was conducted by Radhakumary et al. [76]. They tested the toxicity of poly (ethylene glycol monomethacrylate)-g-chitosan (PEG Mw 0.4 kDa, DS 36%, 164%, 320%) films using different methods, such as direct contact assay, MTT assay, and live
dead assay. All copolymer films showed very high cytocompatability by showing 100% metabolically active cells (L929 cells) when compared to the controls. Zhang et al. [112] reported that PEGylated chitosans were associated with a cell viability (evaluated by MTT) of .70% in HeLa and A549 cells. A clear correlation between the DS of PEG and molecular weight of PEG with cell viability has been reported by Casettari et al. [67]. They have examined the cytotoxic profile of the Calu-3 cell line and found a positive effect on cell viability with the increase of DS and PEG molecular weight. In all cases cell viability was higher compared to unmodified chitosan. This phenomenon could be attributed to the steric effect of the conjugated PEG chains which provides shielding against positive charges on the chitosan molecule [100], as the extent of toxicity is considered to be related to the charge density and the spatial arrangement of the cationic residues on the chitosan molecule [113]. Mao et al. [100] also reported a linear correlation between the degree of substitution and IC50 value (after 3 h incubation) in cytotoxicity (L929 cell line) of trimethyl chitosan (TMC) with PEGylation, using the MTT and LDH assays (measuring metabolic activity and membrane integrity, respectively). All polymers exhibited a time-and dose-dependent cytotoxic response that increased with molecular weight. TMCs of 100 and 50 kDa demonstrated decreased cytotoxicities of more than 10fold after PEGylation (DS B6%). An increase in molecular weight of PEG showed a decrease in the toxicity of the copolymers. Eighty percent of the cells were observed to be still viable after 24 h incubation with a

Handbook of Chitin and Chitosan

3.5 Characterization of PEGylated chitosan and chitin derivatives

85

500 mg/mL polymer solution. Jiang et al. [86] reported on the toxicity effect of PEG-g-chitosan. In their study complexes of unmodified chitosan/DNA and PEG-g-chitosan/DNA, with varying DS (3.6% and 9.6%) were studied. By shielding the positive charge on the surface of the chitosan/DNA complex PEG lowered the acute toxicity, improved the particle stability, and mediated three times higher luciferase expression in the liver compared to unmodified chitosan.

3.5.6 Aggregation The modification of chitosan by PEG leads to the synthesis of chemically grafted biomaterial which can be soluble not only in water, but also in organic solvents. This enhanced solubility provides the biomaterial with more applicability. The introduction of PEG chains onto chitosan dissolve the modified glucosamine units of the chitosan molecule by interacting with water or organic solvents but the unmodified glucosamine units of PEG-g-chitosan still possess strong inter- or intramolecular interaction with other unmodified glucosamine units by means of hydrogen bonds, as found in native chitosan. Derivatives with a low PEG/chitosan ratio, corresponding to high positive charges, have more H-bonding connections, resulting in the formation of highly compact structures [114]. In a study, depending on the intermolecular hydrogen bonds interaction the formation of a new type of PEG-g-chitosan aggregate was reported and the aggregation phenomena was investigated in aqueous solution by measuring transmittance, light scattering, and uptake and release behavior of N-phenyl-L-naphthylamine (PNA) by the aggregates [62]. PEG-g-chitosan was prepared by the coupling reaction of 6-O-triphenylmethyl-chitosan with MeO-PEG acid in the presence of the water-soluble carbodiimide (WSC)-HOBOt in N,N-dimethylformamide (DMF) to give PEG-grafted 6-O-triphenylmethyl-chitosan [56]. Different degrees of PEG-g-chitosan were prepared by varying the feed ratio of MeO-PEG acid to 6-O-triphenylmethyl-chitosan. The degree of PEG-g-chitosan (DEPG) values were estimated by a colloidal titration method with a polyanionic solution of potassium polyvinyl sulfate [115]. The solution properties of PEG-g-chitosan were investigated by transmittance measurements. The transmittance of PEG-g-chitosan aqueous solutions was drastically changed with an increase in DPEG value of around 10
20 mol%/sugar unit. Transmission increases with the increase of DPEG of PEG-g-chitosan. These results suggest that the solubilization effect of PEG chains became predominant and the hydrogen bonds between chitosan moieties were inhibited by the static hindrance of PEG chain at the DPEG range over 20 mol%/sugar unit. The presence of intermolecular hydrogen bonds interaction is examined by the

Handbook of Chitin and Chitosan

86

3. PEGylated chitin and chitosan derivatives

FIGURE 3.9 Formation of intramolecular aggregate of PEG-g-chitosan by hydrogen bonds in aqueous solution.

addition effect of a small amount of a hydrogen bond breaker, dichloroacetic acid. It was found that the addition of dichloroacetic acid results in smaller particles which proves the presence of H-bonding in the moieties. PEG-g-chitosans did not show the formation of particles with obvious hydrodynamic diameters in DMSO. These results suggest the derivatives formed nanometer-sized particles in water by hydrogen bond formation at the chitosan moiety. To confirm the self-aggregate process, the particle size distribution of PEG-g-chitosan was investigated. In DMSO solution, there was no aggregation formation, however, after dialysis in water the formation of nanometer-sized particles with 70 nm of average hydrodynamic diameter was observed, which suggested that PEG-g-chitosans could form the aggregates spontaneously by strong intermolecular hydrogen bonds between chitosan moieties in water. The aggregation number of PEG-g-chitosan was evaluated. This number revealed that higher DPEG results in a lower aggregation number and thus formed looser aggregates or unimolecular micelles in aqueous solution, while derivatives consist of lower DPEG, have higher aggregation number, and more compact aggregates. These PEG-g-chitosan aggregates are able to take up a small hydrophobic molecule, such as PNA, that can be released from the aggregates by changing the pH to an acidic condition. So this PEG-g-chitosan aggregate can be expected to be used as a pH-dependent drug carrier for drug delivery systems (Fig. 3.9).

3.6 Applications of PEGylated derivatives of chitosan 3.6.1 Medical applications Intrinsic properties of native chitosan, such as aggregation by means of inter- and intramolecular hydrogen bonds, and complexation by

Handbook of Chitin and Chitosan

3.6 Applications of PEGylated derivatives of chitosan

87

electrostatic interaction with negatively charged compounds are usually expected to be maintained by the free amino groups of PEGylated chitosan. Such prominent properties allow the derivatized copolymer to be used in major medicinal applications. PEGylated chitosans, formed only by grafting PEG onto the amino or hydroxyl groups, may be employed during the transportation of peptides, proteins, oligonucleotides, and anionic drugs (e.g., insulin, heparin with the formation of polycomplexes) in the mucosal system. The use of nanoparticles in a drug delivery system depends frequently on their size and surface charge for cellular uptake and trafficking. Some groups have described selfaggregation (due to hydrogen bonding) of the derivatives in aqueous solution affording such activities. An investigation by Ouchi et al. [62] described the aggregation behavior of mPEG-g-chitosan (CS Mw 150 kDa and mPEG Mw 5 kDa) in an aqueous medium. In their study, they proposed nanosized (,120 nm) aggregates of final copolymer due to strong intermolecular hydrogen bonding between chitosan moieties. Similar to the previous study, Bae et al. synthesized stable PEG-g-chitosan/heparin (CS Mw 4 kDa, mPEG 5 kDa and a DS of 12.8% cal. by NMR) complex nanocarriers, applying a self-assembly process to induce apoptotic death of cancer cells in vitro. The aggregates (spherical shape of approximately 200 nm) were stable and well-dispersed throughout the serum under a physiological environment owing to steric stabilization of the PEG shell layer. Similar studies by Yang et al. reported the same phenomenon forming nanocarriers with an anticancer drug (methotrexate) using an mPEG-g-chitosan (CS Mw 200 kDa and mPEG 2 kDa). They found the aggregates (,300 nm) to have a drug absorption efficiency ranging from 21% to 95%, depending on DS and carrier to drug ratio. A release study, conducted at pH 7.4 C and 37 C, demonstrated an initial rapid release of drug by 40% in 4 h and a second slower release of the remaining drug over 48 h. Another study by Jeong et al. [116] produced and utilized mPEG-gchitosan (CS Mw 65 kDa and mPEG Mw 5 kDa) forming core
shell type polyion complex micelles to encapsulate all-trans retinoic acid (ATRA). In their study, the micelles were prepared following a simple procedure, followed by mixing of ATRA in PEGylated chitosan aqueous solution by ultrasonication, and then purification via dialysis. The micellar systems were of spherical shape with sizes in the range of 50
200 nm with a degree of PEGylation from 5.8% to 14.3% and had a drug loading efficiency greater than 80% (w/w). The structure of the polyion complex comprising an inner core and a hydrophilic outer shell was confirmed by 1H-NMR spectroscopy, whilst a cytotoxicity study using U87MG cells showed similar cytotoxicity to that of the free drug and a more effective tumor cell migration inhibition than free ATRA. The same

Handbook of Chitin and Chitosan

88

3. PEGylated chitin and chitosan derivatives

micelles were synthesized by Kim and coworkers [99] to encapsulate retinoic acid, where the degree of PEGylation of the copolymers ranged from 4.3% to 19.8% with sizes between 200
400 nm and a loading efficiency of 79.3%
96.4%. Numerous studies have also found nanoparticles comprising chitosan or PEGylated chitosan, obtained through utilizing the sodium tripolyphosphate (TPP) ionic gelation technique, as effective drug carriers [84,103,105,117,118]. In their studies, the nanosized PEGylated chitosan showed improved stability in gastrointestinal (GI) fluid, efficient surface interaction with the absorptive epithelium, and provided a protective effect for the associated peptide [119,120]. A study by Prego et al. [120] reported the capability of PEG-g-chitosan/TPP nanoparticles, not only enhancing and extending the intestinal absorption of salmon calcitonin, but also giving improved stability of the nanocapsules in GI fluids.

3.6.2 Thermoresponsive PEGylated chitosan hydrogels Chitosan and its derivatives in medicinal applications can not only be implemented as nanoparticles but also can be used as hydrogels. An earlier study patented by Bently et al. [121] reported the use of PEGylated chitosan for the formation of a gelling delivery system where gels were prepared by adopting various techniques and were characterized based on the release of lysosome and BSA from such a gel depot system. Later, Huh et al. [122] reported the preparation of hydrogels consisting of PEG-g-chitosan complexes with β 2 cyclodextrin. Such polymer inclusion complexes were suggested to be an effective hydrogel system, easily applicable in biomedical applications owing to the thermoreversible gel
sol transition, pH dependency of the gel, and microdomains having channel-type crystalline structures. Ganji and coworkers [123,124] produced an injectable thermoresponsibe hydrogel system in their studies where the PEGylated chitosan underwent a thermosensitive transition from solution at room temperature to a gel (studied by the vial inversion method and viscosity measurements) at approximately 36 C. The reaction involved the presence of potassium persulfate as a free radical initiator which led to a block copolymerized with PEG. On another study, Bhattarai et al. [125] found mPEG-g-chitosan copolymers for the production of thermoresponsibe polymer hydrogels which underwent sol
gel transition in response to temperature changes. They implemented the PEG-aldehyde method proposed by Harris and coworkers to graft PEG onto chitosan, obtaining the effectiveness of an injectable hydrogel with short-term protein release. According to the previous studies, natural as well as synthetic crosslinkers were found useful while producing hydrogels with improved

Handbook of Chitin and Chitosan

3.6 Applications of PEGylated derivatives of chitosan

89

properties. In a study, Lee et al. [126] reported a procedure to produce hydrogel based on PEG and chitosan following the simultaneous IPN method. UV radiation was utilized in achieving IPN formation using glutaraldehyde as a cross-linker. Similar to this, Wu et al. [127] proposed using quaternized chitosan in a thermosensible hydrogel formation for nasal drug delivery. In their proposed method, the quaternized chitosan chloride was simply mixed with PEG and a small amount of α-β-glycerophosphate, in a solution that could easily be transformed to hydrogel at around 37 C. In the in vivo experiments on male Sprague Dawley rats hydrogel formulations were found to decrease the blood glucose concentration by approximately 40%
50% of the initial blood glucose concentration for at least 4
5 h after administration, maintaining a biocompatible profile throughout. In a study by Khurma et al. [128], chitosan and PEG were utilized in the formulation of hydrogels while using genipin as a crosslinker. The swelling behavior of the hydrogel showed a dependency on temperature, pH, and PEG content. The swelling ratio was found to be increased with temperature (from 25 C to 37 C
45 C) due to the weakening of secondary interactions. On the other hand, the swelling ratio of the hydrogels was found to decrease with the increase of pH (from 1.2 to 7.4). Bhattarai and coworkers [125] also reported a simple procedure of formulating hydrogels using genipin as a cross-linker at physiological conditions (pH 7.4; PBS), which has been of great interest due to its excellent tissue compatibility in tissue engineering [129]. The hydrogel showed prolonged release behavior of the model protein, BSA, in a quasilinear long-term drug release. Such injectable, thermosensitive hydrogels were produced utilizing PEGylated chitosan where DS ranged from 45% to 55%. At higher DS values no gel was found even at 37 C. Recently, a study by El-Sherbiny et al. [130] reported the use of PEGylated chitosan (PEGylation of O-carboxymethyl chitosan) with 2,2-dimethoxy-2-phenyl acetophenone (DMPA) and methylene bisacrylamide (MBA) as crosslinking agents at numerous proportions (1% and 2%) for the formation of hydrogels. In their study, they loaded the hydrogels with 5-fluorouracil with a loading capacity from 71.0 6 1.1 to 86.6 6 0.9. Rafat et al. [131] reported the use of chitosan and collagen as cross-linkers activated by PEG/EDC/NHS to form hydrogels for corneal tissue engineering implants. The resulted scaffold attained promising enhancement in both mechanical strength and elasticity, while maintaining all the biological characteristics and optical clarity. Kiuchi et al. [75] and Tanuma et al. [74] also reported the use of chitosan for the formation of hydrogels using diepoxy PEG as cross-linker. The swelling behavior of the hydrogels showed their dependency on the molecular weight of PEG and its weight percentage. An increase in the equilibrium swelling ratio was attained by increasing the molecular weight of PEG while reducing its amount.

Handbook of Chitin and Chitosan

90

3. PEGylated chitin and chitosan derivatives

Formulated films of such hydrogels showed similar reversible temperature-dependent swelling behavior, whereas hydrogel samples with lower molecular weight and higher content of PEG showed more sensitivity to enzymatic degradation.

3.6.3 Gene delivery of chitosan derivatives Conventionally, gene delivery systems are of two kinds: viral vector and nonviral vector. Due to risk factors in viral vectors, nonviral delivery systems are effectively proposed to be safer alternatives [132,133]. In line with this, nonviral vectors are also considered to have the potential to be administered frequently with a minimal host immune response, with specific targeting, be stable during storage, and be easy to yield in bulk quantities. As the polycationic polymer
DNA complexes are more stable compared to other kinds of gene carriers, cationic polymers have been considered as prominent carriers among nonviral vectors, particularly while considering liposome or cationic
lipid systems [134]. In recent times, a number of cationic polymers have been studied as gene vectors, including polylysine and polyethyleneimine. However, an ideal gene vector with a higher efficiency of gene transfer, targeting ability, and good biocompatibility, particularly high stability and the ability to extend gene transfer, is assumed to be a significant feature for pharmaceutical application. Chitosan is an abundant and renewable carbohydrate polymer which can be biodegraded in vivo and has minimal toxicity [135]. Recently, the potential of chitosan as a polycation gene carrier has been reported by several researchers working with gene delivery system [136]. Chitosan, protonated under acidic conditions, forms complexes with anionic DNA through electrostatic interaction, and such complexes are appropriate for endocytosis by cells as chitosan has a better permeability through the cell membrane. Consequently, these complexes can easily be released from endosomes and enter into the nucleus [137]. pDNA (plasmid DNA) in such complex forms remains protected from enzymatic degradation, thus increasing the efficacy of gene delivery within the cell. So as to obtain optimum gene expression, pDNA has to hold its supercoiled circular and open circular forms. Hence, nanoparticle-based gene delivery systems should be implemented. PEG [138] is considered to be a good candidate owing to its high solubility in water, low cytotoxicity, and high cell permeability. It has been reported that PEG is able to increase plasma half-lives and stability while reducing the immunogenicity of proteins in vivo after being used to modify them, as especially indicated by the finding that PEGmodified tumor necrosis factor-α had a higher antitumor activity than

Handbook of Chitin and Chitosan

3.6 Applications of PEGylated derivatives of chitosan

91

did the unmodified factor in the murine Meth-A fibrosarcoma model [139]. A study by Choi and coworkers [140] reported that attaching PEG in lactose-PEG-grafted poly-L-lysine provided better solubility properties to the gene carrier complex, due to the steric hindrance caused by the attachment to PEG. To develop and characterize a novel PEGylation of chitosan nanoparticles for gene delivery, a study by Zhang et al. [141] prepared chitosan nanoparticles (following the polymer dispersion method) and chitosan
DNA nanoparticles with pEGFP-C1 through the complex coacervation of the cationic polymer (a plasmid that encodes EGFP; enhanced green fluorescent protein), whereas MSS-PEG (α-methoxyω-succinimidylpoly(ethylene glycol)) was conjugated to the surface of the chitosan
DNA complexes utilizing an active ester scheme; after that, the morphology of the nanoparticles were studied by transmission electronmicroscopy and atomic force microscopy. In addition, their grain distribution and zeta potentials were also determined using a laser grain analyzer. The cytotoxicities of the chitosan nanoparticles to HepG2 and L-02 cell lines were examined using an MTT (3-(4,5dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide) assay. DNase I was used to study the gene-protection effects of complexes to pDNA. Finally, the transfection activities of the chitosan
DNA complexes and PEG-grafted chitosan
DNA complexes were qualitatively evaluated in vitro and in vivo. In those experiments different doses of transferred pDNA were tested to determine the prominent conditions for gene delivery and compare the gene expression of different complexes in various tissues of rats. Wu and coworkers [142] synthesized a carboxymethyl chitosan with Mw of 46 kDa and conjugated to a modified PEI-g-PEG on the carboxyl end leaving its amino groups free. More recently, Xu et al. [143] reported an alternative approach to create the same copolymer. The method they followed involved the iodination of a hydroxyl group of chitosan with N-phthaloyl protection to conjugate chitosan with mPEGg-PEI. Such mPEG-g-PEI-grafted chitosan, with an intrinsic viscosity of 0.446 dL/g, was found to be water soluble with mPEG content of 51.3% (w/w) and 28.9% (w/w), respectively. mPEG-g-PEI-grafted chitosan was found to have good condensation capability, retaining the nucleic acid, with the formation of nanocomplexes having an average size of 155 nm and a zeta potential of 17.5 mV. Transfection experiments on human embryonic kidney cells described gene expression with little evidence of toxicity, pointing to mPEG-g-PEI-grafted chitosan as a prominent nonviral gene carrier. Another study by Kievit et al. [144] reported the use of PEGylated chitosan grafted with PEI as a potential carrier with stable binding and protection while delivering pDNA. In their study, they found PEI to be essential for effective DNA binding and

Handbook of Chitin and Chitosan

92

3. PEGylated chitin and chitosan derivatives

transfection in tumor cells while the PEGylated moiety effectively reduced the toxicity of PEI. A new approach by Noh et al. [145] also proposed the nanosized complexes of poly-L-arginine (PLR) and PEGylated chitosan derivatives carrying siRNA. In the proposed study, activated carboxylic groups of PLR were linked to amino groups of chitosan leading to the formation of amide bonds. Both the PEGylated chitosan and PLR-PEG-g-chitosan were utilized to form nanosized polymer/siRNA complexes with positive zeta potentials. PLR-PEG-gchitosan was found to increase the cellular delivery of siRNA, efficiently quieting the target gene expression in vivo, with low cytotoxicity and enhanced serum stability. PEGylation of PLR-chitosan did not show any interference with the cell membrane at the same time.

3.6.4 Formation of nanofibres PEG is considered as an ideal graft-forming polymer due to its promising solubility in both the water and organic solvents as well as having low toxicity, good biocompatibility, and biodegradability [146]. It has prominent applications in food, cosmetics, pharmaceuticals, and personal care products [147]. Similar to this, PEGylated chitosan exhibits improved affinity towards water and other organic solvents where the reaction is carried out commonly by reductive amination of chitosan with PEG-aldehyde in aqueous organic acid [7,125,148,149]. Water solubility was attained at a lower DS (even as low as 0.2%) for any derivative, whereas the PEG-N,O-chitosan with DS of 1.5 was found to be soluble in organic solvents, including CHCl3, DMF, DMSO, and THF. Electrospinning of all the aqueous solutions of PEGylated chitosan produced beads, whereas sprayed droplets were observed with aqueous solutions of all PEG-N-chitosan through reductive amination. The main challenge of fiber formation utilizing PEGylated chitosan is most likely having the presence of chain entanglement due to intermolecular interactions. To overcome such problem, the high surface tension of water needs high voltage, therefore nonionic surfactants (mostly Triton X-100) are used to reduce the surface tension while passing the aqueous solutions through the spinneret. However, even with lower surface tension, beads were still observed in the electrospinning process. Hence, organic solvents and cosolvents were employed. Electrospinning of 15% PEG-N, O chitosan with 75/25(v/v) THF/DMF cosolvents and 0.5% Triton X100 produced identical nanofibers with diameters ranging from 40 to 360 nm and having an average diameter of 162 nm, whereas electrospinning of PEG-N,O-chitosan at 25% in DMF solvent produced fibrous structures fused with beads [150]. Ultrafine fibers with diameters ranging from 40 to 360 nm with an average diameter of 162 nm were also

Handbook of Chitin and Chitosan

References

93

attained successfully through electrospinning of 15% PEG-N,O-chitosan in 75/25 (v/v) THF/DMF cosolvents with 0.5% Triton X-100 surfactant [149]. None of the aqueous solutions of PEG-N-chitosan or PEG-N,Ochitosan alone could be electrospun into fibers.

3.7 Conclusions PEGylated chitin/chitosan can be realized as potential derivatives that offer superior physical and chemical properties compared with the base polymer. The modification of chitin/chitosan is usually done by simple grafting of PEG on the backbone of chitin/chitosan, which drastically improves its solubility (in both polar and nonpolar solvents system), stability, ductility, cytocompatibility, thermal stability, aggregation behavior, and lowers cytotoxicity. The complete characterization of modified chitin/chitosan is important for appropriate applications. PEGylated chitin/chitosan are opening a vast avenue of applications in various emerging fields such as drug delivery, antiobesity agent, biofilm, hydrogel, nanofiber, and antimicrobial systems.

References [1] M.N.V.R. Kumar, et al., Chitosan chemistry and pharmaceutical perspectives, Chem. Rev. 104 (12) (2004) 6017
6084. [2] M.N.V. Ravi Kumar, A review of chitin and chitosan applications, Reactive Funct. Polym. 46 (1) (2000) 1
27. [3] O. Felt, P. Buri, R. Gurny, Chitosan: a unique polysaccharide for drug delivery, Drug. Dev. Ind. Pharm. 24 (11) (1998) 979
993. [4] R.A. Muzzarelli, Human enzymatic activities related to the therapeutic administration of chitin derivatives, Cell Mol. Life Sci. 53 (2) (1997) 131
140. [5] M.M. Bashar, M.A. Khan, An overview on surface modification of cotton fiber for apparel use, J. Polym. Environ. 21 (1) (2013) 181
190. [6] H. Sashiwa, Y. Shigemasa, Chemical modification of chitin and chitosan 2: preparation and water soluble property of N-acylated or N-alkylated partially deacetylated chitins, Carbohydr. Polym. 39 (2) (1999) 127
138. [7] M. Sugimoto, et al., Preparation and characterization of water-soluble chitin and chitosan derivatives, Carbohydr. Polym. 36 (1) (1998) 49
59. [8] K. Kurita, et al., Preparation of nonnatural branched chitin and chitosan, Chem. Lett. 27 (4) (1998) 317
318. [9] T. Sridhari, P. Dutta, Synthesis and characterization of maleilated chitosan for dye house effluent, IJCT 07 (4) (2000) 198
201. [10] N. Terada, et al., Synthesis of water-soluble oxidized chitosan derivatives and their biological activity, Chem. Lett. 28 (12) (1999) 1285
1286. [11] V.A. Alexandrova, et al., Modification of chitosan for construction of efficient antioxidant biodegradable macromolecular systems, Macromol. Symp. 144 (1) (1999) 413
422. [12] V.O. Sheftel, Indirect Food Additives and Polymers: Migration and Toxicology, CRC Press, 2000.

Handbook of Chitin and Chitosan

94

3. PEGylated chitin and chitosan derivatives

[13] P.C. Nalam, et al., Macrotribological studies of poly(L-lysine)-graft-poly (ethylene glycol) in aqueous glycerol mixtures, Tribol. Lett. 37 (3) (2010) 541
552. [14] N. Gorochovceva, et al., Synthesis and study of chitosan and poly(ethylene glycol) graft copolymers containing triazine moiety, Chemija, 15 (1) (2004) 22
27. [15] Y. Hu, et al., Preparation and characterization of poly (ethylene glycol)-g-chitosan with water-and organosolubility, Carbohydr. Polym. 61 (4) (2005) 472
479. [16] L. Liu, et al., Regioselective grafting of poly(ethylene glycol) onto chitosan and the properties of the resulting copolymers, Macromol. Biosci. 6 (10) (2006) 855
861. [17] V.S. Yeul, S.S. Rayalu, Unprecedented chitin and chitosan: a chemical overview, J. Polym. Environ. 21 (2) (2013) 606
614. [18] M.R. Bhuiyan, et al., A novel approach of dyeing jute fiber with reactive dye after treating with chitosan, Open. J. Org. Polym. Mater. 3 (4) (2013) 87. [19] S. Islam, M.R. Bhuiyan, M. Islam, Chitin and chitosan: structure, properties and applications in biomedical engineering, J. Polym. Environ. 25 (3) (2017) 854
866. [20] T.U. Rashid, et al., A new approach for the preparation of chitosan from γ-irradiation of prawn shell: effects of radiation on the characteristics of chitosan, Polym. Int. 61 (8) (2012) 1302
1308. [21] P.K. Dutta, M.N.V. Ravikumar, J. Dutta, Chitin and chitosan for versatile applications, J. Macromol. Sci. Part C. 42 (3) (2002) 307
354. [22] K. Kurita, Controlled functionalization of the polysaccharide chitin, Prog. Polym. Sci. 26 (9) (2001) 1921
1971. [23] R.A.A. Muzzarelli, Natural Chelating Polymers; Alginic Acid, Chitin, and Chitosan, Pergamon Press, Oxford; New York, 1973. [24] K.M. Rudall, W. Kenchington, The chitin system, Biol. Rev. 48 (4) (1973) 597
633. [25] K.M. Rudall, Chitin and its association with other molecules, J. Polym. Sci. Part C. Polym. Symp. 28 (1) (1969) 83
102. [26] E. Atkins, Conformations in polysaccharides and complex carbohydrates, J. Biosci. 8 (1
2) (1985) 375
387. vol. 8, 375
387. [27] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (7) (2006) 603
632. [28] H. Tamura, T. Hamaguchi, S. Tokura, Destruction of rigid crystalline structure to prepare chitin solution, 7 (2004) 84
87. [29] R.C. Capozza, Spinning and shaping poly-(N-acetyl-D-glucosamine), Google Patents, 1976. [30] A. Einbu, et al., Solution properties of chitin in alkali, Biomacromolecules 5 (5) (2004) 2048
2054. [31] M. Vincendon, Regenerated chitin from phosphoric acid solutions, Carbohydr. Polym. 32 (3) (1997) 233
237. [32] D. Gagnaire, J. Saint-Germain, M. Vincendon, NMR studies of chitin and chitin derivatives, Die Makromol. Chem. 183 (3) (1982) 593
601. [33] M. Vincendon, 1H NMR study of the chitin dissolution mechanism, Die Makromol. Chem. 186 (9) (1985) 1787
1795. [34] T. Sannan, K. Kurita, Y. Iwakura, Studies on chitin, 2. Effect of deacetylation on solubility, Die Makromol. Chem. 177 (12) (1976) 3589
3600. [35] K. Kurita, Chemistry and application of chitin and chitosan, Polym. Degrad. Stab. 59 (1) (1998) 117
120. [36] Z. Guo, et al., The influence of molecular weight of quaternized chitosan on antifungal activity, Carbohydr. Polym. 71 (4) (2008) 694
697. [37] R.A. Muzzarelli, C. Jeuniaux, G.W. Gooday, Chitin in Nature and Technology, vol. 385, Springer, 1986. [38] N. Cartier, A. Domard, H. Chanzy, Single crystals of chitosan, Int. J. Biol. Macromol. 12 (5) (1990) 289
294.

Handbook of Chitin and Chitosan

References

95

[39] N. Kubota, Y. Eguchi, Facile preparation of water-soluble N-acetylated chitosan and molecular weight dependence of its water-solubility, Polym. J. 29 (1997) 123. [40] S.-i Aiba, Studies on chitosan: 3. Evidence for the presence of random and block copolymer structures in partially N-acetylated chitosans, Int. J. Biol. Macromol. 13 (1) (1991) 40
44. [41] M. Rinaudo, G. Pavlov, J. Desbrie`res, Influence of acetic acid concentration on the solubilization of chitosan, Polymer 40 (25) (1999) 7029
7032. [42] M. Rinaudc, G. Pavlov, J. Desbrie`res, Solubilization of chitosan in strong acid medium, Int. J. Polym. Anal. Charact. 5 (3) (1999) 267
276. [43] A. Domard, pH and c.d. measurements on a fully deacetylated chitosan: application to CuII—polymer interactions, Int. J. Biol. Macromol. 9 (2) (1987) 98
104. [44] A. Chenite, et al., Novel injectable neutral solutions of chitosan form biodegradable gels in situ, Biomaterials 21 (21) (2000) 2155
2161. [45] A. Chenite, et al., Rheological characterisation of thermogelling chitosan/glycerolphosphate solutions, Carbohydr. Polym. 46 (1) (2001) 39
47. [46] G. Molinaro, et al., Biocompatibility of thermosensitive chitosan-based hydrogels: an in vivo experimental approach to injectable biomaterials, Biomaterials 23 (13) (2002) 2717
2722. [47] J. Cho, et al., Physical gelation of chitosan in the presence of β-glycerophosphate: the effect of temperature, Biomacromolecules 6 (6) (2005) 3267
3275. [48] J. Murata, et al., Inhibitory effect of chitin heparinoids on the lung metastasis of B16BL6 melanoma, Japanese J. Cancer Res. 80 (9) (1989) 866
872. [49] J.V. Jokerst, et al., Nanoparticle PEGylation for imaging and therapy, Nanomedicine 6 (4) (2011) 715
728. [50] K. Knop, et al., Poly(ethylene glycol) in drug delivery: pros and cons as well as potential alternatives, Angew. Chem. Int. Ed. 49 (36) (2010) 6288
6308. [51] F.M. Veronese, A. Mero, The impact of PEGylation on biological therapies, BioDrugs 22 (5) (2008) 315
329. [52] F.M. Veronese, Peptide and protein PEGylation: a review of problems and solutions, Biomaterials 22 (5) (2001) 405
417. [53] F.M. Veronese, G. Pasut, PEGylation, successful approach to drug delivery, Drug. Discov. Today 10 (21) (2005) 1451
1458. [54] J.M. Harris, R.B. Chess, Effect of pegylation on pharmaceuticals, Nat. Rev. Drug. Discov. 2 (2003) 214. [55] S.M. Ryan, et al., Advances in PEGylation of important biotech molecules: delivery aspects, Expert. Opin. Drug. Deliv. 5 (4) (2008) 371
383. [56] M. Yalpani, et al., Synthesis of poly(3-hydroxyalkanoate) (PHA) conjugates: PHAcarbohydrate and PHA-synthetic polymer conjugates, Macromolecules 24 (22) (1991) 6046
6049. [57] N. Gorochovceva, R. Makuˇska, Synthesis and study of water-soluble chitosan-O-poly (ethylene glycol) graft copolymers, Eur. Polym. J. 40 (4) (2004) 685
691. [58] W.-J. Lin, H.-G. Lee, Design of a microporous controlled delivery system for theophylline tablets, J. Control. Release 89 (2) (2003) 179
187. [59] W.J. Lin, H.K. Lee, D.M. Wang, The influence of plasticizers on the release of theophylline from microporous-controlled tablets, J. Control. Release 99 (3) (2004) 415
421. [60] S. Nishimura, et al., Chemospecific manipulations of a rigid polysaccharide: syntheses of novel chitosan derivatives with excellent solubility in common organic solvents by regioselective chemical modifications, Macromolecules 24 (17) (1991) 4745
4748. [61] M. Malhotra, et al., A novel method for synthesizing PEGylated chitosan nanoparticles: strategy, preparation, and in vitro analysis, Int. J. Nanomed. 6 (2011) 485
494. [62] T. Ouchi, H. Nishizawa, Y. Ohya, Aggregation phenomenon of PEG-grafted chitosan in aqueous solution, Polymer 39 (21) (1998) 5171
5175.

Handbook of Chitin and Chitosan

96

3. PEGylated chitin and chitosan derivatives

[63] Y.-I. Jeong, et al., Preparation and spectroscopic characterization of methoxy poly (ethylene glycol)-grafted water-soluble chitosan, Carbohydr. Res. 343 (2) (2008) 282
289. [64] H. Saito, et al., Graft copolymers of poly (ethylene glycol)(PEG) and chitosan, Macromol. Rapid Commun. 18 (7) (1997) 547
550. [65] J.M. Harris, et al., Synthesis and characterization of poly (ethylene glycol) derivatives, J. Polym. Sci. Polym. Chem. Ed. 22 (2) (1984) 341
352. [66] F.Q. Hu, et al., PEGylated chitosan-based polymer micelle as an intracellular delivery carrier for anti-tumor targeting therapy, Eur. J. Pharm. Biopharm. 70 (3) (2008) 749
757. [67] L. Casettari, et al., Effect of PEGylation on the toxicity and permeability enhancement of chitosan, Biomacromolecules 11 (11) (2010) 2854
2865. [68] M. Ravin˜a, et al., Hyaluronic acid/chitosan-g-poly (ethylene glycol) nanoparticles for gene therapy: an application for pDNA and siRNA delivery, Pharm. Res. 27 (12) (2010) 2544
2555. [69] Y.I. Jeong, et al., Polyion complex micelles composed of all-trans retinoic acid and poly (ethylene glycol)-grafted-chitosan, J. Pharm. Sci. 95 (11) (2006) 2348
2360. [70] R. Yoksan, et al., Controlled hydrophobic/hydrophilic chitosan: colloidal phenomena and nanosphere formation, Colloid. Polym. Sci. 282 (4) (2004) 337
342. [71] M. Amidi, et al., Preparation and characterization of protein-loaded N-trimethyl chitosan nanoparticles as nasal delivery system, J. Control. Release 111 (1
2) (2006) 107
116. [72] M.-K. Jang, J.-W. Nah, Characterization and modification of low molecular watersoluble chitosan for pharmaceutical application, Bull. Korean Chem. Soc. 24 (9) (2003) 1303
1307. [73] W.J. Lin, et al., Synthesis of lactobionic acid-grafted-pegylated-chitosan with enhanced HepG2 cells transfection, Carbohydr. Polym. 83 (2) (2011) 898
904. [74] H. Tanuma, et al., Preparation and characterization of PEG-cross-linked chitosan hydrogel films with controllable swelling and enzymatic degradation behavior, Carbohydr. Polym. 80 (1) (2010) 260
265. [75] H. Kiuchi, W. Kai, Y. Inoue, Preparation and characterization of poly (ethylene glycol) crosslinked chitosan films, J. Appl. Polym. Sci. 107 (6) (2008) 3823
3830. [76] C. Radhakumary, et al., Chitosan-comb-graft-polyethylene glycol monomethacrylate—synthesis, characterization, and evaluation as a biomaterial for hemodialysis applications, J. Appl. Polym. Sci. 114 (5) (2009) 2873
2886. [77] H. Wang, et al., Folate-PEG coated cationic modified chitosan
cholesterol liposomes for tumor-targeted drug delivery, Biomaterials 31 (14) (2010) 4129
4138. [78] H. Peng, et al., Methoxy poly (ethylene glycol)-grafted-chitosan based microcapsules: Synthesis, characterization and properties as a potential hydrophilic wall material for stabilization and controlled release of algal oil, J. Food Eng. 101 (1) (2010) 113
119. [79] G. Cai, et al., A facile route for regioselective conjugation of organo-soluble polymers onto chitosan, Macromol. Biosci. 9 (3) (2009) 256
261. [80] F. Lebouc, et al., Different ways for grafting ester derivatives of poly (ethylene glycol) onto chitosan: related characteristics and potential properties, Polymer 46 (3) (2005) 639
651. [81] M.M. Amiji, Synthesis of anionic poly (ethylene glycol) derivative for chitosan surface modification in blood-contacting applications, Carbohydr. Polym. 32 (3
4) (1997) 193
199. [82] R. Makuˇska, N. Gorochovceva, Regioselective grafting of poly (ethylene glycol) onto chitosan through C-6 position of glucosamine units, Carbohydr. Polym. 64 (2) (2006) 319
327.

Handbook of Chitin and Chitosan

References

97

[83] M.D. Bentley, M.J. Roberts, J.M. Harris, Reductive amination using poly (ethylene glycol) acetaldehyde hydrate generated in situ: applications to chitosan and lysozyme, J. Pharm. Sci. 87 (11) (1998) 1446
1449. [84] L. Deng, et al., Investigation on the properties of methoxy poly (ethylene glycol)/ chitosan graft co-polymers, J. Biomater. Sci. Polym. Ed. 18 (12) (2007) 1575
1589. [85] A. Dal Pozzo, et al., Preparation and characterization of poly (ethylene glycol)-crosslinked reacetylated chitosans, Carbohydr. Polym. 42 (2) (2000) 201
206. [86] X. Jiang, et al., Chitosan-g-PEG/DNA complexes deliver gene to the rat liver via intrabiliary and intraportal infusions, J. Gene Med. 8 (4) (2006) 477
487. [87] R. Yoksan, S. Chirachanchai, Amphiphilic chitosan nanosphere: studies on formation, toxicity, and guest molecule incorporation, Bioorg. Med. Chem. 16 (5) (2008) 2687
2696. [88] J. Fangkangwanwong, et al., One-pot synthesis in aqueous system for water-soluble chitosan-graft-poly (ethylene glycol) methyl ether, Biopolym. Orig. Res. Biomol. 82 (6) (2006) 580
586. [89] L. Casettari, et al., PEGylated chitosan derivatives: synthesis, characterizations and pharmaceutical applications, Prog. Polym. Sci. 37 (5) (2012) 659
685. [90] M. Huang, et al., Preparation of chitosan derivative with polyethylene glycol side chains for porous structure without specific processing technique, Int. J. Biol. Macromol. 38 (3
5) (2006) 191
196. [91] Y. Ohya, et al., Preparation of PEG-grafted chitosan nanoparticles as peptide drug carriers, STP Pharm. Sci. 10 (1) (2000) 77
82. [92] X. Yang, et al., Self-aggregated nanoparticles from methoxy poly (ethylene glycol)modified chitosan: synthesis; characterization; aggregation and methotrexate release in vitro, Colloid. Surf. B Biointerf. 61 (2) (2008) 125
131. [93] G. Cai, H. Jiang, pH-sensitive nanoparticles self-assembled from a novel class of biodegradable amphiphilic copolymers based on chitosan, J. Mater. Sci. Mater. Med. 20 (6) (2009) 1315
1320. [94] K. Shantha, D. Harding, Synthesis and characterisation of chemically modified chitosan microspheres, Carbohydr. Polym. 48 (3) (2002) 247
253. [95] X. Kong, et al., Synthesis and characterization of a novel MPEG
chitosan diblock copolymer and self-assembly of nanoparticles, Carbohydr. Polym. 79 (1) (2010) 170
175. [96] R. Yoksan, S.J.B. Chirachanchai, Amphiphilic chitosan nanosphere: studies on formation, toxicity, and guest molecule incorporation, Bioorg Medic. Chem. 16 (5) (2008) 2687
2696. [97] P. Kolhe, R.M.J.B. Kannan, Improvement in ductility of chitosan through blending and copolymerization with. [98] W.H. Jiang, S.J. Han, Study of interaction between polyethylene glycol and chitosan by viscosity method, J. Polym. Sci. Part B Polym. Phys. 36 (8) (1998) 1275
1281. [99] D.-G. Kim, Y.-I. Jeong, J.-W. Nah, All-trans retinoic acid release from polyioncomplex micelles of methoxy poly(ethylene glycol) grafted chitosan, J. Appl. Polym. Sci. 105 (6) (2007) 3246
3254. [100] S. Mao, et al., Synthesis, characterization and cytotoxicity of poly (ethylene glycol)graft-trimethyl chitosan block copolymers, Biomaterials, 26 (32) (2005) 6343
6356. [101] Z. Yao, et al., A series of novel chitosan derivatives: synthesis, characterization and micellar solubilization of paclitaxel, Carbohydr. Polym. 68 (4) (2007) 781
792. [102] J. Wu, et al., Water soluble complexes of chitosan-g-MPEG and hyaluronic acid, J. Biomed. Mater. Res. A 80 (4) (2007) 800
812. [103] X. Zhang, et al., PEG-grafted chitosan nanoparticles as an injectable carrier for sustained protein release, J. Mater. Sci. Mater. Med. 19 (12) (2008) 3525
3533.

Handbook of Chitin and Chitosan

98

3. PEGylated chitin and chitosan derivatives

[104] K.H. Bae, et al., Intracellular delivery of heparin complexed with chitosan-g-poly (ethylene glycol) for inducing apoptosis, Pharma, Res. 26 (1) (2009) 93
100. [105] X. Zhang, et al., Nasal absorption enhancement of insulin using PEG-grafted chitosan nanoparticles, Eur. J. Pharm. Biopharm. 68 (3) (2008) 526
534. [106] G. Avigad, A simple spectrophotometric determination of formaldehyde and other aldehydes: application to periodate-oxidized glycol systems, Anal. Biochem. 134 (2) (1983) 499
504. [107] K.W. Yang, et al., Novel polyion complex micelles for liver-targeted delivery of diammonium glycyrrhizinate: in vitro and in vivo characterization, J. Biomed. Mater. Res. A 88 (1) (2009) 140
148. [108] V. Alekseev, et al., Structure and mechanical properties of chitosan-poly (ethylene oxide) blend films, Poly. Sci, B 43 (9
10) (2001) 281
284. [109] V. Alexeev, et al., Improvement of the mechanical properties of chitosan films by the addition of poly (ethylene oxide), Polym. Sci. Engg. 40 (5) (2000) 1211
1215. [110] P. Chan, et al., Synthesis and characterization of chitosan-g-poly (ethylene glycol)folate as a non-viral carrier for tumor-targeted gene delivery, Biomaterials, 28 (3) (2007) 540
549. [111] T.-C. Yang, C.-C. Chou, C.-F.J.F.R.I. Li, Preparation, Food Res. Int. 35 (8) (2002) 707
713. [112] W. Zhang, et al., In vitro study on polyethylene glycol-chitosan copolymer as a gene delivery vector, Acta Pharm. Sini. 43 (8) (2008) 848
854. [113] A. Dyer, et al., Nasal delivery of insulin using novel chitosan based formulations: a comparative study in two animal models between simple chitosan formulations and chitosan nanoparticles, Pharm. Res. 19 (7) (2002) 998
1008. [114] Y. Chen, et al., Evaluation of the PEG density in the PEGylated chitosan nanoparticles as a drug carrier for curcumin and mitoxantrone, Nanomaterials, 8 (7) (2018) 486. [115] K. Kina, K. Tamura, I.N.J.J. Analyst, Use of 8-anilino-1-naphthalene sulfonate as a fluorescent indicator in colloid titration, Japan Analyst, 23 (9) (1974) 1082
1083. [116] C.-M. Lee, et al., Temperature-induced release of all-trans-retinoic acid loaded in solid lipid nanoparticles for topical delivery, Macromol. Res. 16 (8) (2008) 682
685. [117] Y. Akta¸s, et al., Development and brain delivery of chitosan 2 PEG nanoparticles functionalized with the monoclonal antibody OX26, Bioconjugate Chem. 16 (6) (2005) 1503
1511. [118] M. Kaloti, H. Bohidar, Kinetics of coacervation transition versus nanoparticle formation in chitosan
sodium tripolyphosphate solutions, Colloid. Surf. B Biointerf. 81 (1) (2010) 165
173. [119] N. Csaba, et al., Ionically crosslinked chitosan nanoparticles as gene delivery systems: effect of PEGylation degree on in vitro and in vivo gene transfer, J. Biomed. Nanotechnol. 5 (2) (2009) 162
171. [120] C. Prego, et al., Chitosan
PEG nanocapsules as new carriers for oral peptide delivery: effect of chitosan pegylation degree, J. Control. Release 111 (3) (2006) 299
308. [121] M.D. Bentley, X. Zhao, Hydrogels derived from chitosan and poly (ethylene glycol) or related polymers, Google Patents, 2003. [122] K.M. Huh, et al., Supramolecular hydrogel formation based on inclusion complexation between poly (ethylene glycol)-modified chitosan and α-cyclodextrin, Macromol. Biosci. 4 (2) (2004) 92
99. [123] F. Ganji, M. Abdekhodaie, Synthesis and characterization of a new thermosensitive chitosan
PEG diblock copolymer, Carbohydr. Polym. 74 (3) (2008) 435
441. [124] F. Ganji, M. Abdekhodaie, The effects of reaction conditions on block copolymerization of chitosan and poly (ethylene glycol), Carbohydr. Polym. 81 (4) (2010) 799
804.

Handbook of Chitin and Chitosan

References

99

[125] N. Bhattarai, et al., PEG-grafted chitosan as an injectable thermosensitive hydrogel for sustained protein release, J. Control. Release 103 (3) (2005) 609
624. [126] S.J. Lee, S.S. Kim, Y.M. Lee, Interpenetrating polymer network hydrogels based on poly (ethylene glycol) macromer and chitosan, Carbohydr. Polym. 41 (2) (2000) 197
205. [127] J. Wu, et al., A thermosensitive hydrogel based on quaternized chitosan and poly (ethylene glycol) for nasal drug delivery system, Biomaterials 28 (13) (2007) 2220
2232. [128] J.R. Khurma, A.V. Nand, Temperature and pH sensitive hydrogels composed of chitosan and poly (ethylene glycol), Polym. Bull. 59 (6) (2008) 805
812. [129] H.-W. Sung, et al., In vitro evaluation of cytotoxicity of a naturally occurring crosslinking reagent for biological tissue fixation, J. Biomater. Sci. Polym. Ed. 10 (1) (1999) 63
78. [130] I.M. El-Sherbiny, H.D. Smyth, Poly (ethylene glycol)
carboxymethyl chitosan-based pH-responsive hydrogels: photo-induced synthesis, characterization, swelling, and in vitro evaluation as potential drug carriers, Carbohydr. Res. 345 (14) (2010) 2004
2012. [131] M. Rafat, et al., PEG-stabilized carbodiimide crosslinked collagen
chitosan hydrogels for corneal tissue engineering, Biomaterials 29 (29) (2008) 3960
3972. [132] S.R. Winn, et al., Gene therapy approaches for modulating bone regeneration, Adv. Drug. Deliv. Rev. 42 (1-2) (2000) 121
138. [133] Y. Omidi, J. Barar, S. Akhtar, Toxicogenomics of cationic lipid-based vectors for gene therapy: impact of microarray technology, Curr. Drug. Deliv. 2 (4) (2005) 429
441. [134] M. Thanou, J. Verhoef, H. Junginger, Chitosan and its derivatives as intestinal absorption enhancers, Adv. Drug. Deliv. Rev. 50 (2001) S91
S101. [135] S. Mansouri, et al., Characterization of folate-chitosan-DNA nanoparticles for gene therapy, Biomaterials 27 (9) (2006) 2060
2065. ¨ . Guliyeva, et al., Chitosan microparticles containing plasmid DNA as potential [136] U oral gene delivery system, Eur. J. Pharm. Biopharm. 62 (1) (2006) 17
25. [137] G. Borchard, Chitosans for gene delivery, Adv. Drug. Deliv. Rev. 52 (2) (2001) 145
150. [138] P.C. Ross, S.W. Hui, Polyethylene glycol enhances lipoplex-cell association and lipofection, Biochim. Biophys. Acta 1421 (2) (1999) 273
283. [139] Y. Tsutsumi, et al., Molecular design of hybrid tumour necrosis factor alpha with polyethylene glycol increases its anti-tumour potency, Br. J. Cancer 71 (5) (1995) 963. [140] Y.H. Choi, et al., Lactose-poly (ethylene glycol)-grafted poly-L-lysine as hepatoma cell-targeted gene carrier, Bioconjugate Chem. 9 (6) (1998) 708
718. [141] Z.-S. Yang, et al., Synthesis and immunosuppressive activity of new artemisinin derivatives. 1.[12 (β or α)-dihydroartemisininoxy] phen(ox)yl aliphatic acids and esters, J. Med. Chem. 48 (14) (2005) 4608
4617. [142] Y. Wu, et al., A new biodegradable polymer: PEGylated chitosan-g-PEI possessing a hydroxyl group at the PEG end, J. Polym. Res. 15 (3) (2008) 181
185. [143] Z. Xu, et al., Synthesis of biodegradable polycationic methoxy poly (ethylene glycol)
polyethylenimine
chitosan and its potential as gene carrier, Carbohydr. Polym. 78 (1) (2009) 46
53. [144] F.M. Kievit, et al., PEI
PEG
chitosan-copolymer-coated iron oxide nanoparticles for safe gene delivery: synthesis, complexation, and transfection, Adv. Funct. Mater. 19 (14) (2009) 2244
2251. [145] S.M. Noh, et al., Pegylated poly-l-arginine derivatives of chitosan for effective delivery of siRNA, J. Control. Release 145 (2) (2010) 159
164.

Handbook of Chitin and Chitosan

100

3. PEGylated chitin and chitosan derivatives

[146] J.M. Harris, Introduction to biotechnical and biomedical applications of poly (ethylene glycol), Poly (Ethylene Glycol) Chemistry, Springer, 1992, pp. 1
14. [147] N. Tirelli, et al., Poly (ethylene glycol) block copolymers, Rev. Mol. Biotechnol. 90 (1) (2002) 3
15. [148] N. Gorochovceva, et al., Chitosan
N-poly (ethylene glycol) brush copolymers: Synthesis and adsorption on silica surface, Eur. Polym. J. 41 (11) (2005) 2653
2662. [149] J. Du, Y.-L. Hsieh, PEGylation of chitosan for improved solubility and fiber formation via electrospinning, Cellulose 14 (6) (2007) 543
552. [150] M.Z. Elsabee, H.F. Naguib, R.E. Morsi, Chitosan based nanofibers, review, Mater. Sci. Eng. C. 32 (7) (2012) 1711
1726.

Handbook of Chitin and Chitosan

C H A P T E R

4 Solubility, chain characterization, and derivatives of chitin Mi Feng1,2, Xingmei Lu1,2, Danfeng Hou1 and Suojiang Zhang1,2 1

CAS Key Laboratory of Green Process and Engineering, State Key Laboratory of Multiphase Complex Systems, Beijing Key Laboratory of Ionic Liquids Clean Process, Institute of Process Engineering, Chinese Academy of Sciences, Beijing, P.R. China, 2School of Chemical and Engineering, University of Chinese Academy of Sciences, Beijing, P.R. China O U T L I N E 4.1 Solubility of chitin 4.1.1 NaOH/urea aqueous solution 4.1.2 CaCl2/methanol solvent 4.1.3 N,N-dimethylacetamide/lithium chloride solvent 4.1.4 Ionic liquid

102 102 103 104 104

4.2 Chain characterization of chitin 4.2.1 The configuration of chitin 4.2.2 The length of chitin chain

107 108 111

4.3 Derivatives of chitin 4.3.1 The etherification of chitin 4.3.2 The graft of chitin 4.3.3 The O-acylation of chitin

112 112 115 119

Acknowledgment

125

References

125

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00004-3

101

© 2020 Elsevier Inc. All rights reserved.

102

4. Solubility, chain characterization, and derivatives of chitin

4.1 Solubility of chitin Chitin is a polymer[(1
4)-linked N-acetyl-β-D-glucosamine], which has three different crystalline polymorphic forms: α-chitin, β-chitin, and γ-chitin. The chitin chains are organized in sheets that are tightly held by a number of intrasheet hydrogen bonds, which leads to a denser structure and results in a difficulty in dissolution due to the difficult infiltration of the solvent. It is important to dissolve chitin in solvents for the preparation of functional materials. Although the compact structure of chitin makes it hard to dissolve, there are some solvent systems that address the destruction of its structure to dissolve it.

4.1.1 NaOH/urea aqueous solution As a novel green solvent, NaOH/urea aqueous solution was first used for the rapid dissolution of cellulose at low temperature by Zhang et al. [1]. Due to its excellent performance for the dissolution of biomacromolecules, Hu et al. first used NaOH/urea aqueous solution for the dissolution of chitin [2]. They found that 8 wt.% NaOH/4 wt.% urea was the suitable concentration and
20 C was the appropriate temperature. In addition, urea was considered to play an important role in the stability of chitin aqueous solution. Meanwhile, the chitin aqueous solution was sensitive to temperature and was transformed to gel when the temperature increased. Subsequently, Hu et al. used NaOH/urea aqueous solution to dissolve four kinds of natural chitin originating from crab, shrimp, silkworm chrysalis, and fly shells, and the dilute solution behavior of chitin was examined by laser light scattering and viscometry [3]. The results indicated that chitin molecules exist in a random coil chain conformation, and chain flexibility increases from C1 to C4 in NaOH/urea aqueous system. Zhang and her coworkers prepared transparent chitin solution without derivatization by completely dissolving chitin in 8 wt.% NaOH/4 wt.% urea aqueous solution via the freezing/thawing method [4]. The chitin solution was directly used for the preparation of hydrogel, which maintained the attractive structure and properties of chitin for the first time. Cai et al. used 1 M NaOH solution and 0.3% NaClO2 solution to pretreat the raw chitin powder. Then the purified chitin powder in the desired amount was dispersed into a mixed solution of NaOH
urea
H2O of 11:4:85 by weight to form a suspension (3
8 wt.%). Subsequently, a transparent and viscous chitin solution was obtained by freezing the suspension at
20 C overnight and then thawing at 5 C with stirring [5]. The results

Handbook of Chitin and Chitosan

4.1 Solubility of chitin

103

indicated that the most critical factors in improving solubility were the concentrations of NaOH and urea, and dissolution temperature. The best chitin solution was obtained by using a 11 wt.% NaOH/4 wt.% urea aqueous solution. The dissolution mechanism of chitin in NaOH/urea aqueous solution at low temperature was studied by Zhang et al. [6]. The results revealed that the formation of the hydrogen bond network between NaOH and chitin chains played a key role in the chitin dissolution, whereas urea had no direct interaction with chitin. The OH2 of NaOH and the amino group of urea formed the hydrogen bond, which made the urea hydrate clusters attach to the surface of the NaOH hydrogen-bonded chitin to form a soluble complex, leading to the dissolution of chitin.

4.1.2 CaCl2/methanol solvent A solution of calcium chloride dihydrate in methanol had excellent performance for the dissolution of chitin, and a viscous chitin solution was obtained [7]. The degradation of chitin was avoided due to the mild solvent system. Tokura et al. reported that the amount of water and the number of calcium ions had crucial effects on the dissolution of chitin in calcium chloride dihydrate-saturated methanol. Moreover, the solubility of chitin was also influenced by the degree of N-acetylation and the molecular weight (Mw) of chitin [8]. For the dissolution mechanism, it was proposed that calcium attacked the amide bond in the chitin side chains and broke the rigid crystalline structure [9]. To get the chitin hydrogels, both α-chitin powder and nanofiber, as the raw materials, were dissolved in calcium chloride dehydratesaturated methanol under mild conditions [10]. It was found that nanofibrillation of powder occurred during the treatment. To investigate the crystalline structure of β-chitin from squid pen, X-ray powder diffraction (XRD), studies were used on β-chitin which was treated with a calcium chloride dihydrate/methanol solvent system at different conditions [11]. The results showed that the CaCl2/methanol solvent influenced the plane (010) of β-chitin structure especially. Based on the calcium chloride dihydrate/methanol solvent, Kadokawa et al. developed a novel solvent system, in which calcium chloride dihydrate was substituted with calcium bromide dihydrate, to dissolve chitin for the gel [12]. In the study, the resulting gel was converted into a porous chitin by a regeneration technique. Moreover, the result revealed that the amounts of calcium bromide dihydrate in the gel had an important effect on both the porosities and mechanical properties of the porous chitin.

Handbook of Chitin and Chitosan

104

4. Solubility, chain characterization, and derivatives of chitin

4.1.3 N,N-dimethylacetamide/lithium chloride solvent In 1976 it was first reported that a mixture of lithium chloride (LiCl) in N,N-dimethylacetamide (DMAc) was used as a solvent for the dissolution of chitin under moderate conditions [13]. The Mw of chitin had effects on the content of chitin in the solvent, and solution containing up to 15 wt.% chitin was obtained. To determine the dissolution mechanism of chitin in the solvent, a chitin concentration of 1
3 wt.% was obtained by the use of N,Ndimethylacetamide containing 5 wt.% of lithium chloride [14] The 1H NMR result revealed that the strong interaction of the LiCl molecule with intermolecularly hydrogen-bonded hydroxyl and acetamide groups enabled the dissolution of chitin in DMA/LiCl. This interaction broke the intermolecular hydrogen bond and allowed chitin to swell and then dissolve in the solvent via the fixation of one LiCl molecular per labile proton. When it was used for the dissolution of synthetic polyamide, cellulose, and protein, it also had effective performance [15,16]. Due to its advantage, the solvent system was often used for the analysis of the polymer product by several techniques, such as Fourier-transform infrared spectroscopy (FT-IR), nuclear magnetic resonance (NMR), and gel permeation chromatography (GPC), which made it become an important solvent in the field of polymer analysis. Meanwhile, it also has potential in the application of the acetylation of chitin, in which it was regarded as the reaction medium.

4.1.4 Ionic liquid As novel and promising media, ionic liquids (ILs) have a number of favorable advantages, such as near-zero vapor pressure, high thermal stability, wide electrochemical window, low flammability, and tunable properties. These properties are responsible for their reputation as environment-friendly solvents and catalysts, and have been used extensively in various applications, such as electrochemistry [17
19], separations [20
22], biomass [23
25], and gas absorption [26
28]. Zhang et al. first reported that the ionic liquid 1-butyl-3-methyl-imidazolium chloride ([Bmim][Cl]) had excellent performance for the dissolution of chitin and chitosan, and a clear and viscous solution of 10 wt. % chitin/IL was easily obtained in 5 h by an oil bath at 110 C [29]. Kadokawa et al. used 1-allyl-3-methyl-imidazolium bromide ([Amim] [Br]) to dissolve chitin, and a clear mixture of 5% (w/w) and 7% (w/w) chitin with IL was obtained by heating at 100 C for 48 h [30]. They confirmed that degradation and a decrease of chitin’s Mw did not frequently occur during the clear liquid formation. Based on these results,

Handbook of Chitin and Chitosan

4.1 Solubility of chitin

105

they studied the chemical modification of chitin in IL by dissolving α-chitin in [Amim][Br] (2%, w/w) at 100 C for 24 h, which confirmed that [Amim][Br] was an effective solvent for the modification of α-chitin [31]. A chitin solution (10 wt.%) was obtained by mixing chitin with [Amim][Br] at 100 C for 24 h, which was used for the preparation of cellulose-chitin hybrid gel [32]. Sakurai et al. synthesized a novel biomass
ionic liquid 1-butyl-3-methylimidazolium acetate ([Bmim][OAc]) for the utilization of chitin with different origins and Mws [33]. They found that the chitin could be dissolved in [Bmim][OAc] at elevatedtemperature (more than 85 C) regardless of its origins and Mws, while a transparent viscous solutions could be obtained at satisfactory concentration at 110 C depending on the origins and Mws. The results revealed that the dissolution of chitin in [Bmim][OAc] was strongly related to its Mw, while the effect of the crystal forms of the chitin samples was ignored. A solution (6 wt.%) of chitin with a relatively lower Mw (β-chitin-L) could be obtained by [Bmim][OAc], while it could only dissolve 3 wt.% of the chitin sample with a relatively higher Mw (β-chitin-H). Simone and his coworkers prepared a chitin solution with [Bmim] [OAc] at 90 C
95 C for 5 h under stirring, and 3% w/v of the highest chitin concentration of the solution was obtained [34]. They found that the viscosity of the chitin/IL solution as well as dissolution time increased with increasing polymer concentration. To have a better understanding of the nature of dissolving chitin in ILs, Wang et al. used a series of ILs including 1-allyl-3-methylimidazolium chloride ([Amim][Cl]), [Bmim][Cl], 1-(2-hydroxyethyl)-3-methyl-imidazolium chloride ([C2OHmim][Cl]), 1,3-dimethylimidazolium dimethyl phosphate ([Mmim][Me2PO4]), 1-ethyl-3-methylimidazolium dimethyl phosphate ([Emim][Me2PO4]), and [Amim][OAc] to dissolve chitin with different degrees of deacetylation (DA) and Mw [35]. In their study, native chitin could be dissolved in [Amim][Cl], [Mmim][Me2PO4], and [Emim] [Me2PO4] at lower temperature, and the chitin concentration in [Amim] [OAc] was up to 5 wt.% at 110 C. The results showed that the dissolution behavior of chitin in ionic liquids was affected by the degree of acetylation (DA), the crystallinity, and the Mw, as well as the nature of the anion of the ionic liquid. ILs with stronger hydrogen bond acceptor ability could dissolve chitin more efficiently. Rogers et al. compared the ability of [Emim][OAc], [Bmim][Cl], and [Emim][Cl] to dissolve a given mass of pure chitin (chitin content of 94.7%
96.4%) and PG-chitin (practical grade) [36]. The results revealed that much more of the pure chitin sample (80.0%) could be dissolved in [Emim][OAc], compared to [Bmim][Cl] (24.4%) and [Emim][Cl] (13.9%). When PG-chitin was used as a raw material, [Emim][OAc] (15.2%)

Handbook of Chitin and Chitosan

106

4. Solubility, chain characterization, and derivatives of chitin

could dissolve more PG-chitin samples than [Bmim][Cl] (6.8%) and [Bmim][Cl] (4.2%). However, the dissolved load of PG-chitin was distinctly lower than pure chitin, which was presumably caused by the higher mineral content of PG-chitin. In another study, they reported that practical grade chitin was dissolved in [Emim][OAc] using microwaves, and a 1.5 wt.% solution was obtained [37]. Subsequently, they directly used [Emim][OAc] to extract chitin from dried shrimp shells via microwaves, and a chitin concentration of 0.67 wt.% was obtained. Buchmeiser et al. synthesized a series of phosphonium- and imidazolium-based ILs to dissolve chitin, and the influence of the cation on the dissolution process was compared [38]. To measure the maximum solubility of chitin in different ILs, a purified and depolymerized chitin was used. The results of the solubility studies of different ILs to dissolve chitin are summarized in Table 4.1. To verify the effect of cation in ILs on the dissolution of chitin, Jaworska et al. synthesized six ionic liquids—1-ethyl-3-methylimidazolium chloride ([Emim][Cl]), 1-butyl-3-methylimidazolium chloride ([Bmim][Cl]), 1-hexyl-3-methylimidazolium chloride ([Hmim][Cl]), 1-octyl-3-methylimidazolium chloride ([Omim][Cl]), 1-allyl-3-methylimidazolium chloride ([Amim][Cl]), and 1-butyl-2,3-dimethylimidazolium chloride ([Bdmim][Cl])—to dissolve chitin at 105 C for 48
72 h [39]. The measured solubility of chitin was less than 5% for all the ILs tested.

TABLE 4.1 Maximum solubility of chitin in different ILs [38]. IL name

Maximum solubility (wt.%)

Tetrabutylphosphonium glycinate

3

Tetrabutylphosphonium lysinate

2

Tetrabutylphosphonium valinate

4

Tetrabutylphosphonium propionate

3

1-ethyl-3-methylimidazolium gylcinate

6

1-ethyl-3-methylimidazolium lysinate

6

1-ethyl-3-methylimidazolium formiate




1-ethyl-3-methylimidazolium acetic

5

1-ethyl-3-methylimidazolium propionate

14

1-ethyl-3-methylimidazolium butanoate

13

1-ethyl-3-methylimidazolium pentanoate

4

1-ethyl-3-methylimidazolium octanoate

2

Handbook of Chitin and Chitosan

4.2 Chain characterization of chitin

107

The FT-IR spectra indicated that interactions between chitin and the used ILs were physical rather than chemical. To investigate the dissolution mechanism of chitin in ILs, Kadokawa et al. adopted a molecular dynamics (MD) approach to study the dissolution of chitin crystals in imidazolium-based ILs [40]. The MD simulation for [Amim][Br] demonstrated that the dissolution process involved peeling of chitin chains from the crystal surface, with Br2 cleaving the chitin hydrogen bonds, and [Amim]1 preventing a return to the crystalline phase after the peeling (as shown in Fig. 4.1). The bromide ions played a critical role in forming bridging via hydrogen bonds with chitin (NH Br2 HO) to induce the twisting and peeling of the molecular chain.





4.2 Chain characterization of chitin As described above, chitin is obtained by linking N-acetyl-2-amino-2deoxy-D-glucose with β-1.4-glycosidic bonds, as shown in Fig. 4.2. There

FIGURE 4.1 The mechanism of chitin solubility in the IL.

FIGURE 4.2 The structure of chitin.

Handbook of Chitin and Chitosan

108

4. Solubility, chain characterization, and derivatives of chitin

is a quaternary structure in chitin. The primary structure is a sugar chain formed by a β-1,4-glycosidic bonding to N-acetylglucosamine, excluding the interaction between chains. The secondary structure is a sugar chain of chitin. The interactions between the different molecules are formed by intramolecular or intermolecular hydrogen bonding between a hydroxyl group, an N-acetylamino group, and an amino group. This structure involves the conformation of the main chain of the chitin molecule, and excludes the spatial arrangement of the side chain. The tertiary structure is a spatially regular conformation formed by the primary structure and the secondary structure. The fourth-order structure refers to the aggregate formed by the noncovalent bond between the long chains. According to the structure of chitin, the characterization of the chitin chain mainly includes the configuration and the length [41].

4.2.1 The configuration of chitin As a long-chain polymer compound, chitin has a regular arrangement of rigid sugar chains and a large number of intermolecular and intramolecular hydrogen bonds, which is very favorable for the formation of a crystalline state. Thus there are crystalline regions and amorphous regions in chitin. The crystallization zone includes three crystal forms, namely α, β, and γ. The most common configuration is α-chitin, which is mainly found in the crustacean shell, the stratum corneum of arthropods, and the cell walls of certain fungi. β-Chitin is a less common configuration, mainly found in the scales of carp. γ-Chitin is a rare configuration found in the stomach of beetles and calamari [42]. The literature is mainly focused on α- and β-chitin, because γ-chitin is rare. However, the determination of the crystal structure of α- and β-chitin has undergone a long research process. Studies have shown that α-chitin is an orthorhombic system with a lattice constant of a 5 0.474, b 5 1.886, c 5 1.032, γ 5 90 degrees with P212121 spatial lattice [43]. The β-chitin is monoclinic, and its lattice constant is a 5 0.485, b 5 0.926, c 5 1.038, γ 5 97.5 degrees with P212121 spatial lattice [44]. In addition, it is generally believed that α-chitin is composed of two antiparallel sugar chains, β-chitin is composed of two parallel sugar chains, and γ-chitin is composed of two in the same direction and one opposite. The results showed that both α- or β-chitin have a layered structure formed by hydrogen bonding, and the distance between the molecular chains is maintained at 0.47 nm. Among them, α-chitin also has interlayer hydrogen bonding, but β-chitin does not have this effect. It can be seen that the crystal structure of α-chitin is tighter and more difficult to destroy than β-chitin [45]. Several characterization can be used to determine the configuration of the chitin, including FT-IR, XRD, and 13C NMR. The details are as followings.

Handbook of Chitin and Chitosan

4.2 Chain characterization of chitin

109

4.2.1.1 Fourier-transform infrared spectroscopy Generally, FT-IR spectroscopy is used to determine the characteristic groups of chitin, which can be used to distinguish the configurations of chitin. The FT-IR spectrum [42] of α-chitin is shown in Fig. 4.3, which is basically the same as the spectrum of β-chitin except for the peak of Amide I, OH, and NH. The characteristic peaks of Amide I in α-chitin are 1656 and 1619 cm21, while β-chitin only has one peak at 1656 cm21. The difference between the two spectra is due to their different structure. In the α-chitin, the CQO of Amide I exists in two forms, namely CQO. . .HaN and CQO. . .HOCH2a(side chain), corresponding to the absorption peaks at 1660 and 1627 cm21, respectively. Additionally, the characteristic peaks assigned to OH and NH (3600
3000 cm21) of α-chitin are a more detailed structure compared with β-chitin. A shoulder peak exists at 3284 cm21 in α-chitin, which corresponds to the OH (6). . .OQC intramolecular hydrogen bond. This peak cannot be found in β-chitin. The peak at 3444 cm21 is attributed to O(3)H. . .O(5) from the ring in the α-chitin, which is at a higher frequency than β-chitin (3426 cm21). Regarding the asymmetric stretching vibration of NH, the peak of α-chitin is at 3259 cm21 and β-chitin is at 3290 cm21. The details of other characteristic peaks in α- and β-chitin are as follows. 3108, 1255, and 697 cm21 correspond to the stretching, deformation, and out-plane bending vibration of NH, respectively. 2957 and 2886 cm21 are assigned to the stretching vibration of CH3. The 2928, 1414, and 747 cm21 peaks are attributed to the stretching, deformation, and in-plane bending vibration of CH2, respectively. The 1112, 1072, 1022, and 610 cm21 peaks are assigned to the stretching and out-plane bending vibration of CaO, respectively.

FIGURE 4.3 The FT-IR spectrum of α-chitin.

Handbook of Chitin and Chitosan

110

4. Solubility, chain characterization, and derivatives of chitin

4.2.1.2 X-ray powder diffraction spectroscopy The XRD patterns can be used to evaluate the crystallinity degree of chitin. The XRD pattern [46] of α-chitin is shown in the Fig. 4.4, which includes the 2θ at 9.25, 12.79, 19.27, 22.98, 26.19 degrees assigned to (020), (101), (040)/(110), (130), and (013), respectively. While the XRD pattern of β-chitin is different, including 2θ at 8.64, 10.42, 12.55, 18.78, 23.94, and 26.24 degrees, attributed to (010), (122), (102), (110)/(020), (114), and (121), respectively. Obviously, the α- and β-chitin can be distinguished using the XRD patterns. Moreover, the crystalline index (CI) can be calculated from the XRD patterns as follows: CI% 5 ½ðI110 2 Iam Þ 3 100

(4.1)

where I110 is the maximum peak at 19 degrees, and Iam is the intensity of the amorphous diffraction at 12.6 degrees. 4.2.1.3 Solid

13

C nuclear magnetic resonance spectroscopy

The solid C NMR of α-chitin is shown in Fig. 4.5, which involves eight carbon signals. In particular, the signals of C3 and C5 exist at 72.6 and 75.4 ppm, respectively. However, in the spectrum of β-chitin the signals of C3 and C5 overlap, at 75.2 ppm [46]. Additionally, the DA of the chitin can be obtained from the 13C NMR spectrum. However, as suggested by Rinaudo et al. [47], the DA value obtained from 13C NMR has a systematic error of about 4%, because the contact time of each carbon during magnetization is different. Generally, the DA value can be calculated as follows,   ICH3 DA% 5 3 100 (4.2) IC1 1 IC2 1 IC3 1 IC4 1 IC5 1 IC6 =6 13

FIGURE 4.4 The XRD spectrum of α-chitin.

Handbook of Chitin and Chitosan

4.2 Chain characterization of chitin

111

FIGURE 4.5 The solid 13C NMR spectrum of α-chitin.

where the ICH3, IC1, IC2, IC3, IC4, IC5, and IC6 are the intensity of each carbon.

4.2.2 The length of chitin chain The length of the chitin chain can be expressed by Mw. However, the Mw of chitin is a homologous mixture of different Mws that are the same as other polymer compounds, which is only statistically significant. During the process of chitin extraction, the chitin chain is destroyed leading to lower Mw. The literature has noted that different raw materials and methods for chitin preparation can lead to different Mw [48]. Generally, the viscosity method is used to determine the Mw of the chitin, and the lithium chloride/dimethylacetamide (LiCl/DMAc) is the normal solvent for dissolving chitin. As reported by Poirier [49], the Mw of the chitin can be obtained from the viscosity of LiCl/DMAc 5% (w/w) chitin solution. However, this method can only obtain the Mw of chitin dissolved in the LiCl/ DMAc (5%, w/w). The details of the measurement method are as follows. The LiCl was dried before preparing the 5% LiCl/DMAc solution. Then, the fixed mass of chitin was weighed to prepare different concentrations of chitin LiCl/DMAc solution between 0.03 and 0.05 g/dL. It is worth noting that the dissolution time needs more than 1 week, because the chitin is difficult to dissolve. After dissolution, the solution was

Handbook of Chitin and Chitosan

112

4. Solubility, chain characterization, and derivatives of chitin

filtered, then the viscosity was measured. The Mw can be calculated using the following equation, ½η 5 7:6 3 1025 M0:95 w where [η] is the intrinsic viscosity of the solvent.

4.3 Derivatives of chitin Although chitin has many advantages, there are a large number of intermolecular and intramolecular hydrogen bonds in the chitin structure. These make chitin difficult to dissolve in common organic solvents, which greatly limits the application of chitin. Therefore chitin is usually chemical modified to obtain chitin derivatives that have high solubility and that can maintain its advantages, including biodegradability and biocompatibility. This method achieves a high value of chitin. As shown in Fig. 4.2, the C6 and C3 positions of aOH in the chitin structure are the two reactive sites with higher activity. Due to the steric hindrance effect, C6-OH is more active than C3-OH [50]. The chemical reactions that can be carried out at these two sites are mainly etherification, grafting, and acylation reactions, which will be described in detail.

4.3.1 The etherification of chitin The etherification of chitin is usually carried out at the C6-OH site. The common etherified chitins are carboxymethyl-chitin and quaternized chitin, which are water soluble above a fixed degree of substitution (DS). 4.3.1.1 The carboxymethyl chitin Tokura et al. [51] reported the method for obtaining carboxymethyl chitin (CM chitin, as shown in Fig. 4.6) in 1983. In this process, the chitin was first prepared as alkali chitin by putting the chitin powder in 60% NaOH aqueous solution at 4 C and
20 C for 1 h and 1 night, respectively. Then the alkali chitin was used directly for CM chitin preparation. The obtained alkali chitin was dispersed in the isopropyl alcohol at room temperature, to which monochloroacetic acid was added until neutral. After separation, the residue was extracted with water. Acetone was added to the water extract to precipitate the CM chitin Na salt, which was treated with HCl (2 N) to give the CM chitin with DS of 0.6. In the FT-IR spectrum, the additional peak at 1730 cm21 attributed to carbonyl proved the carboxymethyl chitin was produced. The CM chitin is already available on the market, and has been studied and applied widely. According to Kong et al. [52], the CM chitin showed an antiobesity effect by inhibiting adipogenesis and inducing

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

113

FIGURE 4.6 The structure of carboxymethyl chitin.

lipolysis of 3T3-L1 cells through affecting the aquaporin-7 and adenosine monophosphate-activated protein kinase (AMPK) pathways. Moreover, CM chitin can be used to make nanomaterial. According to Dev et al. [53], CM chitin nanoparticles were prepared in a crosslinking process using FeCl3 and CaCl2; the obtained diameters were 200
250 nm. The CM chitin nanoparticle was nontoxic, and showed a good inhibitory effect on Staphylococcus aureus. What’s more, the cancer drug 5-Fluorouracil (5-F)-loaded CM chitin nanoparticle can be prepared by emulsion cross-linking with FeCl3 and CaCl2 as cross-linking agent. The in vitro studies showed that the loaded CM chitin nanoparticle can release 5-F sustainably. The CM chitin also can be used in other composite material. Shalumon et al. [54] synthesized a CM chitin/poly(vinyl alcohol) (PVA) nanofibrous scaffold by electrospinning. In this method, 7% CM chitin aqueous solution was mixed with PVA aqueous solution (8%) in different mass ratios to produce nanofiber scaffold using electrospinning. Then the watersoluble scaffold was cross-linked with glutaraldehyde into waterinsoluble products. The material has potential for tissue engineering applications, because calcium phosphate can be formed on the surface and human mesenchymal stem cells can be attached on and in it. 4.3.1.2 The quaternization of chitin Quaternization is also a common method to improve the solubility of chitin. The quaternized chitin could be soluble in water, and it has

Handbook of Chitin and Chitosan

114

4. Solubility, chain characterization, and derivatives of chitin

FIGURE 4.7 The structure of quaternized chitin.

antibacterial ability because of its high positive charge. The details of quaternized chitin preparation are as follows (Fig. 4.7). The β-chitin was used first to prepare quaternized chitin due to its loose structure compared with α-chitin. Chen et al. [55] prepared quaternized β-chitin, 2’-O-hydroxypropyltrimethylammoniumchitin chloride (2’-O-HTACCt), using β-chitin and 3-chloro-2-hydroxypropyltrimethylammonium chloride (CTA) in a NaOH aqueous solution with 2-propanol as dispersant. Their study pointed that excessive temperature, NaOH concentration, and experiment time were not conducive to the quaternization reaction, because of the deetherification reaction and side reactions. Therefore the optimal conditions were selected to be 40 wt.% NaOH aqueous solution, 40 C, 6 h, and mole ratio 4:1 (n:n, CTA:β-chitin unit), and 2’-O-HTACCt with DS of 85% and yield of 77% was obtained. Subsequently, Zhang et al. [56] used the similar method to prepare O-(2-hydroxy-3-trimethylammonium) propyl chitin (OHT-chitin) and evaluated its antioxidant activity. In this report, the method for the preparation of OHT-chitin is basically the same as that reported above, except the chitin used was in its α-form. According to the β-carotene bleaching test and evaluation of stable free radical α,α-diphenylβ-picrylhydrazyl, the OHT-chitin showed better antioxidant activity than chitosan, and has potential for preservation and cosmetic applications.

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

115

Moreover, the quaternized chitin also has an antibacterial effect. A report [57] from the same group noted that the OHT-chitin could inhibit the growth of bacteria Salmonella choleraesuis and Bacillus subtilis, of which the minimum inhibitory concentration (MIC) was 20.48 and 2.56 mg/mL, respectively, and the minimum bactericidal concentration (MBC) was 40.96 and 10.24 mg/mL, respectively. In addition to the above heterogeneous preparation methods, quaternized chitin could be synthesized in a homogeneous system. Peng et al. [58] dissolved chitin in a NaOH (8 wt.%)
urea (4 wt.%)
water (88 wt.%) solvent using the freeze
thaw method, and obtained a uniform solution. Then, they added (3-chloro-2-hydroxypropyl) trimethylammonium chloride (CHPTAC) drop by drop, which was reacted at 4 C. The solution was dialyzed to neutral using water, giving quaternized chitin after lyophilizing. During the process, the concentration of the chitin solution was between 1 and 2 wt.% with the molar ratio of CHPTAC to chitin unit from 8 to 16 for a reaction time of 24
74 h. Under different conditions, the DS of quaternized chitin was between 0.27 to 0.54. Among the products, quaternized chitin with DS of 0.36 and 0.54 were used for gene delivery. The selected quaternized chitin can load the gene because of the proper size, positive surface, low cytotoxicity, and high transfection efficiency. The homogeneous reaction also can be carried out with a KOH/urea solution. According to Xu et al. [59], β-chitin was first dissolved in the KOH/urea solution with a concentration of 1 wt.%. Then the solution was reacted with 2,3-epoxypropyltrimethylammonium chloride (EPTMAC) to produce the quaternized β-chitin. During the process, the KOH was also the catalyst. Under different molar ratios of EPTMAC to chitin unit (4B6), the DS of quaternized chitin ranged from 0.20 to 0.33. Among them, quaternized chitin with DS of 0.43 showed inhibition of Escherichia coli, S. aureus, Candida albicans, and Rhizopus oryzae, of which MICs were 8, 12, 60, and 40 μg/mL, respectively. Importantly, the quaternized β-chitin could promote the formation of new blood vessels, granulation tissue, and collagen fibers, and reduced inflammatory cell infiltration. It can be applied to treat wounds infected with E. coli and S. aureus.

4.3.2 The graft of chitin The grafting of chitin is also a method for modifying chitin to improve its solubility and hydrophilicity. According to the literature, chitin and chitin nanofiber film are the common raw materials for the grafting reaction. The details are as follows (Fig. 4.8). 4.3.2.1 The graft of chitin powder In 1979 Kojima et al. [60] used tributylborane (TBB) as the initiator to graft methyl methacrylate onto chitin in a water system. The results

Handbook of Chitin and Chitosan

116

4. Solubility, chain characterization, and derivatives of chitin

FIGURE 4.8 The structure of grafted chitin.

showed that the percentage of grafting was up to 5%, and the efficiency of grafting reached 25.2% at 37 C for 2 h with MMA (5 g), chitin (1.0 g), TBB (0.05 mL), and water (15 mL). They showed that the water is the key factor in the graft reaction, and this reaction can be carried out in the usual organic solvents including n-hexane, tetrahydrofuran (THF), and cyclohexanone. During the process, the complex formed from water-solvated chitin and TBB produced free radicals to initiate the graft reaction. In the subsequence experiments, the ceric ion was used as an initiator to prepare grafted chitin. In the literature from Kurita [61] the ceric ammonium nitrate was used as the initiator to prepare acrylamide and acrylic acid-grafted chitin with 240% and 200% grafting percentage, respectively, under the optimal conditions. The results showed that the amount of the cerium had a significant effect on the grafting percentage. Moreover, the chitin-g-grafted (sodium acrylate) (grafting 190%) and chitin-g-polyacrylamide (grafting 240%) showed about 18% and 8% higher hygroscopicity than chitin, respectively. Furlan et al. [62] also used the ceric ammonium nitrate nitric acid solution to synthesize poly (acrylic acid)-grafted chitin with 45% grafting efficiency, which performed the maximum absorption of calcium ion at 0.195 6 0.005 mmol/ g. Ren et al. [63] also showed that poly(methyl methacrylate) (MMA)grafted chitin can be obtained with ceric ammonium nitrate as the initiator. The results showed that a longer reaction time, a higher initiator concentration below B47, a higher temperature below 40 C, and a higher molar ratio of MMA added to chitin are beneficial for grafting yield.

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

117

The chitin-g-poly (MMA) with grafting yield of 620% showed the highest degree of swelling (12.0) in the N,N-DMF solvent. Moreover, Tanodekaew et al. [64] prepared chitin-g-poly (acrylic acid) hydrogels in an acid system. Chitin powder was mixed with acrylic acid with different mass ratio in a small amount of sulfuric acid, and then the mixture was reacted at 70 C for 1 h. After reaction, a sticky chitin-g-acrylic acid mixture was obtained, which had the active grafting sites to produce the polymer. Then the chitin-g-acrylic acid mixture was added to potassium peroxodisulfate and water. The mixture was poured into a petri-dish and maintained at 65 C for 4 h to produce film. The chitin-g-poly (acrylic acid) film was neutralized and washed. In this process, the configuration of chitin is β-form which is easier to modify. The experiments for testing the properties of film were carried out, showing that the degree of swelling was 30
60 times that of their original weights, and L929 cells could attach to it after incubation for 14 days. Literature suggested that the ammonium peroxy disulphate (APS) also can be an initiator to prepare chitin-g-polypyrrole. In the Ramaprasad report [65], chitin was first soaked in the pyrrole and 0.1 M HCl for 24 h. Then, the treated chitin was added to the APS HCl solution, and the mixture was reacted at 75 C for 12 h. With the 0.5 M concentration of pyrrole loading, the grafting percentage of the chitin-g-polypyrrole was up to 138%. In this process, the grafting mechanism was also radical copolymerization. With the APS stimulation, polypyrrole and chitin radicals were formed, which initiated the polymerization. In addition to the above compounds, literature has shown that chitin derivates also can be the initiator. Yamamoto et al. [66] prepared chiting-polystyrene through atom transfer radical polymerization (ATRP). During the process, the chitin macroinitiator was first prepared from chitin and 2-bromopropionyl bromide in 1-allyl-3-methylimidazolium bromide ([Amim][Br]). Obviously, the [Amim][Br] was the solvent to dissolve chitin to promote the reaction. Then, the chitin macroinitiators DMSO solution, styrene, PMDETA, and CuBr were mixed, and stirred at 60 C for 10 h. After reaction, the mixture was poured intto a large amount of water to get the chitin-g-polystyrene, which was washed with water, methanol, and acetone. With feed ratios of styrene per initiating site of chitin macroinitiator (100 equiv.), the yield and conversion of chitin-g-polystyrene can be up to 1432.2% and 50.0%, respectively. The GPC results showed that the Mw was 178,000. Furthermore, Liang et al. [67] reported a facile one-step method for preparing chitin-g-poly(aminoethyl) without an additional initiator. In this pathway, chitin was first prepared as alkaline chitin to improve its activity. Then, 2-chloroethylamine hydrochloride aqueous solution was introduced to the alkaline chitin drop by drop whilst stirring at 80 rpm.

Handbook of Chitin and Chitosan

118

4. Solubility, chain characterization, and derivatives of chitin

After this addition the solution was left to react for 18 h. The obtained solution was dialyzed, concentrated, and freeze-dried, producing chiting-poly(aminoethyl). Characterization of the chitin-g-poly(aminoethyl) showed that its Mw was 851.0 kDa, and DS was 1.77. It also showed high antibacterial effect on six bacteria: Staphylococcus epidermidis, S. aureus, Pseudomonas aeruginosa, E. coli, Bacillus proteus, and Klebsiella pneumoniae. Detailed experiments showed that the membrane damage of bacteria was the reason for chitin-g-poly(aminoethyl)’s antibacterial effect. 4.3.2.2 The graft of chitin nanofiber film Yamamoto’s group used the chitin nanofiber film (CNF) as raw material to develop a series of CNF-g-polymer composite films by surfaceinitiated graft polymerization. Firstly, the CNF was synthesized as follows [68]. Chitin powder was immersed in the [Amim][Br] at room temperature for 24 h, then the mixture was heated at 100 C for 48 h. After reaction, methanol was added to generate chitin dispersion, which was filtered to give a raw CNF. The raw CNF was washed with methanol by Soxhlet extraction, and dried to produce the final CNF. Before preparing the grafted CNF, the CNF was soaked in 40% (w/v) NaOH aqueous solution at 80 C for 7 h to give the partial deacetylation of chitin nanofiber (PDA-CNF). The obtained surface-grafted CNF showed a higher mechanical property than CNF. In 2012 Yamamoto’s group [69] prepared chitin nanofiber-graft-poly (ι-lactide-co-ε-caprolactone) films by surface-initiated ring-opening graft copolymerization. The CNF was pretreated with water before use, then the pretreated CNF was soaked in the toluene of ι-lactide (LA) or ε-caprolactone (CL) at room temperature for 12 h. After adding tin(II) 2-ethylhexanoate, the mixture was heated at 80 C for 48 h. The CNF-gpoly (LA-co-CL) film was obtained by adding chloroform, washing, and drying. The SEM, XRD, and FT-IR results all proved that the grafting was reacting on the surface of CNF. Additionally, the CNF-g-poly (LA-co-CL) films had a higher elastic ability than the CNF. In 2013 Yamamoto’s group [70] put PDA-CNF in an ethyl acetate solution of γ-benzyl ι-glutamate N-carboxyanhydride (BLG-NCA) at 0 C for 24 h, producing chitin nanofiber-g-poly(c-benzyl L-glutamate) (CNF-gPBLG) film. The SEM and XRD results showed that the polymerization occurred on the surface of the PDA-CNF. The CNF-g-PBLG was further converted to CNF-g-poly (c-L-glutamic acid sodium salt) (CNF-g-PLGA) under 1.0 mol/L NaOH aqueous solution treatment. Finally, the CNF-gPLGA network film was obtained using N-hydroxysuccinimide (NHS) and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) aqueous solution at room temperature for 12 h. The film showed a

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

119

higher flexible nature than the chitin nanofiber film or the CNF-g-PBLG film. In 2014 Yamamoto’s group [71] used the ATRP method. The CNF was pretreated with LiCl/N,N’-dimethylacetamide at 80 C for 14 h. Then CNF macroinitiator film was obtained after pretreated CNF film, α-bromoisobutyryl bromide, and pyridine reacted at 60 C for 24 h. Finally, the CNF-g-poly (2-hydroxyethyl acrylate) (HEA) was obtained by ATRP. The FT-IR, XRD, and SEM results showed that the HEA was grafted on the surface of the CNF. Then, Yamamoto’s group [72] prepared CNF-graft-poly MMA film in the same way. In this process, the CNF macroinitiator was first prepared using a mixture of pyridine, CNF dispersion with DMF, and α-bromoisobutyryl bromide reacting at room temperature for 12 h. Subsequently, MMA, CuBr, and 2,20 -bipyridine (Bpy) were added to the DMF solution of the CNF macroinitiator dispersion, which was heated at 60 C for 24 h and filtered. The CNF-graft-poly MMA film was obtained after washing with chloroform, methanol, and acetone, and dried. Obviously, the characterization of the film also suggested that the grafting was on the surface. The tensile testing showed that the elongation at break value of the CNF-g-PMMA film is 0.2% larger than that of the original CNF film. In 2018 the Yamamoto group [73] prepared CNF-g-poly(2-methyl-2oxazoline) (PMeOx) by putting PDA-CNF in poly(2-methyl-2-oxazoline) solution at 80 C for 24 h. The degree of grafting was up to 89.5% under this condition, which can synthesize hydrogel by gelling in the DMSO solution. Importantly, the grafting occurred with the aNH because of the use of PDA-CNF.

4.3.3 The O-acylation of chitin The O-acylation reaction of chitin is a common derivatization treatment. The obtained O-acylated chitin not only has high solubility in some organic solvents, but also has functions, such as antibacterial and coagulation, which allow it to be used as a medical material and a pharmaceutical carrier. According to the literature, the preparation of Oacylated chitin can be carried out in heterogeneous and homogeneous systems (Fig. 4.9). 4.3.3.1 Preparation of O-acylated chitin in heterogeneous system Since chitin is insoluble in common organic solvents, the conventional preparation method of O-acylated chitin is mostly carried out in heterogeneous system. Generally, the heterogeneous system is a strong acid or strong alkali environment, and the reactants are not sufficiently

Handbook of Chitin and Chitosan

120

4. Solubility, chain characterization, and derivatives of chitin

FIGURE 4.9 The structure of O-acylated chitin.

contacted. This method usually results in the degradation of the raw materials, and the low and uneven degree of acylation of the obtained product. Importantly, the heterogeneous method has the advantages of simple operation and low cost, and is suitable for easy scaling up of production. Common heterogeneous methods are listed below. 4.3.3.1.1 Acid catalyst method

In the early days, Schorigin and Hait [74] used hydrochloric acid and acetic anhydride as a catalyst and an acylating reagent, respectively, to produce O-acylated chitin. After reacting with chitin at 23 C for 120 h, full O-acetylated chitin was obtained. Subsequently, Norio et al. [75] used this method to produce O-acylated chitin. Firstly, the chitin was dispersed in the acetic anhydride solution, then it was poured into a hydrochloric acid solution. Finally, the solution was reacted at 0 C for 10 days to obtain an O-acetylated chitin with DS of 1.6. Although this acid catalyst method is effective, it is time consuming, poorly reproducible, and its harsh conditions lead to the degradation of chitin. Hence, this method has rarely been used. Subsequently, Norio et al. [75] prepared O-acylated chitin using perchloric acid as a catalyst. In this method, perchloric acid is dissolved in glacial acetic acid first, to inhibit the hydrolysis of chitin, then acetic anhydride and chitin were added in different molar ratios. Under the conditions at 0 C for 5 h, O-acylated chitin with different DS was

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

121

obtained. Although the reaction is a heterogeneous reaction, perchloric acid with higher acidity allows the production of the acylated chitin in a shorter time compared with the hydrochloric acid system. 4.3.3.1.2 Activation of chitin

Activation of chitin can destroy its hydrogen bond network, promoting the acylation reaction. Currently, there are two methods for activating chitin. One is to soak the chitin in sodium hydroxide aqueous solution to synthesize the alkalized chitin. Due to the intrusion of sodium hydroxide, the crystallinity of chitin is lowered, and the intermolecular and intramolecular hydrogen bonds of chitin are destroyed. This is beneficial to the acylation of chitin. Norio et al. [75] used alkalized chitin to directly react with acetic anhydride at room temperature for 24 h, producing acetylated chitin with a DS of 0.3. By carrying out the secondary acylation reaction at 95 C, an acetylated chitin with a DS of 1.1 can be obtained. Obviously, the use of alkalized chitin to prepare acylated chitin is not an efficient method, although it is easier to carry out compared with using raw chitin. The other method is to soak chitin in a specific solvent to loosen its structure. Vasnev et al. [76] mixed chitin, hexafluoroisopropanol, and pnitrobenzoic acid under argon atmosphere to produce a highly viscous solution. Then, the hexafluoroisopropanol was removed, and pyridine was added. The mixture was reacted at room temperature for 2 h to obtain O-acylated chitin with a DS of 0.22. The results showed that the addition of hexafluoroisopropanol can loosen the structure of chitin, reduce its crystallinity, and destroy hydrogen bonds, leading to the promotion of the acylation reaction. Although this method has an improved DS compared with raw chitin, it is still not a highly efficient method for chitin acylation. Additionally, according to the literature of Yoshifuji et al. [77], the β-chitin is activated by inserting dimethyl sulfoxide, hexylamine, and n-octanol between its layers. Then, the activated β-chitin was reacted with the anhydride via the host
guest reaction, giving acylated chitin with a DS of 1.0. However, this intercalation method can only be applied to β-chitin because its molecular chains are arranged in parallel, and there is no hydrogen bond between the sugar chain layers. Therefore it is easy to form an intercalation intermediate. However, α-chitin has a large number of interlayer hydrogen bonds and cannot be activated by this method. 4.3.3.2 Preparation of O-acylated chitin in homogeneous system The preparation of O-acylated chitin in a homogeneous system can be roughly divided into two types: (1) the homogeneous system is formed during the reaction process, such as methanesulfonic acid or trifluoroacetic anhydride systems; and (2) the homogeneous system is

Handbook of Chitin and Chitosan

122

4. Solubility, chain characterization, and derivatives of chitin

formed before reaction by dissolving chitin in specific solvents to prepare a chitin homogeneous solution, such as the lithium chloride/ dimethylacetamide (LiCl/DMAc) and ionic liquids (ILs) systems. 4.3.3.2.1 Methanesulfonic acid system

Methanesulfonic acid with high acidity is not only the catalyst in the process of chitin acylation, but also is a good solvent for partially acylated chitin. Thus homogeneous acylation of chitin can be achieved in the methanesulfonic acid system. Norio et al. [75] mixed chitin, methanesulfonic acid, and glacial acetic acid according to different molar ratios, and reacted it at 0 C overnight to obtain acetylated chitin with different DS. In this experiment, a homogeneous phase is gradually formed as the reaction proceeds, which contributes to further acylation. This reaction should be kept at a low temperature to prevent degradation of chitin in acidic conditions. The acylating agent is not limited to carboxylic acid but also acid chloride. Furthermore, Kaifu et al. [78] mixed the chitin, methanesulfonic acid, and acid chloride first, then the mixture was reacted at 0 C for 2 h, followed by an overnight reaction at
20 C to obtain acylated chitin. By changing the kind and molar amount of acid chloride, hexanoylation, oxime acylation, and dodecyl acylation of chitin with different DS can be obtained, of which DS can be up to 1.9. In this process, the crystallinity of chitin can be effectively destroyed by further acylation by reacting at
20 C overnight. In general, the acylation ability of the acid chloride is higher than that of the carboxylic acid. The larger the acylation group, the greater the damage to the crystalline region of chitin. 4.3.3.2.2 Trifluoroacetic anhydride system

Compared with the methanesulfonic acid system, the chitin acylation reaction in the trifluoroacetic anhydride system is easier to carry out. According to Mine et al. [79], chitin was reacted with carboxylic acid in the trifluoroacetic acid system at 70 C for a fixed time, which produced butyl-, hexyl-, octyl-, acetyl-, and palmitoyl chitin with DS of 1
2. In this process, after only 30 min. of reaction, a homogeneous system can be obtained, which is much faster than the methanesulfonic acid system. The obtained acylated chitin can be dissolved in dimethylformamide (DMF) solvent to prepare a film with high flexibility and high strength. The tensile modulus of this film can reach 1.5
5.8 Gpa, which is comparable to commercial polyphenylene. Subsequently, Bhatt et al. [80] used the same method to prepare cyclopropyl, cyclobutyl, cyclopentane, and cyclohexanoyl chitin with DS of 1.1
1.4. The obtained O-acylated chitin was soluble in methanol, ethanol, dimethylformamide (DMF), and tetrahydrofuran (THF) solvents. Moreover, as the volume of the intercalating group increases, the solubility gradually increases. It is seen that the

Handbook of Chitin and Chitosan

4.3 Derivatives of chitin

123

trifluoroacetic anhydride system can be used to prepare O-acylated chitin with different groups, and it is easier to carry out than the mesylate system. Furthermore, the acylation reaction of chitin also can be carried out in the phosphoric acid/trifluoroacetic anhydride mixed system. As reported by Bhatt et al. [81], chitin and butyric acid are reacted at 50 C for 3 h in the trifluoroacetic anhydride/phosphoric acid mixed system, producing butylated chitin with DS of 1.9
2.38. The results showed that the DS of the obtained O-acylated chitin was gradually reduced with the amount of phosphoric acid decrease. It is obvious that the phosphoric acid plays a key role in this system. Bhatt pointed out that the reaction processed as follows: trifluoroacetic acid was first reacted with butyric acid to form the acid anhydride, which was in turn reacted with chitin to form acylated chitin. Thus the phosphoric acid and trifluoroacetic anhydride was a synergistic catalyst in this system. Then, the Bhatt group [82] further extended the trifluoroacetic anhydride/phosphoric acid system to prepare other kinds of O-acylated chitin. The benzoylated chitin with the DS of 1.17
1.83 was obtained using this catalyst, which was soluble in dimethyl sulfoxide, dimethylformamide, benzyl alcohol, and formic acid. 4.3.3.2.3 The lithium chloride/dimethylacetamide system

As early as 1978 it was reported that the lithium salt organic solution can dissolve a polyamide and a polysaccharide containing a large amount of hydrogen bonds. Based on this report, McCormick et al. [16] first used the lithium chloride/dimethylacetamide (LiCl/DMAc) system to achieve the homogeneous acylation of chitin. In this process, chitin was first dissolved in a 5% LiCl/DMAc solution to form a clear solution, then acid chloride was added. The mixture was reacted at 90 C for a period of time under nitrogen protection conditions, which produced a variety of O-acylated chitin with DS of 0.91
1.2. Yamamoto et al. [83] also used this method to prepare 17 kinds of chiral active phenyl carbamate chitin, which further expanded the application field of this method. The resulting chiral O-acylated chitin can be used as a stationary phase in a high-performance liquid phase for separating chiral substances. Vasnev et al. [76] added the catalytic pyridine to the LiCl/DMAc system in order to facilitate the chitin acylation reaction. In this system, chitin and p-nitrobenzoic acid are reacted at room temperature for 4 h to obtain p-nitrobenzoyl chitin with a DS of 0.4. The literature pointed out that since Li1 had a shielding effect on the nucleophilic action of pyridine, so it is necessary to add an excess of pyridine. Further, pyridine also neutralized HCl generated in the reaction, preventing degradation of chitin.

Handbook of Chitin and Chitosan

124

4. Solubility, chain characterization, and derivatives of chitin

Yoshikuni et al. [84] further investigated the preparation of different alkyl-acylated chitins in this system. Firstly, the 0.3 wt.% solution of chitin Li/DMAc (5%) was prepared. Secondly, a DMAc solution of pyridine and a DMAc solution of carboxylic acid were added. Finally, the mixture was reacted at 50 C for 100 h to obtain nine kinds of O-acylated chitin with DS of 1.65
1.84. Results showed that the crystallinity of alkyl-acylated chitin gradually decreased with the change of alkyl chain length, and exhibited different thermodynamic properties. The glass transition temperature does not appear when the carbon chain length is 4
10. The glass transition temperature is
70 C to
40 C when the carbon chain length increases to 12 and 14. A phase transition phenomenon is observed between 10 C and 30 C. It can be seen that the insertion of the alkyl chain causes a change in the structure of the chitin. Sugimoto et al. [85] used this method to synthesize alkyl-acylated chitin with a DS of 1.62
3.89 using the acid chloride as an acylating agent. Among them, an N-acylation reaction also occurred with DS above 2, indicating that the acylation ability of the acid chloride is higher than that of the carboxylic acid. In this work, the obtained acylated chitin was blended with polycaprolactone (PCL) to prepare different transparency films using DMF as solvent. The results showed that as the total DS of acylated chitin increases, the miscibility increases. Among them, O-acylation has a greater influence on miscibility than N-acylation. 4.3.3.2.4 The ionic liquid system

As above, the IL can dissolve the chitin. In the field of chitin utilization, ILs play the role of a solvent to dissolve chitin. Hence, the preparation of O-acylated chitin in the ionic liquid is an homogeneous system. In 2009 Mine et al. [79] prepared acetylated chitin using [Amim][Br]. The chitin was dissolved in [Amim][Br] first, then the uniform solution was reacted with acetic anhydride, producing acetylated chitin. The results showed that the DS of the acetylated chitin increased with the temperature and the increase in the amount of acetic anhydride. Subsequently, the research group further expanded the substrate for the acylation of chitin in [Amim][Br]. In their report in 2018 [86], the chitin was reacted with acid chloride, octadecanoyl chloride, and oleoyl chloride in [Amim][Br] with pyridine and N,N-dimethyl-4-aminopyridine as catalysts, which gave acylated chitin with a DS of 1.6
2.0. It is indicated that the [Amim][Br] system can also be used for the preparation of long-chain acylated chitin, demonstrating the universality of this method. In 2019 Feng et al. [87] reported a green, effective, and simple method for the one-step preparation of O-acylated chitin from shrimp shells, which included homogeneous and heterogeneous acylation processes. In their work, 11 kinds of multifunctional deep eutectic solvents (DESs),

Handbook of Chitin and Chitosan

References

125

a new generation of IL, were synthesized from choline chloride (ChCl) and organic acid. These DESs held the ability of decalcification, deproteinization, and acylation. The results showed that O-malate chitin with a purity of 98.6 wt.% and DS of 0.46 was obtained with ChCl/DL-malic acid (1:2, ChCl 1-DL Mal 2) treatment at 150 C for 3 h. Moreover, the obtained O-malate chitin showed a good inhibitory effect on a Grampositive bacterium and C6 glioma cells. Feng et al. suggested that the H1 and the hydrogen bond formation were the main reasons for the preparation of O-acylated chitin. Obviously, Feng et al. provide a novel and meaningful alternative for O-acylation preparation.

Acknowledgment This research was supported financially by National Basic Research Program of China (973 Program, 2015CB251401), Major Program of National Natural Science Foundation of China (21890762), the National Natural Scientific Fund of China (No. 21878292, No. 21878314, No. 21606240, NO. 21476234), K. C. Wong Education Foundation (No. GJTD-2018-04), and the Strategic Priority Research Program of Chinese Academy of Science (No. XDA21060300).

References [1] J. Zhou, L. Zhang, Solubility of cellulose in NaOH/urea aqueous solution, Polym. J. 32 (10) (2000) 866
870. [2] X. Hu, Y. Du, Y. Tang, et al., Solubility and property of chitin in NaOH/urea aqueous solution, Carbohydr. Polym. 70 (4) (2007) 451
458. [3] G. Li, Y. Du, Y. Tao, et al., Dilute solution properties of four natural chitin in NaOH/ urea aqueous system, Carbohydr. Polym. 80 (3) (2010) 970
976. [4] C. Chang, S. Chen, L. Zhang, Novel hydrogels prepared via direct dissolution of chitin at low temperature: structure and biocompatibility, J. Mater. Chem. 21 (11) (2011) 3865
3871. [5] B. Ding, J. Cai, J. Huang, et al., Facile preparation of robust and biocompatible chitin aerogels, J. Mater. Chem. 22 (12) (2012) 5801
5809. [6] Y. Fang, B. Duan, A. Lu, et al., Intermolecular interaction and the extended wormlike chain conformation of chitin in NaOH/Urea aqueous solution, Biomacromolecules 16 (4) (2015) 1410
1417. [7] H. Tamura, Y. Tsuruta, S. Tokura, Preparation of chitosan-coated alginate filament, Mater. Sci. Eng. C 20 (1) (2002) 143
147. [8] H. Tamura, H. Nagahama, S. Tokura, Preparation of chitin hydrogel under mild conditions, Cellulose 13 (4) (2006) 357
364. [9] H. Tamura, M. Sawada, H. Nagahama, et al., Influence of amide content on the crystal structure of chitin, Holzforschung 60 (5) (2006) 480
484. [10] C. Chen, H. Yano, D. Li, et al., Preparation of high-strength α-chitin nanofiber-based hydrogels under mild conditions, Cellulose 22 (4) (2015) 2543
2550. [11] H. Nagahama, T. Higuchi, R. Jayakumar, et al., XRD studies of β-chitin from squid pen with calcium solvent, Int. J. Bio. Macromol. 42 (4) (2008) 309
313. [12] R. Tajiri, A. Mihata, K. Yamamoto, et al., Facile preparation of chitin gels with calcium bromide dihydrate/methanol media and their efficient conversion into porous chitins, RSC Adv. 4 (11) (2014) 5542
5546.

Handbook of Chitin and Chitosan

126

4. Solubility, chain characterization, and derivatives of chitin

[13] P. Austin, Chitin solution. 4059457[P], United States, 1976. [14] M. Vincendon, 1H NMR study of the chitin dissolution mechanism, Die Makromol. Chem. 186 (9) (1985) 1787
1795. [15] T.R. Dawsey, C.L. McCormick, The lithium chloride/dimethylacetamide solvent for cellulose: a literature review, J. Macromol. Sci. C 30 (3-4) (1990) 405
440. [16] C.L. McCormick, D.K. Lichatowich, Homogeneous solution reactions of cellulose, chitin, and other polysaccharides to produce controlled-activity pesticide systems, J. Polym. Sci. Polym. Lett. Ed. 17 (8) (1979) 479
484. [17] Q. Yang, Z. Zhang, X.-G. Sun, et al., Ionic liquids and derived materials for lithium and sodium batteries, Chem. Soc. Rev. 47 (6) (2018) 2020
2064. [18] L. Feng, K. Wang, X. Zhang, et al., Flexible solid-state supercapacitors with enhanced performance from hierarchically graphene nanocomposite electrodes and ionic liquid incorporated gel polymer electrolyte, Adv. Funct. Mater. 28 (4) (2018) 1704463 (1
9). [19] H. Beitollahi, S.G. Ivari, M. Torkzadeh-Mahani, Application of antibody
nanogold
ionic liquid
carbon paste electrode for sensitive electrochemical immunoassay of thyroidstimulating hormone, Biosens. Bioelectron. 110 (2018) 97
102. [20] S. Dezhang, F. Huisheng, X. Feng, et al., Feasibility of ionic liquid as extractant for biobutanol extraction: experiment and simulation, Sep. Purif. Tech. 215 (2019) 287
298. [21] S.P. Ventura, F.A. e Silva, M.V. Quental, et al., Ionic-liquid-mediated extraction and separation processes for bioactive compounds: past, present, and future trends, Chem. Rev. 117 (10) (2017) 6984
7052. [22] A.E. Visser, R.P. Swatloski, W.M. Reichert, et al., Task-specific ionic liquids for the extraction of metal ions from aqueous solutions, Chem. Commun. 1 (2001) 135
136. [23] P. Moyer, M.D. Smith, N. Abdoulmoumine, et al., Relationship between lignocellulosic biomass dissolution and physicochemical properties of ionic liquids composed of 3-methylimidazolium cations and carboxylate anions, Phys. Chem. Chem. Phys. 20 (4) (2018) 2508
2516. [24] A. Brandt, J. Gra¨svik, J.P. Hallett, et al., Deconstruction of lignocellulosic biomass with ionic liquids, Green Chem. 15 (3) (2013) 550
583. [25] T.-A.D. Nguyen, K.-R. Kim, S.J. Han, et al., Pretreatment of rice straw with ammonia and ionic liquid for lignocellulose conversion to fermentable sugars, Bioresour. Technol. 101 (19) (2010) 7432
7438. [26] S. Zeng, L. Liu, D. Shang, et al., Efficient and reversible absorption of ammonia by cobalt ionic liquids through Lewis acid
base and cooperative hydrogen bond interactions, Green Chem. 20 (9) (2018) 2075
2083. [27] F.T. Kohler, S. Popp, H. Klefer, et al., Supported ionic liquid phase (SILP) materials for removal of hazardous gas compounds
efficient and irreversible NH3 adsorption, Green Chem. 16 (7) (2014) 3560
3568. [28] E.D. Bates, R.D. Mayton, I. Ntai, et al., CO2 capture by a task-specific ionic liquid, J. Am. Chem. Soc. 124 (6) (2002) 926
927. [29] H. Xie, S. Zhang, S. Li, Chitin and chitosan dissolved in ionic liquids as reversible sorbents of CO2, Green Chem. 8 (7) (2006) 630
633. [30] K. Prasad, M.-a Murakami, Y. Kaneko, et al., Weak gel of chitin with ionic liquid, 1-allyl-3-methylimidazolium bromide, Int. J. Bio. Macromol. 45 (3) (2009) 221
225. [31] S. Mine, H. Izawa, Y. Kaneko, et al., Acetylation of α-chitin in ionic liquids, Carbohydr. Res. 344 (16) (2009) 2263
2265. [32] S. Yamazaki, A. Takegawa, Y. Kaneko, et al., An acidic cellulose
chitin hybrid gel as novel electrolyte for an electric double layer capacitor, Electrochem. Commun. 11 (1) (2009) 68
70. [33] Y. Wu, T. Sasaki, S. Irie, et al., A novel biomass-ionic liquid platform for the utilization of native chitin, Polymer 49 (9) (2008) 2321
2327.

Handbook of Chitin and Chitosan

References

127

[34] S.S. Silva, A.R.C. Duarte, A.P. Carvalho, et al., Green processing of porous chitin structures for biomedical applications combining ionic liquids and supercritical fluid technology, Acta Biomater. 7 (3) (2011) 1166
1172. [35] W.-T. Wang, J. Zhu, X.-L. Wang, et al., Dissolution behavior of chitin in ionic liquids, J. Macromol. Sci. B 49 (3) (2010) 528
541. [36] Y. Qin, X. Lu, N. Sun, et al., Dissolution or extraction of crustacean shells using ionic liquids to obtain high molecular weight purified chitin and direct production of chitin films and fibers, Green Chem. 12 (6) (2010) 968
971. [37] P.S. Barber, C.S. Griggs, J.R. Bonner, et al., Electrospinning of chitin nanofibers directly from an ionic liquid extract of shrimp shells, Green Chem. 15 (3) (2013) 601
607. [38] P. Walther, A. Ota, A. Mu¨ller, et al., Chitin foils and coatings prepared from ionic liquids, Macromol. Mater. Eng. 301 (11) (2016) 1337
1344. [39] M.M. Jaworska, A. Gorak, Modification of chitin particles with chloride ionic liquids, Mater. Lett. 164 (2016) 341
343. [40] T. Uto, S. Idenoue, K. Yamamoto, et al., Understanding dissolution process of chitin crystal in ionic liquids: theoretical study, Phys. Chem. Chem. Phys. 20 (31) (2018) 20669
20677. [41] T.D. Jiang, Chitin. Chemical Industry Press Environmental Science and Engineering Publishing Center, 2003. [42] M.K. Jang, B.G. Kong, Y.I. Jeong, et al., Physicochemical characterization of α-chitin, β-chitin, and γ-chitin separated from natural resources, J. Polym. Sci. A: Polym. Chem. 42 (14) (2004) 3423
3432. [43] P. Sikorski, R. Hori, M. Wada, Revisit of α-chitin crystal structure using high resolution X-ray diffraction data, Biomacromolecules 10 (5) (2009) 1100
1105. [44] K. Gardner, J. Blackwell, Refinement of the structure of β-chitin, Biopolym. Orig. Res. Biomol. 14 (8) (1975) 1581
1595. [45] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (7) (2006) 603
632. [46] G. Ca´rdenas, G. Cabrera, E. Taboada, et al., Chitin characterization by SEM, FTIR, XRD, and 13C cross polarization/mass angle spinning NMR, J. Appl. Polym. Sci. 93 (4) (2004) 1876
1885. [47] J. Brugnerotto, J. Desbrieres, L. Heux, et al., Overview on structural characterization of chitosan molecules in relation with their behavior in solution, Macromolecular Symposia, Wiley Online Library, 2001. [48] Y. Kim, R.-D. Park, Progress in bioextraction processes of chitin from crustacean biowastes, J. Korean Soc. Appl. Bio. Chem. 58 (4) (2015) 545
554. [49] M. Poirier, G. Charlet, Chitin fractionation and characterization in N, N-dimethylacetamide/lithium chloride solvent system, Carbohydr. Polym. 50 (4) (2002) 363
370. [50] F.I. Khan, S. Rahman, A. Queen, et al., Implications of molecular diversity of chitin and its derivatives, Appl. Microbiol. Biotechnol. 101 (9) (2017) 3513
3536. [51] S. Tokura, N. Nishi, A. Tsutsumi, et al., Studies on chitin VIII: some properties of water soluble chitin derivatives, Polym. J. 15 (6) (1983) 485
489. [52] C.S. Kong, J.A. Kim, S.S. Bak, et al., Anti-obesity effect of carboxymethyl chitin by AMPK and aquaporin-7 pathways in 3T3-L1 adipocytes, J. Nutri. Biochem. 22 (3) (2011) 276
281. [53] A. Dev, J.C. Mohan, V. Sreeja, et al., Novel carboxymethyl chitin nanoparticles for cancer drug delivery applications, Carbohydr. Polym. 79 (4) (2010) 1073
1079. [54] K.T. Shalumon, N.S. Binulal, N. Selvamurugan, et al., Electrospinning of carboxymethyl chitin/poly(vinyl alcohol) nanofibrous scaffolds for tissue engineering applications, Carbohydr. Polym. 77 (4) (2009) 863
869. [55] Q. Chen, Y. Wu, Y. Pu, et al., Synthesis and characterization of quaternized betachitin, Carbohydr. Res. 345 (11) (2010) 1609
1612.

Handbook of Chitin and Chitosan

128

4. Solubility, chain characterization, and derivatives of chitin

[56] X. Zhang, X. Geng, H. Jiang, et al., Synthesis and characteristics of chitin and chitosan with the (2-hydroxy-3-trimethylammonium)propyl functionality, and evaluation of their antioxidant activity in vitro, Carbohydr. Polym. 89 (2) (2012) 486
491. [57] X. Geng, R. Yang, J. Huang, et al., Evaluation antibacterial activity of quaternarybased chitin/chitosan derivatives in vitro, J. Food Sci. 78 (1) (2013) M90
M97. [58] N. Peng, Z. Ai, Z. Fang, et al., Homogeneous synthesis of quaternized chitin in NaOH/urea aqueous solution as a potential gene vector, Carbohydr. Polym. 150 (2016) 180
186. [59] H. Xu, Z. Fang, W. Tian, et al., Green fabrication of amphiphilic quaternized betachitin derivatives with excellent biocompatibility and antibacterial activities for wound healing, Adv. Mater. 30 (29) (2018) 1801100 (1
11). [60] K. Kojima, M. Yoshikuni, T. Suzuki, Tributylborane-initiated grafting of methyl methacrylate onto chitin, J. Appl. Polym. Sci. 24 (7) (1979) 1587
1593. [61] K. Kurita, M. Kawata, Y. Koyama, et al., Graft copolymerization of vinyl monomers onto chitin with cerium (IV) ion, J. Appl. Polym. Sci. 42 (11) (1991) 2885
2891. [62] L. Furlan, V.T. de Fa´vere, M.C. Laranjeira, Adsorption of calcium ions by graft copolymer of acrylic acid on biopolymer chitin, Polymer 37 (5) (1996) 843
846. [63] L. Ren, Y. Miura, N. Nishi, et al., Modification of chitin by ceric salt-initiated graft polymerisation—preparation of poly (methyl methacrylate)-grafted chitin derivatives that swell in organic solvents, Carbohydr. Polym. 21 (1) (1993) 23
27. [64] S. Tanodekaew, M. Prasitsilp, S. Swasdison, et al., Preparation of acrylic grafted chitin for wound dressing application, Biomaterials 25 (7-8) (2004) 1453
1460. [65] A.T. Ramaprasad, D. Latha, V. Rao, Synthesis and characterization of polypyrrole grafted chitin, J. Phys. Chem. Solids 104 (2017) 169
174. [66] K. Yamamoto, S. Yoshida, S. Mine, et al., Synthesis of chitin-graft-polystyrene via atom transfer radical polymerization initiated from a chitin macroinitiator, Polym. Chem. 4 (11) (2013) 3384
3389. [67] S. Liang, Q. Dang, C. Liu, et al., Characterization and antibacterial mechanism of poly(aminoethyl) modified chitin synthesized via a facile one-step pathway, Carbohydr. Polym. 195 (2018) 275
287. [68] J.-i Kadokawa, A. Takegawa, S. Mine, et al., Preparation of chitin nanowhiskers using an ionic liquid and their composite materials with poly(vinyl alcohol), Carbohydr. Polym. 84 (4) (2011) 1408
1412. [69] T. Setoguchi, K. Yamamoto, J.-i Kadokawa, Preparation of chitin nanofiber-graft-poly (l-lactide-co-ε-caprolactone) films by surface-initiated ring-opening graft copolymerization, Polymer 53 (22) (2012) 4977
4982. [70] J.-i Kadokawa, T. Setoguchi, K. Yamamoto, Preparation of highly flexible chitin nanofiber-graft-poly(γ-l-glutamic acid) network film, Polym. Bull. 70 (12) (2013) 3279
3289. [71] K Yamamoto, S. Yoshida, J. Kadokawa, Surface-initiated atom transfer radical polymerization from chitin nanofiber macroinitiator film, Carbohydr. Polym. 112 (2014) 119
124. [72] R. Endo, K. Yamamoto, J.-i Kadokawa, Surface-initiated graft atom transfer radical polymerization of methyl methacrylate from chitin nanofiber macroinitiator under dispersion conditions, Fibers 3 (4) (2015) 338
347. [73] J.-i Kadokawa, Y. Obama, J. Yoshida, et al., Gel formation from self-assembled chitin nanofiber film by grafting of poly(2-methyl-2-oxazoline), Chem. Lett. 47 (7) (2018) 949
952. ¨ ber die Acetylierung des Chitins (Vorla¨ufig. Mitteil.), Berichte [74] P. Schorigin, E. Hait, U der deutschen chemischen Gesellschaft (A and B Series) 68 (5) (1935) 971
973.

Handbook of Chitin and Chitosan

References

129

[75] N. Nishi, J. Noguchi, S. Tokura, et al., Studies on chitin. I. Acetylation of chitin, Polym. J. 11 (1) (1979) 27
32. [76] V.A. Vasnev, A.I. Tarasov, G.D. Markova, et al., Synthesis and properties of acylated chitin and chitosan derivatives, Carbohydr. Polym. 64 (2) (2006) 184
189. [77] A. Yoshifuji, Y. Noishiki, M. Wada, et al., Esterification of β-chitin via intercalation by carboxylic anhydrides, Biomacromolecules 7 (10) (2006) 2878
2881. [78] K. Kaifu, N. Nishi, T. Komai, Preparation of hexanoyl, decanoyl, and dodecanoylchitin, J. Polym. Sci.: Polym. Chem. Ed. 19 (9) (1981) 2361
2363. [79] S. Mine, H. Izawa, Y. Kaneko, et al., Acetylation of α-chitin in ionic liquids, Carbohydr. Res. 344 (2009) 2263
2265. [80] L.R. Bhatt, B.M. Kim, C.Y. An, et al., Synthesis of chitin cycloalkyl ester derivatives and their physical properties, Carbohydr. Res. 345 (14) (2010) 2102
2106. [81] L.R. Bhatt, B.M. Kim, K. Hyun, et al., Preparation of chitin butyrate by using phosphoryl mixed anhydride system, Carbohydr. Res. 346 (5) (2011) 691
694. [82] L.R. Bhatt, B.M. Kim, K. Hyun, et al., Preparation and characterization of chitin benzoic acid esters, Molecules 16 (4) (2011) 3029
3036. [83] C. Yamamoto, T. Hayashi, Y. Okamoto, High-performance liquid chromatographic enantioseparation using chitin carbamate derivatives as chiral stationary phases, J. Chromatogr. A 1021 (1-2) (2003) 83
91. [84] Y. Teramoto, T. Miyata, Y. Nishio, Dual mesomorphic assemblage of chitin normal acylates and rapid enthalpy relaxation of their side chains, Biomacromolecules 7 (1) (2006) 190
198. [85] M. Sugimoto, M. Kawahara, Y. Teramoto, et al., Synthesis of acyl chitin derivatives and miscibility characterization of their blends with poly(ε-caprolactone), Carbohydr. Polym. 79 (4) (2010) 948
954. [86] H. Hirayama, J. Yoshida, K. Yamamoto, et al., Facile acylation of alpha-chitin in ionic liquid, Carbohydr. Polym. 200 (2018) 567
571. [87] M. Feng, X. Lu, J. Zhang, et al., Direct conversion of shrimp shells to O-acylated chitin with antibacterial and anti-tumor effects by natural deep eutectic solvents, Green Chem. 21 (1) (2019) 87
98.

Handbook of Chitin and Chitosan

C H A P T E R

5 Solubility, degree of acetylation, and distribution of acetyl groups in chitosan E.I. Akpan1, O.P. Gbenebor2, S.O. Adeosun2 and Odili Cletus2 1

Institute for Composite Materials, Kaiserslautern, Germany, 2Materials and Metallurgical Engineering, University of Lagos, Lagos, Nigeria O U T L I N E 5.1 Introduction

132

5.2 Chemistry and structure of chitosan

132

5.3 Acetylation of chitosan 5.3.1 Methods of deacetylation of chitin to chitosan 5.3.2 Degree of acetylation of chitosan

134 135 139

5.4 Solubility of chitosan 5.4.1 Chitosan solubility: applications and requirements 5.4.2 Solubility of chitosan in solution 5.4.3 Modifications of chitosan for solubility enhancement

145 145 146 147

5.5 Conclusion

153

References

154

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00005-5

131

© 2020 Elsevier Inc. All rights reserved.

132

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

5.1 Introduction Chitosan competes with cellulose and lignin in terms of abundance in nature. It is a cationic polymer that is obtained from chitin by alkaline N-deacetylation [1]. Structurally, chitosan contains a fraction of β-(1-4)-N-acetyl-D-glucosamine and a fraction of β-(1-4)-D-glucosamine. It has been shown that in chitosan the A-units GlcNAc (b(1/4)-2acetamido-2 deoxy-b-D-glucopyranose) and D-units Glc (b(1/4)-2amino-2-deoxy-b-D-glucopyranose) are randomly distributed along the chains. Chitin is found in several natural sources, including crustaceans, fungi, insects, and some algae. Chitosan is known to be biocompatible, renewable, nontoxic, nonallergenic, and biodegradable. Chitin and chitosan possess antifungal, antibacterial, antitumor, immunoadjuvant, antithrombogenic, and anticholesteremic activities. Currently, chitosan has been used for drug delivery, enzyme immobilization, gene delivery, drug encapsulation, surface modification, bone regeneration, wound healing, dialysis membrane, surgical glove powder, cosmetics, etc. Chitin is converted to chitosan by a series of depolymerization reactions. Controlling these depolymerization reactions can adjust useful properties like viscosity, solubility, and biological activity. In medical applications it is required that chitosan be soluble in water or organic solvents without losing its functionality. Understanding the solubility behavior of chitosan as it relates to its structure and other properties is very important. In this chapter, the solubility of chitosan, the degree of acetylation (DA), and the distribution of acetyl groups are explained in details, along with their correlated effects [2 8].

5.2 Chemistry and structure of chitosan Chitosan is a derivative of chitin with N-deacetylation of typically less than 0.35. It is regarded as a copolymer comprising glucosamine and N-acetylglucosamine. It is a polycationic polymer with more than 5000 D-glucosamine units [9 11]. Since chitosan is derived from chitin it is important to look at the chitin structure in relation to chitosan (Fig. 5.1). The structure of chitin is very close to that of cellulose, consisting of several β-(1 4)-linked D-glucose units (Fig. 5.2). The difference between chitin and cellulose is that in chitin the hydroxyl group at position C-2 (Fig. 5.2) is replaced by an acetamide group. Chitosan, on the other hand, is a chitin derivative in which the attached acetamide groups are transformed into primary amino groups following a set of chemical or biological treatments. They are chemically described as β-(1 4)-linked 2-amino-2-deoxy-β-D-glucopyranose or N-deacetylated derivatives of chitin [13 17].

Handbook of Chitin and Chitosan

5.2 Chemistry and structure of chitosan

133

FIGURE 5.1 Structure of chitin (A) and chitosan (B). Reprinted with permission from S. Islam, M.A.R.R. Bhuiyan, M.N. Islam, J. Polym. Environ. 25 (2017) 854 866 [12].

FIGURE 5.2 (1-4)-β-D-glucan unit.

To form chitosan, chitin is deacetylated using various methods (see Section 5.3). However, deacetylation of chitin is almost always never 100% complete so that chitosan (also called deacetylated chitin) retains a certain amount of the acetamide groups. Chitin and chitosan have 5% 8% nitrogen in the form of acetamide in chitin and in the form of amine groups in chitosan. The presence of these groups makes chitin and chitosan suitable for typical reactions of amines. Chitosan contains primary and secondary hydroxyl groups on each repeat unit and primary amine groups depending on the DA (Fig. 5.3). The presence of these functional groups makes chitosan chemically active compared with chitin. These groups are readily subject to chemical modification and determine the chemical, physical, and mechanical properties of the polymer. The crystal structure of chitin has been studied by various researchers [21 27]. Chitin crystalline structure has been examined using IR spectroscopy, nuclear magnetic resonance (NMR), X-ray scattering, microscopy, sorption techniques, etc. These studies reveal that chitin is characterized by an ordered fibrillar structure, a developed intra- and intermolecular

Handbook of Chitin and Chitosan

134

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

FIGURE 5.3 Major units in chitin and chitosan. Source: Adapted from K. Kurita, Prog. Polym. Sci. 26 (2001) 1921 1971; A. Aljawish, I. Chevalot, J. Jasniewski, J. Scher, L. Muniglia, J. Mol. Catal. B Enzym. 112 (2014) 25 39; N. Berezina, Phys. Sci. Rev. 1 (2016) 1 8 [18 20].

hydrogen bonds system, and a high degree of crystallinity and polymorphism [28]. On the other hand, chitosan has a high degree of deacetylation (DD) with a low content of crystalline regions compared with chitin (Fig. 5.4). Chitosan is hydrated in its usual form with one water molecule per unit of the polysaccharide. The unit cell parameters of chitosan (P212121) are a 5 0.895, b 5 1.698, and c 5 1.034 nm [22,24,29]. Chitosan is known to be biocompatible and nontoxic, with antibacterial, antifungal, and antitumor activity, and possess good metal binding and wound healing capabilities, as well as able to lower cholesterol [30 46].

5.3 Acetylation of chitosan Chitosan is obtained from chitin by deacetylation to varying degrees. The DA which is the balance between the two polymeric residues (amine rich and N-acetyl rich) is what differentiates chitin from chitosan (Fig. 5.3). When the DA is lower than 50%, the polymer is called chitosan, which is soluble in acidic aqueous solutions. The deacetylation process involves the removal of acetyl groups to obtain amino groups. However, a depolymerization reaction also occurs leading to changes in the molecular weight of the polymer. Several methods have been used to convert

Handbook of Chitin and Chitosan

5.3 Acetylation of chitosan

135

FIGURE 5.4 The chemical structure of chitosan. Source: Reprinted with permission from K. Okuyama, K. Noguchi, T. Miyazawa, T. Yui, K. Ogawa, Macromolecules 30 (1997) 5849 5855.

chitin to chitosan, including enzymatic and chemical processes [47 52]. Commercially, the chemical processes are widely utilized because of their simplicity, suitability for mass production, and inexpensive nature.

5.3.1 Methods of deacetylation of chitin to chitosan 5.3.1.1 Chemical deacetylation Deacetylation of chitin involves the transformation of the N-acetyl groups in chitin to (aNH2) amino groups (Fig. 5.3). The process is like the production of citric acid from Aspergillus niger, Mucor rouxii, and Streptomyces, and can be achieved by acid or alkali treatment. The use of alkali is the most prevalent means of deacetylation of chitin because glycosidic bonds are known to be susceptible to acid [48,53]. It is either performed heterogeneously [54], or homogeneously [55]. In heterogeneous deacetylation [54,56 65] chitin is treated with a hot concentrated NaOH solution and chitosan is produced as an insoluble residue. In homogeneous deacetylation [66 69] alkali chitin is prepared after dispersal of chitin in concentrated NaOH followed by dissolution in crushed ice at around 0 C. Homogeneous deacetylation results in chitosan with acetyl groups uniformly distributed along the chains. Although complete removal of the acetyl group has not been reported,

Handbook of Chitin and Chitosan

136

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

FIGURE 5.5 Illustration of the possible reaction sites in chitin and chitosan. Source: Reprinted with permission from C.K.S.S. Pillai, W. Paul, C.P. Sharma, Prog. Polym. Sci. 34 (2009) 641 678.

the repeated use of NaOH on chitin can yield up to 98% deacetylation. The efficiency of the process is dependent on several factors including NaOH concentration, the reaction temperature, and time. Depending on the method and the species used, the DD usually ranges between 56% and 99%. However, a DD of over 85% is required for solubility of chitosan to be achieved [48,70,71]. Several attempts have been made to produce chitosan with a very high DD. To obtain chitosan with 98% DD, Dung et al. [72] applied a reducing agent (NaBH4) and reported that the reducing agent did not have any significant effect on the DA of chitosan but affected the molecular weight. The addition of sodium borohydride also prevented polymer degradation [73]. Younce et al. [74] also investigated the use of sodium borohydride as an oxygen scavenger using statistical methods. With a combination of temperature, time, and atmospheric conditions, the study was able to realize a DDA of 99%. Another method of deacetylation of chitin that has been examined is the use of autoclaving conditions [75]. Deacetylation was successfully achieved by treatment of chitin under elevated temperature and pressure with 45% NaOH for 30 min and a solids/solvent ratio of 1:15. A mixture of glycerol and NaOH has also been used for the deacetylation of chitin using the schematic in Fig. 5.5. The process involved soaking chitin for 4 16 h at temperatures (120 C 180 C) with 10% 40% (w/w) NaOH solution at a ratio of 1:30 1:60 (chitin glycerol, w/w). One percent of water was added to promote the ionization of NaOH and prevent the polymerization of glycerol catalyzed by NaOH at high temperature. The final solution was obtained by centrifugation (7000 r/min for 5 min) and washed thoroughly with distilled water to remove the remaining NaOH and glycerol completely [76]. The study realized chitosan with a DD up to 85.36% 6 1.04%. Mima et al. [71] reported the realization of 100% deacetylation of chitin using a series of treatments according to the scheme in Fig. 5.6. Other authors also applied water-miscible organic solvent [77], dispersing organic liquids, and high temperature (Figs. 5.7 and 5.8).

Handbook of Chitin and Chitosan

5.3 Acetylation of chitosan

137

FIGURE 5.6 Chemical deacetylation of chitin to chitosan.

FIGURE 5.7 Schematic of chitosan preparation by glycerol as reaction solvent. Source: Reprinted with permission from C. Liu, G. Wang, W. Sui, L. An, C. Si, A.C.S. Sustain. Chem. Eng. 5 (2017) 4690 4698.

FIGURE 5.8

Schematic of chitosan preparation.

Alsharabasy [78] proposed a new method for deacetylation of chitin by introducing a controlled cooling step. The study realized chitosan with DD of 98% without degradation of the polymer chains. Some authors also used a combination of alkali and acid to arrive at a high

Handbook of Chitin and Chitosan

138

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

DD [79 82]. Anwar et al. [83] investigated different green methods of deacetylation including grinding, maceration, and sonication. Yuan et al. [84] used acetic acid for deacetylation of chitosan. DD of 99.7% was determined using UV spectroscopy. He et al. [85] investigated the application of compressive pressure (0.11 0.12 MPa) at high temperatures. The study reported the production of chitosan with 100% DD using low concentration alkali conditions. Other methods that have been applied in the deacetylation of chitosan include the use of ethanol [86], n-butyl alcohol sodium hydroxide and amyl alcohol [87], and dimethyl sulfoxide [88]. These methods were able to record an increase in DD to about 99.7%. Microwave-assisted deacetylation has also been applied to shorten the time needed for deacetylation [89]. Although 20 kGy gamma radiation in 60% concentration alkali at 100 C resulted in a 13% increase in DD, it poses serious health challenges [90]. A new route of deacetylation involving the use of freeze pump out thaw cycles was proposed by Guillaume [91]. Although high DD have been obtained from chemical deacetylation, the method suffers from high energy consumption, high effluent waste generation, and produces a broad range of soluble and insoluble products. The use of ultrasound to improve deacetylation at very low concentrations of NaOH has been investigated by Ngo and Ngo [63]. The use of water-miscible organic solvents was also examined by Batista and Roberts [77]. 5.3.1.2 Enzymatic deacetylation The use of chitin deacetylase to prepare chitosan offers the possibility of developing an enzymatic process that overcomes the potential drawbacks of the chemical deacetylation methods. The rationale behind the use of chitin deacetylase is that it catalyzes the hydrolysis of the N-acetamido bonds in chitin to produce chitosan (Fig. 5.9) [92]. The mechanism of deacetylation with chitin deacetylase is a multiple attack mechanism on monomeric acetylglucosamine units of chitin producing high-quality chitosan [93,94]. Key enzymes that have been studied are those from the fungi M. rouxii [47,95,96], Absidia coerulea [97], Aspergillus nidulans [98], and strains of Colletotrichum lindemuthianum [99,100]. Enzymatic deacetylation

FIGURE 5.9 The catalytic action of chitin deacetylases. Source: Reprinted with permission from I. Tsigos, A. Martinou, D. Kafetzopoulos, V. Bouriotis, Trends Biotechnol. 18 (2000) 305 312.

Handbook of Chitin and Chitosan

5.3 Acetylation of chitosan

139

with chitin deacetylases offers a possibility of producing specific chitosan oligomers and polymers. Tokuyasu et al. [99] developed and purified chitin deacetylase from C. lindemuthianum. Results showed that the enzyme was active toward glycol chitin, partially N-deacetylated water-soluble chitin, and chitin oligomers with more than fourfold degrees of polymerization. The enzyme was inactive with N-acetylglucosamine. Zhang et al. [101] studied the optimization and fermentation conditions of Rhizopus japonicas for the production of deacetylase and chitosan. Most of the enzymes that have been used in deacetylation display outstanding thermal stability at their optimal temperatures and exhibit very strong specificity for β-(1,4)-linked N-acetyl-D-glucosamine polymers. Yang et al. [102] investigated the production and purification of protease enzymes from Bacillus subtilis capable of deproteinizing crustacean wastes. Protease produced by Pseudomonas aeruginosa has also been investigated for deproteinization of shrimp and crab shell waste [103]. In another study, protease-producing bacterium has been investigated for the deproteinization of squid to produce β-chitin [104]. Morley et al. [105] showed that acetyl xylan esterase is capable of hydrolyzing N-acetyl groups in chitin to form chitosan of varying DD. Cai et al. [106] also investigated enzymatic preparation of chitosan. Using chitin deacetylase, the study recorded DD of 73.6% which was 2% lower than chemical deacetylation. Martinou et al. [107] studied the mode of action of deacetylase extracted from M. rouxii on partially N-acetylated chitosan. Kohlhoff et al. [108] also investigated the functionality of chitinosanase enzyme on the generation of partially acetylated chitosan. Aspras et al. [109] studied the kinetics of chitin deacetylase in the presence of ionic liquids. Ionic liquids were found to increase the activity of the enzyme [110]. Pareek et al. [111] investigated the use of chitin deacetylase from Penicillium oxalicum for bioconversion of chitin to chitosan using a two-stage chemical and enzymatic process. Results showed that pretreatment and optimization of the process variables led to a 3.2-fold increase in DD of chitosan. Martinou et al. [112] developed a monitoring process for deacetylation using chitin deacetylase. Other studies on the use of enzymes for deacetylation of chitin have been reported [113 116]. The major disadvantage of this method is that the enzyme is ineffective in deacetylating insoluble chitin substrates. Many researchers report that the crystallinity and insolubility of chitin are the major barriers to the development of the enzymatic deacetylation process [47,51,52,117,118].

5.3.2 Degree of acetylation of chitosan The most important characteristic of chitosan that influences all the important properties is the DA. It has been employed to

Handbook of Chitin and Chitosan

140

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

differentiate chitin from chitosan and serves as a quantitative means of characterizing chitosan. Several characterization methods have been used for quantitative estimation of the DA of chitosan (DD of chitin). Chitosan is a linear heteropolysaccharide composed of β-1,4-linked D-glusosamine (GlcN) and N-acetyl-D-glucosamine (GlcNAc) with varying compositions of these two monomers. The DA is defined as the portion of GlcNAc units to the total number of units. Mathematically DA is defined as DA 5 100 DD, and DD can be calculated as given in Eq. 5.1. DD 5 100 3

nGlcN nGlcN 1 nGlcNAc

(5.1)

where nGlcN is average number of D-glucosamine units, nGlcNAc is average number of N-acetylglucosamine units [119]. The determination of DD or DA is therefore hinged on the accurate measurement of the number of D-glucosamine units and number of N-acetylglucosamine units present in the polymer. The accuracy of the measurement therefore relies on the accuracy of the instrument to quantitatively measure these two units. Methods that have been used in the determination of DD and DA of chitosan over the years include elemental analysis [120,121], titration [122 129], hydrolytic methods [120,130 133], high-performance liquid chromatography (HPLC) ultraviolet [134,135], infrared [136 141], 1H NMR [124,142 145], CP-MAS 13C NMR [125,146,147], CP-MAS 15N NMR [148,149], steric exclusion chromatography [142], nitrous acid deamination [150], thermal analysis [151], and gas chromatography [152]. 5.3.2.1 Measurement of degree of acetylation of chitosan 5.3.2.1.1 Elemental analysis

Determination of DA or DD with elemental analysis involves the use of appropriate equipment to determine the composition of C, H, N, and O in the sample. Basic measurement of DA with elemental analysis involves the direct measurement of the ratio of C/N [153 157]. The DA can be calculated from Eq. 5.2. The values 5.145 and 6.861 are C/N of fully N-deacetylated chitosan (repeat unit C6H11NO4) and C/N of fully N-acetylated polymer (repeat unit C8H13ON5) in chitin. The limitations of elemental analysis lie in the fact that samples with varying DA possess relatively small variations in nitrogen content giving rise to imprecise results, especially if contaminants are present. DA 5 100 3

C=N 2 5:145 6:861 2 5:145

Handbook of Chitin and Chitosan

(5.2)

5.3 Acetylation of chitosan

141

5.3.2.1.2 Titration methods

Titration methods are the most common and affordable methods of measuring DA of chitosan. Several titration methods have been used by several researchers including acid base titration, potentiometric titration, colloidal titration, and conductometric titration. In alkali titration, chitosan is dissolved in an organic acid and then titrated with dilute alkali solution with continuous measurement of the pH of the solution. Plotting pH of the solution against the volume of added alkali will yield two points of inflection, which are indicators of the amount of alkali consumed for the conversion of the amine groups into ammonium salts [122]. In acid titration DA is calculated from the amount of acid consumed to neutralize free amino groups of chitosan dissolved in an alkali [55]. The use of colloidal, potentiometric titration, and conductometry has been reported in the literature [125,128,158 165]. The ninhydrin (triketohydrinedene hydrate) method centers on the determination of the free amine group of glucosamine units [166]. In the picric acid method, the acid is used to adsorb free amino groups of chitin/chitosan with a ratio of 1:1. 5.3.2.1.3 Acid hydrolysis/ high-performance liquid chromatography

Acid hydrolysis of chitosan is performed first by digestion of the chitin/chitosan sample by acid to release acetic acid and later analyzing the acetic acid by HPLC. Analysis with HPLC involves the measurement of the peak area and its conversion into molar concentration using a standard curve. The standard curve should be constructed using glucosamine or N-acetylglucosamine as the standard material. The method is relatively valid over a wide range of DA. It is affordable and easy to perform. It can also be used to analyze insoluble chitin. However, a calibration curve is usually required and the measurement is sensitive to the presence of water [130,131,167]. 5.3.2.1.4 Pyrolysis GC-MS

Conventional pyrolysis GC-MS has been used to identify and measure the different fragments present in a chitosan sample and N-acetylD-glucosamine is used as standard to calculate the DA. The method is highly efficient and requires only a small amount of samples but possesses the tendency to overestimate DA if there are contaminants from carbohydrates [132,167]. 5.3.2.1.5 Infrared

Infrared is probably the most used method in the determination of DA of chitosan. It involves recording the infrared spectrum of the chitosan sample and calculating the DA or DD from the absorbance of some

Handbook of Chitin and Chitosan

142

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

specific peaks using specific equations that have been developed over the years. Some of the proposed equations uses absorption ratios. A1560/ A2875, A1655/A2875, A1655/A3450, A1320/A3450, A1655/A1070, A1655/A1030, A1560/ A1160, A1560/A897, and A1320/A1420 have been already proposed to determine the DA of chitosan [137,139,141,168 172]. In most cases baselines and internal reference bands are used to correct the spectra before they are used to calculate DA [71,138,139,171,173 176]. The use of nearinfrared (NIR) was proposed by Rathke and Hudson [177]. The absorption ratios of A7669/A7474 and A6039/A5342 were chosen as the best ratios for chitin and chitosan model compounds, respectively. Another study on the use of NIR was reported by Varum et al. [178]. The method involves several uncertainties including the interference of water, broadening of peaks, and the need for extensive purification before analysis. 5.3.2.1.6 Nuclear magnetic resonance spectroscopy

Quantitative NMR spectroscopy has been used in the determination of DA of chitosan. Liquid and solid-state NMR are the most used methods in the determination of DD of chitosan. They are versatile but they are complex and expensive. 5.3.2.2 Distribution of acetyl groups The distribution of acetylated glucosamine residues in chitin/chitosan is an important parameter in the definition of chitin/chitosan quality for medical applications. Two major distribution patterns have been reported in the literature [67,179 181]. Studies show that moderately Ndeacetylated chitosan obtained by heterogeneous N-deacetylation of chitin possesses blocks of GIcNAc sequences, whereas partially N-acetylated chitosan obtained by homogeneous N-acetylation of highly deacetylated chitosan possesses random-type copolymers of GIcNAc and GIcN [182,183] (Figs. 5.10 and 5.11). The random type arrangement of the partially N-acetylated chitosan is responsible for its unique properties, such as solubility in aqueous acetic acid without forming precipitates or gels. The distribution of acetyl groups can be measured by gel filtration separation and reflective index detection. Han et al. [182] investigated the use of a UV-based detection assay for the determination of N-acetyl distribution in chitosan. 5.3.2.3 Relationship between acetylation and properties The effectiveness of chitosan as an industrial product is dependent on DA. It has been reported that the swelling capacity, stiffness, and the tendency to aggregate is strongly dependent on the DA [153,184,185]. DA also influences the physicochemical characteristics [55,160,186 189], biodegradability [190 192], and immunological activity [193] of chitosan. Wenling [194] showed that the swelling index of chitosan films

Handbook of Chitin and Chitosan

5.3 Acetylation of chitosan

143

FIGURE 5.10 Speculative representation of the structures of partially N-acetylated chitosan. Source: Reprinted with permission from S. Ichi Aiba, Int. J. Biol. Macromol. 13 (1991) 40 44.

decreases with increase in DD, whereas elastic modulus and tensile strength increased with the increase in DD. Hussain showed that chitosan swelling decreased with the increase in DD [195]. It was also shown

Handbook of Chitin and Chitosan

144

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

FIGURE 5.11 (A) Structural formulas of the A-unit N-acetylglucosamine and the Dunit glucosamine. (B) Theoretical patterns of acetylation illustrated as pictographs. A square represents a N-acetylglucosamine and a circle a glucosamine unit. A pattern can adopt the following extreme values: an alternating, a random, and a block-wise distribution. Reprinted with permission from M.X. Weinhold, J.C.M. Sauvageau, J. Kumirska, J. Tho¨ming, Carbohydr. Polym. 78 (2009) 678 684.

that keratinocyte cell proliferation of chitosan increases with a decrease in DA showing that DA influences the cell growth in the same way as cell adhesion [196]. Foster et al. [197] showed a linear increase in olfactory ensheathing cells proliferation as DD increased. Tensile strength and elongation to break also varied with DD. Sorlier et al. [198] also reported changes in electrostatic properties of chitosan in relation to other properties. Hussain et al. also noticed that at low cross-linking concentration, oil loading, loading efficiency, and release rate of essential oil from deacetylated chitosan decreased with the increase in DD but vice versa at higher cross-linking [199]. Wan et al. [200] reported that the release of nitric oxide from chitosan nitric oxide adducts is dependent on the DA of the chitosan. Thermal studies on chitosan/ PA66 blends showed that thermal properties of the blend were greatly influenced by the DD of the chitosan [201]. It has been shown that an increase in DA leads to a reduction in the inhibitory potency of chitosan [202]. Carvalho et al. [203] showed that controlling the DA of chitosan can lead to the realization of tunable properties even at a molecular level that will enable chitosan to be used as a novel nerve conduits.

Handbook of Chitin and Chitosan

5.4 Solubility of chitosan

145

Amaral et al. [204] examined the effect of DA on the attachment, spreading, proliferation, and osteogenic differentiation of rat bone marrow stromal cells. Results showed that a decrease in DA favored the formation of fibronectin on the surface of the chitosan. Tomihata et al. [205] investigated the effect of DA on the biodegradation of chitosan. Varum et al. [206] investigated the degradation rates in human serum of three chitosans with DA. Hidaka et al. [207] showed that DA affects osteogenesis of chitosan implants at the site of their implantation. It has also been shown that DA influences crystallinity, hydrophilicity, degradation, and cell response [208 210]. Lim et al. [211] showed that cell adhesion and growth on chitosan film surfaces decreased with increasing DA.

5.4 Solubility of chitosan 5.4.1 Chitosan solubility: applications and requirements The solubility of chitosan at neutral or high pH region is one of the barriers to the use of chitosan in the biomedical industries. Chitosan molecules have a dense concentration of intermolecular and intramolecular hydrogen bonds which strongly stabilize its packing structure in the three-unit cell directions. This is the reason chitosan exhibits no melting point and it is difficult to dissolve except in the acid pH range [36,212,213]. Chitosan dissolves in formic, acetic, propionic, lactic, citric, succinic, hydrochloric, phosphoric, and nitric acid [214]. The degree of solubility of chitosan in acid is dependent on the concentration and type of acid. The solubility declines with increasing concentration of acid. It has been noted that some aqueous acid solutions such as phosphoric, sulfuric, citric, and sebacic acids are not good solvents for chitosan solubility. The solubility of chitosan is determined by DD, distribution of acetyl groups, and degree of polymerization. These parameters can be used to control and improve the solubility of chitosan [28,215 219]. However, solubility of chitosan also depends on temperature, alkali concentration, time of deacetylation, prior treatments applied during chitin isolation, ratio of chitin to alkali solution, particle size, ionic concentration, pH, the nature of the acid used for protonation, etc. [220]. Chitosan is soluble in dilute acidic solutions below pH 6.0 because it is a strong base as it possesses primary amino groups with a pKa value of 6.3. The presence of the amino groups indicates that pH substantially alters the charged state of the polymer [221]. When the pH is low the amines in chitosan get protonated and become positively charged making it a water-soluble cationic polyelectrolyte [10]. When the pH increases above

Handbook of Chitin and Chitosan

146

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

6, the amines in chitosan become deprotonated and the polymer loses its charge and becomes insoluble. The transition between solubility and insolubility in chitosan occurs at a pKa value around a pH between 6 and 6.5. Because the pKa value is extremely reliant on DD, solubility is likewise dependent on the DD [222]. As the substitution of the amino groups by carboxylic groups in chitosan increases it becomes soluble over the entire pH range. For partially deacetylated chitins the solubility is closely related to their crystal structure, crystallinity, and crystal imperfection as well as the glucosamine content. It has been shown that the crystal structure of chitin changes as the DD changes. Homogeneous deacetylation transforms the crystal structure of chitin from α- to β-form which enables water solubility [222 224]. Water-soluble chitosan is useful in several applications, e.g., antimicrobial, antioxidant, and antitumor activity, control of the progression of diabetes mellitus, and as a potential material for DNA delivery [33,225 228]. Some of them are known for their antibacterial activity [229 233], immunoenhancing effects, and enhancing protective effects against infection with certain pathogens in mice [234,235]. Extensive studies on water-soluble chitosan for antioxidant activities have been reported [236 247]. Extensive studies on the antimicrobial behavior of water-soluble chitosan have also been reported [248 261]. They have also been used in wood dyeing [262,263] and drug and protein antigen delivery systems [264,265].

5.4.2 Solubility of chitosan in solution The solubility behavior of chitosan in various solvents is of the utmost importance in the application of chitosan. Solvents such as organic and inorganic acids, bases, and water are important in the biomedical industries. Chitosan is readily soluble in many inorganic acids, bases, and salts. They are used as solvents for the dissolution of both chitosan and chitin. Chitosan was shown to be soluble in 8 wt.% NaOH and 4 wt.% urea without prior treatment. Chitosan is readily soluble in dilute organic acidic media below its pKa (pH 5 6.5). This is because at low pH values the amino groups in the chitosan backbone enhance ionization, forming 2 NH31, leading to an increase in solubility. In acids with pKa smaller than 6.2 the amino groups (pKa from 6.2 to 7.0) are completely protonated making chitosan soluble. However, at higher pH the polymer precipitates. The solubility of chitosan in organic acids is dependent on the ability of the medium to protonate the chitosan molecule and the solubility of the polyelectrolytes. Chitosan is soluble in acetic acid, formic acid, L-glutamic acid, lactic acid, and succinic acid. Chitosan is depolymerized in organic acids by first depolymerization of the glycosidic linkage via hydrolysis followed by deacetylation of the

Handbook of Chitin and Chitosan

5.4 Solubility of chitosan

147

N-acetyl groups [266 268]. The role of the protonation of chitosan in the presence of acetic acid [269] and hydrochloric acid on solubility [270] reveals that the degree of ionization is dependent on the pH and the pKa of the acid. The most common means of testing solubility of chitosan is dissolving it in 1% or 0.1 M acetic acid. It has been shown that the amount of acid needed to dissolve chitosan is dependent on the quantity of chitosan to be dissolved. The concentration of protons desired should be equal to the concentration of 2 NH2 units involved. Chitosan is generally insoluble in water. However, water solution of chitosan is the most needed property in medical applications.

5.4.3 Modifications of chitosan for solubility enhancement To make chitosan soluble, several researchers have investigated chemical modification methods and the use of chemical entities that enable solubility. Chitosan can be modified using reactions such as etherification [18,271,272], esterification [272,273], cross-linking [274], and graft copolymerization [275,276]. Some other chemical modifications of chitosan have been reviewed by Muzzarelli and Muzzarelli [277]. Fig. 5.12 shows possible derivatization and depolymerization reactions of chitin and chitosan. Yi et al. [221] showed that conjugation (e.g., tyrosinase-initiated enzymatic assembly of proteins onto chitosan) of chitosan can confer chitosan’s pH-responsive solubility. Research has shown that acylation of chitosan with long-chain aliphatic carboxylic acid chlorides, such as hexanoyl, dodecanoyl, and tetradecanoyl chlorides, make it soluble in chloroform [279]. In another study, it was shown that acylation with a cyclic ester (lactone) (β-propiolactone or γ-butyrolactone) in a suitable solvent gives derivatives with N-hydroxyalkanoyl groups [280,281]. It has been noted that alkylation of chitosan can lead to an improvement in solubility. Reacting chitosan with alkyl halides under basic conditions will lead to the introduction of alkyl groups on the N and O atoms of chitosan. Studies have shown that isobutylchitosan synthesized alkylation improved solubility in neutral aqueous solution. This was attributed to the decrease of crystallinity of chitosan due to the introduction of the isobutyl group [282]. It has been reported that enzymatic grafting of phenolic compounds onto chitosan can confer water solubility under basic conditions [283]. The research showed that the use of tyrosinase converted a wide range of phenolic substrates into electrophilic o-quinones which are soluble under basic conditions [284,285]. The reaction path is shown in Fig. 5.13. Engibaryan et al. [287] fabricated a new water-soluble chitosan by sulfonation of chitosan. The water-soluble chitosan (chitosan

Handbook of Chitin and Chitosan

148

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

FIGURE 5.12 Possible derivatization (A) and depolymerization (B) products and cross-linking of chitosan. Source: Reprinted with permission from K.V. Harish Prashanth, R.N. Tharanathan, Trends Food Sci. Technol. 18 (2007) 117 131 [278].

oligoethylene oxide sulfonate) was 100% soluble in water. It has also been shown that anionic side-chain grafting on chitosan will confer water solubility [229]. Chen and Wang [288] studied the fabrication of cyclodextrins-linked chitosan using tosylated β-cyclodextrins for improved solubility. A blend of chitosan and PEG was also found to result in a water-soluble polymer [223]. To improve the solubility of chitosan, the microencapsulation of lactic acid bacteria based on the cross-linking of chitosan by 1,6-diisocyanatohexane has been studied [289] (Fig. 5.14).

Handbook of Chitin and Chitosan

5.4 Solubility of chitosan

FIGURE 5.12

149

(Continued)

Studies have shown that modification of chitosan by N-acylation results in enhanced solubility [224,290 292]. Simple acylation has also been reported to enhance solubility [293,294]. Research has shown that chain extension of chitosan with hexanoyl, decanoyl, and lauroyl chlorides resulted in acylated chitosan which exhibited excellent solubility in organic solvents such as chloroform, benzene, pyridine, and Tetrahydrofuran (THF) [295]. The polymers had four degrees of substitution per monosaccharide ring (disubstitution at amino and monosubstitution each at hydroxyl groups). A series of N-aliphatic-Odicinnamoyl-chitosans has been found to exhibit solubilities strongly related to the length of the flexible side chains. The study showed that

Handbook of Chitin and Chitosan

150

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

FIGURE 5.13 (A) Enzymatic grafting of chitosan with phenol and tyrosinase [283,286]. (B) Enzymatic modification of chitosan [10].

FIGURE 5.13 (Continued)

FIGURE 5.14 Amphiphilic water-soluble chitosan derivatives. Source: Reprinted with permission from B.O. Jung, C.H. Kim, K.S. Choi, Y.M. Lee, J.J. Kim, J. Appl. Polym. Sci. 72 (1999) 1713 1719. Handbook of Chitin and Chitosan

5.4 Solubility of chitosan

151

FIGURE 5.15 Formation of water-soluble N-(n-fatty acyl) chitosan. Source: Reprinted with permission from C. Le Tien, M. Lacroix, P. Ispas-Szabo, M.A. Mateescu, J. Control. Release 93 (2003) 1 13.

increasing length of the flexible side chains reduced the solubility [296]. Chitosan has been modified to improve solubility by the introduction of succinyl groups onto the N-terminal of the glucosamine units of chitosan [228,297 306]. The succinyl chitosan are made soluble by the addition of a long alkyl moiety as a hydrophobic function to the amino group. The succinyl moiety confers a hydrophilic character, whereas the alkyl moiety provides hydrophobic properties. It has been shown that the introduction of hydrophobic moieties with an ester linkage into chitosan benefits hydrophobic groups and contributes to organosolubility [307 310]. Methods that involve the introduction of water-soluble entities, hydrophilic moieties, bulky, and hydrocarbon groups to chitosan chains to enhance solubility can be found in some literature reviews [18,311 313]. Qin et al. obtained water-soluble chitosan by N-acetylation with acetic anhydride [314]. Kennedy et al. also obtained water-soluble chitosan with acetate [315]. Research has shown that N-acylation of chitosan with various fatty acid chlorides improved its hydrophobic character and with significant variations in structural features [316] (Fig. 5.15). Water-soluble chitosan has been obtained with n-fatty acid anhydrides in a homogeneous solution of 2 vol% aqueous acetic acid methanol [317,318]. Introduction of bulky groups, such as butyrylchitin, valeroylchitin, triethyl, butyryl-to-acetyl, and succinyl, has been shown to produce enhanced water solubility [298,319 325]. Water-soluble carboxymethyl derivatives of chitin and chitosan have been reported by some researchers [326 329]. Water-soluble N-carboxymethyl chitosan fabricated by reacting with glyoxylic acid has been reported [330,331]. Water-soluble ethylamine hydroxyethyl chitosan produced by reacting chloroethylamine hydrochloride under alkali conditions has been reported [332]. A method of modifying chitosan, targeted at disrupting the hydrogen bonding between amino groups of chitosan has been found to improve the hydrophilicity of chitosan. This can be achieved by the covalent attachment of a hydrophilic sugar moiety, gluconic acid, through the formation of an amide bond and the N-acetylation of sugar-bearing chitosan [333]. Water-soluble chitosan, such as chitosan maltose,

Handbook of Chitin and Chitosan

152

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

chitosan fructose, chitosan glucosamine, and chitosan-mannose, has been fabricated using saccharides under various operating conditions [334 340]. Adding an alkyl group to a chitosan lactose using potassium borohydride was found to confer excellent water solubility [341]. There have been reports of the use of enzymatic hydrolysis to improve the water solubility of chitosan [33]. Water-soluble chitosan polymers have been obtained when poly(ethyleneglycol) dialdehyde diethyl acetals were cross-linked with partially reacetylated chitosan via Schiff’s reaction and hydrogenation of the aldimines [342]. A method that involves random substitution of partial N-partial O-acetylated in chitosan with an acetylating agent in the presence of a phase transfer reagent has been shown to result in water-soluble products [343,344]. Water-soluble chitosan salts have also been formed by a heterogeneous reaction between chitosan and monohydric alcohol containing an amount of water sufficient to raise the dielectric constant [345]. N-(2-carboxybenzyl) with good water solubility has been reported [346]. The introduction of quaternary ammonium groups, such as N-[(2-hydroxy-3-trimethylammonium) propyl], N-methylenephenyl phosphonic, N-methylene phosphonic, and glycidyltrimethylammonium chloride, to chitosan have been also reported to impart water solubility [347 350]. Grafting copolymerization of chitosan has been found to lead to improvements in solubility. Grafting with polar monomers, nonacrylic monomer (N-vinyl pyrrolidone), and sialic acid has been reported to improve solubility [271,275,276,351 353]. Enzymatic grafting of chitosan with trosinase, mono (2-methacryloyl oxyethyl) acid phosphate, and vinylsulfonic acid sodium salt to improve solubility has been studied [229,283,354 357]. Glycol chitosan that is soluble in water at neutral and acidic pH has been prepared by the conjugation of chitosan with ethylene glycol [358]. Glycerol was also shown to enhance the water solubility of fish gelatin chitosan films [359]. The use of bipolar membrane electroacidification has been shown to solubilize chitosan [360]. It has been reported that hydrophylic moieties can be introduced into chitosan during oxidation resulting in enhanced water solubility [55]. Oxidization of chitosan was performed in water using NaClO and catalytic amounts of 2,2,6,6tetramethylpiperidinyloxy radical and NaBr. Water-soluble chitosan has been prepared by a reaction with epoxy group-containing moieties [361], as shown in Fig. 5.16. Two novel water-soluble O-carboxymethyl chitosan Schiff bases have been synthesized. The polymer was fabricated by a condensation reaction of 2,6-diacetylpyridine with aniline orisopropyl aniline (1:1 mole) [362]. Oxidative degraded chitosan using H2O2 under the catalysis of phosphotungstic acid has been reported to have 94.7% water solubility [363]. Using oxidative degradation assisted by microwave irradiation water-soluble chitosan produced water-soluble chitosan with excellent

Handbook of Chitin and Chitosan

5.5 Conclusion

FIGURE 5.16

153

Water-soluble epoxidized of chitosan [361].

fat- and cholesterol binding capacities [364]. Water-soluble N-(2-hydroxy) propyl-3-trimethylammonium chitosan chloride have been synthesized by the reaction of chitosan with glycidtrimethyl ammonium chloride in neutral aqueous condition [365]. Water-soluble O,N-(2-sulfoethyl) chitosan with degrees of substitution up to 130% have been synthesized by reacting chitosan with sodium 2-chloroethanesulfonate in isopropanol in the presence of NaOH [366]. Ammonium salts of chitosan with different halogens (chitosan bromoacetate, chitosan chloroacetate, chitosan dichloroacetate, chitosan trichloroacetate, and chitosan trifluoroacetate) have been found to be water soluble [367]. Water-soluble chitosan derivatives have been designed using urea groups with quaternary ammonium salt [368]. Water-soluble hydroxyethyl chitosan has been prepared by linking hydroxyethyl groups to the C6 unit in chitosan [369]. The reaction between isoniazid (INH) and N-(3-chloro-2hydroxypropyl) chitosan has been found to yield water-soluble chitin [370]. Reacting acrylic acid and hydroxyethyl acrylate with chitosan under moderate reaction conditions yielded water-soluble chitosan derivatives [371].

5.5 Conclusion The chapter presents basic knowledge of the DA of chitosan, solubility, and distribution of acetyl groups in chitosan. Basic requirements for the application of chitosan in the industry are outlined and discussed. Different methods of measuring the DA are presented.

Handbook of Chitin and Chitosan

154

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

Several modifications that render chitosan water soluble are also presented.

References ´ . Heras, Carbohydr. Polym. 62 (2005) [1] G. Galed, B. Miralles, I. Pan˜os, A. Santiago, A 316 320. [2] M. Kucharska, M.H. Struszczyk, M. Cichecka, K. Brzoza, Prog. Chem. Appl. Chitin Its Deriv. 2012 (2012) 129 136. [3] A.O. Pereira, D.J. Cartucho, A.S. Duarte, M.H. Gil, A.M.S. Cabrita, J.A. Patricio, et al., Curr. Drug Discov. Technol. 2 (2005) 231 238. ¨ zbac¸s-Turan, C. Aral, L. Kabasakal, M. Keyer-Uysal, J. Akbuga, J. Pharm. Pharm. [4] S. O Sci. 6 (2003) 27 32. [5] A. Alishahi, M. Aı¨der, Food Bioprocess Tech. 5 (2012) 817 830. [6] K. Ohkawa, K.-I. Minato, G. Kumagai, S. Hayashi, H. Yamamoto, Biomacromolecules 7 (2006) 3291 3294. [7] H. Honarkar, M. Barikani, Monatsh. Chem. 140 (2009) 1403 1420. [8] S.M. Kuo, G.C.-C. Niu, S.J. Chang, C.H. Kuo, M.S. Bair, J. Appl. Polym. Sci. 94 (2004) 2150 2157. [9] T.J. Madera-Santana, C.H. Herrera-Me´ndez, J.R. Rodrı´guez-Nu´n˜ez, Green Mater. 6 (2018) 131 142. [10] C.K.S.S. Pillai, W. Paul, C.P. Sharma, Prog. Polym. Sci. 34 (2009) 641 678. [11] V. Zargar, M. Asghari, A. Dashti, ChemBioEng Rev. 2 (2015) 204 226. [12] S. Islam, M.A.R.R. Bhuiyan, M.N. Islam, J. Polym. Environ. 25 (2017) 854 866. [13] M.A. Rahman Bhuiyan, A. Shaid, M.A. Khan, Chem. Mater. Eng. 2 (2014) 96 100. [14] D.B. Zekovi´c, S. Kwiatkowski, M.M. Vrvi´c, D. Jakovljevi´c, C.A. Moran, Crit. Rev. Biotechnol. 25 (2005) 205 230. [15] T. Jiang, R. James, S.G. Kumbar, C.T. Laurencin, in: S.G. Kumbar, C.T. Laurencin, M. B.T.-N. and S.B.P. Deng (Eds.), Natural and Synthetic Biomedical Polymers, Elsevier, Oxford, 2014, pp. 91 113. [16] K. Ogawa, T. Yui, K. Okuyama, Int. J. Biol. Macromol. 34 (2004) 1 8. [17] K.V. Harish Prashanth, F.S. Kittur, R.N. Tharanathan, Carbohydr. Polym. 50 (2002) 27 33. [18] K. Kurita, Prog. Polym. Sci. 26 (2001) 1921 1971. [19] A. Aljawish, I. Chevalot, J. Jasniewski, J. Scher, L. Muniglia, J. Mol. Catal. B Enzym. 112 (2014) 25 39. [20] N. Berezina, Phys. Sci. Rev. 1 (2016) 1 8. [21] L. Geng, D.W. Hutmacher, H. Tong Loh, W. Feng, J.Y.H. Fuh, Y. San Wong, Rapid Prototyp. J. 11 (2005) 90 97. [22] K. Okuyama, K. Noguchi, T. Miyazawa, T. Yui, K. Ogawa, Macromolecules 30 (1997) 5849 5855. [23] K. Mazeau, W.T. Winter, H. Chanzy, Macromolecules 27 (1994) 7606 7612. [24] K. Okuyama, K. Noguchi, M. Kanenari, T. Egawa, K. Osawa, K. Ogawa, Carbohydr. Polym. 41 (2000) 237 247. [25] K. Okuyama, K. Noguchi, Y. Hanafusa, K. Osawa, K. Ogawa, Int. J. Biol. Macromol. 26 (1999) 285 293. [26] B. Ma, A. Qin, X. Li, X. Zhao, C. He, Int. J. Biol. Macromol. 64 (2014) 341 346. [27] K. Ogawa, S. Hirano, T. Miyanishi, T. Yui, T. Watanabe, Macromolecules 17 (1984) 973 975. [28] E.L. Mogilevskaya, T. a Akopova, A.N. Zelenetskii, A.N. Ozerin, Polym. Sci. Ser. A 48 (2006) 116 123.

Handbook of Chitin and Chitosan

References

[29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] [41] [42] [43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54] [55] [56] [57] [58] [59] [60] [61] [62] [63] [64]

155

G.L. Clark, A.F. Smith, J. Phys. Chem. 40 (2005) 863 879. G. Molinaro, J.C. Leroux, J. Damas, A. Adam, Biomaterials 23 (2002) 2717 2722. S.B. Rao, C.P. Sharma, J. Biomed. Mater. Res. 34 (1997) 21 28. M. Hasegawa, K. Yagi, S. Iwakawa, M. Hirai, Japanese J. Cancer Res. 92 (2001) 459 466. C. Qin, Y. Du, L. Xiao, Z. Li, X. Gao, Int. J. Biol. Macromol. 31 (2002) 111 117. A. Park, A. Ro-Dong, K.-J. Jo, Y.-Y. Jo, Y.-L. Jin, K.-Y. Kim, et al., J. Microbiol. Biotechnol. 12 (2002) 84 88. S. Roller, N. Covill, Int. J. Food Microbiol. 47 (1999) 67 77. S.S. Koide, Nutr. Res. 18 (1998) 1091 1101. Y.C. Chung, H.L. Wang, Y.M. Chen, S.L. Li, Bioresour. Technol. 88 (2003) 179 184. P.J. VandeVord, H.W.T. Matthew, S.P. DeSilva, L. Mayton, B. Wu, P.H. Wooley, J. Biomed. Mater. Res. 59 (2002) 585 590. M. Ko¨ping-Ho¨gga˚rd, I. Tubulekas, H. Guan, K. Edwards, M. Nilsson, K.M. Va˚rum, et al., Gene Ther. 8 (2001) 1108 1121. H.K. No, S.H. Lee, N.Y. Park, S.P. Meyers, J. Agric. Food Chem. 51 (2003) 7659 7663. M. Ishihara, K. Ono, M. Sato, K. Nakanishi, Y. Saito, H. Yura, et al., Wound Repair Regen. 9 (2001) 513 521. H. Bokura, S. Kobayashi, Eur. J. Clin. Nutr. 57 (2003) 721 725. D.D. Gallaher, C.M. Gallaher, G.J. Mahrt, T.P. Carr, C.H. Hollingshead, R. Hesslink, et al., J. Am. Coll. Nutr. 21 (2002) 428 433. B.O. Schneeman, K. Ebihara, J. Nutr. 119 (1989) 1100 1106. I.M.N. Vold, K.M. Va˚rum, E. Guibal, O. Smidsrød, Carbohydr. Polym. 54 (2003) 471 477. A. Domard, E. Piron, Adv. Chitin Sci. 4 (2000) 295 301. D. Kafetzopoulos, A. Martinou, V. Bouriotis, Proc. Natl. Acad. Sci. 90 (2006) 2564 2568. H.K. No, S.P. Meyers, J. Aquat. Food Prod. Technol. 4 (1995) 27 52. F. Casting, Makromol. Chem. 195 (1975) 1191 1195. K. Tokuyasu, M. Mitsutomi, I. Yamaguchi, K. Hayashi, Y. Mori, Biochemistry 39 (2000) 8837 8843. A.V. Ilyina, N.Y. Tatarinova, V.P. Varlamov, Process Biochem. 34 (1999) 875 878. S. Ichi Aiba, Carbohydr. Res. 265 (1994) 323 328. S. Hajji, I. Younes, O. Ghorbel-Bellaaj, R. Hajji, M. Rinaudo, M. Nasri, et al., Int. J. Biol. Macromol. 65 (2014) 298 306. K.L.B. Chang, G. Tsai, J. Lee, W.-R. Fu, Carbohydr. Res. 303 (1997) 327 332. T. Sannan, K. Kurita, Y. Iwakura, Die Makromol. Chemie 177 (1976) 3589 3600. J. Bossaer (2011) Water soluble chitosan by heterogeneous deacetylation of chitin, Masters Thesis, University of Gent 1 146. P. Methacanon, M. Prasitsilp, T. Pothsree, J. Pattaraarchachai, Carbohydr. Polym. 52 (2003) 119 123. V. Novikov, I. Konovalova, Kuchina , N. Dolgopyatova and Y Cherkun, MSTU, 20(3), 20017, 515-525. S.P. Campana-Filho, R. Signini, M.B. Cardoso, Int. J. Polym. Mater. Polym. Biomater. 51 (2002) 695 700. B. Bradi´c, D. Bajec, A. Pohar, U. Novak, B. Likozar, React. Chem. Eng. 3 (2018) 920 929. G. Lamarque, G. Chaussard, A. Domard, Biomacromolecules 8 (2007) 1942 1950. A. Wojtasz-Paja˛k, J. Szumilewicz, Progress on Chemistry and Application of Chitin and its Derivatives, XIV, 2009, 15 - 24. T.H.D. Ngo, D.N. Ngo, Int. J. Biol. Macromol. 104 (2017) 1604 1610. K. Liang, B. Chang, G. Tsai, J. Lee, W.-R. Fu, Carbohydr. Res., 1997, 327 332.

Handbook of Chitin and Chitosan

156 [65] [66] [67] [68] [69]

[70] [71] [72] [73] [74] [75] [76] [77] [78] [79] [80] [81] [82] [83] [84] [85] [86] [87] [88] [89] [90] [91] [92] [93] [94] [95] [96] [97] [98]

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

C.J. Jiang, M.Q. Xu, Chem. Eng. Technol. 29 (2006) 511 516. R.K. Mohammad, J. Agric. Food Chem. 57 (2009) 1667 1676. S. Ichi Aiba, Int. J. Biol. Macromol. 13 (1991) 40 44. H. Sashiwa, H. Saimoto, Y. Shigemasa, S. Tokura, Carbohydr. Res. 242 (1993) 167 172. T.T. Khong, Vietnamese Chitin Raw Material, the Chitin de-N-Acetylation Reaction, and a New Chitosan-Alginate Gelling Concept (Ph.D Thesis), Norwegian University of Science and Technology, 2013. C. Pardo-Castan˜o, G. Bolan˜os, J. Supercrit. Fluids 151 (2019) 63 74. S. Mima, M. Miya, R. Iwamoto, S. Yoshikawa, J. Appl. Polym. Sci. 28 (1983) 1909 1917. P. le Dung, M. Milas, M. Rinaudo, J. Desbrie`res, Carbohydr. Polym. 24 (1994) 209 214. A.T. Paulino, J.I. Simionato, J.C. Garcia, J. Nozaki, Carbohydr. Polym. 64 (2006) 98 103. I. Younes, O. Ghorbel-Bellaaj, M. Chaabouni, M. Rinaudo, F. Souard, C.C.C. Vanhaverbeke, et al., Int. J. Biol. Macromol. 70 (2014) 385 390. H.K. No, Y.I. Cho, H.R. Kim, S.P. Meyers, J. Agric. Food Chem. 48 (2000) 2625 2627. C. Liu, G. Wang, W. Sui, L. An, C. Si, A.C.S. Sustain, Chem. Eng. 5 (2017) 4690 4698. I. Batista, G. Roberts, Die Makromol. Chemie 434 (1990) 429 434. A.M. Alsharabasy, Polym. Sci. 03 (2017) 1 8. N. Gagne´, B.K. Simpson, Food Biotechnol. 7 (1993) 253 263. S. Chatterjee, M. Adhya, A.K. Guha, B.P. Chatterjee, Process Biochem. 40 (2005) 395 400. W. Suntornsuk, P. Pochanavanich, L. Suntornsuk, Process Biochem. 37 (2002) 727 729. K.D. Rane, D.G. Hoover, Process Biochem. 28 (1993) 115 118. M. Anwar, A.S. Anggraeni, M.H. Al Amin, AIP Conf. Proc. 1823 (() (2017). Y. Yuan, B.M. Chesnutt, W.O. Haggard, J.D. Bumgardner, Materials (Basel) 4 (2011) 1399 1416. X. He, K. Li, R. Xing, S. Liu, L. Hu, P. Li, Egypt. J. Aquat. Res. 42 (2016) 75 81. A.Q. Wang, X.D. Yu, Fine Chem. 15 (1998) 17 19. Q. Song, Q. Li, G. Li, C. Ding, Chinese J. Synth. Chem. 13 (2005) 187 189. A. Domard, M. Rinaudo, Int. J. Biol. Macromol. 5 (1983) 49 52. F.A.A. Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Carbohydr. Polym. 77 (2009) 410 419. D. Tahtat, C. Uzun, M. Mahlous, O. Gu¨ven, Nucl. Instrum. Meth. B 265 (2007) 425 428. G. Lamarque, M. Cretenet, C. Viton, A. Domard, Biomacromolecules 6 (2005) 1380 1388. I. Tsigos, A. Martinou, D. Kafetzopoulos, V. Bouriotis, Trends Biotechnol. 18 (2000) 305 312. Y. Arakane, T. Taira, T. Ohnuma, T. Fukamizo, Curr. Drug Targets 13 (2012) 442 470. J.A. Liu, Y.H. He, Adv. Mater. Res. 236 238 (2011) 996 999. Y. Araki, E. Ito, Eur. J. Biochem. 56 (1974) 669 675. A. Martinou, D. Kafetzopoulos, V. Bouriotis, J. Chromatogr. A 644 (1993) 35 41. X. Gao, T. Katsumoto, K. Onodera, J. Biochem. 117 (1995) 257 263 C. Alfonso, O.M. Nuero, F. Santamarı´a, F. Reyes, Curr. Microbiol. 30 (1995) 49 54.

Handbook of Chitin and Chitosan

References

157

[99] K. Tokuyasu, M. Ohnishi-Kameyama, K. Hayashi, Biosci. Biotechnol. Biochem. 60 (2009) 1598 1603. [100] V. Bouriotis, I. Tsigos, J. Biol. Chem. 270 (1995) 26286 26291. [101] H. Zhang, S. Yang, J. Fang, Y. Deng, D. Wang, Y. Zhao, Carbohydr. Polym. 101 (2014) 57 67. [102] J. Yang, I. Shih, Y. Tzeng, S. Wang, Enzyme Microb. Technol. 26 (2000) 406 413. [103] Y.S. Oh, I.L. Shih, Y.M. Tzeng, S.L. Wang, Enzyme Microb. Technol. 27 (2000) 3 10. [104] S.L. Wang, T.Y. Kao, C.L. Wang, Y.H. Yen, M.K. Chern, Y.H. Chen, Enzyme Microb. Technol. 39 (2006) 724 731. [105] K.L. Morley, G. Chauve, R. Kazlauskas, C. Dupont, F. Shareck, R.H. Marchessault, Carbohydr. Polym. 63 (2006) 310 315. [106] J. Cai, J. Yang, Y. Du, L. Fan, Y. Qiu, J. Li, et al., Carbohydr. Polym. 64 (2006) 151 157. [107] A. Martinou, V. Bouriotis, B.T. Stokke, K.M. Va˚rum, Carbohydr. Res. 311 (1998) 71 78. [108] M. Kohlhoff, A. Niehues, J. Wattjes, J. Be´ne´teau, S. Cord-Landwehr, N.E. El Gueddari, et al., Carbohydr. Polym. 174 (2017) 1121 1128. [109] I. Aspras, M.M. Jaworska, A. Go´rak, J. Biotechnol. 251 (2017) 94 98. ´ ´ [110] I. Aspras, M. Kaminska, K. Karzynski, M. Kawka, M.M. Jaworska, Chem. Process Eng. Inz. Chem. Proces. 37 (2016) 77 82. [111] N. Pareek, V. Vivekanand, P. Agarwal, S. Saroj, R.P. Singh, Carbohydr. Polym. 96 (2013) 417 425. [112] A. Martinou, D. Kafetzopoulos, V. Bouriotis, Carbohydr. Res. 273 (1995) 235 242. [113] K.N. Aye, R. Karuppuswamy, T. Ahamed, W.F. Stevens, Bioresour. Technol. 97 (2006) 577 582. [114] F. Caufrier, A. Martinou, C. Dupont, V. Bouriotis, Carbohydr. Res. 338 (2003) 687 692. [115] R. Willaert, V.A. Nedovic, J. Chem. Technol. Biotechnol. 1367 (2006) 1353 1367. [116] N. Win, W. Stevens, Appl. Microbiol. Biotechnol. 57 (2002) 334 341. [117] N.N. Win, G. Pengju, W.F. Stevens, Adv. Chitin Sci. 4 (2000) 55 62. [118] Z.E.S.I. Kolodziejska, A. Wojtasz-Pajak, G. Ogonowska, Bull. Sea Fish. Institute 2 (2000) 15 24. ´ ´ [119] R. Czechowska-biskup, D. Jarosinska, B. Rokita, P. Ulanski, J.M. Rosiak, Prog. Chem. Appl. Chitin Its Deriv. 17 (2012) 5 20. [120] D.H. Davies, E.R.B.T.-M. in E. Hayes, Biomass Part B Lignin, Pectin, Chitin, Academic Press, 1988, pp. 442 446. [121] Z.M. dos Santos, A.L.P.F. Caroni, M.R. Pereira, D.R. da Silva, J.L.C. Fonseca, Carbohydr. Res. 344 (2009) 2591 2595. [122] S. Arcidiacono, D.L. Kaplan, Biotechnol. Bioeng. 39 (1992) 281 286. [123] N. Bala´zs, P. Sipos, Carbohydr. Res. 342 (2007) 124 130. [124] E.S. de Alvarenga, C. Pereira de Oliveira, C. Roberto Bellato, Carbohydr. Polym. 80 (2010) 1155 1160. [125] L. Raymond, F.G. Morin, R.H. Marchessault, Carbohydr. Res. 246 (1993) 331 336. [126] T. Hattori, K. Katai, M. Kato, M. Izume, Y. Mizuta, Bull. Chem. Soc. Jpn. 72 (1999) 37 41. [127] J.W. Park, K.H. Choi, K.K. Park, Bull. Korean Chem. Soc. 4 (2) (1983) 68 72. [128] K. Toˆei, T. Kohara, Anal. Chim. Acta 83 (1976) 59 65. [129] Y. Zhang, X. Zhang, R. Ding, J. Zhang, J. Liu, Carbohydr. Polym. 83 (2011) 813 817. [130] F. Niola, N. Basora, E. Chornet, P.F. Vidal, Carbohydr. Res. 238 (1993) 1 9. [131] F. Nanjo, R. Katsumi, K. Sakai, Anal. Biochem. 193 (1991) 164 167. [132] H. Sato, S.I. Mizutani, S. Tsuge, H. Ohtani, K. Aoi, A. Takasu, et al., Anal. Chem. 70 (1998) 7 12.

Handbook of Chitin and Chitosan

158

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

[133] C. Niklasson, M.J. Taherzadeh, A. Jeihanipour, A. Zamani, L. Edebo, J. Agric. Food Chem. 56 (2008) 8314 8318. [134] S. Ichi Aiba, Int. J. Biol. Macromol. 8 (1986) 173 176. [135] R.A.A. Muzzarelli, R. Rocchetti, Carbohydr. Polym. 5 (1985) 461 472. [136] A. Baxter, M. Dillion, K. Taylor, G. Roberts, Int. J. Biol. Macromol. 14 (1992) 166 169. [137] M.L. Duarte, M.C. Ferreira, M.R. Marva˜o, J. Rocha, Int. J. Biol. Macromol. 31 (2002) 1 8. [138] M.R. Kasaai, Carbohydr. Polym. 71 (2008) 497 508. [139] M. Miya, R. Iwamoto, S. Yoshikawa, S. Mima, Int. J. Biol. Macromol. 2 (1980) 323 324. [140] G.K. Moore, G.A.F. Roberts, Int. J. Biol. Macromol. 2 (1980) 115 116. [141] T. Sannan, Polymer (Guildf) 19 (1978) 458. [142] G. Roberts, M. Rinaudo, J. Brugnerotto, J. Desbrie, Polymer (Guildf) 42 (2001) 9921 9927. [143] E. Fernandez-Megia, R. Novoa-Carballal, E. Quin˜oa´, R. Riguera, Carbohydr. Polym. 61 (2005) 155 161. [144] K.M. Va˚rum, M.W. Antohonsen, H. Grasdalen, O. Smidsrød, Carbohydr. Res. 211 (1991) 17 23. [145] M. Lavertu, Z. Xia, A.N. Serreqi, M. Berrada, A. Rodrigues, D. Wang, et al., J. Pharm. Biomed. Anal. 32 (2003) 1149 1158. [146] L. Heux, J. Brugnerotto, J. Desbrie`res, M.F. Versali, M. Rinaudo, Biomacromolecules 1 (2000) 746 751. [147] L. Manni, O. Ghorbel-Bellaaj, K. Jellouli, I. Younes, M. Nasri, Appl. Biochem. Biotechnol. 162 (2010) 345 357. [148] M.R. Kasaai, Carbohydr. Polym. 79 (2010) 801 810. [149] G. Er Yu, F.G. Morin, G.A.R. Nobes, R.H. Marchessault, Macromolecules 32 (1999) 518 520. [150] H. Sashiwa, H. Saimoto, Y. Shigemasa, R. Ogawa, S. Tokura, Carbohydr. Polym. 16 (1991) 291 296. [151] I. Garcia-Alonso, C. Peniche-Covas, J.M. Nieto, J. Therm. Anal. 28 (1983) 189 193. [152] R.A.A. Muzzarelli, F. Tanfani, G. Scarpini, J. Biochem. Biophys. Methods 2 (1980) 299 306. [153] M.H. Ottøy, K.M. Va˚rum, O. Smidsrød, Carbohydr. Polym. 29 (1996) 17 24. [154] M.R. Kasaai, J. Arul, S.L. Chin, G. Charlet, J. Photochem. Photobiol. A Chem. 120 (1999) 201 205. [155] S. Tokura, N. Nishi, A. Tsutsumi, O. Somorin, Polym. J. 15 (1983) 485. [156] Y. Dong, C. Xu, J. Wang, Y. Wu, M. Wang, Y. Ruan, J. Appl. Polym. Sci. 83 (2002) 1204 1208. [157] S.S. Kim, S.H. Kim, Y.M. Lee, J. Polym. Sci. Part B Polym. Phys. 34 (1996) 2367 2374. [158] A. Hirai, H. Odani, A. Nakajima, Polym. Bull. 26 (1991) 87 94. [159] S.-C. Hsu, T.-M. Don, W.-Y. Chiu, Polym. Degrad. Stab. 75 (2002) 73 83. [160] W. Wang, S. Bo, S. Li, W. Qin, Int. J. Biol. Macromol. 13 (1991) 281 285. [161] H. Terayama, Method of colloid titration (a new titration between polymer ions), J. Polym. Sci. (1952). [162] Q. Ke, H. Chen, Huaxue Tongbao 10 (1990) 44 46. [163] S.C. Tan, E. Khor, T.K. Tan, S.M. Wong, Talanta 45 (1998) 713 719. [164] A. Tolaimate, J. Desbri??res, M. Rhazi, A. Alagui, M. Vincendon, P. Vottero, et al., Polymer (Guildf) 41 (2002) 2463 2469. [165] X. Jiang, L. Chen, W. Zhong, Carbohydr. Polym. 54 (2003) 457 463. [166] S. Prochazkova, K.M. Va˚rum, K. Ostgaard, Carbohydr. Polym. 38 (1999) 115 122. [167] Z. Holan, J. Votruba, V. Vlasa´kova´, J. Chromatogr. A 190 (1980) 67 76.

Handbook of Chitin and Chitosan

References

159

[168] S. Sabnis, L.H. Block, Polym. Bull. 39 (1997) 67 71. [169] S. Beil, A. Schamberger, W. Naumann, S. MacHill, K.H. Van Pe´e, Carbohydr. Polym. 87 (2012) 117 122. [170] J. Brugnerotto, J. Lizardi, F.M. Goycoolea, W. Argu¨elles-Monal, J. Desbrie`res, M. Rinaudo, Polymer (Guildf) 42 (2001) 3569 3580. [171] Y. Shigemasa, H. Matsuura, H. Sashiwa, H. Saimoto, Int. J. Biol. Macromol. 18 (1996) 237 242. [172] G.K. Moore, G.A.F. Roberts, in: Proceedings of the First International Conference on Chitin/Chitosan, MIT Sea Grant Program, Cambridge, MA, 1978, pp. 421 425. [173] Y. Shigemasa, H. Matsuura, H. Sashiwa, H. Saimato, Adv. Chitin Sci. 1 (1996) 204 209. [174] C. Peniche, C. Elvira, J. San Roman, Polymer (Guildf) 39 (1998) 6549 6554. [175] S.G. Caridade, C. Monge, J. Almodo´var, R. Guillot, J. Lavaud, V. Josserand, et al., Acta Biomater. 15 (2015) 139 149. [176] K.T. Hwang, J.T. Kim, S.T. Jung, G.S. Cho, H.J. Park, J. Appl. Polym. Sci. 89 (2003) 3476 3484. [177] T.D. Rathke, S.M. Hudson, J. Polym. Sci. Part A Polym. Chem. 31 (1993) 749 753. [178] K.M. Va˚rum, B. Egelandsdal, M.R. Ellekjłr, Carbohydr. Polym. 28 (1995) 187 193. [179] M.X. Weinhold, J.C.M. Sauvageau, J. Kumirska, J. Tho¨ming, Carbohydr. Polym. 78 (2009) 678 684. [180] J.J. Thevarajah, M.P. Van Leeuwen, H. Cottet, P. Castignolles, M. Gaborieau, Int. J. Biol. Macromol. 95 (2017) 40 48. [181] S. Ichi Aiba, Int. J. Biol. Macromol. 11 (1989) 249 252. [182] Z. Han, Y. Zeng, H. Lu, L. Zhang, Carbohydr. Res. 413 (2015) 75 84. [183] S. Ichi Aiba, Int. J. Biol. Macromol. 15 (1993) 241 245. [184] R. Olsen, D. Schwartzmiller, W. Weppner, R. Winandy, Chitin and Chitosan (1989) 813 828. [185] P.A. Sanford, G.P. Hutchings, Prog. Biotechnol. (1987). [186] N. Errington, S. Harding, K. Varum, L. Illum, Int. J. Biol. Macromol. 15 (1993) 113 117. [187] M. Rinaudo, M. Milas, P. Le Dung, Int. J. Biol. Macromol. 15 (1993) 281 285. [188] Y. Dang, K. Bin Li, Q. Luo, H. Wei, H.S. Guo, 2011 Int. Conf. Electr. Inf. Control Eng. ICEICE 2011—Proc. 25 (2011) 4025 4029. [189] N. Zydowicz, L.A. Vachoud, L.A. Domard, Adv. Chitin Sci. 1 (1996) 262. [190] Y. Shigemasa, K. Saito, H. Sashiwa, H. Saimoto, Int. J. Biol. Macromol. 16 (1994) 43 49. [191] N. Hutadilok, T. Mochimasu, H. Hisamori, K. Ichiro Hayashi, H. Tachibana, T. Ishii, et al., Carbohydr. Res. 268 (1995) 143 149. [192] R.J. Nordtveit, K.M. Va˚rum, O. Smidsrød, Carbohydr. Polym. 29 (1996) 163 167. [193] G. Peluso, O. Petillo, M. Ranieri, M. Santin, L. Ambrosic, D. Calabro´, et al., Biomaterials 15 (1994) 1215 1220. [194] C. Wenling, J. Biomater. Appl. 20 (2005) 157 177. [195] R. Hussain, Int. J. Adv. Eng. Appl. 1, 2013, 4 12. [196] C. Chatelet, Biomaterials 22 (2001) 261 268. [197] L.J.R. Foster, S. Ho, J. Hook, M. Basuki, H. Marc¸al, PLoS One 10 (2015) 1 22. [198] P. Sorlier, A. Denuzie`re, C. Viton, A. Domard, Biomacromolecules 2 (2001) 765 772. [199] R. Hussain, T.K. Maji, T.K. Maji, Rev. Te´c. Ing. Univ. Zulia 37 (2014) 69 77. [200] A. Wan, Q. Gao, H. Li, J. Appl. Polym. Sci. 117 (2010) 2183 2188. [201] Z. Zakaria, Z. Izzah, M. Jawaid, A. Hassan, BioResources 7 (2012) 5568 5580. [202] Y. Omura, M. Shigemoto, T. Akiyama, H. Saimoto, Y. Shigemasa, I. Nakamura, et al., Science 8 (2003) 25 30. [203] C. Carvalho, J. Silva-Correia, R.L.R. J.M. Oliveira, Front. Bioeng. Biotechnol. (n.d.).

Handbook of Chitin and Chitosan

160

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

[204] I.F. Amaral, M. Lamghari, S.R. Sousa, P. Sampaio, M.A. Barbosa, J. Biomed. Mater. Res. Part A 75 (2005) 387 397. [205] K. Tomihata, Y. Ikada, Biomaterials 18 (1997) 567 575. [206] K.M. Va˚rum, M.M. Myhr, R.J.N. Hjerde, O. Smidsrød, Carbohydr. Res. 299 (1997) 99 101. [207] Y. Hidaka, I. Michio, K. Mori, H. Yagasaki, A.H. Kafrawy, J. Biomed. Mater. Res. 46 (1999) 418 423. [208] M. Prasitsilp, R. Jenwithisuk, K. Kongsuwan, N. Damrongchai and P. Watts, J. Mater. Sci. Mater. Med. 11 (2000) 773 778. [209] T. Freier, H.S. Koh, K. Kazazian, M.S. Shoichet, Biomaterials 26 (2005) 5872 5878. [210] M. Jaworska, K. Sakurai, P. Gaudon, E. Guibal, Polym. Int. 52 (2003) 198 205. [211] S.M. Lim, D.K. Song, K.J. Cho, S.H. Oh, D.S. Lee-Yoon, E.H. Bae, J.H. Lee, in: F. Ibrahim, N.A.A. Osman, J. Usman, N.A. Kadri (Eds.), Springer Berlin Heidelberg, Berlin, Heidelberg, 2007, pp. 94 97. [212] K. Sakurai, T. Shibano, K. Kimura, T. Takahashi, Sen’i Gakkaishi 41 (1985) T361 T368. [213] M. Fan, Q. Hu, K. Shen, Carbohydr. Polym. 78 (2009) 66 71. [214] T. Wang, M. Turhan, S. Gunasekaran, Polym. Int. 53 (2004) 911 918. [215] T.Q. Yuan, J. He, F. Xu, R.C. Sun, Polym. Degrad. Stab. 94 (2009) 1142 1150. [216] S.I. Nishimura, O. Kohgo, K. Kurita, H. Kuzuhara, Macromolecules 24 (1991) 4745 4748. [217] B. Prathab, T.M. Aminabhavi, J. Polym. Sci. Part B Polym. Phys. 45 (2007) 1260 1270. [218] R. Ravindra, K.R. Krovvidi, A.A. Khan, Carbohydr. Polym. 36 (1998) 121 127. [219] M. Kumar, React. Funct. Polym. 46 (2000) 1 27. [220] R.A.A. Muzzarelli, Carbohydr. Polym. 3 (1983) 53 75. [221] H. Yi, L.Q. Wu, W.E. Bentley, R. Ghodssi, G.W. Rubloff, J.N. Culver, et al., Biomacromolecules 6 (2005) 2881 2894. [222] Y.W. Cho, J. Jang, C.R. Park, S.W. Ko, Biomacromolecules 1 (2000) 609 614. [223] M. Morimoto, H. Saimoto, Y. Shigemasa, Trends Glycosci. Glycotechnol. 14 (2002) 205 222. [224] P. Sorlier, C. Viton, A. Domard, Biomacromolecules 3 (2002) 1336 1342. [225] W. Xie, P. Xu, W. Wang, Q. Liu, Carbohydr. Polym. 50 (2002) 35 40. [226] H.Y. Lin, C.C. Chou, Food Res. Int. 37 (2004) 883 889. [227] L.Y. Zheng, J.F. Zhu, Carbohydr. Polym. 54 (2003) 527 530. [228] T. Oikawa, Chem. Pharm. Bull. (2002) 2091. [229] B.O. Jung, C.H. Kim, K.S. Choi, Y.M. Lee, J.J. Kim, J. Appl. Polym. Sci. 72 (1999) 1713 1719. [230] R.G. Barbosa, M. Trigo, R. Prego, R. Fett, S.P. Aubourg, Int. J. Food Sci. Technol. 53 (2018) 271 281. [231] Z.M. Kochkina, S.N. Chirkov, Microbiology 69 (2000) 217 219. [232] C.H. Kim, J.W. Choi, H.J. Chun, K.S. Choi, Polym. Bull. 38, 1997, 387 393. [233] R. Muzzarelli, R. Tarsi, O. Filippini, E. Giovanetti, G. Biagini, P.E. Varaldo, Antimicrob. Agents Chemother. 34 (1990) 2019 2023. [234] N. Nishi, S. Tokura, K. Nishimura, I. Azuma, Science 146 (80 ) (1986) 251 258. [235] A. Tokoro, M. Kobayashi, N. Tatewaki, T. Mikami, K. Suzuki, Y. Okawa, et al., Microbiol. Immunol. 33 (1989) 357 367. [236] M. Anraku, A. Hiraga, D. Iohara, K. Uekama, H. Tomida, M. Otagiri, et al., Int. J. Biol. Macromol. 70 (2014) 64 69. [237] X. Zhu, X. Zhou, J. Yi, J. Tong, H. Wu, L. Fan, Int. J. Biol. Macromol. 70 (2014) 300 305.

Handbook of Chitin and Chitosan

References

161

[238] J. Liu, C. Guang Meng, S. Liu, J. Kan, C. Hai Jin, Food Hydrocoll. 63 (2017) 457 466. [239] N.S. Chatterjee, S.K. Panda, M. Navitha, K.K. Asha, R. Anandan, S. Mathew, J. Food Sci. Technol. 52 (2015) 7153 7162. [240] M. Liu, L. Min, C. Zhu, Z. Rao, L. Liu, W. Xu, et al., Int. J. Biol. Macromol. 104 (2017) 732 738. [241] I. Khan, S. Ullah, D.H. Oh, Carbohydr. Polym. 152 (2016) 87 96. [242] J. Liu, X. Yuan Wen, J. Feng Lu, J. Kan, C. Hai Jin, Int. J. Biol. Macromol. 65 (2014) 97 106. [243] A.O. Aytekin, S. Morimura, K. Kida, J. Biosci. Bioeng. 111 (2011) 212 216. [244] Q. Hu, T. Wang, M. Zhou, J. Xue, Y. Luo, J. Agric. Food Chem. 64 (2016) 5893 5900. [245] J. Liu, J. Feng Lu, J. Kan, Y. Qing Tang, C. Hai Jin, Int. J. Biol. Macromol. 62 (2013) 85 93. [246] S. Woranuch, R. Yoksan, Carbohydr. Polym. 96 (2013) 495 502. [247] S.H. Yu, F.L. Mi, J.C. Pang, S.C. Jiang, T.H. Kuo, S.J. Wu, et al., Carbohydr. Polym. 84 (2011) 794 802. [248] C.H. Wang, W.S. Liu, J.F. Sun, G.G. Hou, Q. Chen, W. Cong, et al., Int. J. Biol. Macromol. 84 (2016) 418 427. [249] R. Li, Z. Guo, P. Jiang, Carbohydr. Res. 345 (2010) 1896 1900. [250] C.S. Oliveira, C. Airoldi, Carbohydr. Polym. 102 (2014) 38 46. [251] R.C. Chien, M.T. Yen, J.L. Mau, Carbohydr. Polym. 138 (2016) 259 264. [252] Q. Li, W. Tan, C. Zhang, G. Gu, Z. Guo, Int. J. Biol. Macromol. 91 (2016) 623 629. [253] Y. Chen, J. Li, Q. Li, Y. Shen, Z. Ge, W. Zhang, et al., Carbohydr. Polym. 143 (2016) 246 253. [254] X. Feng, K. Zheng, C. Wang, F. Chu, Y. Chen, Fibers Polym. 17 (2016) 371 379. [255] M. Badawy, E. Rabea, Synthesis and characterization of N-(maleoyl) chitosan at different degrees of substitution with antibacterial activity, J. Polym. Mater. 34 (2017) 249 259. [256] Z. Wang, L. Zheng, C. Li, S. Wu, Y. Xiao, J. Appl. Polym. Sci. 134 (2017). [257] N.L. Vanden Braber, L.I. Dı´az Vergara, F.E. Mora´n Vieyra, C.D. Borsarelli, M.M. Yossen, J.R. Vega, et al., Int. J. Biol. Macromol. 102 (2017) 200 207. [258] M.J. Moreno-Va´squez, E.L. Buitimea-Valenzuela, M. Plascencia-Jatomea, J.C. Encinas-Encinas, F. Rodrı´guez-Fe´lix, S. Sa´nchez-Valdes, et al., Carbohydr. Polym. 155 (2017) 117 127. [259] T.M. Tamer, M.A. Hassan, A.M. Omer, K. Valachova´, M.S.M. Eldin, M.N. Collins, et al., Carbohydr. Polym. 169 (2017) 441 450. [260] Z. Li, F. Yang, R. Yang, Int. J. Biol. Macromol. 75 (2015) 378 387. ¨ . Vidar Ru´narsson, J. Holappa, C. Malainer, H. Steinsson, M. Hja´lmarsdo´ttir, T. [261] O Nevalainen, et al., Eur. Polym. J. 46 (2010) 1251 1267. [262] R. Tang, Y. Zhang, Y. Zhang, Z. Yu, Carbohydr. Polym. 139 (2016) 191 196. [263] R. Tang, Z. Yu, Y. Zhang, C. Qi, Cellulose 23 (2016) 1741 1749. [264] G. Srivastava, S. Walke, D. Dhavale, W. Gade, J. Doshi, R. Kumar, et al., Int. J. Biol. Macromol. 91 (2016) 381 393. ´ lvarez, L.C.C. Iturbe, I. Katime, Carbohydr. Polym. 101 [265] M.A. Pujana, L. Pe´rez-A (2014) 113 120. [266] S. Hunt, T.N. Huckerby, Comp. Biochem. Physiol. Part B Biochem 88 (1987) 1107 1116. [267] G. Romanazzi, F.M. Gabler, D. Margosan, B.E. Mackey, J.L. Smilanick, Phytopathology 99, 2009, 1028 1036 [268] B.J. Smith, J. Am. Chem. Soc. 119 (1997) 2699 2706. [269] M. Rinaudo, G. Pavlov, J. Desbrie`res, Polymer. 40 (1999) 7029 7032 [270] M. Rinaudc, G. Pavlov, J. Desbrie`res, Int. J. Polym. Anal. Charact. 5 (1999) 267 276.

Handbook of Chitin and Chitosan

162

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

[271] N. Gorochovceva, R. Makuˇska, Eur. Polym. J. 40 (2004) 685 691. [272] M. Ren, Z. Huixia, Chinese J. Tissue Eng. Res. 11 (2007) 9817 9820. [273] A. Yoshifuji, Y. Noishiki, M. Wada, L. Heux, S. Kuga, Biomacromolecules 7 (2006) 2878 2881. [274] Muzzarelli, R.A.A. (1991), ChemInform Abstract: Modified Chitosans and Their Chemical Behavior. ChemInform, 22. doi:10.1002/chin.199111346 [275] D.W. Jenkins, S.M. Hudson, Chem. Rev. 101 (2001) 3245 3273. [276] R. Jayakumar, M. Prabaharan, R.L. Reis, J.F. Mano, Carbohydr. Polym. 62 (2005) 142 158. [277] R.A.A. Muzzarelli, C. Muzzarelli, Adv. Polym. Sci 186 (2005) 151 209. [278] K.V. Harish Prashanth, R.N. Tharanathan, Trends Food Sci. Technol. 18 (2007) 117 131. [279] S. Fuji, H. Kumagai, M. Noda, Carbohydr. Res. 83 (1980) 389 393. [280] E. Loubaki, M. Ourevitch, S. Sicsic, Eur. Polym. J. 27 (1991) 311 317. [281] G.K. Moore, G.A.F. Roberts, Int. J. Biol. Macromol. 2, 1980, 73 77. [282] D.H. Li, L.M. Liu, K.L. Tian, J.C. Liu, X.Q. Fan, Carbohydr. Polym. 67 (2007) 40 45. [283] G. Kumar, P.J. Smith, G.F. Payne, Biotechnol. Bioeng. 63 (1999) 154 165. [284] C. Muzzarelli, R.A.A. Muzzarelli, Trends Glycosci. Glycotechnol. 14 (2002) 223 229. [285] L. Shao, G. Kumar, J.L. Lenhart, P.J. Smith, G.F. Payne, Enzyme Microb. Technol. 25 (1999) 660 668. [286] T. Chen, G. Kumar, M.T. Harris, P.J. Smith, G.F. Payne, Biotechnol. Bioeng. 70 (2000) 564 573. [287] L.G. Engibaryan, A.I. Chernukhina, G.A. Gabrielyan, L.S. Gal’braikh, Fibre Chem. 37 (2005) 285 288. [288] S. Chen, Y. Wang, J. Appl. Polym. Sci. 82 (2001) 2414 2421. [289] A.F. Groboillot, C.P. Champagne, G.D. Darling, D. Poncelet, R.J. Neufeld, Biotechnol. Bioeng. 42 (1993) 1157 1163. [290] N. Kubota, Y. Eguchi, Polym. J. 29 (2005) 123 127. [291] K. Kurita, M. Kamiya, S.I. Nishimura, Carbohydr. Polym. 16 (1991) 83 92. [292] G.A. Vikhoreva, L.S. Gal, Rev. Lit. Arts Am. 31 (1999) 25 29. [293] H. Sashiwa, Y. Shigemasa, Carbohydr. Polym. 39 (1999) 127 138. [294] H. Sashiwa, N. Kawasaki, A. Nakayama, E. Muraki, N. Yamamoto, S. Aiba, Biomacromolecules 3 (2002) 1126 1128. [295] Z. Zong, Y. Kimura, M. Takahashi, H. Yamane, Polymer (Guildf) 41 (2000) 899 906. [296] Y. Wu, T. Seo, T. Sasaki, S. Irie, K. Sakurai, Carbohydr. Polym. 63 (2006) 493 499. [297] X. Xiangyang, L. Ling, Z. Jianping, L. Shiyue, Y. Jie, Y. Xiaojin, et al., Colloids Surfaces B Biointerfaces 55 (2007) 222 228. [298] Y. Kato, Biomaterials 25 (2004) 907 915. [299] L. Yan, Y.B. Zheng, F. Zhao, S. Li, X. Gao, B. Xu, et al., Chem. Soc. Rev. 41 (2012) 97 114. [300] Z. Aiping, C. Tian, Y. Lanhua, W. Hao, L. Ping, Carbohydr. Polym. 66 (2006) 274 279. [301] Y. Uchida, M.Izume, and A. Ohtakara, In: Chitin and Chitosan : Skjak-Braek, G., Anthonsen, T. and Sandford (Eds.), (1989) 373 382. London: Elsevier. [302] M. Tajima, M. Izume, T. Fukuhara, T. Kimura, Y. Kuroyanagi, J.-Japanese Soc. Biomater. 18 (2000) 220 226. [303] Y. Kuroyanagi, A. Shiraishi, Y. Shirasaki, N. Nakakita, Y. Yasutomi, Y. Takano, et al., Wound Repair Regen. 2 (1994) 122 129. [304] Y. Kato, H. Onishi, Y. Machida, Carbohydr. Res. 337 (2002) 561 564. [305] Y. Kato, H. Onishi, Y. Machida, Macromol. Res. 11 (2003) 382 386. [306] T.A.I. Moriyasu, Carbohydr Res.92 (1981) 323 327. [307] H. Sashiwa, N. Kawasaki, A. Nakayama, E. Muraki, N. Yamamoto, H. Zhu, et al., Biomacromolecules 3 (2002) 1120 1125.

Handbook of Chitin and Chitosan

References

163

[308] H. Sashiwa, Y. Shigemasa, R. Roy, Macromolecules 34 (2001) 3211 3214. [309] T. Seo, Y. Ikeda, K. Torada, Y. Nakata, Y. Shimomura, Chitin Chitosan Res. 7 (2001) 212 213. [310] K. Inoue, K. Yoshizuka, K. Ohto, H. Nakagawa, Chem. Lett. 30 (2001) 698 699. [311] S. Hirano, Polym. Int. 48 (2002) 732 734. [312] K. Kurita, Polym. Degrad. Stab. 59 (1998) 117 120. [313] H. Baumann, C. Liu, V. Faust, Cellulose 10 (2003) 65 74. [314] C. Qin, H. Li, Q. Xiao, Y. Liu, J. Zhu, Y. Du, Carbohydr. Polym. 63 (2006) 367 374. [315] Y. Li, X.G. Chen, N. Liu, C.S. Liu, C.G. Liu, X.H. Meng, et al., Carbohydr. Polym. 67 (2007) 227 232. [316] C. Le Tien, M. Lacroix, P. Ispas-Szabo, M.A. Mateescu, J. Control. Release 93 (2003) 1 13. [317] S. Hirano, Y. Yamaguchi, M. Kamiya, Carbohydr. Polym. 48 (2002) 203 207. [318] S. Hirano, Y. Yamaguchi, M. Kamiya, Macromol. Biosci. 3 (2003) 629 631. [319] H. Sashiwa, Y. Makimura, Y. Shigemasa, R. Roy, Chem. Commun. 061 (2000) 909 910. [320] K.G.P.C. De Mello, L.D.C. Bernusso, R.N.D.M. Pitombo, B. Polakiewicz, Brazilian Arch. Biol. Technol. 49 (2006) 665 668. [321] R. Yamaguchi, Y. Arai, T. Itoh, S. Hirano, Carbohydr. Res. 88 (1981) 172 175. [322] M.R. Avadi, M.J. Zohuriaan-Mehr, P. Younessi, M. Amini, M.R. Tehrani, A. Shafiee, J. Bioact. Compat. Polym. 18 (2003) 469 479. [323] L. Szosland, Adv. Chitin Sci. 1 (1995) 297 302. [324] L. Szosland, Chitin Handb. (1997) 53 60. [325] D. Van Luyen, V. Rossbach, J. Appl. Polym. Sci. 55 (1995) 679 685. [326] J.M. Wasikiewicz, H. Mitomo, N. Nagasawa, T. Yagi, M. Tamada, F. Yoshii, J. Appl. Polym. Sci. 102 (2006) 758 767. [327] S. Santhosh, P.T. Mathew, J. Appl. Polym. Sci. 107 (2008) 280 285. [328] Z. Li, X. Zhuang, X. Liu, Y. Guan, K. Yao, J. Appl. Polym. Sci. 88 (2003) 1719 1725. [329] T.K. Sini, S. Santhosh, P.T. Mathew, Polymer (Guildf) 46 (2005) 3128 3131. [330] R.A.A. Muzzarelli, P. Ilari, M. Petrarulo, Int. J. Biol. Macromol. 16 (1994) 177 180. [331] M. Rinaudo, P. Le Dung, C. Gey, M. Milas, Int. J. Biol. Macromol. 14 (1992) 122 128. [332] Y. Xie, X. Liu, Q. Chen, Carbohydr. Polym. 69 (2007) 142 147. [333] J.H. Park, Y.W. Cho, H. Chung, I.C. Kwon, S.Y. Jeong, Biomacromolecules 4 (2003) 1087 1091. [334] D. Hall, (n.d.) 1153 1154. [335] M. Yalpani, L.D. Hall, Macromolecules 17 (1984) 272 281. [336] Y.C. Chung, C.L. Kuo, C.C. Chen, Bioresour. Technol. 96 (2005) 1473 1482. [337] Y.C. Chung, C.F. Tsai, C.F. Li, Fish. Sci. 72 (2006) 1096 1103. [338] K. Kurita, H. Akao, J. Yang, M. Shimojoh, Biomacromolecules 4 (2003) 1264 1268. [339] K. Kurita, K. Shimada, Y. Nishiyama, M. Shimojoh, S.I. Nishimura, Macromolecules 31 (1998) 4764 4769. [340] K. Kurita, S. Nishimura, Y. Koyama, S. Inoue, O. Kohgo, in: Abstr. Pap. Am. Chem. Soc. (1990) 295. [341] W. Xie, P. Xu, W. Wang, Q. Liu, J. Appl. Polym. Sci. 85 (2002) 1357 1361. [342] A. Dal Pozzo, L. Vanini, M. Fagnoni, M. Guerrini, A. De Benedittis, R.A.A. Muzzarelli, Carbohydr. Polym. 42 (2000) 201 206. [343] A.C. Wilson, M.A. Us, H. Zhang, S.B. Glazer, 2 (2016). [344] E. Bombardelli, R.U.S.A. Data, J.T.H. Book, K. Tebib, E. Bombardelli, F. Application, P. Data, P.E.P. Weber, 1 (2001) 4 6. [345] U. Wagner, H.J. Limburgerhof, K. Henrici, H. Kuessner, K. Volkamer, E. Fuerst, BASF Aktiengesellschaft (1982) 0 4.

Handbook of Chitin and Chitosan

164

5. Solubility, degree of acetylation, and distribution of acetyl groups in chitosan

[346] Y. Lin, Q. Chen, H. Luo, Carbohydr. Res. 342 (2007) 87 95. [347] R. Jayakumar, R.L. Reis, J.F. Mano, J. Macromol. Sci. Part A 44 (2007) 271 275. [348] J. Cho, J. Grant, M. Piquette-Miller, C. Allen, Biomacromolecules 7 (2006) 2845 2855. [349] A. Heras, N.M. Rodrı´guez, V.M. Ramos, E. Agullo´, Carbohydr. Polym. 44 (2001) 1 8. [350] S.H. Lim, S.M. Hudson, Carbohydr. Res. 339 (2004) 313 319. [351] M. Yazdani-Pedram, J. Retuert, J. Appl. Polym. Sci. 63 (1997) 1321 1326. [352] J. Du, Y. Lo Hsieh, Cellulose 14 (2007) 543 552. [353] X. Guan, D. Quan, X. Shuai, K. Liao, K. Mai, J. Polym. Sci. Part A Polym. Chem. 45 (2007) 2556 2568. ˇ ´ s, E. MacHova´, Ultrason. [354] D. Misloviˇcova´, J. Masa´rova´, K. Bendˇza´lova´, L. Solte Sonochem. 7 (2000) 63 68. [355] H. Feng, C.M. Dong, Carbohydr. Polym. 70 (2007) 258 264. [356] K. Aoi, A. Takasu, M. Okada, T. Imae, Macromol. Chem. Phys. 203 (2002) 2650 2657. [357] A.V. Ilyina, V.E. Tikhonov, A.I. Albulov, V.P. Varlamov, Process Biochem. 35 (2000) 563 568. [358] E.S. Lee, K.H. Park, I.S. Park, K. Na, Int. J. Pharm. 338 (2007) 310 316. [359] I. Kołodziejska, B. Piotrowska, Food Chem. 103 (2007) 295 300. [360] F.L.T. Shee, J. Arul, S. Brunet, A.M. Mateescu, L. Bazinet, J. Agric. Food Chem. 54 (2006) 6760 6764. [361] Y.W. Cho, Y.N. Cho, S.H. Chung, G. Yoo, S.W. Ko, Biomaterials 20 (1999) 2139 2145. [362] T. Baran, A. Mente¸s, Int. J. Biol. Macromol. 79 (2015) 542 554. [363] Z. Xia, S. Wu, J. Chen, Int. J. Biol. Macromol. 59 (2013) 242 245. [364] Q. Jin, H. Yu, X. Wang, K. Li, P. Li, PeerJ 5 (2017) e3279. [365] L. Mivehi, S. Hajir Bahrami, R.M.A. Malek, J. Appl. Polym. Sci. 109 (2008) 545 554. [366] V. Petrova, D.D. Chernyakov, Y.E. Moskalenko, E.R. Gasilova, I. Strelina, O.V. Okatova, et al., Carbohydr. Polym. 157 (2017) 866 874. [367] W. Tan, Q. Li, F. Dong, L. Wei, Z. Guo, Int. J. Biol. Macromol. 92 (2016) 293 298. [368] J. Zhang, W. Tan, Z. Zhang, Y. Song, Q. Li, F. Dong, et al., Int. J. Biol. Macromol. 109 (2018) 1061 1067. [369] K. Shao, B. Han, J. Gao, F. Song, Y. Yang, W. Liu, J. Ocean Univ. China 14, 2015, 703 709 [370] A.S. Berezin, Y.A. Skorik, Carbohydr. Polym. 127 (2015) 309 315. [371] C. Sheng, B. Wenting, T. Shijian, W. Yuechuan, Polymer (Guildf) 109 (2008) 1093 1098.

Handbook of Chitin and Chitosan

C H A P T E R

6 Chitin nanomaterials: preparation and surface modifications Abul K. Mallik, Md. Nurus Sakib, Md. Shaharuzzaman, Papia Haque and Mohammed Mizanur Rahman Department of Applied Chemistry and Chemical Engineering, Faculty of Engineering and Technology, University of Dhaka, Dhaka, Bangladesh O U T L I N E 6.1 Introduction

166

6.2 Structure and properties of chitin

168

6.3 Chitin-based nanomaterials 6.3.1 Chitin nanofiber 6.3.2 Chitin nanowhisker 6.3.3 Chitin nanocomposite 6.3.4 Polymer/chitin bionanocomposite 6.3.5 Chitin nanogel 6.3.6 Crab chitin-based two-dimensional soft nanomaterials

169 170 172 174 175 176 177

6.4 Preparation of chitin-based nanomaterials 6.4.1 Electrospinning of chitin 6.4.2 Aqueous counter collision method 6.4.3 Self-assembly 6.4.4 Microcontact printing 6.4.5 Mechanical treatment

178 178 180 180 181 181

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00006-7

165

© 2020 Elsevier Inc. All rights reserved.

166

6. Chitin nanomaterials: preparation and surface modifications

6.4.6 6.4.7 6.4.8 6.4.9 6.4.10

Ultrasonication TEMPO-mediated oxidation Extraction of chitin nanowhisker Gelation method Casting and evaporating technique

182 183 184 184 185

6.5 Surface modification of chitin 6.5.1 Chemical modification of chitin surface 6.5.2 Hydrophobization of chitin surface 6.5.3 Hydrophilization of chitin surface 6.5.4 Physical modification of chitin surface 6.5.5 Ultrasound-assisted surface modification of chitin 6.5.6 Plasma treatment

185 186 187 188 188 188 188

6.6 Conclusions

189

References

189

6.1 Introduction Chitin is a very important natural polysaccharide that was first recognized in 1884. After cellulose, it is the second most abundant polysaccharide. Polysaccharides (cellulose, chitin/chitosan, starch, alginate) are a group of natural macromolecules, usually derived by various biotechnological methods from crustacean shell wastes and/or agricultural feedstock and can have very bioactive properties. The annual natural production rate of chitin has been assessed to be around 1011 tons [1,2]. Cellulose is usually present in wood cell walls, existing as cellulose nanofibers (or microfibrils) with hugely crystalline structures. Chitin is like an analogue of cellulose with a (1,4)-β-N-acetyl glycosaminoglycan repeating structure (Fig. 6.1). By deacetylation of chitin one can get the most important derivative of it, known as chitosan [3], and one of the reasons for this deacetylation is the low solubility of chitin in many common solvents (Fig. 6.1). Chitin can be converted to various nanomaterials known as chitin nanomaterials. Chitin is mostly crystalline with strong hydrogen bonding due to its linear structure with two hydroxyl groups and an acetamide group. Although, chitin has huge applicability, pure chitin has some limitations regarding functionality, properties, uniformity, durability, etc. to its use in high-performance materials. Therefore scientists have taken into account the highly crystalline regions that compose these natural fibers called nanofibrils or nanoscaled polymeric assemblies [4]. Chitin nanofibrils’ surfaces could be chemically modified to increase their

Handbook of Chitin and Chitosan

6.1 Introduction

167

FIGURE 6.1 The structures of cellulose, chitin, and chitosan.

applicability. Moreover, they are anisotropic particles with a high level of biodegradability and along their axes they show very high modulus and strength [5]. They are also called nanocrystals or nanowhiskers [6]. Chitin also can be regenerated as nanogels by a controlled regeneration method. Chitin nanogels benefit from their ligand binding capacity, functionalities, and improved biomineralization related to synthetic polymer-based nanogels [7]. Moreover, two-dimensional (2D) soft nanomaterials can be produced from crab chitin with excellent aqueous dispersibility and structural constancy during cycles of centrifugation and redispersion [8]. There are many methods/techniques used for the preparation of chitin-based nanomaterials. For example, electrospinning is one of the most important methods to prepare chitin nanofibers [5,9]. This technique could be used to prepare polymer nanofibers with a diameter in the range of micrometers to nanometers depending on the processing conditions and polymer properties. Nanofibers have many encouraging properties like distinctive dimensional, mechanical, optical, and other characteristics. They have a very high surface to volume ratio [10] and form a highly porous mesh [11], and therefore their properties are unlike from those of microsized fibers. There are many other techniques also available for the preparation of chitin nanofibers, like selfassembly, phase separation, microcontact printing, mechanical treatment, ultrasonication, TEMPO-mediated oxidation, extraction of chitin nanowhisker, sol
gel method, and controlled regeneration method [9]. High-quality chitin nanofibers can be prepared from crab shells, prawn shells, mushrooms, dry chitin powder [12], etc. Surface modification of chitin is an approach to improve and diversify the applications of chitin-based nanomaterials. The lower solubility of chitin is one of the limitations of chitin. Therefore to improve the solubility characteristics of chitin, chemical modification is possible. For example, incorporating a carboxymethyl group is the most advantageous method of increasing the solubility of chitin/chitosan at neutral and alkaline pH [13]. Other modifications are also available for chitin, such as acetylation of chitin, acetylation of chitin nanofibers, chemical

Handbook of Chitin and Chitosan

168

6. Chitin nanomaterials: preparation and surface modifications

modification of chitin nanowhiskers, and disintegrating chitin nanofibrils using calcium ion [14,15]. In the following sections, we will discuss in detail the chitin-derived nanomaterials, their preparation, and their surface modification processes.

6.2 Structure and properties of chitin The Greek word “chiton” is the origin of the name “chitin,” which means a coat of mail [16]. Chitin [β-(1
4)-N-acetyl-D-glucosamine] is a natural polysaccharide of foremost prominence [17]. Chitin (C8H13O5N)n has a close resemblance to cellulose except the monomer units (2-acetamido-2-deoxy-β-D-glucose) connect to each other through β(1-4) linkages (Fig. 6.1). It is generally a white-colored hard and inelastic nitrogenous polysaccharide that occurs in nature as semicrystalline to crystalline microfibrils that can produce complex hierarchical architectures within multiple length scales (nano to micro), from simple molecules to composites. In the exoskeletons of arthropods and fungal cell walls it forms structural elements. It is also formed by other organisms in the lower plant and animal kingdoms to increase the strength of their protection systems [17,18]. Chitin is a biomaterial which has special properties, such as nontoxicity, antimicrobial activity, biocompatibility, bioabsorbability, and low antigenicity with the capability to induce healing effects, and therefore could be applied in various biomedical fields [19]. Chitin synthesis is promoted by chitosomes through polymerization of glucosamine units. Depending on the species and crystallinity, there are three types: α-, β-, and γ-forms of chitin [20,21] 1. α-Chitin is the commonest form of chitin and exists in shrimps and crabs. It is also the most stable one, where the polymer units are arranged antiparallel (like cellulose type II) and allow the polymer chains to be connected by hydrogen bonds with maximum molecular aspect ratio [22]. Both intramolecular and intermolecular hydrogen bonds are possible and therefore it forms quite stable structures and microfibrils are self-assembled in a highly crystalline orientation (higher than 80%) [23]. 2. β-Chitin is a less common form of chitin and usually is found in squids. It has its molecular units aligned parallel (similar to cellulose type I) which form intramolecular hydrogen bonding instead of intermolecular hydrogen bonding. It is more flexible than the antiparallel arrangement in α-chitin, but it has sufficient strength [24]. The crystallinity of the fibrils are less than in α-chitin (about 70%). It is more reactive and susceptible to dissolving in solvents.

Handbook of Chitin and Chitosan

6.3 Chitin-based nanomaterials

169

3. γ-Chitin is a mixture of α- and β-chitins with an orientation of parallel and antiparallel chains and found in mushrooms [25,26]. In nature, α-chitin is more plentiful compared with squid pen β-chitin and γ-chitin. Commercially α-chitin is also more readily available [27]. The aggregation of 18
25 native chitin chains can form fibrils with a rod-like or spindle-like morphology with a length to diameter ratio of approximately 300 to 2
5 nm, as shown in Fig. 6.2. Acid hydrolyzes of chitin can produce highly crystalline chitin, by dissolving the amorphous region, that is often called chitin nanofibrils, chitin whiskers, or chitin nanocrystals. Chitin can be changed to various derivatives, which will be discussed in the later sections.

6.3 Chitin-based nanomaterials Chitin is a biomaterial often found in the exoskeletons of crabs, shrimps, and insects. It is the second most abundant after cellulose and are produced in more than 10 billion tons each year [29]. Although most often chitin is discarded as waste, it has huge potential as a flexible biopolymer due to the reactive groups present in the chitin structure. In recent years, chitin-based biomaterials have been often used in lieu of

FIGURE 6.2 Schematic illustration of the exoskeleton structure of crab shells. Source: Reproduced from S. Ifuku, H. Saimoto, Chitin nanofibers: preparations, modifications, and applications, Nanoscale 4 (2012) 3308
3318 [28] by permission of The Royal Society of Chemistry.

Handbook of Chitin and Chitosan

170

6. Chitin nanomaterials: preparation and surface modifications

plastics because of their biodegradability and biocompatibility. This section will highlight the different types of biomaterials, along with their applications, that are being produced with chitin or its derivatives. Chitin can be converted to various nanomaterials with special properties, especially for the use in biomedical applications. Following are the chitin-based nanomaterials commonly prepared.

6.3.1 Chitin nanofiber Chitin nanofibers are crystalline in nature and have been found in nature in a hierarchical assembly. The nanofibers produced from chitin have high mechanical properties such as stiffness and strength. Fibers from chitin can be produced in different ways, such as the highpressure homogenization route [30], and the synthesis process varies with the source. 6.3.1.1 Pure chitin nanofiber Pure chitin nanofibers are prepared from crab shells, prawn shells [29,31,32], etc. Pure chitin from crab shells is prepared via various chemical treatments. The shells are first demineralized using HCl and then treated with NaOH to remove proteins. Then the pigments and lipids are removed by means of ethanol. The degree of N-acetylation is very high and in this way pure chitin fiber is synthesized from crab shells. Chitins from prawn shells have been also similarly derived. The demineralization and removal of proteins were done according to the usual method; using HCl and NaOH. After this chemical treatment, the fibrillation process is somewhat mechanical, such as grinding the solution to form nanofibers [33] Chitin nanofibers from prawn shells show approximate uniform width when the same methods of extraction are used. However, for the crab shells, the width size varies even if the methods applied are the same. The crab shell chitin nanofibers have a width range of about 10
100 nm [34]. Furthermore, chitin from prawn shells can be extracted in a neutral pH medium, whereas crab shells chitin extraction is done at a lower pH. This phenomenon has been explained by Ifuku et al. who have stated that crab shells consist of more endocuticles which are coarser and thicker in nature, whereas for prawn shells the amount of fine exocuticle is higher in comparison with the crab shells. Due to this physical difference, prawn chitin nanofibers can be produced in neutral pH conditions [33]. Pure chitin nanofibers have many applications such as sutures, gas barriers, etc. Wu et al. have prepared a translucent gas barrier from the purified chitin derived from crab shells. The prepared nanofiber fiber

Handbook of Chitin and Chitosan

6.3 Chitin-based nanomaterials

171

barriers have the lowest oxygen and carbon dioxide permeability. In the process, the crab shells were dried and ground and then refluxed. After further treatment and removal of proteins and other residues, nanofibers were prepared with homogenization and drying. The pure chitin nanofiber film showed good mechanical characteristics and exceeded the commercially used PET as a barrier terms of oxygen and carbon dioxide barrier properties [30]. Chitin nanofibers have also been tested against inflammatory bowel disease (IBD). The study shows chitin nanofibrils showed antiinflammatory effects and a reduction of disease activity index when compared to other chitin suspensions. In the experiment, chitin nanofibrils and chitin suspensions were administered orally to mice with IBD and observed for several days. Mouse that were administered with chitin fibers showed significantly lower disease activity index on days 4
6. A study showed that the colon weight to length ratio was less than in other study groups. The same group showed lower rate of erosions, crypt destruction, and edema. In addition, tissue damage was markedly lower in the nanofibrils group on day 6 [12]. A proof of concept presented by [35] shows that chitin nanofibers can be used as a tissue engineering substrate. In the experiment, replicas of PDMS were used to make different microscale substrates. Both supported and freestanding substrates were prepared and studies found that both types are stable and biocompatible. This shows potential for preparing substrates that can be used in tissue engineering. 6.3.1.2 Chitin-based blended nanofiber Generally, polymer blends are used to enhance the spinnability in nanofiber production. Park et al. have prepared biodegradable and biomimetic nanostructured scaffolds. The poly(glycolic acid) (PGA) and chitin-blended nanofibers were produced via an electrospinning method. The scaffold with 25% PGA and 75% chitin showed great potential in cell adhesion and proliferation [36]. 6.3.1.3 Modified chitin nanofiber Often the nanofibers are chemically modified. In the most common cases, surface modifications take place changing their physical and chemical characteristics. This is done by introducing functional groups to the hydroxyl groups of chitin. Acetylation is the simple modification of surfaces carried out on chitin nanofibers. Other chemical modifications that can be done on chitin nanofibers are the following: (1) partial deacetylation; (2) surface maleylation, phthaloylation, and napthaloylation; (3) surface N-halamine modification; and (4) graft polymerization onto chitin nanofibers [37].

Handbook of Chitin and Chitosan

172

6. Chitin nanomaterials: preparation and surface modifications

6.3.1.4 Nanofiber from chitin derivatives Chitosan is the deacetylated form of chitin. Chitosan can be dissolved in a wide array of solvents, unlike chitin. Hence, chitosan has many variants of nanofibers, nanocomposites, nanogels, etc. In general, chitosan nanofibers are generally difficult to produce using the electrospinning technique since the surface tension of the chitosan solution is relatively very high [37]. Applications of chitin-derived chitosan nanofibers include filtration, enzyme immobilization, and wound healing [38]. For example, chitosan nanofibers produced by electrospinning by means of trifluoroacetic acid show the adsorption of lead and copper. Nanofiber mats around 235 nm in diameter showed great affinity for the heavy metals [39]. Zhou et al. have prepared carboxyethyl chitosan and PVA nanofiber which is biocompatible and can be used as a scaffolding material for fibroblast cells. The nanofibrous mats showed good cell adhesion and growth [40].

6.3.2 Chitin nanowhisker Nanowhiskers of chitin have earned a special attention in preparing polymeric matrices, especially as fillers. The crystalline structure of the chitin is broken down to its nanofragments and resulting rod-like nanosized structures are termed as nanowhiskers. Such rod-shaped structures are able to form a hard structure by forming in line next to one another. The nanowhiskers are mostly used as polymer or composite reinforcing additives [14]. Furthermore, they have found their use in the synthesis of nanocomposite materials such as bionanocomposites and drug delivery, as well as in tissue engineering, because of their high surface area, high adsorption ability, biodegradability, and nontoxicity [41]. Chitin’s structure is similar to the structure of cellulose. It exists as crystalline fibrils and composite, hence nanowhiskers can be synthesized from chitin similar to cellulose. The chitin whisker preparation is also similar to the cellulose whisker preparation process. In the process the chitins are purified and then acid-treated under reflux condition. The treated solution is then decanted and the suspension shows a spontaneous dispersion of whisker-shaped particles [42]. The hydrolysis depends on parameters such as the concentration of the acids and consequently the hydrolytic extent that the chitin undergoes. Breaking down the process, the acid dissolves the amorphous phases faster because the chitin crystalline domains have tighter bonds and are difficult to be hydrolyzed. However, after the amorphous domains are removed, excess acids start to hydrolyze the surface of the chitin crystallites. This part is important as the extent of hydrolysis determines the

Handbook of Chitin and Chitosan

6.3 Chitin-based nanomaterials

173

yield and size of the nanocrystals [42]. Li et al. have prepared the chitin whiskers by hydrolyzing the chitin with 3 M HCl and then centrifuging the suspension, followed by extensive removal of the supernatant by decantation. The resulting size of the rods is around 200 6 20 nm in length [43]. Chitin nanowhiskers can be prepared from chitins originating from different sources such as squid pen, crab shell, shrimp shell, and Riftia tubes [6]. A common and advantageous method of preparation of chitin nanowhiskers is the TEMPO-mediated oxidation method which is discussed in a later part of this book. Fan et al. have devised a surface cationization method where the nanowhiskers are synthesized from partially deacetylated chitins. In this process, the surface is charged with cations and the individualization occurs because of the electrical repulsion [44]. There are many examples of chitin nanowhiskers as fillers since they add mechanical strength to composites. Reinforcing a composite requires consideration of several parameters such as the aspect ratio, glass transition temperature of the matrix, and the path of synthesis. In addition, surface modification can enhance the mechanical property as well. In an experiment, crab shell-derived nanowhiskers underwent surface modification with succinic anhydride (ASA), phenyl isocyanate (PI), and isopropenyl-α,α0 -dimethylbenzyl isocyanate (TMI). Then they were added to the natural rubber matrix. The results show that nanowhiskers and rubber interact with each other more after the surface modification. But interestingly the surface-modified nanowhiskers/rubber composite showed lower mechanical properties and this was attributed to the absence of the whisker network formation [45]. Wongpanit et al. have added nanowhiskers with silk fibroin prepared from silkworm. The addition of whiskers has improved the stability and compression strength [46]. Another use of chitin nanowhiskers as fillers was with acrylic polymer matrix. The chitin source for this experiment was squid pen. Aqueous suspensions of both nanowhiskers and polymer were mixed to obtain nanocomposite films with improved mechanical and thermal properties [47]. Kadokawa et al. have prepared chitin nanowhiskers using ionic liquid and further prepared a composite of chitin nanowhiskers and poly (vinyl alcohol). The researchers soaked the chitin in the ionic liquid at room temperature. Then the soaked chitin was heated and further soaked in methanol. After sonication, the chitin nanowhiskers were prepared. The composite showed a good mechanical property when the ratio of PVA to chitin was increased [48]. Chitin nanowhisker composites with synthetic polymers show special optical properties. Nge et al. incorporated chitin whiskers in poly (acrylic acid) and prepared a composite that showed uniplanar orientation and the long axes of molecules were perpendicular to magnetic

Handbook of Chitin and Chitosan

174

6. Chitin nanomaterials: preparation and surface modifications

field [49]. Ifuku et al. have also prepared optically transparent nanocomposites made from nanowhiskers and acrylic resin with optical losses smaller than 2% [50]. Chitin nanowhiskers can be turned into highly porous, thermally stable mesoporous aerogels with low densities. At first the whiskers are sonicated and formed into hydrogels. After ethanol solvent exchange and drying, the aerogels are prepared. The aerogels were crystalline like the nanowhiskers, showed good mechanical characteristics, and had the minimum shrinkage when drying [51]. Chitin nanowhiskers have also found their applications in biomedical materials. A process of preparing chitosan nanoscaffold was devised by Phongying et al. from chitin nanowhiskers. In the process, the chitin nanowhiskers are deacetylated in the presence of a strong alkaline solution and the resulting solution contained chitosan. The scaffolds produce a fibrous nanoporous arrangement with pore diameter of around 200 nm [52]. Because of their biocompatibility, biodegradability, and nontoxicity, chitosan nanowhiskers and the derivatives of the nanowhiskers have huge potential in the fields of biomedicine and food. The only thing that holds back the chitin nanowhiskers applications at the large scale is their production cost. Hence it is imperative to come up with a costeffective production method for chitin and its derivatives and the smart synthesis of high-value products.

6.3.3 Chitin nanocomposite Nanocomposites have seen a wide range of applications from biomedicine to waste treatment. In recent years, nanocomposites have been generally synthesized using materials that are biologically compatible and environment-friendly, since synthetic components that are used may pose threats to us and to the environment, even though they can be synthesized using comparatively inexpensive methods. Similar to nanowhiskers, chitin nanocomposites’ chitin part is mostly used for reinforcements. Novel biomaterials with totally different characteristics are often observed in the nanocomposites. The characteristics that chitin bring about in nanocomposites are mostly in terms of physical, mechanical, optical, and sometimes electrical properties [53]. Marcin and colleagues have developed a chitin nanocomposite with a photoluminescent property. In this research, they have used a marine sponge skeleton as a template and this process is also known as biomimetics. In the experiment, germanium(IV) ethoxide is used as a precursor. The germanium
chitin composite showed enhanced photoluminescence compared with other germanium oxide
based materials [54].

Handbook of Chitin and Chitosan

6.3 Chitin-based nanomaterials

175

Shams et al. have prepared a similar optically transparent nanocomposite from chitin and resin. The nanofibers were able to hold the transparency of resin due to their small size. The researchers could lower the time required for vacuum filtering the chitin suspension compared with the time usually required for cellulose suspension. The chitin acrylic resin composite is optically more transparent than the cellulose one due to the affinity formed between less hydrophilic chitin with less hydrophobic resin. The addition of chitin give the composite characteristics such as enhanced thermal expansion and mechanical properties. Its potential application fields include optoelectric devices and solar cells [55]. Another nanocomposite was made with chitin nanowhisker dispersed in the matrix of poly(caprolactone). The chitin nanowhiskers were prepared from Riftia tubes and poly(styrene-co-butyl acrylate) was used as a model matrix. The composite shows higher thermal stabilization over a large temperature range [56]. Another example of chitin nanocomposite is the use of chitin nanofibrils with the polylactic acid. In general, the extrusion process is disadvantageous because the nanofibrils tend to aggregate and it is difficult to control the extrusion process. Hence a precomposite of poly(ethylene glycol) is prepared and then added in the extruder machine. The results show that the addition of up to 12% chitin does not change the thermal and morphological properties and therefore chitin fibrils can be used as reinforcing fillers and be applied in areas such as bioplastics due to chitin’s bioconformity [57]. In most of cases, chitins are used as reinforcement for the composites. However, other examples of chitin nanocomposites are available. For example, chitin can be used in composites to enhance the biocompatibility. Such examples are given in the next section.

6.3.4 Polymer/chitin bionanocomposite Chitin bionanocomposites have been getting extensive attention recently because of their biocompatibility and lower toxicity. Singh and coworkers have prepared a bionanocomposite from chitin and carbon nanotubes where the composite has incorporated the biocompatible properties of chitin. The mixing of carbon nanotubes and chitin was done in imidazolium-based ionic liquid. The nanocomposite was used for stem cell growth, and it has been observed that the cell adhesion and proliferation occurred in different ratios. The scaffold for the stem cell growth is biocompatible, electrically conductive, and suitable for mesenchymal cells [58]. Another example of a polymer bionanocomposite involved the use of chitin nanowhiskers as reinforcing agents. The polysaccharides of a red

Handbook of Chitin and Chitosan

176

6. Chitin nanomaterials: preparation and surface modifications

algae-based composite showed better mechanical property and has been stated to be a more environment-friendly composite. The mechanical properties of the chitin nanocomposite improved with the increase in chitin percentage [59]. Another chitin-based bionanocomposite was prepared using bentonite nanoclay and chitin on polyurethane. The composite showed a cation exchange capacity of 74 meq/100 g. The layers of nanoclay bentonite dispersed in an ordered intercalated fashion in the polyurethane matrix. The study also shows that the morphology formation mechanism is different for this composite than the usual polymer solution. The whole composite-making process was carried out by the emulsion polymerization method [60].

6.3.5 Chitin nanogel Nanogels are nanosized hydrogels. Hydrogels are known for their water retention capabilities and are an effective source of drug release. Chitins are biocompatible and hence chitin nanogels have found application in drug delivery and nanotherapeutics. Drugs or molecules are encapsulated within the nanogels and when given an outer stimulus the drug molecules are released. Use of biobased carriers such as chitin and chitosan in drug delivery are increasing day-by-day, replacing the synthetic polymers [61]. Nanogels as drug carriers have an added advantage since they help the drugs to be channeled through intracellular space and thus it becomes easier for the cells to absorb them. Furthermore, they are more hydrophilic and hence are safe from the phagocytic action [62]. Zhang et al. have showed a facile way to prepare chitin nanogels straight from the chitin solution. In the process, the chitin was dissolved in NaOH and urea and the resulting chitin solution is then subjected to vigorous stirring. The heat from the stirring lowered chitin chain regeneration and the nanogels were well dispersed. The prepared nanogels have spherical shapes and showed biocompatibility when tested against L929 cells [63]. Nanogels have been used as a drug delivery medium through the transdermal route. Sabitha et al. have prepared a chitin nanogel loaded with curcumin to act as an anticancer drug. The prepared nanogels show that they have toxicity toward the human melanoma cells but are somewhat compatible with the human fibroblast cells. The curcumin was delivered transdermally and showed four times more curcumin flow than the control curcumin solution. Furthermore, the porcine skin that was treated with the nanogels did not show inflammatory signs. The nanogels showed pH dependency on the release rate. At acidic pH

Handbook of Chitin and Chitosan

6.3 Chitin-based nanomaterials

177

FIGURE 6.3 Chitin nanogels in biomedical applications. Source: Reprinted from M. Vishnu Priya, M. Sabitha, R. Jayakumar, Colloidal chitin nanogels: a plethora of applications under one shell, Carbohydrate Polym. 136 (2016) 609
617 [65] with permission from Elsevier.

the release rate was higher than that of neutral pH. Hence the nanogels can be used for the treatment of melanoma through transdermal route [64] (Fig. 6.3). Development of antipsoriatic drugs (acitretin and aloe-emodin)loaded chitin nanogels have given a good result against psoriasis when applied transdermally. Similar to the previous nanogel, this nanogel is also pH dependent and showed higher swelling at acidic pH. The nanogels are pseudoplastic in nature and show non-Fickian release pattern. The antipsoriatic drugs aggregate highly at deeper layers of skin. The nanogel is biocompatible and hence shows good candidacy for the psoriasis treatment [66]. Dexorubicin is a hydrophilic drug used for human lung cancer treatment. A nanogel of chitin
poly(caprolactone) has been prepared for the delivery of doxorubicin anticancer drug, which showed cyctotoxicity toward adenocarcinomic human alveolar basal epithelial cells. For the treatment of lung cancer, the nanogel successfully released the drug and was found to be pH dependent. At lower pH, that is, acidic condition, the release rate and higher swelling along with degradation was observed. The spherical nanogel was shown to be biocompatible and can serve as a promising drug carrier [67].

6.3.6 Crab chitin-based two-dimensional soft nanomaterials Two-dimensional materials such as graphene, carbon nitride, etc. have been in the research spotlight due to their extensive application fields (optics, sensing, energy, electronics, etc.). 2D materials are

Handbook of Chitin and Chitosan

178

6. Chitin nanomaterials: preparation and surface modifications

different in dimensions and their optical and electrical properties differ due to electron confinement. Furthermore, their high surface to bulk ratio is an equally important characteristic [68]. Even though 2D materials have vast application fields their use is rather limited due to the high synthesis cost, cytotoxicity, and nondegradability [69,70]. So continuous research is ongoing to make the 2D production cost-effective as well as biocompatible. Using crab chitin-based 2D soft materials help to make the process sustainable and biocompatible. A study shows the preparation of nanosheets from marine source-derived chitin. You et al. have prepared crab chitin-based 2D material by means of hydrophobization-induced interfacial assembly method. Two layers of chitin formed the nanosheets. When emulsified and carbonized jointly, the sheets can form carbon nanosheets that have a minimum thickness of 3.8 nm. Such hybrid films offer inexpensive and environmentfriendly approaches to make 2D materials with applications in electronics and biological devices [8].

6.4 Preparation of chitin-based nanomaterials Chitin nanofibers have been prepared mainly from crab shells, squid pens, prawn shells, tubes of Riftia pachyptila worms, and shrimp shells. Different types of synthetic [poly(ethylene oxide) (PEO), poly(vinyl alcohol) (PVA), poly(L-lactide) (PLA), poly(glycolic acid) (PGA), and polyvinylpyrrolidone (PVP)] as well as natural polymers (silk, cellulose, collagen, alginate, zein, and agarose) have been blended with chitin fibers to form chitin-based nanomaterials. There are many methods such as electrospinning, printing, self-assembly, phase separation, and template synthesis to be used for the preparation of chitin-based nanomaterials. In addition some new methods are also available, including microcontact printing, simple mechanical treatment, ultrasonication, and 2,2,6,6-tetra-methylpiperidinooxy (TEMPO)-mediated oxidization, for the preparation of chitin-based nanomaterials.

6.4.1 Electrospinning of chitin Electrospinning, namely utilizing electrostatic forces is one of the most important methods to prepare chitin nanofibers for more than six decades [71]. It is a technique from which polymer nanofibers can be produced. The range of diameter of the polymer is from several micrometers down to tens of nanometers, depending on the polymer and processing conditions. Electrospinning still attracts interest for their tremendous advantages such as low cost, uniform nanofibers, inexpensive

Handbook of Chitin and Chitosan

6.4 Preparation of chitin-based nanomaterials

179

purification, and continuous nanofibers. There are many parameters that influence the electrospinning process: (1) system parameters, such as the types of the polymer and properties of the polymer solution (conductivity, viscosity, and surface tension); (2) process parameters, such as electric potential, the distance between the tip and the collector, flow rate, and concentration; and (3) ambient parameters, such as humidity, solution temperature, and air velocity in the chamber. The schematic diagram of the polymer nanofiber fabrication process using the electrospinning method is shown in Fig. 6.4. Min et al. [73] reported an electrospinning method to fabricate chitin nanofibers. To improve the solubility of chitin, the chitin powder (100
500 nm) was depolymerized and packed in polyethylene bags and then 60Co gamma-irradiated for 3 days. Chitin solutions with concentrations in the range from 3%
6% were obtained. 1,1,1,3,3,3-hexafluoro-2propanol (HFIP) was used as a spinning solvent for electrospinning of chitin. The electrospun chitin nanowhiskers were collected on a target drum. The drum was placed at a distance of B7 cm from the syringe tip and a voltage of 15 kV was applied by using a high-voltage power supply. The average diameter of chitin nanofibers was found to be 110 nm. Electrospinning is the most frequently used method for the preparation of chitin-based nanomaterials. Many attempts have been made in the last few decades to prepare chitin-based nanomaterials. In order to

FIGURE 6.4 Schematic diagram of polymer nanofibers formation using the electrospinning method. Source: Reprinted from H.S. Yoo, T.G. Kim, T.G. Park, Surface-functionalized electrospun nanofibers for tissue engineering and drug delivery, Adv. Drug Deliv. Rev. 61 (2009) 1033
1042 [72] with permission from Elsevier.

Handbook of Chitin and Chitosan

180

6. Chitin nanomaterials: preparation and surface modifications

improve the nanofiber formation capacity of chitin, the chitin nanofibers are always combined with other substances such as synthetic polymers, biopolymers, and inorganic particles. Chitin-based binary blend nanofibers are normally fabricated using HFIP as solvent. Park et al. reported the preparation of chitin/PGA blend nanofibers by using HFIP as the solvent via the electrospinning method [74]. The resulting blend nanofibers had an average diameter of 140 nm. Park et al. also reported chitin/silk fibroin nanofibers using the electrospinning method. The average diameters of chitin/silk blend fibers decreased from 920 to 340 nm with the increase of chitin content in blend compositions. Recently, chitin-based ternary composite nanofiber scaffolds are produced by blending nanofibers with synthetic polymers, biopolymers, and inorganic substances [75,76]. Chitin derivatives such as carboxymethyl chitin and dibutyrylchitin have also been blended with other substances to form nanofibers [77,78].

6.4.2 Aqueous counter collision method Kose and Kondo [79] reported an aqueous counter collision (ACC) method for the preparation of chitin nanofibers in an aqueous dispersion state. In the ACC system liquid suspension of the sample is ejected from a pair of nozzles under a high pressure of 200 Mpa which forms a pair of jets. Though chitin fibers are not soluble in water, they can be dispersed in water by the ACC method. The number of ejecting steps and ejecting pressure is fixed to focus the sample to an appropriate degree of pulverization. Phase separation of chitin powder was done before the ACC treatment. The chitin samples became turbid after the ACC treatment at 0, 1, 5, 10, 30, 60, and 120 ejecting steps, respectively. The sample is supposed to be more downsized by changing the number of ejecting steps and the desired pressure. Polarizing light microscopy was used to observe chitin particles having microsize diameter. After chitin samples obtained by ACC treatment, the chitin nanofibers exhibited a favorable aggregation as a three-dimensional network formation.

6.4.3 Self-assembly Molecular self-assembly is an influential technique to fabricate nanomaterials mainly from biomolecules including peptides and proteins. It is facilitated by notable hydrogen bonds, weak noncovalent bonds, ionic bonds, van der Waals interactions, and hydrophobic interactions [80]. Chitin nanofibers with diameters of 3 nm have been fabricated by a facile self-assembly method using HFIP solvent [81
83]. Fig. 6.5 shows the

Handbook of Chitin and Chitosan

6.4 Preparation of chitin-based nanomaterials

181

FIGURE 6.5 (A) Schematic of chitin nanofiber formation using the self-assembly method and (B) Topographic AFM image of the chitin nanofibers. Source: Reproduce from F. Ding, H. Deng, Y. Du, X. Shi, Q. Wang, Emerging chitin and chitosan nanofibrous materials for biomedical applications, Nanoscale 6 (2014) 9477
9493 by permission of The Royal Society of Chemistry.

schematic of chitin nanofiber formation using the self-assembly method which was initiated by the evaporation of HFIP. The diameter of the chitin nanofiber is independent of the concentration of the solution and was found to be 3 nm. Though HFIP is a toxic solvent, it is still used to prepare chitin nanofibers using the self-assembly method. However, it is better to use green solvents such as ionic liquids and urea
NaOH mixture to develop chitin nanofibers [9].

6.4.4 Microcontact printing Microcontact printing is a great technique for the fabrication of nanostructured macromolecules, such as dendrimers, peptides, and conducting polymers [84,85]. Chitin is dissolved in HFIP to prepare chitin nanofibers using the microcontact printing method. The high evaporation rate of HFIP allowed the formation of chitin nanofibers with a width of 30 nm and a height of 20 nm. The procedure of microcontact printing is shown in Fig. 6.6. Chitin nanofibers prepared by the microcontact printing method have the following advantages: 1. 2D and 3D chitin nanofibers can be manufactured ranging from micrometers to sub-50 nm in one step. 2. the condition used is very simple and no heating, vacuum, or ultrasonication is required. 3. it can be combined with other fabrication technologies to produce more complex structures.

6.4.5 Mechanical treatment Mechanical treatment is known as a “green” method because there is no involvement of toxic organic solvents during fabrication. Recently,

Handbook of Chitin and Chitosan

182

6. Chitin nanomaterials: preparation and surface modifications

FIGURE 6.6 (A) Process of chitin nanofibers formation using the micro- contact printing method, (B) AFM image of chitin nanofibers prepared from 0.05% (w/v) chitin solution. Scale bar: 4 mm, (C) AFM image of two chitin nanofibers. Scale bar 1/4 400 nm. Source: Reproduced from F. Ding, H. Deng, Y. Du, X. Shi, Q. Wang, Emerging chitin and chitosan nanofibrous materials for biomedical applications, Nanoscale 6 (2014) 9477
9493 by permission of The Royal Society of Chemistry.

many attempts have been made to prepare chitin nanofibers through mechanical treatment [28,86]. The crab shell is made up of an aggregation of chitin nanofibers. The aggregated chitin nanofibers can be disintegrated by using a grinder. First proteins and minerals were removed from chitin by treating chitin with NaOH and HCl. Then the purified chitin with 1 wt.% concentration in water was passed through a specially designed grinder. After the grinder treatment, the chitin slurry formed a gel that suggested the accomplishment of the disintegration. Highly uniform chitin nanofibers with a width of 10
20 nm were successfully obtained through this process. The key step in the mechanical treatment process is the disaggregation of original chitin nanofibers under acidic conditions. This simple method offers a way to obtain chitin nanofibers in large amounts.

6.4.6 Ultrasonication Ultrasonication is a powerful approach that has been widely used to prepare individualized chitin nanofibers. This method is simple because no chemical modification is required during the sonication process. Ultrasonication produces individualized chitin nanofibers with a width of 3
4 nm in cross section and at least a few microns in length [87]. The optimal conditions for the formation of chitin nanofibers were as follows: (1) β
chitin was used as raw material; (2) chitin was dispersed in

Handbook of Chitin and Chitosan

6.4 Preparation of chitin-based nanomaterials

183

water with a pH of 3
4; (3) the concentration was 0.1%
0.3%; and (4) the time was 2 min. It was reported that the most important factor for preparing the nanofibers via ultrasonication was the protonation of amino groups under acidic conditions. Zhao et al. [88] developed an ultrasonication technique which is a simple, versatile and environment-friendly approach for extracting bionanofibers from various natural and synthetic materials. Among bionanofibers, chitin nanofibers have been fabricated from various materials such as chitin fibers, spider and silkworm silks, collagen, cotton, bamboo, and ramie and hemp fibers. The diameter is very uniform, in the range of 25
120 nm, which is a useful size range for many tissue engineering and filtration applications. The ultrasonic shock waves cause erosion of the surface of the fibers that gradually disintegrate the micron-sized natural fibers into nanowhiskers, splitting along the axial direction. The speed of disassembly depends on the intensity and frequency of the ultrasonic wave. Recently, Lu et al. reported the fabrication of chitin nanofibers via a simple high-intensity ultrasonication treatment under neutral conditions (60 kHz, 300 W, pH 1/4 7) using α-chitin as the raw material [89]. Chitin nanofibers with diameters of 20
200 nm were successfully prepared by adjusting the ultrasonication time for 30 min.

6.4.7 TEMPO-mediated oxidation Chitin nanomaterials were prepared by 2,2,6,6-tetramethylpiperidine-1-oxyl radical (TEMPO)-mediated oxidation of α-chitin in water at pH 10. NaClO was added as a cooxidant in the reaction. Chitin was transformed into water-soluble polyuronic acid and waterinsoluble chitin nanowhiskers. The crystallinity of TEMPO-oxidized chitin is as high as that of the original α-chitin. In the TEMPO oxidized chitin, carboxylate content or the amount of NaClO are the significant factors that affect the transparency of the dispersions, the resultant weight ratio of water-insoluble fractions, shape, length, and width of the chitin nanofibers. The addition of 5.0 mmol of NaClO per gram of chitin seems to be optimal for the preparation of mostly individualized nanofibers, with fiber widths smaller than 15 nm, and average widths of 8 nm. Fan et al. [90] reported the preparation of chitin nanofibers from squid pen β-chitin by TEMPO-mediated oxidation of native chitins followed by mild mechanical agitation in water at pH 3
4.8. Chitin nanofibers prepared by the TEMPO-mediated oxidation method were not continuous nanofibers. For that reason, the chitin nanofibers produced in this method can be used as injectable nanomaterials for controlled drug release.

Handbook of Chitin and Chitosan

184

6. Chitin nanomaterials: preparation and surface modifications

6.4.8 Extraction of chitin nanowhisker The extraction of chitin nanowhiskers from different raw sources have been studied by many researchers [29,56,90,91]. Morin et al. [56] reported the synthesis of chitin nanowhiskers. First, the commercial raw chitin was dispersed into a 5 wt.% KOH aqueous solution, and then boiled for 6 h with mechanical stirring to remove most of the proteins. The resultant suspension was kept at ambient temperature overnight under continuous mechanical stirring. The product was filtered and washed with distilled water several times. Subsequently, 17 g of NaClO2 in 1 L of water containing 0.3 M sodium acetate buffer was used to bleach the crude product for 6 h at 80 C, and then fully rinsed with distilled water. Finally, residual proteins were removed by dispersing the crude product into a 5 wt.% KOH aqueous solution for 48 h followed by centrifugation to produce proteinfree chitin. The resulting suspension was centrifuged at 3000 rpm for 20 min. Chitin whisker suspensions were prepared by hydrolyzing the purified chitin sample with boiling solution of 3 N HCl for 1.5 h under stirring in the ratio of 30 mL/g. After acid hydrolysis, the suspensions were diluted with distilled water followed by centrifugation (10,000 trs/min for 5 min). This process was repeated three times. Next, the suspensions were transferred to a dialysis bag and dialyzed for 24 h against distilled water. The pH of the suspension was 6 and subsequently changed to 3.5 by adding HCl. The dispersion of whiskers was done by ultrasonication using B12 Branson sonifier for 2.5 min for every 40 mL aliquot. Chitin nanowhiskers from Riftia were also prepared by Morin and Dufresne [56] with the diameter of 18 nm and length around 120 nm. In another study, Gopalan and Dufresne [29] reported successful extraction of chitin nanowhiskers from crab shell with the length of 100
600 nm and width 4
40 nm. Moreover, Zhang and his coworkers [90] prepared spindle-shaped chitin nanowhiskers from crab shell with a broad distribution in length ranging from 100 to 650 nm and a diameter ranging from 10 to 80 nm. Ifuku et al. [50,86] used mechanical disassembly to prepare highly uniform chitin nanowhiskers.

6.4.9 Gelation method Chitin nanofibers were easily prepared by the gelation of commercial chitin powder with ionic liquid 1-allyl-3-methylimidazolium bromide (AMIMBr) at room temperature, followed by heating at 100 C. The resulting gel was soaked in methanol and subsequent sonication gave chitin dispersion [48]. This processing technique for the preparation of the chitin nanofibers is considered to have great advantages because special equipment and chemical modifications are not necessary for this method.

Handbook of Chitin and Chitosan

6.5 Surface modification of chitin

185

6.4.10 Casting and evaporating technique Casting and evaporation is an important technique which has an influence on the final properties of nanocomposites. The technique describes the intrinsic properties of CNWs, the nature of polymer matrix, and the desired final properties of the composites. To prepare high-performance polymer/CNW nanocomposites, good dispersibility of CWs in polymer matrix is the prerequisite. CNWs are homogenously dispersed in water to confirm uniform composition and are usually obtained as aqueous suspensions. Though water is the best standard for preparation of CNWs-reinforced nanocomposites, most of the investigators preferred water-soluble, water-dispersible, and latex-form polymers as the polymer matrixes for making nanocomposites. In this process, first the polymer aqueous solution or dispersion was mixed with CNW aqueous suspension to obtain homogenous dispersion. Then the dispersion was cast onto a container. And finally the evaporation of water results in the nanocomposites. For that reason, this technique is renamed as the casting and evaporating technique. Most of the recent reported polymer/CNW nanocomposites were prepared by this technique. The reported polymer matrixes contain poly(styrene-co-butyl acrylate), poly(caprolactone), natural rubber, soy protein isolate, poly (vinyl alcohol), chitosan, silk fibroin, alginate, starch, hyaluronan
gelatin, and waterborne polyurethane [6,56,92,93].

6.5 Surface modification of chitin There are multiple methods of surface treatment of polysaccharides, each with their own pros and cons. The arena covers the following known modification procedures: chemical modification, enzymatic modification, radiation-induced modification, ultrasonic modification, plasma treatment, laser treatment, etc. However, all the processes of surface modifications are not suitable for chitin and have not been tested with chitin so far. The surface energy of chitin is very high due to stronger hydrogen bonds between the OH and NH groups and between the NH and the carbonyl moieties, hence, it shows low solubility in water. Chitin, similar to other polysaccharides, may undergo many of the typical reactions of hydroxyl groups, like esterification, etherification, urethane formation, cross-linking with polyfunctional reagents, and graft copolymerization. The water contact angle of chitin (50 degrees) has been altered to 62
84 degrees after several modification processes [94]. The increased hydrophobicity, however, does not impart mechanical enhancement.

Handbook of Chitin and Chitosan

186

6. Chitin nanomaterials: preparation and surface modifications

Surface modification of chitin has made it possible to adjust some of its physical properties (solubility, viscosity, molecular weight, etc.) and chemical properties for different applications.

6.5.1 Chemical modification of chitin surface 6.5.1.1 Acetylation of chitin Chitin nanofibers were dispersed in water at a 0.1 wt.% to produce a nanofibers sheet of 60 μm thickness after filtering the dispersion with a hydrophilic polytetrafluoroethylene filter membrane. The sheets were cut into rectangle size with a dimension of 3 3 4 cm and dried in an oven at 65 C overnight. The sheets were placed in a Petri dish containing a mixture of 5.0 mL of acetic anhydride and 0.1 mL of 60% perchloric acid. The mixture was stirred for the desired time at room temperature. After acetylation, the chitin sheets were washed by Soxhlet extraction with methanol overnight [95]. In another method, acylation was achieved by using an acylation mixture of HClO4 and butyric acid (BA) at (217) C with chitin powder on ice at a molar ratio of 1:10:1 (chitin: BA:HClO4). The reaction was continued for 30 min and was then kept at room temperature and terminated after 3 h by adding diethyl ether. The precipitate was then collected, washed, and neutralized [96]. Water-soluble chitin was made by dissolving 4 g of chitosan in 2.8% (v/v) aqueous acetic acid. 100 mL ethanol and 32 mL pyridine was added in turn and the mixture was stirred until transparent. A given amount of anhydrous acetic acid was added and the mixture was stirred vigorously for 2 h. Then the mixture was poured into ethanol and filtrated to obtain a white solid. The crude product was dissolved in deionized water, and precipitated out by adding ethanol for purification. Finally, the product was filtered, rinsed thrice with acetone and anhydrous ether, and vacuum-dried at room temperature for 24 h [97]. 6.5.1.2 Surface modification of chitin nanowhiskers Nair and Dufresne investigated the surface chemical modification of chitin nanowhiskers using a small molecule chemical reaction between hydroxyl groups of chitin and isocyanate groups from phenyl isocyanate and isopropenyl-α,α’-dimethylbenzyl isocyanate [29]. 6.5.1.3 Oxidative modification Oxidative modification of chitin was achieved by directly grafting poly(3-hexylthiophene) (P3HT) to their surfaces in the presence of FeCl3. Introducing P3HT to the surface of chitin improved significantly

Handbook of Chitin and Chitosan

6.5 Surface modification of chitin

187

the electrical property of it with an increase in conductivity from 1029 to 1027 S/cm and water contact angle up to 97.7 degrees [98]. The C6 primary hydroxyl groups on the chitin nanowhiskers surfaces are selectively oxidized to carboxylate groups via the aldehyde structure by using 2,2,6,6-tetramethylpiperidine-1-oxyl radical (TEMPO)-mediated oxidation. The total contents of carboxylate and aldehyde groups at 5.0 and 10 mmol of NaClO per gram of chitin were 12% and 24% of the C6 primary hydroxyl groups of the original chitin to be oxidized to either carboxylate or aldehyde groups. When a sufficient amount of NaClO is added to the chitin/water slurries in the oxidation, chitin can be converted to the corresponding water-soluble polyuronic acid with partial depolymerization at pH 10
11 [14]. In the presence of anionic functional groups (carboxylates, sulfates, phosphates, etc.), the chitin surface gains a negative charge under basic conditions resulting in a stable dispersion similar to that of TEMPOoxidized chitin nanocrystals. The grafting reaction was then carried out with potassium persulfate as an initiator, which allows a facile radical grafting from polymerization in aqueous media, where radicals are formed along the chitin polymer backbone followed by a free radical polymerization of the acrylic acid monomer. Thus graft copolymerization of acrylic acid on chitin nanofibers was carried out with potassium persulfate as a free radical initiator in an aqueous medium [13]. Reductive alkylation and direct alkylation are the methods of synthesis of the carboxymethylated forms of chitin and chitosan. OCarboxymethyl chitin was prepared by reacting chitin with NaOH and SDS (sodium dodecyl sulfate) at 20 C for 12 h, and then adding isopropanol and monochloroacetic acid at 25 C [99].

6.5.2 Hydrophobization of chitin surface Feng et al. prepared new thermoformable bionanocomposites of chitin whisker-graft-polycaprolactone (CHW-g-PCL) by the ring-opening polymerization of caprolactone and grafting onto the CHW surface under microwave radiation. This modification imparted a higher hydrophobicity to the ensuing surface, and, depending on the thickness of the PCL coating, water contact angles in the range of 86
107 degrees were attained [100]. In an oil-in-water emulsification/evaporation method, the hydrophobization of chitin was conducted by the addition of labile butyryl groups onto chitin, disrupting intermolecular hydrogen bonds and enabling solubility in the organic solvent used as the oil phase during fabrication [96]. Hydrophobization of sheet-like chitin was obtained after dissolving chitin molecules in aqueous NaOH/urea solution and chitin molecules

Handbook of Chitin and Chitosan

188

6. Chitin nanomaterials: preparation and surface modifications

then get a solvated sheath-like structures and extended wormlike conformations. The addition of CH2CH2CN imparted hydrophobization on chitin chains, which thus depressed H-bonding between the chitin chain core and NaOH/urea hydrate shell. The presence of hydrophobic CH2CH2CN groups also drove molecular aggregation of unstable chitin chains, analogous to most amphiphilic polymers [8].

6.5.3 Hydrophilization of chitin surface Hydrophilicity of 2-hydroxyethylmethacrylate (HEMA) and 4vinylpyridine (4-VP) grafted chitin was investigated using cerium(IV) ammonium nitrate as the redox initiator. The intractable nature of chitin, which is one of its primary drawbacks as a grafting substrate was overcome by applying a CaCO3 treatment during bead preparation. The maximum grafting percentage of poly(HEMA) and poly(4-VP) onto chitin bead with CaCO3 treatment was found to be 515% and 380%, respectively, at optimum conditions [101].

6.5.4 Physical modification of chitin surface A corrugated surface of chitin particles along with increased porosity was obtained after applying several ionic liquids (with imidazolium tetracyanoborates and with pyridinium tetracyanoborates) in chitin. The methods applied only changed the surface physically. Approximately 2 g ionic liquid and 5% w/w of chitin were mixed at 200 rpm in a test tube, incubated at 105 C for 48 h. Then the chitin was precipitated adding the chitin suspension in the IL dropwise into hot (95 C) water and left in the solution until full sedimentation. The final surface modified chitin was precipitated after decantation and washing with an ethanol/water mixture followed with 3
7 times further washing with ethanol and water separately [102].

6.5.5 Ultrasound-assisted surface modification of chitin Raw chitin of 5.00 g was mixed with 300 mL of deionized water in an ultrasonic processor of 400 W, equipped with a titanium sonotrode. The sonication was performed at 24 kHz, cycle of 1.00, amplitude of 60% for 1 h. The final chitin slurry was separated by filtration and dried at 40 C for 24 h [103].

6.5.6 Plasma treatment The cellulose and chitin mix fibers were activated for 15 min in highfrequency plasma using a 0.4 mbar (40 Pa) vacuum. After plasma

Handbook of Chitin and Chitosan

References

189

activation, the fibers were modified with (N-isopropylacrylamide) NIPAAm or poly(N-isopropylacrylamide) (PNIPAAm) by immersing the fibers for 30 min in a solution of NIPAAm or PNIPAAm (10 wt.%) along with two coupling agents used separately. One is 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide (EDC) and the other is N-hydroxysuccinimide (NHS). The fibers were dried at 60 C, and then extracted for 25 h in a Soxhlet extractor with methanol in order to remove the physically adsorbed unreacted chemicals [104].

6.6 Conclusions Due to the advances in nanotechnology and biomedicine the demand for chitin nanomaterials has been increasing and they are expected to have great potential to be used as biomedical materials. This chapter focused on the preparation and surface modification of chitin nanomaterials. To diversify the applicability of chitin-based nanomaterials, various preparation techniques and surface modification have been reported and are discussed here. As the nanomaterials of chitin have numerous useful properties, like high specific surface area and high porosity, they have prospects for application in biomedical fields. Although before the age of nanotechnology, chitin-based materials have been applied in many biomedical fields, the development of nanotechnology has enhanced the range of applicability. Because of the promising biological properties of chitin nanomaterials, like biocompatibility, nontoxicity, biodegradability, and antibacterial activity, these nanomaterials are potential candidates for the improvement of absorption of drugs, cell proliferation, enzyme immobilization, wound healing, and other biological applications. Nevertheless, most of these uses are still at the laboratory scale. Further studies are important to develop new preparation techniques of chitin nanomaterials and their surface modification to improve their solubility as well as new fields of application. We hope this chapter will help to find out new chitin-based nanomaterials and their surface modification techniques to widen their fields of application in the near future.

References [1] I.A. Hoell, G. Vaaje-Kolstad, V.G.H. Eijsink, Structure and function of enzymes acting on chitin and chitosan, Biotechnol. Genet. Eng. 27 (2010) 331
366. [2] H. Merzendorfer, L. Zimoch, Chitin metabolism in insects: structure, function and regulation of chitin synthases and chitinases, J. Exp. Biol. 206 (2003) 4393
4412. [3] C.K.S. Pillai, W. Paul, C.P. Sharma, Chitin and chitosan polymers: chemistry, solubility and fiber formation, Prog. Polym. Sci. 34 (2009) 641
678.

Handbook of Chitin and Chitosan

190

6. Chitin nanomaterials: preparation and surface modifications

[4] R.J. Moon, A. Martini, J. Nairn, J. Simonsen, J. Youngblood, Cellulose nanomaterials review: structure, properties and nanocomposites, Chem. Soc. Rev. 40 (2011) 3941
3994. [5] R. Jayakumar, M. Prabaharan, S.V. Nair, H. Tamura, Novel chitin and chitosan nanofibers in biomedical applications, Biotechnol. Adv. 28 (2010) 142
150. [6] J.-B. Zeng, Y.-S. He, S.-L. Li, Y.-Z. Wang, Chitin whiskers: an overview, Biomacromolecules 13 (2012) 1
11. [7] N.S. Rejinold, A. Nair, M. Sabitha, K.P. Chennazhi, H. Tamura, S.V. Nair, et al., Synthesis, characterization and in vitro cytocompatibility studies of chitin nanogels for biomedical applications, Carbohydr. Polym. 87 (2012) 943
949. [8] J. You, M. Li, B. Ding, X. Wu, C. Li, Crab chitin-based 2D soft nanomaterials for fully biobased electric devices, Adv. Mater. 29 (2017) 1606895. [9] F. Ding, H. Deng, Y. Du, X. Shi, Q. Wang, Emerging chitin and chitosan nanofibrous materials for biomedical applications, Nanoscale 6 (2014) 9477
9493. [10] Z.-M. Huang, Y.Z. Zhang, M. Kotaki, S. Ramakrishna, A review on polymer nanofibers by electrospinning and their applications in nanocomposites, Compos. Sci. Technol. 63 (2003) 2223
2253. [11] S. Ramakrishna, K. Fujihara, W.-E. Teo, T. Yong, Z. Ma, R. Ramaseshan, Electrospun nanofibers: solving global issues, Mater. Today 9 (2006) 40
50. [12] K. Azuma, S. Ifuku, T. Osaki, Y. Okamoto, S. Minami, Preparation and biomedical applications of chitin and chitosan nanofibers, J. Biomed. Nanotechnol. 10 (2014) 2891
2920. [13] D. Narayanan, R. Jayakumar, K.P. Chennazhi, Versatile carboxymethyl chitin and chitosan nanomaterials: a review, Wiley Interdiscip. Rev. Nanomed. Nanobiotechnol. 6 (2014) 574
598. [14] M. Mincea, A. Negrulescu, V. Ostafe, Preparation, modification, and applications of chitin nanowhiskers: a review, Rev. Adv. Mater. Sci. 30 (2012) 225
242. [15] D.X. Oh, Y.J. Cha, H.-L. Nguyen, H.H. Je, Y.S. Jho, D.S. Hwang, et al., Chiral nematic self-assembly of minimally surface damaged chitin nanofibrils and its load bearing functions, Sci. Rep. 6 (2016) 23245. [16] E. El-Diasty, N. Eleiwa, A. Aideia, Using of chitosan as antifungal agent in Kariesh cheese, N.Y. Sci. J. 5 (2012) 5
10. [17] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (2006) 603
632. [18] Y. Noishiki, Y. Nishiyama, M. Wada, S. Okada, S. Kuga, Inclusion complex of β-chitin and aliphatic amines, Biomacromolecules 4 (2003) 944
949. [19] D.L. Wise, Biomaterials and Bioengineering Handbook, Marcel Dekker, 2000. [20] N.S. Gupta, Chitin: Formation and Diagenesis, Springer Science & Business Media, 2010. [21] I. Younes, M. Rinaudo, Chitin and chitosan preparation from marine sources. Structure, properties and applications, Mar. Drugs 13 (2015) 1133
1174. [22] A.J. Friedman, J. Phan, D.O. Schairer, J. Champer, M. Qin, A. Pirouz, et al., Antimicrobial and anti-inflammatory activity of chitosan-alginate nanoparticles: a targeted therapy for cutaneous pathogens, J. Invest. Dermatol. 133 (2013) 1231
1239. [23] G.W. Gooday, The ecology of chitin degradation, in: K.C. Marshall (Ed.), Advances in Microbial Ecology, Springer US, Boston, MA, 1990. [24] M.-T. Yen, J.-H. Yang, J.-L. Mau, Physicochemical characterization of chitin and chitosan from crab shells, Carbohydr. Polym. 75 (2009) 15
21. [25] S. Ospina, S.P. Lvarez, D.A. Ramirez, D.M. Escobar Sierra, C.P. Ossa Orozco, D.F. Rojas Vahos, et al., Comparison of extraction methods of chitin from ganoderma lucidum mushroom obtained in submerged culture, BioMed. Res. Int. 2014 (2014) 7. [26] S.M. Bowman, S.J. Free, The structure and synthesis of the fungal cell wall, BioEssays 28 (2006) 799
808. [27] M. Peesan, R. Rujiravanit, P. Supaphol, Characterisation of beta-chitin/poly(vinyl alcohol) blend films, Polym. Test. 22 (2003) 381
387.

Handbook of Chitin and Chitosan

References

191

[28] S. Ifuku, H. Saimoto, Chitin nanofibers: preparations, modifications, and applications, Nanoscale 4 (2012) 3308
3318. [29] K. Gopalan Nair, A. Dufresne, Crab shell chitin whisker reinforced natural rubber nanocomposites. 1. Processing and swelling behavior, Biomacromolecules 4 (2003) 657
665. [30] J. Wu, K. Zhang, N. Girouard, J.C. Meredith, Facile route to produce chitin nanofibers as precursors for flexible and transparent gas barrier materials, Biomacromolecules 15 (2014) 4614
4620. [31] J.N. BeMiller, R.L. Whistler, Alkaline degradation of amino sugars, J. Org. Chem. 27 (1962) 1161
1164. [32] S. Ifuku, M. Nogi, K. Abe, M. Yoshioka, M. Morimoto, H. Saimoto, et al., Preparation of chitin nanofibers with a uniform width as α-chitin from crab shells, Biomacromolecules 10 (2009) 1584
1588. [33] S. Ifuku, A.N. Nakagaito, H. Saimoto, Chitin nanofibers with a uniform width of 10 to 20 nm and their transparent nanocomposite films, Nanofibers-Production, Properties and Functional Applications (2011). IntechOpen. [34] S. Ifuku, Z. Shervani, H. Saimoto, Chitin nanofibers, preparations and applications, Advances in Nanofibers, InTech, 2013. [35] M. Rolandi, R. Rolandi, Self-assembled chitin nanofibers and applications, Adv. Colloid Interf. Sci. 207 (2014) 216
222. [36] K.E. Park, S.Y. Jung, S.J. Lee, B.-M. Min, W.H. Park, Biomimetic nanofibrous scaffolds: preparation and characterization of chitin/silk fibroin blend nanofibers, Int. J. Biol. Macromol. 38 (2006) 165
173. [37] S. Ifuku, Chitin and chitosan nanofibers: preparation and chemical modifications, Molecules 19 (2014) 18367
18380. [38] K. Sun, Z. Li, Preparations, properties and applications of chitosan based nanofibers fabricated by electrospinning, Exp. Polym. Lett. (2011) 5. [39] S. Haider, S.-Y. Park, Preparation of the electrospun chitosan nanofibers and their applications to the adsorption of Cu(II) and Pb(II) ions from an aqueous solution, J. Memb. Sci. 328 (2009) 90
96. [40] Y. Zhou, D. Yang, X. Chen, Q. Xu, F. Lu, J. Nie, Electrospun water-soluble carboxyethyl chitosan/poly(vinyl alcohol) nanofibrous membrane as potential wound dressing for skin regeneration, Biomacromolecules 9 (2008) 349
354. [41] M.E. Villanueva, A. Salinas, L.E. Dı´az, G.J. Copello, Chitin nanowhiskers as alternative antimicrobial controlled release carriers, New J. Chem. 39 (2015) 614
620. [42] J.F. Revol, R.H. Marchessault, In vitro chiral nematic ordering of chitin crystallites, Int. J. Biol. Macromol. 15 (1993) 329
335. [43] J. Li, J.F. Revol, R.H. Marchessault, Rheological properties of aqueous suspensions of chitin crystallites, J. Colloid Interf. Sci. 183 (1996) 365
373. [44] Y. Fan, T. Saito, A. Isogai, Individual chitin nano-whiskers prepared from partially deacetylated α-chitin by fibril surface cationization, Carbohydr. Polym. 79 (2010) 1046
1051. [45] K. Gopalan Nair, A. Dufresne, A. Gandini, M.N. Belgacem, Crab shell chitin whiskers reinforced natural rubber nanocomposites. 3. Effect of chemical modification of chitin whiskers, Biomacromolecules 4 (2003) 1835
1842. [46] P. Wongpanit, N. Sanchavanakit, P. Pavasant, T. Bunaprasert, Y. Tabata, R. Rujiravanit, Preparation and characterization of chitin whisker-reinforced silk fibroin nanocomposite sponges, Eur. Polym. J. 43 (2007) 4123
4135. [47] M. Paillet, A. Dufresne, Chitin whisker reinforced thermoplastic nanocomposites, Macromolecules 34 (2001) 6527
6530. [48] J.-I. Kadokawa, A. Takegawa, S. Mine, K. Prasad, Preparation of chitin nanowhiskers using an ionic liquid and their composite materials with poly(vinyl alcohol), Carbohydr. Polym. 84 (2011) 1408
1412.

Handbook of Chitin and Chitosan

192

6. Chitin nanomaterials: preparation and surface modifications

[49] T.T. Nge, N. Hori, A. Takemura, H. Ono, T. Kimura, Phase behavior of liquid crystalline chitin/acrylic acid liquid mixture, Langmuir 19 (2003) 1390
1395. [50] S. Ifuku, M. Nogi, M. Yoshioka, M. Morimoto, H. Yano, H. Saimoto, Fibrillation of dried chitin into 10
20 nm nanofibers by a simple grinding method under acidic conditions, Carbohydr. Polym. 81 (2010) 134
139. [51] L. Heath, L. Zhu, W. Thielemans, Chitin nanowhisker aerogels, ChemSusChem 6 (2013) 537
544. [52] S. Phongying, S.-I. Aiba, S. Chirachanchai, Direct chitosan nanoscaffold formation via chitin whiskers, Polymer 48 (2007) 393
400. [53] K.-K. Yang, X.-L. Wang, Y.-Z. Wang, Progress in nanocomposite of biodegradable polymer, J. Industr. Eng. Chem. 13 (2007) 485
500. [54] M. Wysokowski, M. Motylenko, J. Beyer, A. Makarova, H. Sto¨cker, J. Walter, et al., Extreme biomimetic approach for developing novel chitin-GeO2 nanocomposites with photoluminescent properties, Nano Res. 8 (2015) 2288
2301. [55] M.I. Shams, S. Ifuku, M. Nogi, T. Oku, H. Yano, Fabrication of optically transparent chitin nanocomposites, Appl. Phys. A 102 (2011) 325
331. [56] A. Morin, A. Dufresne, Nanocomposites of chitin whiskers from riftia tubes and poly (caprolactone), Macromolecules 35 (2002) 2190
2199. [57] M.-B. Coltelli, P. Cinelli, V. Gigante, L. Aliotta, P. Morganti, L. Panariello, et al., Chitin nanofibrils in poly(lactic acid) (PLA) nanocomposites: dispersion and thermomechanical properties, Int. J. Mol. Sci. 20 (2019) 504. [58] N. Singh, K.K.K. Koziol, J. Chen, A.J. Patil, J.W. Gilman, P.C. Trulove, et al., Ionic liquids-based processing of electrically conducting chitin nanocomposite scaffolds for stem cell growth, Green Chem. 15 (2013) 1192
1202. [59] S.A. Rodriguez, E. Weese, J. Nakamatsu, F. Torres, Development of biopolymer nanocomposites based on polysaccharides obtained from red algae chondracanthus chamissoi reinforced with chitin whiskers and montmorillonite, Polym.-Plast. Technol. Eng. 55 (2016) 1557
1564. [60] M. Zuber, K.M. Zia, S. Mahboob, M. Hassan, I.A. Bhatti, Synthesis of chitin
bentonite clay based polyurethane bio-nanocomposites, Int. J. Biol. Macromol. 47 (2010) 196
200. [61] A.K. Mallik, M. Shahruzzaman, M.N. Sakib, A. Zaman, M.S. Rahman, M.M. Islam, et al., Benefits of renewable hydrogels over acrylate- and acrylamide-based hydrogels, in: M.I.H. Mondal (Ed.), Cellulose-Based Superabsorbent Hydrogels, Springer International Publishing, Cham, 2018. [62] K. Raemdonck, J. Demeester, S. De Smedt, Advanced nanogel engineering for drug delivery, Soft Matter 5 (2009) 707
715. [63] J. Zhang, P. Wu, Y. Zhao, S. Xue, X. Zhu, J. Tong, et al., A simple mechanical agitation method to fabricate chitin nanogels directly from chitin solution and subsequent surface modification, J. Mater. Chem. B 7 (2019) 2226
2232. [64] S. Mangalathillam, N.S. Rejinold, A. Nair, V.-K. Lakshmanan, S.V. Nair, R. Jayakumar, Curcumin loaded chitin nanogels for skin cancer treatment via the transdermal route, Nanoscale 4 (2012) 239
250. [65] M. Vishnu Priya, M. Sabitha, R. Jayakumar, Colloidal chitin nanogels: a plethora of applications under one shell, Carbohydr. Polym. 136 (2016) 609
617. [66] G. Divya, R. Panonnummal, S. Gupta, R. Jayakumar, M. Sabitha, Acitretin and aloeemodin loaded chitin nanogel for the treatment of psoriasis, Eur. J. Pharm. Biopharm. 107 (2016) 97
109. [67] T.R. Arunraj, N. Sanoj Rejinold, N. Ashwin Kumar, R. Jayakumar, Doxorubicin-chitin-poly(caprolactone) composite nanogel for drug delivery, Int. J. Biol. Macromol. 62 (2013) 35
43. [68] R. Mas-Balleste´, C. Go´mez-Navarro, J. Go´mez-Herrero, F. Zamora, 2D materials: to graphene and beyond, Nanoscale 3 (2011) 20
30.

Handbook of Chitin and Chitosan

References

193

[69] Q. Lu, Y. Yu, Q. Ma, B. Chen, H. Zhang, 2D transition-metal-dichalcogenidenanosheet-based composites for photocatalytic and electrocatalytic hydrogen evolution reactions, Adv. Mater. 28 (2016) 1917
1933. [70] X. Zhuang, Y. Mai, D. Wu, F. Zhang, X. Feng, Two-dimensional soft nanomaterials: a fascinating world of materials, Adv. Mater. 27 (2015) 403
427. [71] A. Baji, Y.-W. Mai, S.-C. Wong, M. Abtahi, P. Chen, Electrospinning of polymer nanofibers: effects on oriented morphology, structures and tensile properties, Compos. Sci. Technol. 70 (2010) 703
718. [72] H.S. Yoo, T.G. Kim, T.G. Park, Surface-functionalized electrospun nanofibers for tissue engineering and drug delivery, Adv. Drug Deli. Rev. 61 (2009) 1033
1042. [73] B.-M. Min, S.W. Lee, J.N. Lim, Y. You, T.S. Lee, P.H. Kang, et al., Chitin and chitosan nanofibers: electrospinning of chitin and deacetylation of chitin nanofibers, Polymer 45 (2004) 7137
7142. [74] K.E. Park, H.K. Kang, S.J. Lee, B.-M. Min, W.H. Park, Biomimetic nanofibrous scaffolds: preparation and characterization of pga/chitin blend nanofibers, Biomacromolecules 7 (2006) 635
643. [75] H. Deng, P. Lin, S. Xin, R. Huang, W. Li, Y. Du, et al., Quaternized chitosan-layered silicate intercalated composites based nanofibrous mats and their antibacterial activity, Carbohydr. Polym. 89 (2012) 307
313. [76] W.-W. Hu, H.-N. Yu, Coelectrospinning of chitosan/alginate fibers by dual-jet system for modulating material surfaces, Carbohydr. Polym. 95 (2013) 716
727. [77] S.I. Jang, J.Y. Mok, I.H. Jeon, K.-H. Park, T.T.T. Nguyen, J.S. Park, et al., Effect of electrospun non-woven mats of dibutyryl chitin/poly(lactic acid) blends on wound healing in hairless mice, Molecules (Basel, Switzerland) 17 (2012) 2992
3007. [78] S. Xin, Y. Li, W. Li, J. Du, R. Huang, Y. Du, et al., Carboxymethyl chitin/organic rectorite composites based nanofibrous mats and their cell compatibility, Carbohydr. Polym. 90 (2012) 1069
1074. [79] R. Kose, T. Kondo, Favorable 3D-network formation of chitin nanofibers dispersed in water prepared using aqueous counter collision, 繊維学会誌 67 (2011) 91
95. [80] R.M. Capito, H.S. Azevedo, Y.S. Velichko, A. Mata, S.I. Stupp, Self-assembly of large and small molecules into hierarchically ordered sacs and membranes, Science 319 (2008) 1812
1816. [81] A. Cooper, C. Zhong, Y. Kinoshita, R.S. Morrison, M. Rolandi, M. Zhang, Selfassembled chitin nanofiber templates for artificial neural networks, J. Mater. Chem. 22 (2012) 3105
3109. [82] J. Jin, P. Hassanzadeh, G. Perotto, W. Sun, M.A. Brenckle, D. Kaplan, et al., A biomimetic composite from solution self-assembly of chitin nanofibers in a silk fibroin matrix, Adv. Mater. 25 (2013) 4482
4487. [83] C. Zhong, A. Kapetanovic, Y. Deng, M. Rolandi, A chitin nanofiber ink for airbrushing, replica molding, and microcontact printing of self-assembled macro-, micro-, and nanostructures, Adv. Mater. 23 (2011) 4776
4781. [84] S. Alom Ruiz, C.S. Chen, Microcontact printing: a tool to pattern, Soft Matter 3 (2007) 168
177. [85] A.P. Quist, E. Pavlovic, S. Oscarsson, Recent advances in microcontact printing, Anal. Bioanal. Chem. 381 (2005) 591
600. [86] S. Ifuku, M. Nogi, K. Abe, M. Yoshioka, M. Morimoto, H. Saimoto, et al., Simple preparation method of chitin nanofibers with a uniform width of 10
20nm from prawn shell under neutral conditions, Carbohydr. Polym. 84 (2011) 762
764. [87] Y. Fan, H. Fukuzumi, T. Saito, A. Isogai, Comparative characterization of aqueous dispersions and cast films of different chitin nanowhiskers/nanofibers, Int. J. Biol. Macromol. 50 (2012) 69
76.

Handbook of Chitin and Chitosan

194

6. Chitin nanomaterials: preparation and surface modifications

[88] H.-P. Zhao, X.-Q. Feng, H. Gao, Ultrasonic technique for extracting nanofibers from nature materials, Appl. Phys. Lett. 90 (2007) 073112. [89] Y. Lu, Q. Sun, X. She, Y. Xia, Y. Liu, J. Li, et al., Fabrication and characterisation of α-chitin nanofibers and highly transparent chitin films by pulsed ultrasonication, Carbohydr. Polym. 98 (2013) 1497
1504. [90] X. Wu, F.G. Torres, F. Vilaseca, T. Peijs, Influence of the processing conditions on the mechanical properties of chitin whisker reinforced poly(caprolactone) nanocomposites, J. Biobased Mater. Bio 1 (2007) 341
350. [91] Y. Lu, L. Weng, L. Zhang, Morphology and properties of soy protein isolate thermoplastics reinforced with chitin whiskers, Biomacromolecules 5 (2004) 1046
1051. [92] P.R. Chang, R. Jian, J. Yu, X. Ma, Starch-based composites reinforced with novel chitin nanoparticles, Carbohydr. Polym. 80 (2010) 420
425. [93] J. Huang, J. Zou, P. Chang, J. Yu, A. Dufresne, New waterborne polyurethane-based nanocomposites reinforced with low loading levels of chitin whisker, Express Polym. Lett. (2011) 5. [94] A.G. Cunha, A. Gandini, Turning polysaccharides into hydrophobic materials: a critical review. Part 2. Hemicelluloses, chitin/chitosan, starch, pectin and alginates, Cellulose 17 (2010) 1045
1065. [95] S. Ifuku, S. Morooka, M. Morimoto, H. Saimoto, Acetylation of chitin nanofibers and their transparent nanocomposite films, Biomacromolecules 11 (2010) 1326
1330. [96] B. Blanco-Fernandez, S. Chakravarty, M.K. Nkansah, E.M. Shapiro, Fabrication of magnetic and fluorescent chitin and dibutyrylchitin sub-micron particles by oil-inwater emulsification, Acta Biomater. 45 (2016) 276
285. [97] J. Wang, C. Liu, P. Chi, Aggregate formation and surface activity of partially deacetylated water-soluble chitin, Res. Chem. Intermediate. 34 (2008) 169
179. [98] T.A.P. Hai, R. Sugimoto, Surface modification of chitin and chitosan with poly(3hexylthiophene) via oxidative polymerization, Appl. Surf. Sci. 434 (2018) 188
197. [99] L. Feng, Z. Zhou, A. Dufresne, J. Huang, M. Wei, L. An, Structure and properties of new thermoforming bionanocomposites based on chitin whisker-graftpolycaprolactone, J. Appl. Polym. Sci. 112 (2009) 2830
2837. [100] R. Salah, D. Tazdaı¨t, N. Mameri, Tumoricidal effect of O-carboxymethyl chitin, N,Ocarboxymethyl chitosan and 2-phtalimido chitin evaluation with human tumour cell line, International Conference on Chemical, Civil and Environmental Engineering (CCEE-2015) June 5-6, 2015 Istanbul (Turkey). [101] Z. Yalinca, D.A.K. Mohammed, J.M. Hadi, E. Yilmaz, Effect of CaCO3/HCl pretreatment on the surface modification of chitin gel beads via graft copolymerization of 2hydroxy ethyl methacrylate and 4-vinylpyridine, Int. J. Biol. Macromol. 82 (2016) 208
216. [102] M.M. Jaworska, A. Go´rak, New ionic liquids for modification of chitin particles, Res. Chem. Intermediat. 44 (2018) 4841
4854. [103] D.S.P. Franco, J.S. Piccin, E.C. Lima, G.L. Dotto, Interpretations about methylene blue adsorption by surface modified chitin using the statistical physics treatment, Adsorption 21 (2015) 557
564. [104] A. Sdrobi¸s, G.E. Ioanid, T. Stevanovic, C. Vasile, Modification of cellulose/chitin mix fibers with N-isopropylacrylamide and poly(N-isopropylacrylamide) under cold plasma conditions, Polym. Int. 61 (2012) 1767
1777.

Handbook of Chitin and Chitosan

C H A P T E R

7 Importance of electrospun chitosan-based nanoscale materials for seafood products safety ¨ zogul3 and Zafer Ceylan1, Raciye Meral2, Fatih O Mustafa Tahsin Yilmaz4,5 1

Faculty of Fisheries, Department of Seafood Processing Technology, Van Yuzuncu Yil University, Van, Turkey, 2Faculty of Engineering, Department of Food Engineering, Van Yuzuncu Yil University, Van, Turkey, 3Faculty of Fisheries, Department of Seafood Processing Technology, C ¸ ukurova University, Adana, Turkey, 4Faculty of Engineering, Department of Industrial Engineering, King Abdulaziz University, Jeddah, Saudi Arabia, 5 Chemical and Metallurgical Engineering Faculty, Department of Food Engineering, Yıldız Technical University, Istanbul, Turkey O U T L I N E 7.1 Optimization 7.1.1 Definition of chitosan molecular weight 7.1.2 Determination of concentrations 7.1.3 Solvent system 7.1.4 Preparation of electrospinning dope solutions

196 198 199 200 202

7.2 Determination of electrospinning parameters 7.2.1 Applied voltage 7.2.2 Adjustment proper distance 7.2.3 Flow rate

202 202 203 204

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00007-9

195

© 2020 Elsevier Inc. All rights reserved.

196

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

7.2.4 Environmental conditions 7.2.5 Selection of collector

205 206

7.3 Characterization of fabricated nanoscale material(s) 7.3.1 Definition of morphological characteristics of chitosan-based nanomaterial 7.3.2 Encapsulation efficiency 7.3.3 Controlled release property of chitosan-based nanomaterial 7.3.4 Thermal decomposition of chitosan-based nanomaterial 7.3.5 Zeta potential and size of chitosan-based nanomaterial

207

7.4 Use of electrospun nanomaterials for seafood products safety 7.4.1 Chitosan-based nanofiber coating 7.4.2 Bio/active material-loaded chitosan-based nanofiber coating 7.4.3 Chitosan nanoparticles

212 212 214 215

7.5 Conclusion

216

References

217

207 209 210 210 211

7.1 Optimization Optimization is defined to be a procedure of improving a process, system, or a product to obtain maximum output from it [1]. In the fabrication of nanoscale material, optimization plays a unique role. Optimization is not only important for the electrospinning method but also for the other techniques that allow nanoscale materials to be obtained. Fig. 7.1 shows an electrospinning unit. Electrospinning is a process that creates nanofibers through an electrically charged jet of polymer solution or melt [2]. A simple electrospinning unit consists of a high-voltage unit, collector, and syringe pump [1]. In this unit, the determination of the molecular weight of the substance, concentrations of the polymer, solvent systems, and preparation of electrospinning dope solutions are highly significant for successfully obtaining a nanoscale material. There are some studies related to the optimization of electrospun nanoscale chitosan. According to Jacobs et al. [2], the interaction effect between electric field strength and concentration of chitosan and the ratio of solvents and electric field strength had the most important roles in obtaining chitosan nanofibers (Fig. 7.2). Qasim et al. [3] reported that the viscosity of the chitosan solution played one of the most important roles in the optimization. Amiri et al. [4] revealed that response surface

Handbook of Chitin and Chitosan

7.1 Optimization

197

FIGURE 7.1 A sample electrospinning unit.

FIGURE 7.2 (A) Electrospun chitosan-based nanofiber after well optimization, (B) halogen lamp, (C) jet, (D) distance between the collector and Taylor cone, (E) needle, and (F) flat collector.

Handbook of Chitin and Chitosan

198

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

methodology (RSM) was employed to develop a statistical model and also to define the optimum condition for the fabrication of a chitosan collagen nanofiber with minimum diameter. Thus the optimization process indicated that the chitosan collagen nanofiber diameter of 156.05 nm could be obtained at 9 kV (high voltage), 0.2 mL/h (flow rate), and 25 cm (distance), which was confirmed by experiment (155.92 6 18.95 nm). As stated by Pillai and Sharma (2009), RSM was used in order to model and optimize the electrospinning parameters for chitosan PVA nanofibers, and the predicted nanofiber diameters were determined to be in agreement with the experimental results of chitosan PVA electrospun nanofibers. Additionally, the use of the RSM model for the electrospinning technique could reveal the antimicrobial properties of the chitosan-based nanofibers [5]. Therefore RSM for the electrospinning process should be developed/discovered prior to the more time-consuming and expensive experimental processes. However, besides RSM, the main parameters in the electrospinning technique had an important role, thus preliminary studies can define the high voltage, flow rate, distance, and concentration. These parameters affect the antimicrobial and antioxidant properties of the electrospun chitosan-based nanoscale materials.

7.1.1 Definition of chitosan molecular weight The molecular weight (MW) of chitosan is widely classified as low, medium, or high, thus the MW can change between 50,000 and .375,000 Da. In nanotechnological application, the MW of chitosan can affect some properties of the nanoscale material. Maximum encapsulation efficiencies (EE) can be also affected depending on the MW of chitosan used in nanotechnological application. Kouchak et al. [6] reported that the maximum EE of chitosan nanoparticles with low, medium, and high MWs was found to be 61.88%, 70.89%, and 53.73%, respectively. The MW of the obtained nanoscale material can also affect antibacterial efficiency; in this sense, the antibacterial effect of nanochitosan with low MW was determined to be significant [7]. The MW of chitosan can have a key role in obtaining thinner or thicker nanofibers. Qinna et al. [8] noted that the use of higher MWs generates a thicker nanofiber, while lower MWs of chitosan can provide thin nanofibers. The average diameter of nanoscale material can be affected, for example, the average diameter of electrospun chitosan and PEO nanofibers were revealed to be lower than 147 6 28 nm, when 102 kg/mol average MW was used to obtain nanofiber [9]. In terms of seafood products quality, nanofiber obtained from low-molecular-weight chitosan successfully delayed the rapid increase in bacterial growth in fish fillets stored at

Handbook of Chitin and Chitosan

7.1 Optimization

199

6 C [10]. In fact, different studies related to the MW of chitosan nanoscale material and seafood quality should be carried out to provide safer aquatic food products for consumers.

7.1.2 Determination of concentrations The processing parameters directly affect the morphology of electrospun chitosan fibers. The diameter of the fibers depends on different factors, such as concentration of polymer in the solution, type of solvent used, and feeding rate of the solution [11,12]. One of the most effective variables to determine the fiber morphology and diameter is the concentration of the electrospinning solution [13]. The concentration of the chitosan solution has a significant effect on the final size and distribution of nanomaterials [14]. Susanto et al. [15] found that increasing the concentration increased the average diameter of the poly(vinyl alcohol) (PVA)/chitosan-based nanofiber. These researchers also showed that when the concentration increased, larger beads formed. There is a critical need to produce uniform and bead-free nanomaterial, so that the electrospun chitosan nanomaterials can penetrate the food surface [16]. The solution concentration plays a significant role in the preparation of bead-free electrospun mats (Li and Wang, 2013). Jia et al. [13] used SEM to observe the morphology of the PVA/chitosan electrospun fibers as a function of concentration. They found that the morphology of the fibers changed gradually from the more beads structure to the uniform fiber structure as a result of increasing concentration of the solution. Ohkawa et al. [17] prepared electrospun chitosan nanofibers and they studied the solvent effect on the morphology of these nanofibers. They demonstrated that when the concentration of chitosan increased, the morphology of electrospun chitosan fibers changed from spherical beads to an interconnected fibrous system. Van der Schueren et al. [18] prepared poly(ε-caprolactone) (PCL)/chitosan nanofiber by using a novel solvent system (acetic acid/formic acid). It was observed that the diameter of nanofibers at a constant PCL concentration increased with increasing chitosan concentration. At 6 wt.% PCL, when chitosan concentration increased from 5% to 20%, the diameter increased from 114 6 27 to 196 6 47 nm. This increase in fiber diameter was related to the increase in viscosity of solution. According to [19], when the concentration of poly(ethyleneoxide) (PEO)/chitosan solution reached 8 wt.%, the electrospinning process could not perform because of the higher viscosity of the PEO/chitosan solution. [20] stated that the average diameter increased with increasing concentrations of gelatin in mixed solutions. [21] found that average diameter of collagen/chitosan complex was in the range 434 691 nm, whereas the average diameters of pure chitosan

Handbook of Chitin and Chitosan

200

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

and collagen were 810 and 415 nm, respectively. In the electrospinning process, the mass ratio of polymers is as important as the concentration in generating bead-free nanofibers. Duan et al. [19] also reported that when the mass ratio of chitosan to PEO was increased to 5:1, beads and small amount of bead-fibers were formed without continuous ultrafine fibers. Thus the diameter of electrospun chitosan nanomaterials is affected by electrospinning process parameters. Although viscosity and concentration of solution are important parameters changing the diameter of nanomaterials, the viscosity of the solution alone is not effective for the fabricatation of nanofibers with lower diameter. Crucial factors affecting the diameter and characteristics of the nanomaterials are likewise discussed in this chapter.

7.1.3 Solvent system Chitosan is usually insoluble in aqueous solutions above a pH of B7. However, it becomes water soluble in acidic conditions because of NH2 protonation which facilitates the solubility of the molecule, becoming therefore highly soluble in acid pH ([22]; Lemma et al., 2016). Also chitosan is soluble in dilute acid solutions such as acetic, lactic, malic, formic, or succinic acid [22,23]. The electrospinnability of chitosan is limited since chitosan has a rigid chemical structure, specific inter- and intramolecular interactions and high viscosity. In the traditional spinning processes, chitosan is dissolved in an acetic acid solution due to the amine functions present in the chemical structure [19,24]. It is difficult to fabricate uniform nanofibers from pure chitosan by electrospinning because of the sensitivity of the process to humidity [20]. A suitable solvent is the key to obtaining electrospun chitosan. Trifluoroacetic acid (TFA) is frequently used to improve the electrospinning of chitosan since TFA can form stable salts with chitosan preventing interchain interactions. Additionally, TFA contributes faster fiber formation due to its low boiling point which enables a faster evaporation during the electrospinning process [17,25]. The addition of dichloromethane (DCM) to the chitosan/TFA solution improved the homogeneity of the electrospun chitosan nanofibers [17]. Similarly, Sangsanoh et al. [26] focused on the manufacture of electrospun chitosan nanofibers using TFA/DCM (70:30 v/v). The bead-free, smooth nanofibers with average diameters of 126 6 20 nm were obtained by the electrospinning process. Gu et al. [27] obtained chitosan nanofibers from a solution consisting of concentrated TFA and DCM (7:3 v/v). Chitosan nanofibers have also been electrospun using 1,1,1,3,3,3-hexafluoro-2-

Handbook of Chitin and Chitosan

7.1 Optimization

201

propanol as a solvent (HFIP). Min et al. [28] employed the solvent HFIP to produce pure chitosan nanofibers through deacetylation. Chen et al. [29] fabricated poly(L-lactic acid-co-ε-caprolactone) (P (LLA-CL))/chitosan blend nanofibers. They used HFIP and TFA as solvents. Results demonstrated that the average fiber diameter increased with increasing polymer concentration and decreasing blend ratio of chitosan to P(LLA-CL). In another study conducted [21], a collagen and chitosan mixture (in a HFIP/TFA, v/v, 90/10) was electrospun. The electrospun collagen chitosan fibers had different average diameters, with different chitosan content in the fibers. They were 691 6 376, 515 6 253, and 434 6 263 nm, with a chitosan content of 20%, 50%, and 80%, respectively. Fiber diameters decreased with the increase in chitosan content. In addition to TFA and HFIP, another solvent that has been shown to effectively produce chitosan nanofibers is concentrated acetic acid [30]. In addition, a novel solvent system composed of acetic acid and formic acid was employed to facilitate the electrospinning process of PCL/chitosan blend nanofibers [18]. A strong solvent, such as concentrated acetic acid solution (90%), is necessary for chitosan electrospinning [24]. Likewise, Geng et al. [31] demonstrated the production of electrospun chitosan nanofibers in concentrated acetic acid solution. Kriegel et al. [32] reported that an increase in acetic acid concentration affects the surface tension. Therefore it is difficult to fabricate pure chitosan nanofibers [30]. Moreover, TFA and HFIP are environmentally harmful, very toxic, corrosive, and relatively costlier than conventional solvent systems [25,33]. This makes the use of TFA very limited for food and biomedical applications. An alternative approach both to facilitate the electrospinnability of chitosan and to improve the properties of chitosan nanofibers is blending chitosan with a second natural or synthetic polymeric phase called the cospinning agent [25]. However, cospinning agents are usually biodegradable, nontoxic, and easily electrospinnable polymers, such as PEO [14,25,34], PVA [35,36], PLA [37], zein [38] gelatin [20], and collagen [21]. Bhattarai et al. [14] used PEO to reduce the viscosity of the chitosan solution so that the solution was spinnable at high polymer concentrations. They stated that the maximum chitosan/PEO ratio for making a spinnable solution is 90/10, above which the spun product exhibited a nonuniform structure or droplets. Likewise, Pakravan et al. [25] obtained bead-free nanofibers with a 90 nm mean diameter from a highly deacetylated chitosan grade blended with PEO. Gu et al. [20] investigated the synergy effect between the hydrophilic gelatin and sonicated nanofiber mats to improve hemostatic function on chitosan nanofiber mats. The electrospinning process was improved by introducing hydrophilic gelatin.

Handbook of Chitin and Chitosan

202

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

In addition, there are a few reports investigating the use of inorganic ions as doping agents for producing electrospun fibers. The addition of salt greatly changes the morphology of the electrospun fibers from a beads-on-fiber structure to a uniform fiber structure [39]. Su et al. [40] aimed to investigate the doping effects of monovalent, bivalent, and trivalent metal ions on the morphological appearance of the electrospun chitosan/PEO blend nanofibers. They stated that the calcium and iron ions demonstrated a similar behavior on reducing fiber diameter and the number of beads in fibers, whereas NaCl or KCl nanocrystals occurred in the fibers containing the alkali metal ions.

7.1.4 Preparation of electrospinning dope solutions The electrospinning dope solutions can be prepared after the first steps of the optimization have been successfully completed. Chitosan is added into the test tubes and then it is dissolved by the proper solvent in the same tubes. In this step, the heating process can be used to dissolve the chitosan or it can be dissolved by only using continuous stirring at room temperature. In this process, time is highly changeable. For example, while the preparation of electrospinning dope solution by the heating process can just take a few hours. depending on the concentration of the chitosan, in the case of dissolving the material under room temperature conditions, it takes at least 2 days.

7.2 Determination of electrospinning parameters 7.2.1 Applied voltage In the electrospinning process, the applied voltage is a very crucial factor. Once the applied voltage is high enough to overcome the surface tension of the electrospinning dope solution, the jets continue drying to form nanofibers by the time they are collected on the collector surface. In this sense, the applied voltage can vary depending on the material type used in any nanotechnological application. Mengistu Lemma et al. [9] found that a 20 kV voltage could be applied to successfully obtain pure and stable chitosan nanofibers by electrospinning in the presence of poly(ethylene oxide). However, Moridi et al. [41] reported that the applied voltage could be fixed at 18 24 kV depending on tip-tocollector distance (4 10 cm). Once the applied voltage was low, the beads were deposited on the collector and the average diameter of nanofibers prepared by 18 kV was measured to be 307 nm. When the applied voltage increased, the average nanofiber diameters increased. Terada et al. [42] reported that the applied voltage was found to be

Handbook of Chitin and Chitosan

7.2 Determination of electrospinning parameters

203

12.5 kV, once the flow rate was 0.3 mL/h. In these conditions, 100 mm diameter pure chitosan nanofibers were fabricated in an aqueous solution. Bizarria et al. [43] stated that electric fields of 2 3 kV were estimated/adjusted for each 1 cm tip-to-collector distance to obtain nanofiber chitosan/PEO membranes. As described by Teepoo et al. [44], the voltage for obtaining electrospun chitosan gelatin biopolymer composite nanofibers was increased up to 20 kV from 15 kV. The optimum voltage to fabricate stable and high-quality chitosan nanofibers was determined to be 17 kV [45]. In a study [24], when the applied voltage was 20 kV, the diameter of the chitosan nanofibers was 70 6 45 nm. The voltage had a small effect on the electrospinnability of chitosan/PVA [35]. They discovered that when the voltage increased from 10 to 20 kV, the morphology of the electrospun chitosan nanofibers showed a slight change. The nanofiber diameters were varied from 50 to 200 nm, and the average fiber diameter was 99 6 26 nm when the voltage was 15 kV. As indicated in previous studies, the applied voltage was the important parameter to get very thin nanofiber or nanostructure. The applied voltage has a critical role on the formation of fiber that varies according to the polymer types. A higher voltage also leads to greater stretch of the solution. This directly affects the morphology and diameter of the electrospun fiber. Increasing the applied voltage not only decreases and refines nanofibers diameters, but also improves the quality of electrospun nanofibers.

7.2.2 Adjustment proper distance The tip-to-collector distance affects the electrospun nanofibers diameter, as tip-to-collector distance has a direct effect on jet flight time and electric field strength. A decrease in the tip-to-collector distance shortens flight and solvent evaporation time, and increases the electric field strength, resulting in more bead formation [45]. In this sense, optimum parameters should be provided to fabricate the defect-free nanomaterials. Jabur et al. [46] found that the average PVA-based nanofiber diameter decreased from 875 nm at a distance of 4 cm to 250 nm at a distance of 20 cm, but the study showed that distances over 20 cm increased the nanofiber diameter again. Aliabadi et al. [47] reported that a shorter distance had the opposite effect on the electrospinning process, as decreasing the tip-to-collector distance increased the average diameter of electrospun chitosan hydroxyapatite nanofibers. A shorter distance (7.5 cm) caused the bead fibers on the collector, because of higher electrical forces on the jet that resulted in it becoming unstable and caused it to produce the bead fibers. However, the stability of the jet solution was increased by an increased distance

Handbook of Chitin and Chitosan

204

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

TABLE 7.1 Optimization parameters in some studies. Polymers

Distance (cm)

Diameter (nm)

References

PVA/chitosan

15

330

[17]

PEO/chitosan

20

80 180

[19]

Chitosan

10

70

[24]

PEO/chitosan

10

80

[34]

Chitosan

16

140

[45]

PVA/chitosan

15

PCL/chitosan

12.5

203 267

[18]

Chitosan

15

302

[27]

Gelatin/chitosan

15

269

[20]

PEO/chitosan

8

200

[40]

PEO/chitosan

10

10 240

[32]

PVA/chitosan

9

,600 nm

[49]

Thymol/chitosan

10

135.94 nm

[50]

[36]

(13.75 cm). As a result, homogeneous fibers with lower diameters were formed. When the distance was further increased (20 cm), the strength of electrical force on the spinning solution was decreased and caused to form fibers with higher diameter. In the literature, there are reports revealing an increase in nanomaterial diameter with increasing distance and also there are studies showing a decrease in nanomaterial diameter with increasing spinning distance. Moreover, there are some cases in which distance did not have a significant influence on nanomaterial diameter [48]. Table 7.1 summarizes the tip-to-collector distance effects on the diameter of electrospun chitosan nanomaterial.

7.2.3 Flow rate Flow rate of the polymeric solution is another parameter which affects the size and the shape of the nanomaterials. Generally, a lower flow rate is more recommended, as sufficient time is required for the formation and maintenance of the Taylor cone [51]. However at a lower flow rate, there is a possibility of drying of the electrospinning solution [47]. At lower flow rates, a small amount of dope solution is removed from the capillary tip leading to the formation of nanomaterials with a lower diameter. On the contrary, when increasing the flow rate, a

Handbook of Chitin and Chitosan

7.2 Determination of electrospinning parameters

205

greater amount of solution volume is removed from the needle tip, which results in the jet of the solution to be electrospun without any sufficient extension [52]. Increasing the flow rate tends to increase nanomaterial diameter and bead formation, and at higher flow rates the fiber nanomaterial is rougher [53]. Dhandayuthapani et al. [51] reported that when the flow rate was 0.005 mL/min, defect-free chitosan nanofibers were fabricated. When the flow rate decreased to 0.002 mL/min, the results obtained for chitosan were more reproducible. Applied voltage and flow rate to generate electrospun chitosan nanofibers with a diameter of about 70 6 45 nm were 20 kV and 0.3 mL/h (0.005 mL/min), respectively [24]. Similarly, Aliabadi et al. [54] showed that PEO/chitosan electrospun nanofiber membrane with an average diameter of 98 nm was obtained when the applied voltage and flow rate were 20 kV and 0.5 mL/h, respectively. Although many previous studies emphasized that flow rate had an important role in the electrospinning of chitosan, Zhang et al. [35] reported that flow rate did not remarkably influence the morphology of the electrospun chitosan/PVA fibers. Therefore the previous studies showed that within the electrospinning process not only electrospinning parameters and polymer properties but also their interactions had a crucial role in the fabrication of homogenous and defect-free nanomaterials.

7.2.4 Environmental conditions The effect of temperature and humidity on electrospinning is important in the successful fabrication of nanoscale materials. Jabur et al. [46] studied the ambient temperature (25 C, 30 C, 35 C, 40 C, 45 C, and 50 C) and needle tip-to-collector distance (4, 8, 12, 15, 20, and 22 cm) to determine the influence of these technological parameters on the formation of fibers. In the electrospinning process, with increasing temperature, the solvent evaporation rate can be increased and the viscosity of the polymer solution can be decreased [55]. Low relative humidity (,50%) resulted in fiber breakage for polymers (poly(ethylene glycol), polycaprolactone, and poly(carbonate urethane)), in contrast to high relative humidity ( . 50%) where three distinct effects were observed [56]. Additionally, Garcia et al. [57] observed that biomaterials based on electrospun chitosan nanofiber could be left in ambient conditions to evaporate an excess of solvent. A heating process between 25 C and 70 C was applied to obtain electrospun chitosan nanofiber during the electrospinning process [34]. Nevertheless, there are some studies that used ambient temperature in order to fabricate electrospun chitosan nanofibers during the electrospinning process [9,58]. Consequently, air conditions, seasons, temperature, and humidity, dependent on the location where

Handbook of Chitin and Chitosan

206

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

the electrospinning process is applied, play a key role in successfully fabricating chitosan-based nanofibers. Therefore environmental conditions in food nanotechnological applications should be clearly identified due to the fact that the conditions may affect the antimicrobial or antioxidant properties of the food products treated with chitosan-based nanoscale material.

7.2.5 Selection of collector Following the Taylor cone and the distance, (chitosan-based) nanofibers can be obtained by electrospinning dope solutions. During this step, the solutions evaporated turn into nanofibers and the nanofibers are collected on the surface of the collectors, which are usually flat or drum (mandrel) (Fig. 7.3). The dimension of the flat or drum collector is changeable, depending on the aim, thus a knife-edge (flat) collector could be used to generate highly aligned fibers. However, in order to fabricate patterned nanofibrous mats, collectors with grids or charged needles were used [59]. Mandrels or drum collectors were preferred to obtain oriented fibers and the mat morphology can be influenced by the collector geometry [60]. Valizadeh et al. [61] also emphasized that the collector was an important parameter in the electrospinning process. Rodoplu et al. [62] noted that the collector chosen was an iron plate, which had a 15 3 15 cm2 surface area on which to collect the nanofibers. Nevertheless, the mandrel collector was used for the preparation of electrospun chitosan polyethylene oxide/fibrinogen composite scaffolds with a lower diameter than 351.1 6 101.7 nm [63]. Ceylan et al. [64] used a flat collector to coat the fish fillets using chitosan-based electrospun nanofibers. Actually, the aim of the study was to choose the optimal collector in the electrospinning process. For example, in a dough

FIGURE 7.3 Drum collector used in electrospinning.

Handbook of Chitin and Chitosan

7.3 Characterization of fabricated nanoscale material(s)

207

study, while the drum collector can be a better option, a flat collector can be more useful for fish and meat or their products.

7.3 Characterization of fabricated nanoscale material(s) 7.3.1 Definition of morphological characteristics of chitosan-based nanomaterial The electron microscopic techniques are most frequently used for the analysis of nanomaterials. Although there are several microscopy techniques used to observe nanomaterials, scanning electron microscopy (SEM) and transmission electron microscopy (TEM) are the most popular [65]. The electrons in the beam interact with the specimen generating various signals that can be used to obtain information about the surface topography and composition [66,67]. The morphology of the nanomaterials is evaluated by using SEM that allows the measurement of the diameter of the nanomaterials. The structure of nanomaterials can be observed by SEM without further sample preparation [68]. The SEM technique is nowadays widely used for the evaluation of the structure, morphology, and diameter of nanomaterials, thus it is a highly popular microscopic technique for the ultrastructural investigation of different kinds of materials [38,68]. The effect of electrospinning parameters on the diameter and the morphology of the electrospun chitosan nanomaterials have been reported by several researchers. Majd et al. [36] investigated PVA polymer blended with chitosan in different proportions and the electrospinning parameters were analyzed using SEM that showed the fiber dimensions were nanoscale. Sencadas et al. [22] studied the effect of the main processing parameters (solvent concentration, flow rate, applied voltage, feed rate, and inner needle diameter) on the chitosan fiber characteristics and sample morphology. They reported that the processing parameters have a strong influence on the electrospinning process and thus in the final fiber morphology. Desai et al. [34] blended PEO with chitosan to improve the spinnability of chitosan. SEM images showed that nanometer-sized fibers with fiber diameter as low as 80 6 35 nm without beads were made by electrospinning highmolecular-weight chitosan/PEO (95:5) blends. They also stated that by increasing the percentage of PEO, the fiber diameter increased. Zhang et al. [35] investigated the electrospinnability of chitosan/PVA by using SEM, which revealed that when the concentration increased, drastic morphological changes were found. Han et al. [69] synthesized methoxy poly(ethylene glycol)-grafted chitosan (PEG-g-CS). SEM images showed that when the mass ratio of PEG-g-CS/PEO ranged from 4/1 to 1/2, uniform ultrafine fibers could be produced. Lee et al. [70] prepared and

Handbook of Chitin and Chitosan

208

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

characterized silver-loaded chitosan nanofibers and determined their antibacterial efficacy in a zone of inhibition antibacterial test system. SEM images of nanomaterials showed that the average diameters of chitosan nanofibers containing various ratios of silver nanoparticles (AgNP) were 460 6 80, 126 6 28, 238 6 46, 337 6 49, and 349 6 56 nm. They indicated that the diameters of these nanofibers decreased due to the increase in the content of the AgNPs. Gu et al. [27] observed the morphology of the electrospun chitosan nanofibers by using field emission SEM (FESEM). FESEM images of the surface morphology of the electrospun chitosan nanofibers showed a smooth and bead-free structure, with the average nanofiber diameter measured to be 301.9 6 30.4 nm. Another technique which is used to determine the morphology of electrospun chitosan is TEM, which produces accurate 2D images of particles and it requires extremely thin specimens for penetration and transmission of electrons [70,71]. Both TEM and SEM can be used to directly measure the size, size distribution, and shape of nanomaterials. TEM has advantages over SEM in providing better spatial resolution [70]. Nirmala et al. [72] reported the morphological and electrical characteristics of polyamide-6/chitosan composite nanofibers fabricated by electrospinning. They further carried out TEM analysis to study the bonding of ultrafine nanofibers. TEM images clearly showed that these ultrafine nanofibers were bound in between the main fibers. Nguyen et al. [37] studied biodegradable nonwoven mats of poly(lactic acid) (PLA) and chitosan fabricated by the coaxial electrospinning process. TEM images of the coaxially electrospun composite nanofibers showed that a high contrast difference between the core and shell was obtained due to the difference between the density of core (PLA) and shell (chitosan) materials. TEM images also revealed that the core layer of PLA was completely encapsulated by the outer layer of chitosan. Han et al. [69] fabricated the blend nanofibers of PEG-g-CS and PEO. The core shell structure nanofibers were observed by TEM. Li et al. [67] developed PLA (core)/chitosan (shell) nanofibrous membrane. They used TEM to characterize the core shell structure of the nanofibers. Surucuu et al. [73] worked to combine synthetic PCL and natural chitosan polymers to develop three-dimensional (3D) PCL/chitosan core shell scaffolds for tissue engineering applications. The scaffolds were fabricated by the coaxial electrospinning technique. Images of the TEM revealed that the core layer of PCL was completely encapsulated by chitosan at the shell flow rate of 2 μL/min. Song et al. [74] prepared electrospun chitosan/PEO nanofibrous membrane containing silver nanoparticles. The in situ formation of AgNPs inside the nanofibers was confirmed by TEM. In addition to SEM and TEM, advanced techniques like energy dispersive X-ray spectroscopy (EDX) are employed for further observation

Handbook of Chitin and Chitosan

7.3 Characterization of fabricated nanoscale material(s)

209

of nanomaterials. The EDX is of great interest for the analysis of nanomaterials. In this technique, TEM equipped with an X-ray probe which is used for the elemental analysis [65]. The electrospinnability of chitosan/PVA was discussed by [35]. TEM-EDX analysis was performed to determine the elemental composition of the electrospun chitosan/PVA fibers and beads. According to the results of TEM-EDX, both electrospun chitosan/PVA fibers and bead included nitrogen and showed the existence of chitosan in both fibers and beads. TEM-EDX was used for elemental analysis of the shell layer component of the electrospun PLA/chitosan nanofiber [67]. The results indicated that chitosan was the major component of the shell layer. Song et al. [74] observed the formation of AgNPs using TEM and TEM-EDX. Elemental analysis of the fibers using TEM-EDX confirmed that the electrospun chitosan/PEO nanofibrous membrane was composed of silver. It was also shown that there was homogeneous distribution of silver inside the organic matrix of the nanofibers.

7.3.2 Encapsulation efficiency The EE is generally defined as the remaining amount of the material encapsulated in polymer electrospun nanofibers compared with the starting amount [75]. Fig. 7.4 shows a sample of nanoencapsulated material. Wen et al. [76] reported that electrospinning can be evaluated to be a novel nanoencapsulation approach for bioactive compounds. For

FIGURE 7.4 A sample of nanoencapsulated material.

Handbook of Chitin and Chitosan

210

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

instance, this efficient strategy can protect unsaturated fatty acids against oxidation when incorporated in foods. Additionally, this approach allows the avoidance of both the loss of nutritional value and the formation of unpleasant off-flavors [76]. Moomand [77] reported that zein nanofibers loaded with fish oil were more oxidatively stable when increasing EE from 91.19% 6 1.09% to 95.88% 6 0.23%. In another study, the EE of fish oil in electrospun nanofibers of 560 nm diameter was found to be 96.9% [78]. The electrospun nanofibers possess different structural and functional advantages, for example, they provide a large surface-to-volume ratio, high EE, and great stability of encapsulated bioactive compounds[76,78]. Ceylan et al. [79] successfully coated fish fillets using nanoencapsulated thymol in chitosan-based nanofibers by using the electrospinning technique. Compared with the control group, fish samples and fish fillets coated only with electrospun chitosan-based nanofiber, the nanoencapsulated thymol provided a shelf life extension for the fish fillets.

7.3.3 Controlled release property of chitosan-based nanomaterial Chitosan has been widely used for the release of bioactive substances [80]. The chitosan concentration and loading material have important roles in the release of the loading material. For example, increasing the chitosan concentration decreased the release of loading material [81]. The stability of vitamin-loaded chitosan nanoparticles could be affected by the applied temperature [82]. In this respect, thermal decomposition of the chitosan-based nanomaterials is also important for the their controlled release property. Ceylan et al. [50] found that the use of loading of thymol into chitosan-based nanofiber protected the vitamin B content of the fish fillets stored in cold storage. Therefore it can be concluded that the controlled release property could not only play a key role in encapsulated materials but also in food products.

7.3.4 Thermal decomposition of chitosan-based nanomaterial Thermogravimetric analysis is mostly used to investigate the thermal stability of nanoscale materials. The first thermal decomposition (50 C 100 C) for electrospun nanofibers from mixed poly(vinyl alcohol)/chitosan solutions (known to be hydrophilic polymers) was attributed to the loss of moisture (about 10% mass) [80]. However, destruction of electrospun chitosan/poly(vinyl alcohol) nanofibers begins at 200 C [81]. In this sense, The TGA thermograms were used to define the content of electrospun PEO/chitosan nanofibers, considering the prospective humidity and

Handbook of Chitin and Chitosan

7.3 Characterization of fabricated nanoscale material(s)

211

residue level in the nanomaterials. The data of the same study revealed that a sharp thermal decomposition of the electrospun nanofiber obtained from PEO/chitosan was observed after 250 C [82]. Kohsari et al. [83] reported that the weight loss in thermal stability of the nanofibers could be delayed due to the loading of different materials into chitosan. Choo et al. [84] also reported that thermal decomposition of chitosan was found to be higher than the group loading polyvinyl alcohol depending on the loading of polyvinyl alcohol to chitosan. In another study, there was a reduction in mass (17%) of liquid smoke-loaded chitosan-based nanofibers and thymol and liquid smoke-loaded chitosan-based nanofibers at temperatures below 147 C, which clearly started to thermally decompose. Moreover, Ceylan et al. [10] determined that pasteurization, sterilization, and cooking temperature of food products are important between 65 C and 150 C. Thus the thermal stability of the nanofibers, especially up to 150 C, should be investigated in further studies that can combine nanotechnology and the cooking process. In order to define the thermal decomposition or stability of the electrospun chitosan-based nanofibers, the diameter of the obtained nanomaterials and the concentration of the polymer can also play key roles in cooked seafood products.

7.3.5 Zeta potential and size of chitosan-based nanomaterial In the electrospinning process, zeta potential (ZP) is one of the most important parameters determining the uniformity and stability of nanomaterials, as ZP has been used for predicting and controlling the stability of colloidal solutions [85]. ZP is a scientific term that describes electrical potential in the interfacial double layer of a dispersed particle. The most known used theory for calculating ZP is that developed by Marian Smoluchowski in 1903. Smoluchowski’s theory is powerful as it is valid for dispersed particles [86]. ZP values give useful information about the stability of colloidal systems and adhesion, surface coating, filtration, lubrication, and corrosion [85]. Losso et al. [87] reported that the surface properties were associated with their half-life stability. Higher and lower ZP (44.4 mV or 245.27) values indicate a high stability [88,89]. Also, higher ZP values indicate bigger repulsion force and dispersion stability against aggregation [10]. Lu et al. [90] reported that the particles in the suspension with high absolute ZP are electrically stabilized and the ZP is affected not only by the properties of nanomaterials, but also by the properties of the medium, such as pH, ionic strength, the concentration of any additives, and temperature. Lu et al. [90] also reported that ZP of chitosan solution and particles, which had different pH values, were found to be between 10 and 50 mV. There was a linear correlation between pH value and ZP. In a previous study, ZP

Handbook of Chitin and Chitosan

212

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

measurements showed the pH dependence of a chitosan nanofiber fabricated by electrospinning [91]. At low pH the ZP of nanoparticles presented higher surface charges than those at high pH [92]. In a study concluded by Liu et al. [92], pH values of chitosan suspensions were adjusted to 4.12, 4.92, 5.5, 6.21, 7.00, and 8.1 to investigate ZP of suspensions. Positive charges on the surface of chitosan increased significantly as the NH2 functional group on chitosan was protonated. SEM images of the particles at pH values of 4.92, 5.50, 7.00, and 8.11 were observed. The original chitosan suspension (pH 6.21) showed uniform nanofiber morphology; at a neutral pH of 7.00, nanofibers exhibited a similar morphology. When the pH value decreased to 5.50, some branched nanofibers were determined. When the pH was reduced to 4.92, the spherical particles with diameters in the range of 100 200 nm displayed a relatively uniform size. The ZP also affects the antimicrobial behavior of electrospun chitosan nanomaterials. For example, Zhao et al. [88] fabricated chitosan/sericin composite nanofibers with diameter between 240 and 380 nm by electrospinning. Antibacterial properties of nanofibers against Gram-negative bacteria (Escherichia coli) and Gram-positive bacteria (Bacillus subtilis) were tested. The ZP of nanofibers were 6.69, 28.9, and 44.4 mV, when the concentrations of the composite nanofibers were 0.1, 0.2, and 0.4 mg/mL, respectively. The antimicrobial activity of the nanofibers against Gramnegative bacteria was better than the Gram-positive bacteria, since the positively charged electrospun nanofibers can easily interact with the negatively charged phospholipid of Gram-negative bacteria. For tissue engineering, materials having a positive ZP have been used, since a positive ZP provides the attachment to anionic cell surfaces, which consequently facilitates cell adhesion on its surface [33].

7.4 Use of electrospun nanomaterials for seafood products safety 7.4.1 Chitosan-based nanofiber coating Electrospun chitosan-based nanofibers affect meat, fish spoilage, and pathogenic bacteria [79,93]. The use of chitosan-based nanofibers as bioactive meat packaging materials could reduce bacterial viability in red meat by 95% [94]. Fish fillets without skin were successfully coated with an electrospun chitosan-based nanofiber by [16,64]. The study results found that the fish fillets coated with electrospun nanofiber (Fig. 7.5) had lower TBA, TVBN, and TMA accumulation as compared to the uncoated (control group) samples during the cold storage.

Handbook of Chitin and Chitosan

7.4 Use of electrospun nanomaterials for seafood products safety

213

FIGURE 7.5 Photographic and SEM images of electrospun chitosan-based nanofibers.

FIGURE 7.6 Flow chart of the fish fillets coated with chitosan-based electrospun nanofiber.

In addition to the limitation of chemical deterioration in fish fillets, Ceylan et al. [50] reported that the use of electrospun chitosan-based nanofibers provided better vitamin B (thiamin, riboflavin, nicotinic acid, nicotinamide, pyridoxal, pyridoxine, pyridoxamine) stability when comparing the control group samples in fish fillets during cold storage. Ceyland et al. [10] also reported that chitosan-based electrospun nanofibers could be used to delay the microbial spoilage (total mesophilic bacteria, yeast and mold, and psychrophilic bacteria growth) for the fish fillets stored at 6 C (Fig. 7.6). In addition to coating fish fillets with electrospun chitosan-based nanofiber, Garcia-Moreno et al. [95] reported that fish oil could be successfully encapsulated, thus the electrospun fibers presented a higher content of hydroperoxides and secondary oxidation products (e.g., 1penten-3-ol, hexanal, octanal, and nonanal) compared with emulsified and unprotected fish oil. There are limited studies related to the use of

Handbook of Chitin and Chitosan

214

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

electrospun chitosan-based nanofiber on seafood product quality. Compared with the micro or microscale material, nanomaterials such as nanofiber can successfully coat the surface of the fish fillets and limit the rapid sensory, physical deterioration in the fish fillets as well. The nanofibers used as a surface coating material can provide a larger contact area on the surface of the fish fillets, so this limits the rapid penetration of oxygen into the surface of the fish fillets. Actually, the limitation of the oxygen penetration with chitosan-based nanofibers can be associated with the limitation of bacterial growth, which can also cause an increase in the rapid chemical, physical, and sensory deterioration in fish meat [96]. All processes related to seafood product safety can be more effectively provided by using cost-effective nanofiber material.

7.4.2 Bio/active material-loaded chitosan-based nanofiber coating Loading of different active or bioactive materials into chitosan-based nanofibers has been applied to improve the antimicrobial or antioxidant activity of the electrospun chitosan-based nanofibers. Recent studies show that active material-loaded chitosan-based nanofibers are used to provide an antimicrobial agent for tissue engineering, drug delivery, wound healing, and wound dressing [30,97,98]. Most of the studies related to loading of the material, such as oil into the chitosan nanofiber, are based on antimicrobial activity (pathogenic bacteria) and antioxidant capacity (DPPH) [99]. There are very limited studies on the quality of seafood products using this approach. In this respect, electrospun chitosan/poly(vinyl alcohol) nanofiber mats, encapsulating fish-purified antioxidant peptides, were successfully fabricated by an electrospinning technique to obtain nanofiber-based bioactive packaging materials [100]. First an active material-loaded chitosan-based nanofibers application devoted to seafood product safety was carried out [79]. In this sense, liquid smoke was successfully loaded into chitosan nanofibers in order to be used as a surface coating material for the fish fillets. The results of the study surprisingly showed that loading of liquid smoke into chitosan enhanced the antimicrobial property of the electrospun chitosanbased nanofiber in the fish fillets. Another study is in agreement that the controlled release property of liquid smoke improved the quality of fish fillets during the cold storage [101]. Kohsari et al. [83] reported that the silver release from nanofiber mats was sharply increased within the first 8 hours for both antimicrobial chitosan polyethylene oxide mats and then the Ag nanoparticles were released slowly. Therefore the release profile affected the bactericidal activities against both Staphylococcus aureus and E. coli. In addition,

Handbook of Chitin and Chitosan

7.4 Use of electrospun nanomaterials for seafood products safety

215

FIGURE 7.7 Coating of fish fillets with electrospun chitosan-based nanofiber.

bacterial effects of the bioactive materials-loaded nanofibers were observed in vitro studies. Ceyland et al. [96] reported that compared with coating with chitosan-based nanofibers (Fig. 7.7), thymol-loaded chitosan-based nanofibers effectively delayed the rapid color changes, sensory deterioration, and the increase in pH value of the fish fillets stored at 4 C. Besides protecting against chemical, physical, sensory deterioration and microbiological spoilage of fish fillets, loading of bioactive material into the chitosan nanofibers can provide better stability in terms of nutritional quality of fish stored at 4 C. Ceyland et al. [64] found that the amounts of lysine, phenylalanine, leucine, isoleucine, methionine, valine, tyrosine, proline, alanine, threonine, histidine, glycine, serine, and glutamic acid were higher in fish fillets coated with thymol-loaded chitosan-based nanofibers than the control group samples after 11 days. EPA (eicosapentaenoic acid) and DHA (docosahexaenoic acid) values of fish fillets coated with nanoencapsulated thymol-loaded chitosanbased nanofibers were found to be more stable when compared with those of uncoated fish fillets during cold storage [102]. There is no doubt that chitosan-based nanofibers can successfully protect the stability of fish fillets. Moreover, the loading of active materials into chitosanbased nanofibers can more effectively protect the stability of seafood products.

7.4.3 Chitosan nanoparticles Chitosan nanoparticles are currently used in various formations as a novel food additive. Khan et al. [103] reported that the particle sizes of the chitosan nanoparticles with antimicrobial potential against foodborne pathogens were found to be 134.3 and 207.9 nm. However, ursolic acid-loaded chitosan-folate nanoparticles were 160 nm in diameter while

Handbook of Chitin and Chitosan

216

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

the average diameter of chitosan folate nanoparticles was found to be 122 nm [104]. Oil-loaded electrospun chitosan nanoparticles embedded gelatin nanofibers were used for food packaging against Listeria monocytogenes and S. aureus on cheese [105]. One of the first studies related to seafood quality and chitosan nanoparticles was reported by Abdou et al. [105]. In this study, chitosan nanoparticle application effectively limited the total bacterial count, coliform bacteria, proteolytic bacteria, and psychrophilic bacteria in fish fingers compared with chitosan application in the fish finger. The reason is that the antimicrobial activity of nanoparticles can increase with decreasing particle size [35]. Tapilatu et al. [106] also reported that nanochitosan treatment could significantly suppress the bacterial activity in the freshly caught fish samples and prolong the storage period. Chitosan nanoparticles provided the effective stability for quality attributes (total volatile nitrogen, trimethylamine, thiobarbituric acid) of fish fingers stored at 218 C [107]. Ceylan [49] reported that chitosan nanoparticles possessing an average diameter of 328.1 nm provided a slower decline in sensory deterioration of fish balls compared with the control group samples (P , .05). TVBN values of control group samples and the fish ball samples treated with chitosan nanoparticles (0.02 g) were found to be 25.20 mg/100 g and 22.12 mg/100 g after 9 days of cold storage (P , .05), respectively. The results of chitosan nanoparticle applications indicated that nanochitosan coating was effective in terms of pH, TVBN, sensory, TBARS values, and bacterial activity in silver carp fillets during refrigerated storage [108]. Ghorabi and Khodanazary [109] also reported that free fatty acids in fish fillets treated with nanochitosan particles indicated an increasing trend, but the total sulfhydryl decreased in the fish fillets during the storage period. The literature studies showed that chitosan nanoparticle applications have recently been used more effectively in order to provide food safety for seafood products.

7.5 Conclusion Seafood products could be preserved using electrospun chitosanbased nanoscale materials. Optimization and characterization processes are very crucial issues to prove whether chitosan-based nanoscale materials are obtained or not. Also, electrospinning parameters such as high voltage should be clearly defined. Morphological properties, EE, thermal stability properties, ZP, and size of the electrospun chitosan-based nanoscale materials play key roles in limiting the microbiological spoilage, and physical and chemical deterioration in seafood, while the nutritional quality of seafood products has been preserved by using nanotechnology applications. Chitosan has already been used as a food

Handbook of Chitin and Chitosan

References

217

additive for the safety of different kinds of aquatic food products, but it is not cost-effective. Thus this nanotechnology application related to chitosan-based nanomaterials provides a cost-effective method for seafood products. Consequently, the use of characterized chitosan-based nanoscale material(s) could be promising for maintaining the safety of seafood products.

References [1] A. Nazir. Modelling and Optimization of Electrospun Materials for Technical Applications. Other. Universite´deHauteAlsace-Mulhouse; National Textile University (Pakistan), 2016. English. NNT: 2016MULH0598 [2] K. Garg and G. L. Bowlin, Electro spinning jets and nanofibrous structures. Biomicrofluidics 5, 13403 (2011) [3] R. Scaˆrlet, L.-R. Manea, B. Cramariuc, Equipment for obtaining nanofibers through the electrospinning system, Int. J. Mod. Manuf. Technol. (2011). ISSN 2067 3604, Vol. IIII. [4] V. Jacobs, A. Patanaik, R.D. Anandjiwala, M. Maaza, Optimization of electrospinning parameters for chitosan nanofibres, Curr. Nanosci. 7 (2011) 396 401. [5] S. Qasim, M. Zafar, S. Najeeb, Z. Khurshid, A. Shah, S. Husain, et al., Electrospinning of chitosan-based solutions for tissue engineering and regenerative medicine, Int. J. Mol. Sci. 19 (2018) 407. [6] N. Amiri, A. Moradi, S.A.S. Tabasi, J. Movaffagh, Modeling and process optimization of electrospinning of chitosan-collagen nanofiber by response surface methodology, Mater. Res. Express 5 (2018) 4. [7] A. Gholipour, S.H. Bahrami, M. Nouri, Nanofibers from chitosan-poly vinyl alcohol blend and their biological and antimicrobial properties and process optimization using response surface methodology, VDI Berichte 2027 (2008) 199 203. [8] M. Kouchak, M. Avadi, M. Abbaspour, A. Jahangiri, S. Kargar Boldaji, Effect of different molecular weights of chitosan onpreparation and characterization of insulin loaded nanoparticles by ion gelation method, Int. J. Drug. Dev. Res. 4 (2) (2012) 271 277. [9] A. Hebeish, S. Sharaf, A. Farouk, Utilization of chitosan nanoparticles as a green finish in multifunctionalization of cotton textile, Int. J. Biol. Macromol. 60 (2013) 10 17. [10] N.A. Qinna, Q.G. Karwi, N. Al-Jabour, M.A. Al-Remawi, T.M. Alhussainy, K.A. AlSo’ud, et al., Influence of molecular weight and degree of deacetylation of low molecular weight chitosan on the bioactivity of oral insulin preparations, Mar. Drugs 13 (2015) 1710 1725. [11] S. Mengistu Lemma, F. Bossard, M. Rinaudo, Preparation of pure and stable chitosan nanofibers by electrospinning in the presence of poly (ethylene oxide), Int. J. Mol. Sci. 17 (2016) 1 16. [12] Z. Ceylan, G.F.U. Sengor, M.T. Yilmaz, Nanoencapsulation of liquid smoke/thymol combination in chitosan nanofibers to delay microbiological spoilage of sea bass (Dicentrarchus labrax) fillets, J. Food Eng. 229 (2018) 43 49. [13] A. Baji, Y. Mai, S. Wong, M. Abtahi, P. Chen, Electrospinning of polymer nanofibers: effects on oriented morphology, structures and tensile properties, Compos. Sci. Technol. 70 (2010) 703 718. [14] R. Sedghi, A. Shaabani, Z. Mohammadi, Z.Y. Samadi, F. Isae, Biocompatible electrospinning chitosan nanofibers: a novel delivery system with superior local cancer therapy, Carbohydr. Polym. 159 (2017) 1 10.

Handbook of Chitin and Chitosan

218

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

[15] Y.T. Jia, J. Gong, X.H. Gu, H.Y. Kim, J. Dong, X.Y. Shen, Fabrication and characterization of poly (vinyl alcohol)/chitosan blend nanofibers produced by electrospinning method, Carbohydr. Polym. 67 (2007) 403 409. [16] N. Bhattarai, D. Edmondson, O. Veiseh, F.A. Matsen, M. Zhang, Electrospun chitosan-based nanofibers and their cellular compatibility, Biomaterials 26 (2005) 6176 6184. [17] H. Susanto, M. Faz, M. Rani, M. Robbani, The Influence of photoinitiator on the characteristics of electrospun poly(vinyl alcohol)/chitosan based nanofibers, in: IOP Conference Series: Materials Science and Engineering 367, 2018, pp. 1 6. [18] Z. Ceylan, G.F.U. Sengor, M.T. Yilmaz, A novel approach to limit chemical deterioration of gilthead sea bream (Sparus aurata) fillets: coating with electrospun nanofibers as characterized by molecular, thermal, and microstructural properties, J. Food Sci. 82 (2017) 1163 1170. [19] K. Ohkawa, D. Cha, H. Kim, A. Nishida, H. Yamamoto, Electrospinning of chitosan, Macromol. Rapid Commun. 25 (18) (2004) 1600 1605. [20] L. Van der Schueren, I. Steyaert, B. De Schoenmaker, K. De Clerck, Polycaprolactone/chitosan blend nanofibres electrospun from an acetic acid/formic acid solvent system, Carbohydr. Polym. 88 (2012) 1221 1226. [21] B. Duan, C. Dong, X. Yuan, K. Yao, Electrospinning of chitosan solutions in acetic acid with poly(ethylene oxide), J. Biomater. Sci. Polym. Ed. 15 (6) (2004) 797 811. [22] B.K. Gu, S.J. Park, M.S. Kim, Y.J. Lee, J.I. Kim, C.H. Kim, Gelatin blending and sonication of chitosan nanofiber mats produce synergistic effects on hemostatic functions, Int. J. Biol. Macromol. 82 (2016) 89 96. [23] Z.G. Chen, P.W. Wang, B. Wei, X.M. Mo, F.Z. Cui, Electrospun collagen chitosan nanofiber: a biomimetic extracellular matrix for endothelial cell and smooth muscle cell, Acta Biomater. 6 (2010) 372 382. [24] V. Sencadas, D.M. Correia, A. Areias, G. Botelho, A.M. Fonseca, I.C. Neves, et al., Determination of the parameters affecting electrospun chitosan fiber size distribution and morphology, Carbohydr. Polym. 87 (2) (2012) 1295 1301. [25] C.A. Vega-Ca´zarez, D.I. Sa´nchez-Machado, J. Lo´pez-Cervantes, Chitin-chitosanmyriad functionalities in science and technology. In: Overview of electrospinned chitosan nanofiber composites for wound dressings, 2018, pp. 157 182 (Chapter 9). [26] S. De Vrieze, P. Westbroek, T. Van Camp, L. Van Langenhove, Electrospinning of chitosan nanofibrous structures: feasibility study, J. Mater. Sci. 42 (19) (2007) 8029 8034. [27] M. Pakravan, M.C. Heuzey, A.A. Ajji, Fundamental study of chitosan/PEO electrospinning, Polymer 52 (21) (2011) 4813 4824. [28] P. Sangsanoh, O. Suwantong, A. Neamnark, P. Cheepsunthorn, P. Pavasant, P. Supaphol, In vitro biocompatibility of electrospun and solvent-cast chitosan substrata towards schwann, osteoblast, keratinocyte and fibroblast cells, Eur. Polym. J. 46 (2010) 428 440. [29] B.K. Gu, S.J. Park, M.S. Kim, C.M. Kang, J.I. Kim, C.H. Kim, Fabrication of sonicated chitosan nanofiber mat with enlarged porosity for use as hemostatic materials, Carbohydr. Polym. 97 (1) (2013) 65 73. [30] B.M. Min, S.W. Lee, J.N. Lim, Y. You, T.S. Lee, P.H. Kang, Chitin and chitosan nanofibers: electrospinning of chitin and deacetylation of chitin nanofibers, Polymer 45 (2004) 7137 7142. [31] F. Chen, X. Li, X. Mo, C. He, H. Wang, Y. Ikada, Electrospun chitosan-P(LLA-CL) nanofibers for biomimetic extracellular matrix, J. Biomater. Sci. Polym. Ed. 19 (5) (2008) 677 691. [32] K. Sun, Z.H. Li, Preparations, properties and applications of chitosan based nanofibers fabricated by electrospinning, Express Polym. Lett. 5 (4) (2011) 342 361.

Handbook of Chitin and Chitosan

References

219

[33] X. Geng, O. Kwon, J. Jang, Electrospinning of chitosan dissolved in concentrated acetic acid solution, Biomaterials 26 (2005) 5427 5432. [34] C. Kriegel, K.M. Kit, D.J. McClements, J. Weiss, Electrospinning of chitosan-poly (ethylene oxide) blend nanofibers in the presence of micellar surfactant solutions, Polymer 50 (2009) 189 200. [35] H. Chen, J. Huang, J. Yu, S. Liu, P. Gu, Electrospun chitosan-graft-poly (ε-caprolactone)/poly (ε-caprolactone) cationic nanofibrous mats as potential scaffolds for skin tissue engineering, Int. J. Biol. Macromol. 48 (2011) 13 19. [36] K. Desai, K. Kit, J. Li, S. Zivanovic, Morphological and surface properties of electrospun chitosan nanofibers, Biomacromolecules 9 (2008) 1000 1006. [37] Y. Zhang, X. Huang, B. Duan, L. Wu, S. Li, X. Yuan, Preparation of electrospun chitosan/poly(vinyl alcohol) membranes, Colloid Polym. Sci. 285 (2007) 855 863. [38] S.A. Majd, M.R. Khorasgani, S.J. Moshtaghian, A. Talebi, M. Khezri, Application of chitosan/PVA nano fiber as a potential wound dressing for streptozotocin-induced diabetic rats, Int. J. Biol. Macromol. 92 (2016) 1162 1168. [39] T.T.T. Nguyen, O.H. Chung, J.S. Park, Coaxial electrospun poly(lactic acid)/chitosan (core/shell) composite nanofibers and their antibacterial activity, Carbohydr. Polym. 86 (4) (2011) 1799 1806. [40] S. Torres-Giner, E. Gimenez, J.M. Lagaro´n, Characterization of the morphology and thermal properties of zein prolamine nanostructures obtained by electrospinning, Food Hydrocoll. 222 (4) (2008) 601 614. [41] X. Zong, K. Kim, D. Fang, S. Ran, B.S. Hsiao, B. Cgu, Structure and process relationship of electrospun bioabsobable nanofiber membranes, Polymer 43 (2002) 4403 4412. [42] P. Su, C. Wang, X. Yang, X. Chen, C. Gao, X. Feng, et al., Electrospinning of chitosan nanofibers: the favorable effect of metal ions, Carbohydr. Polym. 84 (2011) 239 246. [43] Z. Moridi, V. Mottaghitalab, A.K. Haghi, Use of electrospinning technique in production of chıtosan/carbon nanotubes, Cellulose Chem. Technol. 46 (9-10) (2011) 529 532. [44] D. Terada, H. Kobayashi, K. Zhang, A. Tiwari, C. Yoshikawa, N. Hanagata, Transient charge-masking effect of applied voltage on electrospinning of pure chitosan nanofibers from aqueous solutions, Sci. Technol. Adv. Mater. 13 (1) (2012) 1 9. [45] M.T. Bizarria, M.A. Davila, L.H.I. Mei, Non-woven nanofiber chitosan/PEO membranes obtained by electrospinning, Braz. J. Chem. Eng. 31 (1) (2014) 57 68. [46] S. Teepoo, P. Dawan, N. Barnthip, Electrospun chitosan-gelatin biopolymer composite nanofibers for horseradish peroxidase immobilization in a hydrogen peroxide biosensor, Biosensensor 7 (2017) 47. [47] H. Homayoni, S.A.H. Ravandi, M. Valizadeh, Electrospinning of chitosan nanofibers: processing optimization, Carbohydr. Polym. 77 (2009) 656 661. [48] A.R. Jabur, L.K. Abbas, S.M. Muhi Aldain, Effects of ambient temperature and needle to collector distance on pva nanofibers diameter obtained from electrospinning technique, Part. (A) Eng. 35 (4) (2017) 340 347. [49] M. Aliabadi, M. Irani, J. Ismaeili, S. Najafzadeh, Design and evaluation of chitosan/ hydroxyapatite composite nanofiber membrane for the removal of heavy metal ions from aqueous solution, J. Taiwan. Inst. Chem. Eng. 45 (2014) 518 526. [50] K.P. Matabola, R.M. Moutloali, The influence of electrospinning parameters on the morphology and diameter of poly(vinyledene fluoride) nanofibers-effect of sodium chloride, J. Mater. Sci. 48 (16) (2013) 5475 5482. [51] Z. Ceylan, Use of characterized chitosan nanoparticles integrated in poly(vinyl alcohol) nanofibers as an alternative nanoscale material for fish balls, J. Food Saf. 38 (2018) 1 5. [52] Z. Ceylan, M. Yaman, O. Sagdic, E. Karabulut, M.T. Yilmaz, Effect of electrospun thymol-loaded nanofiber coating on vitamin B profile of gilthead sea bream fillets (Sparus aurata), LWT-Food Sci. Technol. 98 (2018) 162 169.

Handbook of Chitin and Chitosan

220

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

[53] B. Dhandayuthapani, U.M. Krishnan, S. Sethuraman, Fabrication and characterization of chitosan-gelatin blend nanofibers for skin tissue engineering, J. Biomed. Mater. Res. Part. B Appl. Biomater. 94 (2010) 264 272. [54] S. Zargham, S. Bazgir, A. Tavakoli, A.S. Rashidi, R. Damerchely, The effect of flow rate on morphology and deposition area of electrospun nylon 6 nanofiber, J. Eng. Fibers Fabr. 7 (2012) 42 49. [55] M. Chowdhury, G. Stylios, Effect of experimental parameters on the morphology of electrospun Nylon 6 fibres, Int. J. Basic. Appl. Sci. 10 (06) (2010) 1609 1624. [56] M. Aliabadi, M. Irani, J. Ismaeili, H. Piri, M.J. Parnian, Electrospun nanofiber membrane of PEO/Chitosan for the adsorption of nickel, cadmium, lead and copper ions from aqueous solution, Chem. Eng. J. 220 (2013) 237 243. [57] S. De Vrieze, T. Van Camp, A. Nelvig, B. Hagstro¨m, P. Westbroek, K. De Clerck, The effect of temperature and humidity on electrospinning, J. Mater. Sci. 44 (2009) 1357 1362. [58] R.M. Nezarati, M.B. Eifert, E. Cosgriff-Hernandez, Effects of humidity and solution viscosity on electrospun fiber morphology, Tissue Eng. Part. C. 19 (2013) 1 10. [59] C.E.G. Garcia, F.A.S. Martı´nez, F. Bossard, M. Rinaudo, Biomaterials based on electrospun chitosan, Relat. Process. Cond. Mech. Prop. Polym. 10 (3) (2018) 257. [60] M. Schreiber, S. Vivekanandhan, P. Cooke, M. Misra, Electrospun green fibres from lignin and chitosan: a novel polycomplexation process for the production of ligninbased fibres, J. Mater. Sci. 49 (2014) 7949 7958. [61] A. Theron, E. Zussman, A.L. Yarin, Electrostatic field-assisted alignment of electrospun nanofibres, Nanotechnology 12 (2001) 384 390. [62] W.E. Teo, S. Ramakrishna, A review on electrospinning design and nanofibre assemblies, Nanotechnology 17 (2006) 89 106. [63] A. Valizadeh, S. Mussa Farkhani, Electrospinning and electrospun nanofibres, IET Nanobiotechnol. 8 (2014) 83 92. [64] D. Rodoplu, M. Mutlu, Effects of electrospinning setup and process parameters on nanofiber morphology intended for the modification of quartz crystal microbalance surfaces, J. Eng. Fibers Fabr. 2 (2012) 118 123. [65] T.T. Yuan, A.M. Di George Foushee, M.C. Johnson, J.M. Stahl, Development of electrospun chitosan-polyethylene oxide/fibrinogen biocomposite for potential wound healing applications, Nanoscale Res. Lett. 13 (1) (2018) 88. [66] Z. Ceylan, M.T. Yilmaz, G.F. S¸ engo¨r, Effect of coating with bioactive material-loaded biopolymer-based electrospun nanofibers on amino acid composition of gilthead sea bream fillets (Sparus aurata). European biotechnology congress, Dubrovnik, Croatia, May 25 27, 2017. [67] G. Singh, C. Stephan, P. Westerhoff, D. Carlander, T.V. Duncan, Measurement methods to detect, characterize, and quantify engineered nanomaterials in foods, Compr. Rev. Food Sci. Food Saf. 13 (2014) 693 704. [68] D. Gupta, D. Singh, N.C. Kothiyal, A.K. Saini, V.P. Singh, D. Pathania, Synthesis of chitosan-g-poly(acrylamide)/ZnS nanocomposite forcontrolled drug delivery and antimicrobial activity, Int. J. Biol. Macromol. 74 (2015) 547 557. [69] Y. Li, F. Chen, J. Nie, D. Yang, Electrospun poly (lactic acid)/chitosan core shell structure nanofibers from homogeneous solution, Carbohydr. Polym. 90 (2012) 1445 1451. [70] V. Klang, N.B. Matsko, C. Valenta, F. Hofer, Electron microscopy of nanoemulsions: an essential tool for characterisation and stability assessment, Micron 43 (2 3) (2012) 85 103. [71] J. Han, J. Zhang, R. Yin, G. Ma, D. Yang, J. Nie, Electrospinning of methoxy poly (ethylene glycol)-grafted chitosan and poly(ethylene oxide) blend aqueous solution, Carbohydr. Polym. 83 (2011) 270 276.

Handbook of Chitin and Chitosan

References

221

[72] S.J. Lee, D.N. Heo, J.H. Moon, W.K. Ko, J.B. Lee, M.S. Bae, et al., Electrospun chitosan nanofibers with controlled levels of silver nanoparticles. Preparation, characterization and antibacterial activity, Carbohydr. Polym. 111 (2014) 530 537. [73] P. Heera, S. Shanmugams, Nanoparticle characterization and application: an overview, Int. J. Curr. Microbiol. Appl. Sci. 4 (2015) 379 386. [74] R. Nirmala, R. Navamathavan, M.H. El-Newehy, H.Y. Kim, Preparation and electrical characterization of polyamide-6/chitosan composite nanofibers via electrospinning, Mater. Lett. 65 (2011) 493 496. [75] S. Surucu, H.T. Sasmazel, Development of core-shell coaxially electrospun composite PCL/chitosan scaffolds, Int. J. Biol. Macromol. 92 (2016) 321 328. [76] P. Wen, M.H. Zong, R.J. Linhardt, K. Feng, H. Wu, Electrospinning: a novel nanoencapsulation approach for bioactive compounds, Trends Food Sci. Technol. (2017). [77] J. Song, J.J.A. Remmers, J. Shao, E. Kolwijck, X.F. Walboomers, J.A. Jansen, et al., Antibacterial effects of electrospun chitosan/poly(ethylene oxide) nanofibrous membranes loaded with chlorhexidine and silver, Nanomed. Nanotechnol. Biol. Med. 12 (2016) 1357 1364. [78] H. Yang, P. Wen, K. Feng, M.H. Zong, W.Y. Lou, H. Wu, Encapsulation of fish oil in a coaxial electrospun nanofibrous mat and its properties, RSC Adv. 7 (24) (2017) 14939 14946. [79] A.A. Nada, A.G. Hassabo, A. Mohamed, S. Zaghloul, Encapsulation of nicotinamide into cellulose based electrospun fibers, J. Appl. Pharm. Sci. 6 (08) (2016) 013 021. [80] C.L. Vela´squez, Chitosan-based nanomaterials on controlled bioactive agents delivery: a review, J. Anal. Pharm Res. 7 (4) (2018) 484 489. [81] Mansouri, M., Khorram, M., Samimi, A. and Osfouri, S., Preparation of bovine serum albumin loaded chitosan nanoparticles using reverse micelle method. http://www. chisa.cz/2010/admin/contrib_get_abstract.asp?id_02=942. [82] Jang, Keum-Il, and H.G. Lee. "Stability of chitosan nanoparticles for L-ascorbic acid during heat treatment in aqueous solution." J. Agric. Food Chem. 56.6 (2008): 1936 1941. [83] P.J. Garcı´a Moreno, K. Boutrup Stephansen, J. Van der Kruijs, A. Guadix, E.M. Guadix, I.S. Chronakis, et al., Encapsulation of fish oil in nanofibers by emulsion electrospinning: physical characterization and oxidative stability, J. Food Eng. 183 (2016) 39 49. [84] K. Moomand, L. Lim, Oxidative stability of encapsulated fish oil in electrospun zein fibres, Food Res. Int. 62 (1) (2014) 523 532. [85] Q. Ma, M. Pyda, B. Mao, P. Cebe, Relationship between the rigid amorphous phase and mesophase in electrospun fibers, Polymer 54 (2013) 2544 2554. [86] Z. Ceylan, M.T. Yilmaz, G.F. S¸ engo¨r, Microbiological stability of sea bass (Dicentrarchus labrax) fillets coated with chitosan based liquid smoke loaded electrospun nanofibers. European Biotechnology Congress, Riga, Latvia, May 5 7, 2016. [87] R.P. Gonc¸alves, W.H. Ferreira, R.F. Gouveˆa, C.T. Andrade, Effect of chitosan on the properties of electrospun fibers from mixed poly(vinyl alcohol)/chitosan solutions, Mater. Res. 20 (4) (2017) 984 993. [88] A.Y. Abdolmaleki, H. Zilouei, S.N. Khorasani, Characterization of electrospinning parameters of chitosan/poly(vinyl alcohol) nanofibers to remove phenol via response surface methodology, Polymer Sci. 4 (2018) 1 5. [89] L.P. Mehdi, Production of chitosan-based non-woven membranes using the electrospinning process. ProQuest Dissertations and Theses, Thesis (Ph.D.)—Ecole Polytechnique, Montreal, Canada, 2012, 152 p. [90] I. Kohsari, Z. Shariatinia, S.M. Pourmortazavi, Antibacterial electrospun chitosan polyethylene oxide nanocomposite mats containing bioactive silver nanoparticles, Carbohydr. Polym. 140 (2016) 287 298.

Handbook of Chitin and Chitosan

222

7. Importance of electrospun chitosan-based nanoscale materials for seafood products safety

[91] K. Choo, Y.C. Ching, C.H. Chuah, J. Sabariah, N.S. Liou, Preparation and characterization of polyvinyl alcohol-chitosan composite films reinforced with cellulose nanofiber, Materials 9 (2016) 644. [92] D. Cho, S. Lee, M.W. Frey, Characterizing zeta potential of functional nanofibers in a microfluidic device, J. Colloid Interf. Sci. 372 (2012) 252 260. [93] B.J. Kirby, E.H. Hasselbrin, Zeta potential of microfluidic substrates: 1. theory, experimental techniques, and effects on separations, Electrophoresis 25 (2002) 187 2002. [94] J.N. Losso, A. Khachatryan, M. Ogawa, J.S. Godber, F. Shih, Random centroid optimization of phosphatidylglycerol stabilized lutein-enriched oil-in-water emulsions at acidic pH, Food Chem. 92 (4) (2005) 737 744. [95] R. Zhao, X. Li, B. Sun, Y. Zhang, D. Zhang, Z. Tang, et al., Electrospun chitosan/ sericin composite nanofibers with antibacterial property as potential wound dressings, Int. J. Biol. Macromol. 68 (2014) 92 97. [96] X. Zhuang, B. Cheng, W. Kang, X. Xu, Electrospun chitosan/gelatin nanofibers containing silver nanoparticles, Carbohydr. Polym. 82 (2010) 524 527. [97] G.W. Lu, P. Gao, Emulsions and microemulsions for topical and transdermal drug delivery (Chapter 3). Non-invasive and minimally-invasive drug delivery systems for pharmaceutical and personal care products, Personal. Care Cosmet. Technol. (2010) 59 94. [98] H. Matsumoto, H. Yako, M. Minagawa, A. Tanioka, Characterization of chitosan nanofiber fabric by electrospray deposition: electrokinetic and adsorption behavior, J. Colloid Interf. Sci. 310 (2007) 678 681. [99] D. Liu, P.R. Chang, M. Chen, Q. Wu, Chitosan colloidal suspension composed of mechanically disassembled nanofibers, J. Colloid Interf. Sci. 354 (2011) 637 643. [100] M. Arkoun, F. Daigle, M.C. Heuzey, A. Ajji, Mechanism of action of electrospun chitosan-based nanofibers against meat spoilage and pathogenic bacteria, Molecule 22 (2017) 85. [101] M. Arkoun, F. Daigle, R.A. Holley, M.C. Heuzey, A. Ajji, Chitosan-based nanofibers as bioactive meat packaging materials, Packag. Technol. Sci. 31 (2018) 185 195. [102] P.J. Garcı´a-Moreno, K. Stephansen, J. Van der Kruijs, A. Guadix, E.M. Guadix, I.S. Chronakis, et al., Encapsulation of fish oil in nanofibers by emulsion electrospinning: physical characterization and oxidative stability, J. Food Eng. 183 (2016) 39 49. [103] Z. Ceylan, G.F.U. Sengor, A. Basahel, M.T. Yılmaz, Determination of quality parameters of gilthead sea bream (Sparus aurata) fillets coated with electrospun nanofibers, J. Food Saf. 38 (2018) 1 7. [104] N. Charernsriwilaiwat, T. Rojanarata, T. Ngawhirunpat, M. Sukma, P. Opanasopit, Electrospun chitosan-based nanofiber mats loaded with Garcinia mangostana extracts, J. Pharm. 452 (2013) 333 343. [105] L. Lin, Y. Gu, H. Cui, Moringa oil/chitosan nanoparticles embedded gelatin nanofibers for food packaging against Listeria monocytogenes and Staphylococcus aureus on cheese, Food Packag. Shelf Life. 19 (2019) 86 93. [106] E. Mirzaei, S. Sarkar, M. Rezayat, R. Faridi-Majidi, Herbal extract loaded chitosanbased nanofibers as a potential wound-dressing, J. Adv. Med. Sci. Appl. Technol. (JAMSAT) 2 (1) (2016) 141. [107] B. Munteanu, L. Sacarescu, A.L. Vasiliu, G. Hitruc, G. Pricope, M. Sivertsvik, et al., Antioxidant/antibacterial electrospun nanocoatings applied onto PLA films, Materials 11 (2018) 1973. [108] S.L. Hosseini, Z. Nahvi, M. Zandi, Antioxidant peptide-loaded electrospun chitosan/poly(vinyl alcohol) nanofibrous mat intended for food biopackaging purposes, Food Hydrocoll. 89 (2019) 637 648.

Handbook of Chitin and Chitosan

References

223

[109] Z. Ceylan, G.F.U. Sengor, O. Sagdic, M.T. Yilmaz, A novel approach to extend microbiological stability of sea bass (Dicentrarchus labrax) fillets coated with electrospun chitosan nanofibers, LWT—Food Sci. Technol. 79 (2017) 367 375. [110] Z. Ceylan, G.F. S¸ engo¨r, R. Meral, M.T. Yılmaz, Effect of coating with thymol-loaded chitosan-based electrospun nanofibers on fatty acid composition of gilthead sea bream fillets (Sparus aurata). European Biotechnology Congress 2018, Athens, Greece, April 26 28, 2018, p. 280. [111] I. Khan, C.N. Tango, S. Miskeen, D.-H. Oh, Evaluation of nisin-loaded chitosanmonomethyl fumaric acid nanoparticles as a direct food additive, Carbohydr. Polym. 184 (2018) 100 107. [112] H. Jin, J. Pi, F. Yang, J. Jiang, X. Wang, H. Bai, Folate-chitosan nanoparticles loaded with ursolic acid confer anti-breast cancer activities in vitro and in vivo, Sci. Rep. 6 (2016) 30782. [113] E.S. Abdou, A.S. Osheba, M.A. Sorour, Effect of chitosan and chitosan-nanoparticles as active coating on microbiological characteristics of fish fingers, Int. J. Appl. Sci. Technol. 2 (2012) 158 169. [114] Y. Tapilatu, P.S. Nugraheni, T. Ginzel, M. Latumahina, G.V. Limmon, W. Budhijanto, Nano-chitosan utilization for fresh yellowfin tuna preservation, Aquat. Procedia 7 (2016) 285 295. [115] A. Osheba, M. Sorour, E.S. Abdou, Effect of chitosan nanoparticles as active coating on chemical quality and oil uptake of fish fingers, J. Agri. Environ. Sci. 2 (1) (2013) 01 14. [116] M. Zarei, Z. Ramezani, S. Ein-Tavasoly, M. Chadorbaf, Coating effects of orange and pomegranate peel extracts combined with chitosan nanoparticles on the quality of refrigerated silver carp fillets, J. Food Process. Pres. 39 (6) (2015) 2180 2187. [117] R.S. Ghorabi, A. Khodanazary, Effects of chitosan and nano-chitosan as coating materials on the quality properties of large scale tongue sole Cynoglossus arel during super-chilling storage, Iran. J. Fish. Sci. (2018). Available from: https://doi.org/ 10.22092/ijfs.2018.119689.

Handbook of Chitin and Chitosan

C H A P T E R

8 Alternative methods for chitin and chitosan preparation, characterization, and application George M. Halla, Claudia H. Barrerab and Keiko Shiraib a

Centre for Sustainable Development, University of Central Lancashire, Preston, United Kingdom, bBiotechnology Department, Laboratory of Biopolymers and Pilot Plant of Bioprocessing of Agro-Industrial and Food By-Products, Autonomous Metropolitan University, Mexico City, Mexico O U T L I N E 8.1 Introduction

226

8.2 Chitin production 226 8.2.1 Synthesis of chitin and enzymes involved in the main commercial sources 227 8.2.2 Raw materials: fungi and insects 229 8.3 Current chitosan production 8.3.1 Alternative methods of production 8.3.2 Quality control of chitin production

231 233 237

8.4 Conclusions

240

References

241

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00008-0

225

© 2020 Elsevier Inc. All rights reserved.

226

8. Alternative methods for chitin and chitosan preparation, characterization, and application

8.1 Introduction The global biomass composition of the biosphere has been estimated to be c. 550 gigatons of carbon (Gt C 5 1015 g of carbon) from the data analysis of hundreds of studies, in which the third major component of biomass is fungi (12 Gt C) and on the penultimate taxon place is animals, in which the terrestrial arthropods represent 0.2 Gt C and the marine arthopods 1 Gt C [1]. Both taxa are composed of chitin as the structural polysaccharide and thus chitin has been claimed as being abundant in nature. In aquatic environments, arthropods are the dominant chitin-producing organisms with c. 2.8 3 107 tons/year in freshwater ecosystems and 1.3 3 109 tons/year in marine ecosystems. In terrestrial environments, the majority of chitin is produced by fungi due to their role in the biogeochemical cycling of carbon and nitrogen [2]. The lower invertebrates such as sponges, coelenterates, nematodes, and mollusks, as well as many microbes, including protists and algae, also produce chitin. In the case of vertebrates, the scales and gut of fish and amphibians contain chitin [3]. The arthropods represent the largest pool of animal biomass today. According to The State of World Fisheries and Aquaculture 2018 published by the Food and Agriculture Organization of the United Nations [4], the total global aquaculture production in 2016, which included both food fish and aquatic plants, was 110.2 million tons. Out of that figure, 80.0 million tons are food fish, of which 7.9 million ton are crustaceans that consist mostly of Whiteleg shrimp (Litopenaeus vannamei), Red swamp crawfish (Procambarus clarkia), Chinese mitten crab (Eriocheir sinensis), Giant tiger prawn (Litopenaeus monodon), Oriental river prawn (Macrobrachium nipponense), and Giant river prawn (Macrobrachium rosenbergii). Similar to other types of food fish, the production of crustaceans depends on inland aquaculture, as it produced 3.03 million tons (live weight) of crustaceans worldwide. Nonetheless, marine and coastal aquaculture are still responsible for the production of 4.8 million tons. From this production, huge amounts of heads and exoskeletons are discarded and very little is utilized as raw matter for chitin extraction. Additionally, in this regard, chitin has also been extracted from fish, but in a very low yield [3]; therefore, chitin from fisheries waste might be an emerging area of scientific and technological opportunity.

8.2 Chitin production Chitin is a linear copolymer of β-(1-4)-linked D-glucosamine and N-acetyl D-glucosamine units in various proportions and sequences.

Handbook of Chitin and Chitosan

8.2 Chitin production

227

Chitin displays highly ordered crystalline structures comprising three allomorphs designated as α, β, and γ [5]. The most abundant allomorph is α-chitin, found in arthropods and fungi, and it is characterized by the antiparallel arrangement of chains with strong hydrogen bonding. In β-chitin, found in cephalopods, the chains are in parallel in the same direction and there is less strong interaction in the network. Less abundant is γ-chitin, present in arthropods’ cocoons and larval peritrophic membranes. It is a combination of the α and β allomorphs, and it presents similar characteristics to α-chitin [6].

8.2.1 Synthesis of chitin and enzymes involved in the main commercial sources Chitin is a structural component of the cell walls of fungi, including Zygomycetes, Ascomycetes, Basidiomycetes, and Deuteromycetes. For almost all eumycota fungi, the cell wall is a branched glucan linked to chitin via a β-1,4 bond [7]. The microfibrils made of α-chitin compensate for the turgor pressure in the inner cell wall [8]. Chitosan is also found naturally in some fungi like Zygomycetes [9]. There is a common biosynthetic pathway for chitin in insects and fungi that employs glucose and its storage compounds, glycogen or trehalose, as sugar substrates. The pathway consists of the formation of the amino sugar N-acetylglucosamine and the activated amino sugar UDP-N-acetylglucosamine by a variant of the Leloir pathway, followed by the polymerization of chitin using UDP-N-acetylglucosamine. The formation of N-acetylglucosamine and UDP-N-acetylglucosamine occurs in the cytoplasm and chitin synthesis occurs in the plasma membrane. The enzymes glutamine-fructose-6-phosphate amidotransferase (EC 2.6.1.16), UDP-Nacetylglucosamine pyrophosphorylase (EC 2.7.7.23), and chitin synthase (EC 2.4.1.16) are highly regulated and limited by the rate of chitin production [8]. Most fungi have multiple chitin synthases that produce chitin at different sites on the cell surface during growth, and as a response of environmental stress to the cell wall [10]. The chitin synthases are integral membrane proteins that are highly conserved that belong to Family 2 of processive polysaccharide synthases. These enzymes are responsible for the synthesis of linear chains of β1,4-N-acetylglucosamine from the substrate UDP-N-acetylglucosamine to the nonreducing end of a growing glycan chain. The glycan chain is concomitantly released from the extracytoplasmic end of the translocation channel, and then once a primer, such as N-acetyl-D-glucosamine, is present, the polymerization of a new chain starts at the catalytic site. The crystalline chitins can be assembled from many relatively short chains that double back on themselves, or

Handbook of Chitin and Chitosan

228

8. Alternative methods for chitin and chitosan preparation, characterization, and application

are reoriented for the formation of antiparallel hydrogen bonding networks. The formation of structural microfibrils contributes to cell wall organization as parts of the chitin β-glucan networks are generated by the action of transglycosylases that cleave chitin chains internally, then transfer the newly generated reducing end to a β-glucan acceptor [11]. The synthesis of fungal chitosan is also the result of chitin synthase and chitin deacetylase (EC 3.5.1.41) activity in a complex synergistic process. Chitin deacetylase cleaves acetic groups from N-acetylglucosamine to produce glucosamine residues in the polymeric chain [12]. Crustaceans are aquatic arthropods, which are a widespread and diverse group that include lobsters, shrimps, and crabs; and isopods, copepods, barnacles, and euphausiids. Crustaceans play several roles in their ecological niches as predators, parasites, and grazers, or as intermediary links in the food chain [13]. The distinctive characteristic of the arthropods as invertebrates is the exoskeleton, which consists of layers: the epi-, exo-, and endocuticles. The cuticle is mineralized with calcium carbonate, and the inner membranous three layers are formed from chitin protein fibrils. There is a hierarchical organization with several structural levels. The molecular level is composed of chitin in antiparallel fashion forming α-chitin crystals. The next structure level is the arrangement of chitin in nanofibrils that are wrapped by proteins [14]. The cluster of chitin nanofibrils and proteins are arranged in a parallel lamellae in a twisted plywood structure or Bouligand pattern that has a constantly changing orientation from layer to layer. The exocuticle is characterized by a very fine woven structure of the chitin protein matrix with high stiffness and hardness, while the endocuticle displays a coarser twisted plywood structure with lower stiffness and hardness than the exocuticle. The exoskeleton is a nanocomposite material with remarkable mechanical resistance that confers protection, support, rigidity, and impermeability to the body [15]. Chitin is also present in the mucous extracellular matrices, such as peritrophic membranes, functioning as protective linings in the intestinal tract. The growth of crustaceans is accompanied by a molting or ecdysis cycle, in which biomineralization of chitinous structures is carried out by degradation and synthesis [16]. The enzyme that gathers monomers of N-acetyl glucosamine into chitin is called chitin synthase (E.C. 2.4.1.16). This is a cell membrane bound enzyme composed of two subunits. Once that chitin is polymerized, it is deposited in the extracellular space. Chitin synthesis is concerned with degradation, which is carried out by chitinases (E.C. 3.2.1.14). Chitinases rupture the glycosidic bond in the middle of the polysaccharide chain to release oligomers of N-acetyl glucosamine. Chitinases are induced prior to molting in the integument of crustaceans and are also present in the hepatopancreas for the digestion of chitinous food [17].

Handbook of Chitin and Chitosan

8.2 Chitin production

229

8.2.2 Raw materials: fungi and insects 8.2.2.1 Fungi Fungi have been claimed to be important alternatives to obtain chitin and chitosan owing to their more facile extraction than crustacean waste, there is no dependence on seasonal variation and production, waste fungal biomass can be utilized, and the products show lower viscosities, molecular weights, and acetylation degree than those obtained by crustacean sources [18]. Chitin/chitosan production can be via bioprocesses that utilize waste fungal biomass [19] or those with the specific purpose to produce chitin. In this regard, chitin has been purified from several fungi such as the Basidiomycete Agaricus bisporus mushroom; three main fruit body parts, that is, pileus, stipes, and gills were employed for alkaline treatment. The extraction yield of chitin was 7.4%, 6.4%, and 5.9% for stipes, pileus, and gills, respectively [20]. Other fungi that have been studied are Aspergillus niger, in which the biopolymer is purified from the chitin glucan complex and the degrading of proteins by alkaline treatment at relatively high temperatures (90 C) [21]. Then, the alkali insoluble residue consists mainly of chitosan, chitin, and β-glucans [12]. The cell walls of yeast are also composed of chitin, which has been successfully extracted from Saccharomyces cerevisiae by alkaline and acid with a purity of 92.80% and a protein content of 3.78% with a degree of acetylation of 54.7% [22], which is lower than those reported for chitin from shrimp [23]. The cell walls of yeast are also composed of chitin which has been successfully extracted by alkaline and acid with a purity of 92.80% and protein content of 3.78% with a degree of acetylation of 54.7% from S. cerevisiae [22]. This is lower than reported for chitin from shrimp [23]. The alkali-insoluble residue consists mainly of chitosan, chitin, and β-glucans [12]. The complexed glucans make it difficult to extract and purify the chitin and chitosan from fungal sources. The cell walls of Zygomycetes such as Absidia, Mucor, Rhizopus, Rhizomucor, and Gongronella, have been reported as being composed mainly of chitin and chitosan (20% 50%) and polymers of glucuronic acid in lower concentrations [12,24 26]. It has been reported that chitosan interacts with other polysaccharides and phosphates forming a complex. This complex is resistant to acidic treatments for the extraction of chitosan from the cell wall such that a considerable amount of chitosan remains in the alkali- and acid-insoluble material [12]. According to the study carried out on Gongronella butleri and Absidia coerulea, fungal chitosans were extracted from mycelia of these fungi by solid substrate fermentation (SSF) and submerged fermentation (SmF). Added 13 C-glucose allowed for the synthesis of 13C-labeled glucosamine for the subsequent production of 13C-labeled chitosan for the further study of

Handbook of Chitin and Chitosan

230

8. Alternative methods for chitin and chitosan preparation, characterization, and application

the fate of chitooligosaccharides in vivo. SSF was the best method for the production of fungal chitosan based on yield. However, A. coerulea was selected for its ability to synthesize 13C-labeled chitosan in SmF [26]. In addition to the production of intracellular chitin deacetylases it is also produced extracellularly in several fungi such as Mucor rouxii [27], Colletotrichum lindemuthianum [28], Colletotrichum gloeosporioides [29] A. coerulea [30], A. nidulans [31], Metarhizium anisopliae [32], Rhizopus nigricans [33], Scopulariopsis brevicaulis [34], and Rhizopus oryzae [25]. 8.2.2.2 Insects Insects are recognized as efficient converters of organic matter into a high-value source of protein and fat; however, in spite of their potential, there are problems for their utilization, such as legislation, their social and psychological acceptance, and their application, which includes high marketing costs of the insect materials and suitable regulations for applying the materials as feed [35,36]. FAO has considered insects as food since 2003 [37]. The insect as food has a long-standing tradition in many parts of the world due to their taste and their importance as a source of proteins of high biological and quality value with a high level of digestibility [38,39]. Edible insects belong to the orders of Lepidoptera, Coleoptera, Hymenoptera, Orthoptera, Hemiptera, Isoptera, Odonata, Megaloptera, Ephemeroptera, Diptera, and Blattaria [39]. In addition to their potential as food, the insects are economically important due to the damage that they inflict on the cultivated plants, such as Cicadas of the order Hemiptera. Therefore it has been proposed to utilize cicadas as a chitin source. Mol et al. [40] have reported that the chitin contents in Cicadatra atra, Cicadatra hyalina, Cicadatra platyptera, Cicada lodosi, Cicada mordoganensis, and Cicadetta tibialis were determined to be 6.7%, 5.51%, 8.84%, 4.97%, 6.49%, and 5.88% on dry weight basis, respectively. The black soldier fly (Hermetia illucens) has been evaluated as feasible source of oil, protein, and chitin within the application of chemical or enzyme-assisted extractions that yielded 32% proteins, 37% lipids, 19% minerals, and 9% chitin [41]. Chitin extracted from Type I and II two-spotted field crickets (Gryllus bimaculatus) by chemical treatment with acid and alkali presented an average yield of chitin of 10.91% on dry weight basis [42]. Mealworms are currently easily reared in captivity and then commercialized as food for insectivores, and also for the preparation of snacks, as an insect protein source, oil, and chitin [38,39,43,44]. The exuvium and whole body of mealworm (Tenebrio molitor) larvae were treated with HCl and NaOH for demineralization (DM) and deproteinization (DP) with average chitin yields of 18.01% and 4.92%, respectively. The chitosan yields were 9.20% and 3.65% of exuviae and whole body, respectively [45]. Chitin is present only in the exocuticle and endocuticle of the insects, while the cuticle is mainly composed of protein and lipids, and most insects contain low

Handbook of Chitin and Chitosan

8.3 Current chitosan production

231

amounts of minerals [43]. Chitin and chitosan produced from insects display advantages, such as low mineral content and a reduction of allergenic reactions due to residual protein; however, chitosan from four insects exhibited lower viscosity and modulus values than that extracted from shrimp shell [46].

8.3 Current chitosan production The global market size of chitosan was assessed at US$ 3.19 billion in 2015 [47] and it was predicted that the global chitin and chitosan market should reach US$ 4.2 billion by 2021, with an annual growth rate of 15.4% from 2016 to 2021 [48]. Moreover, it is expected that this size will increase significantly in the next couple of years due to chitosan having extensive applications across several industries, specifically in water treatment, cosmetics, food and beverages, and agriculture [49]. In 2015 chitosan demand for wastewater treatment was valued at US $ 220 million [47]. However, water treatment is expected to boost the expansion of the global chitosan market size. This is because there has been a reduction in groundwater and an increase in the urban population that is affecting negatively the amount of usable fresh water, especially in Asia Pacific, which is thus pushing for the creation of more policies and an increase in investment in water treatment by private and public agents [47,49]. As a consequence, the use of chitosan in water treatment is increasing as it is accessible, biodegradable, and nontoxic and it has several functions that help remove impurities [47,49]. It is commonly used as a flocculant, but it is also an adequate coagulant of organic and inorganic compounds present in water [49]. Additionally, it also has several functions in industrial wastewaters as it can be used as a chelating agent to bind heavy metals that are highly toxic or to absorb industrial chemicals [49]. Chitosan is also functional in other areas of water treatment, such as dairy, textile, paper and pulp, and chemical and metal cutting [49]. All of this is expected to help to increase the demand for chitosan over the next few years [47,49]. Cosmetics, skincare, and toiletries are other areas that are helping to increase the chitosan market size, which according to Pulidindi and Pandey [49] is expected to grow by over 18.5% in the upcoming years, while Gran View Research [47] mentions a 20% common annual growth rate from 2016 to 2025. The importance of chitosan in these areas is mostly because it has fungistatic and bacteriostatic properties [47]. Furthermore, according to Pulidindi and Pandey [49], chitosan has several applications within the manufacturing industry of antiaging creams, such as (1) when retinol, which is the main active ingredient in

Handbook of Chitin and Chitosan

232

8. Alternative methods for chitin and chitosan preparation, characterization, and application

most antiaging cosmetics, is introduced into a chitosan matrix, it becomes more stable and efficient; and (2) it is a viable alternative for hyaluronic acid, another substance used in antiaging cosmetics. In the case of cosmetics, according to Gran View Research [47], what is boosting the chitosan market size growth is the flourishing industry of biobased color cosmetics, which include lipsticks and eyeshadows. Additionally, chitosan is also used in the industries of hair care and oral care. Pulidindi and Pandey [49] expand on this by mentioning that chitosan, when used in hair care, is able to create an elastic film on the hair, which in consequence increases the softness and the strength of the hair fiber, while when used in oral care, chitosan has antiplaque and antidecay properties. In addition, the pharmaceutical sector and the advances and current studies in the areas of biobased medicine with natural ingredients are likely to increase the demand for chitosan [49]. For example, the growing market for artificial skin and wound dressings manufactured using chitosan is likely to have a beneficial impact on chitosan market growth [47]. Moreover, the importance of this sector for the chitosan market is already visible as the North American chitosan market (led by the United States) is expected to have visible earnings close to 18% in the upcoming years, which is mostly due to the presence of key pharmaceutical companies like Merck & Co., Inc., Pfizer Inc., and Johnson & Johnson [49]. In the food and beverage industry, the chitosan market in Asia Pacific was valued at over US$ 45 million. However, chitosan has been accepted as a natural food additive in Korea, which is prone to pushing the market [47]. As seen the chitosan market is expected to grow because of its numerous uses in different industries. However, there are two major factors that might limit this predicted industry growth: high production costs and not enough high-purity product volume [47,49]. Regarding key players in the chitosan industry, countries like Thailand and China are shrimp-producing countries, and as a result, are in favored positions to become important suppliers of raw materials for the chitosan market in the region. Nevertheless, Japan is regarded as currently dominating the market, which is due to two factors: (1) Japan has great supplies of the raw materials, and (2) early knowledge about the benefits of chitosan [47]. However, China, India, and the United States are becoming emerging players due to the growth of chitosan usage in several industries, like the ones mentioned above, that are likely to increase the competition in the chitosan market [47]. Companies that have an important role in the chitosan market are Yunzhou Biochemistry Co., Advanced Biopolymers AS, Biophrame Technologies, United Chitotechnologies Inc., Koyo World (Hong Kong)

Handbook of Chitin and Chitosan

8.3 Current chitosan production

233

Co. Ltd., Dainichiseika Color & Chemicals Mfg. Co. Ltd., Agratech LLC., Kraeber & Co. GmbH, Foodchem International Corporation, FMC Corporation., GTC Bio Corporation, Panvo Organics Pvt., Ltd., KitoZyme S.A., Xianju Tengwang Chitosan Factory, FMC Health and Nutrition, Golden-Shell Pharmaceutical Co., and PT Biotech Surindo [47,49]. As the chitosan market is becoming more competitive, most of these companies are financing research to increase their product portfolio with the intention of staying ahead of the competition [49].

8.3.1 Alternative methods of production Extraction of chitin by biotechnological means is emerging as a green, cleaner, eco-friendly, and economical process. The chitin recovery microorganisms-mediated fermentation processes are highly desirable due to their easy handling, simplicity, rapidity, controllability through optimization of process parameters, ambient temperature, and negligible solvent consumption, thus reducing the environmental impact and costs [50]. 8.3.1.1 Fermentations Lactic acid fermentation has been studied as an alternative method for chitin recovery, the production of protein hydrolysates and lactic acid, as well as for pigment-extraction from crustacean waste [51 54]. The production of acid has been clearly established as the key factor in fermentation for the inhibition of pathogens and spoilage microorganisms but also for the mineral removal from crustacean waste and also pH affects the protease activity responsible for protein hydrolysis from these shells [55,56]. Thus the DM of shrimp (L. vannamei) hepatopancreas is attained by the solubilization of minerals by the organic acid from lactobacilli, whereas DP is ascribed to digestive and microbial proteases produced during fermentation. In this regard, Pacheco et al. [56] reported that the maximum protein removal was determined at the optimum lactic acid bacteria growth. This demonstrated the key role of bacterial proteases for chitin extraction during lactic acid fermentation of shrimp waste employing the homofermentative Lactobacillus plantarum as the starter. The proteolytic activity of lactic acid bacteria might be limited since it is carried out by cell envelope proteinase and peptidases. However, their contribution is important since they were able to remove 56% of protein from crab waste in sterile SmF of Lactobacillus for a simultaneous chitin and lactic acid production study [54]. Total protein removal from crustacean shells is difficult to achieve in the chitin purification process, therefore several studies have been carried out in order to achieve a high protein removal. Some of these studies involved the combination of microorganisms in more than one stage

Handbook of Chitin and Chitosan

234

8. Alternative methods for chitin and chitosan preparation, characterization, and application

of the process. In this regard, the lactic acid submerged fermentation (SmF) process has been scaled up from 0.25 to 300 L for chitin purification from shrimp shell (Crangon crangon). The process employed an anaerobic, chitinase-deficient, proteolytic-enrichment culture from ground meat for deproteination and a mixed culture of LAB from bioyoghurt for decalcification. Protein removal in the overall process for 40 h presented an efficiency of 89% 91%, while decalcification was 85% and 90% by lactic acid bacteria for another 40 h. These authors reported that chitosan obtained by deacetylation of chitin biological process presented higher viscosities than those determined with chitosan obtained chemically [57]. This is in agreement with a previous report of Pacheco et al. [23] in which the lactic acid solid-state fermentation (SSF) method avoided excessive depolymerization and loss of crystallinity during chitosan production by the freeze pump thaw deacetylation method. Other combinations reported involve the use of bacterium and fungus, such as Kurthia gibsonii and Aspergillus flavus, which were isolated from fermented milk and bread, respectively. In this study, chitin was successfully extracted from the shell of shrimps, Fenneropenaeus semisulcatus and Fenneropenaeus indicus, using a two-step SmF process using a 3-day-old bacterial broth culture for DM over 24 h, followed by 5 days fungal broth culture for 72 h for DP. The yield of chitin achieved under the best condition for the microbial treatments was lower than the chemical method [58]. The fungi bacteria combination has been also studied for chitin extraction from shrimp shells in two-stage SSF by Lactobacillus brevis and Rhizopus oligosporus [59]. The authors claimed the extraction of protein-free chitin using microorganisms GRAS (generally recognized as safe). Therein, the released protein hydrolysates (120.56 mg protein/g) displayed a molecular weight (MW) range between 25 3 103 and 11 3 103 Da. The highest concentration of astaxanthin extracted from liquid was 8.78 μg/g. Protein hydrolysates and astaxanthin showed radical scavenging activity in a DPPH assay with IC50 of 1.13 6 0.03 mg/g and 2.02 6 0.01 μg/g, respectively. The purified chitin presented a molecular weight of 1313 kDa, preserving a high crystalline index (ICR of 87.5%) and 93.67% degree of acetylation [59]. The chitins obtained by successive lactic acid bacterial and fungal fermentations of shrimp waste presented higher molecular weight and crystallinity than the commercial chitins [59]. These characteristics could enhance the properties of nanofibers obtained from chitins extracted using L. brevis with and without further inoculations with R. oligosporus. The nanofibers produced using this biological chitin presented remarkably higher Young’s modulus than that for the commercial product. The method of extraction has a significant impact on the mechanical properties of the nanofibers obtained, which is an important characteristic for polymer reinforcements [60].

Handbook of Chitin and Chitosan

8.3 Current chitosan production

235

Bacillus licheniformis produced proteases when grown in media containing shrimp waste powder as a sole carbon and nitrogen source. The percentage of protein removal after 3 h hydrolysis at 60 C and at an enzyme/substrate ratio of 5 U/mg of protein was about 81% [61]. Ghorbel-Bellaaj et al., [62] studied chitin extraction by proteaseproducing Bacillus pumilus, Bacillus mojavencis, B. licheniformis, Bacillus cereus, Bacillus amyloliquefaciens, and Bacillus subtilis. The DP attained was more than 80% with all the strains tested and produced shrimp waste protein hydrolysates with radical scavenging activity. However, DM was relatively low, 67%, but DM improved when glucose was added to the media. Another bacterium frequently used for the protease producing capacity is Serratia marcescens and along with successive treatment with L. plantarum resulted in the best chitin yield (82.56%) from lobster shell biomass, with total DP of 87.19% and total DM of 89.59% [63]. 8.3.1.2 Enzymic methods The use of proteolytic enzymes for the DP of crustacean waste has been reported as a simple alternative and relatively environment-friendly process compared to chemical chitin extraction methods. A comparison of these methods in terms of sustainability determined that the increase in the chitin yield, the mitigation of the environmental impacts, including process-water recovery, and a reduction in the enzyme production costs are necessary to justify this approach [64]. A crude, solvent-stable protease of Pseudomonas aeruginosa was produced in shrimp shell media and tested for the DP of shrimp waste to produce chitin at different enzyme/substrate ratios. The crude enzyme achieved protein removal up to 85% for 3 h of reaction [65]. In other related work, the protease from B. cereus removed protein from shrimp waste (Metapenaeus monoceros) for chitin extraction. A high level of DP 88.8% was achieved, nevertheless the DM was carried out by treatment with HCl 1.25 M for 6 h at room temperature [66]. Chitin was recovered through enzymatic DP of the shrimp byproducts. The DP rates were about 77% 6 3% and 78% 6 2% using B. mojavensis and Balistes capriscus proteases, respectively, after 3 h of hydrolysis [67]. Furthermore, an extracellular alkaline-stable protease of Bacillus safensis was purified and characterized [68] and was highly efficient in eliminating 93% of protein linked to the chitin after 3 h hydrolysis. The protease showed stability with organic solvents and compatibility with solid and liquid detergents. Other microorganisms have been employed as protease producers, for example, Erwinia chrysanthemi was applied for the DP of crustacean waste in a chitin production process. The highest protein removal

Handbook of Chitin and Chitosan

236

8. Alternative methods for chitin and chitosan preparation, characterization, and application

efficiency of the crude protease was 95% for the mineralized and demineralized wastes [69]. Chitosan in the fungal cell wall has been detected in a free form or bound to glucan. The chitosan glucan complex from mycelia of G. butleri produced by SSF has been successfully hydrolyzed with a heat stable α-amylase. The chitosan produced was characterized as having an increase in the relative average molecular weight with the fermentation time [70]. In addition, the increase of inoculum and addition of phytohormones, such as gibberellic acid and abscisic acid, resulted in significant increases in chitin deacetylase activity of R. oryzae and C. gloeosporioides, respectively [24,71]. The chitin extracted from the cell wall of C. gloeosporioides presented a lower degree of acetylation (75.6%) than that determined with the culture without the addition abscisic acid (DA of 90.6%) [71]. Chitin can be enzymatically deacetylated to chitosan, considering that the latter has at least 50% of D-glucosamine residues in the chain, but usually exceeds 80% [72]. Thus there have been studies on the use of fungal chitin deacetylases for chitosan production, however, the enzymatic deacetylation in crustacean chitin is still challenging, mainly due to the insolubility, high molecular weights, and crystalline nature of this biopolymer [12]. In this regard, chitin deacetylase of C. gloeosporioides achieved up to 25% deacetylation of a chitin biologically extracted from shrimp waste, with 80% DA, MW of 102 3 103 g/mol and an ICR of c. 60% [29]. 8.3.1.3 Combined biological methods The main disadvantage in the fermentation approach for chitin extraction is the culture time, which is considered a time-consuming process. Therefore several authors have proposed a combined method of enzymatic hydrolysis and fermentation that can be carried out simultaneously or in stages to reduce the process time [73]. Chitin has been purified from crayfish shell using Bacillus coagulans and proteinase K. The simultaneous enzymatic hydrolysis and fermentation process was conducted at 50 C with 5% (w/v) 0.5 2.0 mm of crayfish shell, 5% (w/v) glucose, proteinase K of 1000 U/g crayfish shell and inoculum of 10% in a 5-L bioreactor under nonsterile conditions. The DP and DM rates achieved after 48 h fermentation were 93% and 91%, respectively [73]. 8.3.1.4 Other methods The biotechnological method for chitin extraction presents several advantages [23,51,54] compared to other chemical or physicochemical routes. However, their main drawback is the long fermentation times, usually more than 2 days [50]. Alternatively, hydrothermal treatments

Handbook of Chitin and Chitosan

8.3 Current chitosan production

237

to biomass using water under subcritical conditions (Tc of 374 C and Pc of 22.1 MPa) allow the recovery of added-value products in short reaction times with low environmental impact. The use of water treatments under subcritical conditions has been applied for the recovery of calcareous chitin from shrimp cephalothorax to attain a DP up to 96% [74]. Nevertheless, by this method chitin still contains minerals so other methodologies have been proposed based on chitin solubility. In this regard, chitin is very difficult to dissolve, hindering its further application. Chitin exhibits limited affinity toward some solvents capable of destroying the hydrogen bonding interactions [75]. The development of chitin dissolution systems is essential for the fabrication of functional materials. Recent studies have proposed the separation of chitin from crustacean waste based on its dissolution in ionic liquids, which promotes the formation of anion chitin bridging through the cleavage of hydrogen bonds between chitin chains by acetamido groups [76]. The use of deep eutectic solvents, consisting of a mixture of choline chloride malonic acid was also studied for chitin isolation. This mixture produced the elimination of proteins and minerals with a chitin yield (20.63% 6 3.30%) higher than that (16.53% 6 2.35%) of the chemically prepared chitin [77]. Another process has been proposed using hot glycerol on prawn shell waste, which was able to remove protein (perhaps by the formation of low-molecular-weight water-soluble fragments due to dehydration and temperature) that subsequently were removed from the shell matrix by dissolution in water. This method reduced protein content to 0.24% and purified nanofibers of chitin with a higher crystallinity index (80.9%). The recovery of by-products, glycerol, and the simplicity of this method guarantee that it could be a greener alternative to the current chemical method for the industrial-scale production of chitin [78].

8.3.2 Quality control of chitin production One long-recognized drawback of the current acid/alkali process is the random hydrolysis of chitin because of the long, high-temperature exposure to acid or alkali. Although chitin is intractable, in comparison to the protein and calcium components, the nature of the process leads to random hydrolysis and a variable MW and deacetylated product. Microbiological treatment for chitin recovery offers the possibility of a more uniform and higher MW product and recovery of a usable protein fraction. This reflects the more benign processing conditions and the action of native proteases and bacterial proteases in the removal of protein. These proteases are less likely to hydrolyze the chitin component. Reviews of the biotechnical processes to chitin recovery have

Handbook of Chitin and Chitosan

238

8. Alternative methods for chitin and chitosan preparation, characterization, and application

emphasized the range of approaches attempted, and the variable results achieved [64,79]. The lesson to learn from this work is that for a predictable process and product several factors must be controlled as follows: • • • • •

raw material inoculum energy source duration of the process optimization for chitin production.

Raw material: this is usually the by-product from crustacean processing and must be considered a, “raw material”, and not a waste. This infers that it should be kept as fresh as possible before processing for chitin production to preserve protease activity. The crustacean resource can be remarkably consistent because the primary products of processing, themselves, must be reproducible. For example, farmed species such as L. monodon, are grown under controlled conditions to give a uniform weight at harvest and then hand-treated to remove the head and tail shell so that the packaged goods contain a fixed number of clean tails (often quoted as units per kg). All this effort for the consumer also leads to a uniform “waste” for chitin production. Even wild-caught crustaceans are sorted onshore to yield fixed units per kilo, so again the “waste” is reproducible. The proportions of chitin/protein/calcium carbonate in the shell vary considerably across species so the selection of a high chitin species would be useful but not always possible due to local fishing conditions and the fact that chitin production is not the primary concern of the processors. The final consideration for the chitin producer is the existence of a sufficiently large resource over an extended season. Inoculum: a wide variety of microorganisms have been tried for DP and DM [79]. A lactic acid fermentation with species of Lactobacillus and associated organisms has had most impact because they preserve the raw material, in acid conditions which also solubilize calcium carbonate, whilst native proteases break down the protein fraction. The protein hydrolysate has nutritional value in fish and animal feeds. Early work used L plantarum from the dairy industry and, latterly, lactic species isolated from crustaceans have been employed [80]. The challenge for a microbially-based industrial process lies in keeping a pure, active strain of the microorganism over a long period of time, which in turn, demands substantial backroom support from microbiologists. Energy source: laboratory experiments have usually employed pure energy sources such as glucose, although some work has investigated the use of commercially viable sources such as molasses, lactose, and fruit pulp [51,54]. Again, for a successful industrial process, a reliable energy source must be available year-round.

Handbook of Chitin and Chitosan

8.3 Current chitosan production

239

Duration: the duration of the fermentation depends on the overall aim of the process. If chitin is the major product then DP and DM must be optimized to obviate the need for any acid/alkali treatment postfermentation. Under these conditions the protein hydrolysate byproduct is adventitious and not optimized. A batch fermentation involving a single addition of all the elements at the start and following a predictable pattern can lead to predictable products. Optimization: this involves two aspects: firstly, scale-up from the laboratory to the industrial scale, and secondly, process control for reproducibility. Scale-up is often a problem in process engineering because the easy mixing, stirring, and sampling at the smaller scale is not possible at the larger scale. Attempts have been made in lactic acid fermentation to include a solid/liquid separation using rotary perforated drum reactors [81]. The initial pH of the raw material is important here as the final pH achieved for a given combination of inoculum/energy source/ time will depend on the starting point. Following the pH change with time is easy from a process control point of view and has been found to be remarkably consistent if the variables mentioned above are standardized. These two aspects are best achieved using batch processing where elements of the process are reset regularly and monitored throughout. Finally, all elements of the process must be performed to conform with guidelines proposed by the relevant licensing authorities if the chitin and chitosan products are to have food, cosmetic, medical, and pharmaceutical applications. The guidelines are intended to protect the consumer and maintain quality and consistency from batch to batch. These demands have consequences for documentation, staff training, and quality control. This approach is known as Good Manufacturing Practice (GMP) and all chitin manufacturers using the acid/alkali process claim to follow GMP. The development of biological chitin production must also follow GMP and to do so must follow some basic principles: • Manufacturing plant must be designed for hygienic processing with provision for cleaning regimes and controlled environments to prevent cross-contamination. • All process stages must be defined, with clearly written instructions and process control parameters and monitoring protocols. • Records must be kept for each batch and any deviations from normal practice noted and remedial actions recorded. There may be a requirement to keep batch records for a certain time in case of product recall. • Post-production activities such as distribution history, recall procedures, and a complaints procedure must also be in place. • All manufacturing staff must be trained in the nuances of the process, documentation, fault finding, and remediation.

Handbook of Chitin and Chitosan

240

8. Alternative methods for chitin and chitosan preparation, characterization, and application

The principles listed here are all achievable by good process engineering practices such that a chitin/chitosan production industry is possible based on biological agents. What is needed is an economic imperative to change from the traditional acid/alkali process to bioprocessing. The drivers for this change will be environmental awareness and legislation in producing countries and the demand for a range of products where the bioprocesses show better quality characteristics for applications.

8.4 Conclusions It is evident from the references given here, both reviews and individual papers, that there has been a large amount of work done in the last 20 years on the use of bioprocessing for chitin recovery and conversion to other useful products. Papers continue to be written referencing new microorganisms for chitin recovery from a variety of sources. The biomass resource for chitin production is potentially huge but crustacean-processing by-products continue to be the largest easily available and processable resource and are likely to remain so for the foreseeable future. The benefits of bioprocess approaches to chitin recovery are welldocumented with a high MW relatively uniform product and the recovery of valuable physiologically active molecules. This is not possible with the acid/alkali process which holds sway for its simplicity and low cost. A change from the current process to one based on bioprocessing requires a nexus of environmental and economic factors. If the chemical process becomes environmentally unacceptable with added pollution abatement costs and the added-value of other products from chitin recovery become significant then a transfer to the bioprocesses becomes attractive. In the meantime, there is much work to be done to prove any bioprocess at the production scale, although there are precedents and models to use for this to happen. Processes involving microorganisms or enzymes are currently operating and can act as models for chitin recovery by bioprocessing, highlighting the necessary process control, documentation, and legal frameworks at play. With regard to the physical processing plant there is an obvious parallel between bioprocessing for chitin recovery and the production of fish meal. Fish meal production is a wellestablished industry with many elements reflecting those of chitin recovery. Thus in fish meal production a large biomass must be handled; there is a separation of the biomass into solid and liquid fractions, the liquid fraction is very dilute and requires concentration by centrifugation and

Handbook of Chitin and Chitosan

References

241

vacuum evaporation, whilst the solid fraction needs grinding to a uniform size and drying. Once these elements are put together the bioprocessing approach to chitin production can be established at a process-scale—someone needs to grasp the nettle and do this.

References [1] Y.M. Bar-On, R. Phillips, R. Milo, The biomass distribution on earth, Proc. Natl. Acad. Sci. U.S.A. 115 (25) (2018) 6506 6511. Available from: doi:10.1073/ pnas.1711842115. [2] D.S. Gruen, J.M. Wolfe, G.P. Fournier, Paleozoic diversification of terrestrial chitindegrading bacterial lineages, BMC Evol. Biol. 19 (34) (2019). Available from: doi:10.1186/s12862-019-1357-8. [3] W.J. Tang, J.G. Fernandez, J.J. Sohn, C.T. Amemiya, Chitin is endogenously produced in vertebrates, Curr. Biol. 25 (7) (2015) 897 900. Available from: doi:10.1016/j. cub.2015.01.058. [4] Food and Agriculture Organization of the United Nations, The State of World Fisheries and Aquaculture 2018—Meeting the Sustainable Development Goals, Rome, Italy, 2018, Licence: CC BY-NC-SA 3.0 IGO. [5] G.A. Roberts, Chitin Chemistry, first ed., Palgrave Macmillan, London, 1992 (Chapter 1). [6] M. Kaya, M. Mujtaba, H. Ehrlich, A.M. Salaberria, T. Baran, C.T. Amemiya, et al., On chemistry of γ-chitin, Carbohydr. Polym. 176 (2017) 177 186. Available from: doi:10.1016/j.carbpol.2017.08.076. [7] M.D. Lenardon, C.A. Munro, N.A. Gow, Chitin synthesis and fungal pathogenesis, Curr. Opin. Microbiol. 13 (4) (2010) 416 423. Available from: doi:10.1016/j. mib.2010.05.002. [8] H. Merzendorfer, The cellular basis of chitin synthesis in fungi and insects: Common principles and differences, Eur. J. Cell Biol. 90 (9) (2011) 759 769. Available from: doi:10.1016/j.ejcb.2011.04.014. [9] N. Nwe, W.F. Stevens, S. Tokura, H. Tamura, Characterization of chitosan and chitosan glucan complex extracted from the cell wall of fungus Gongronella butleri USDB 0201 by enzymatic method, Enzyme Microb. Technol. 42 (3) (2008) 242 251. Available from: doi:10.1016/j.enzmictec.2007.10.001. [10] J.P. Latge´, The cell wall: a carbohydrate armour for the fungal cell, Mol. Microbiol. 66 (2) (2007) 279 290. Available from: doi:10.1111/j.1365-2958.2007.05872.x. [11] P. Orlean, D. Funai, Priming and elongation of chitin chains: implications for chitin synthase mechanism, Cell Surf. 5 (2018). Available from: doi:10.1016/j. tcsw.2018.100017. [12] G.S. Dhillon, S. Kaur, S. Kaur Brar, M. Verma, Green synthesis approach: extraction of chitosan from fungus mycelia, Crit. Rev. Biotechnol. 33 (4) (2012) 1 25. Available from: doi:10.3109/07388551.2012.717217. [13] R. Eisler, Compendium of trace metals and marine biota, first ed., Plants and Invertebrates, vol. 1, Elsevier, Oxford, 2010 (Chapter 7). [14] R.M. Dillaman, R. Roer, T. Shafer, S. Modla, The crustacean integument: structure and function, in: L. Watling, M. Thiel (Eds.), Functional Morphology and Diversity, Oxford University Press, New York, 2015, pp. 140 166. Available from: doi:10.1093/ acprof:osobl/9780195398038.003.0005.

Handbook of Chitin and Chitosan

242

8. Alternative methods for chitin and chitosan preparation, characterization, and application

[15] D. Raabe, C. Sachs, P. Romano, The crustacean exoskeleton as an example of a structurally and mechanically graded biological nanocomposite material, Acta Mater 53 (15) (2005) 4281 4292. Available from: doi:10.1016/j.actamat.2005.05.027. [16] S. Abehsera, L. Glazer, J. Tynyakov, I. Plaschkes, V. Chalifa-Caspi, I. Khalaila, et al., Binary gene expression patterning of the molt cycle: the case of chitin metabolism, PloS One 10 (4) (2015). Available from: doi:10.1371/journal.pone.0122602. [17] J. Rocha, F.L. Garcia-Carren˜o, A. Muhlia-Almaza´n, A.B. Peregrino-Uriarte, G. Ye´pizPlascencia, J.H. Co´rdova-Murueta, Cuticular chitin synthase and chitinase mRNA of whiteleg shrimp Litopenaeus vannamei during the molting cycle, Aquaculture 330 333 (2012) 111 115. Available from: doi:10.1016/j.aquaculture.2011.12.024. [18] V. Ghormade, E.K. Pathan, M.V. Deshpande, Can fungi compete with marine sources for chitosan production? Int. J. Biol. Macromol. 104 (2017) 1415 1421. Available from: doi:10.1016/j.ijbiomac.2017.01.112. [19] G. Mun˜oz, C. Valencia, N. Valderruten, E. Ruiz-Dura´ntez, F. Zuluaga, Extraction of chitosan from Aspergillus niger mycelium and synthesis of hydrogels for controlled release of betahistine, React. Funct. Polym. 91-92 (2015) 1 10. Available from: doi:10.1016/j.reactfunctpolym.2015.03.008. [20] A. Hassainia, H. Satha, S. Boufi, Chitin from Agaricus bisporus: extraction and characterization, Int. J. Biol. Macromol. 117 (2018) 1334 1342. Available from: doi:10.1016/j. ijbiomac.2017.11.172. [21] K.M. Abdel-Gawad, A.F. Hifney, M.A. Fawzy, M. Gomaa, Technology optimization of chitosan production from Aspergillus niger biomass and its functional activities, Food Hydrocoll. 63 (2017) 593 601 (2017). Available from: doi:10.1016/j. foodhyd.2016.10.001. [22] C. Sun, D. Fub, L. Jina, M. Chena, X. Zhenga, Y. Ting, Chitin isolated from yeast cell wall induces the resistance of tomato fruit to Botrytis cinerea, Carbohydr. Polym. 199 (2018) 341 352. Available from: doi:10.1016/j.carbpol.2018.07.045. [23] N. Pacheco, M. Garnica-Gonza´lez, M. Gimeno, E. Ba´rzana, S. Trombotto, L. David, et al., Structural characterization of chitin and chitosan obtained by biological and chemical methods, Biomacromolecules 12 (9) (2011) 3285 3290. Available from: doi:10.1021/bm200750t. [24] A. Zamani, L. Edebo, B. Sjo¨stro¨m, M.J. Taherzadeh, Extraction and precipitation of chitosan from cell wall of Zygomycetes fungi by dilute sulfuric acid, Biomacromolecules 8 (12) (2007) 3786 3790. [25] S. Chatterjee, S. Chatterjee, B.P. Chatterjee, A.K. Guha, Enhancement of growth and chitosan production by Rhizopus oryzae in whey medium by plant growth hormones, Int. J. Biol. Macromol. 42 (2008) 120 126. Available from: doi:10.1016/j. ijbiomac.2007.10.006. [26] N. Nwe, T. Furuike, I. Osaka, H. Fujimori, H. Kawasaki, R. Arakawa, et al., Laboratory scale production of 13C labeled chitosan by fungi Absidia coerulea and Gongronella butleri grown in solid substrate and submerged fermentation, Carbohydr. Polym. 84 (2) (2011) 743 750. Available from: doi:10.1016/j.carbpol.2010.06.023. [27] Y. Araki, E. Ito, A pathway of chitosan formation in Mucor rouxii: enzymatic deacetylation of chitin, Eur. J. Biochem. 55 (1) (1975) 71 78. [28] K. Tokuyasu, M. Ohnishi-Kameyama, K. Hayashi, Purification and characterization of extracellular chitin deacetylase from Colletotrichum lindemuthianum, Biosci. Biotechnol. Biochem. 60 (10) (1996) 1598 1603. [29] N. Pacheco, S. Trombotto, L. David, K. Shirai, Activity of chitin deacetylase from Colletotrichum gloesporoides on chitinous substrates, Carbohydr. Polym. 96 (1) (2013) 227 232. [30] X.D. Gao, T. Katsumoto, K. Onodera, Purification and characterization of chitin deacetylase from Absidia coerulea, J. Chem. 117 (2) (1995) 257 263.

Handbook of Chitin and Chitosan

References

243

[31] C. Alfonso, O.M. Nuero, F. Santamarı´a, F. Reyes, Purification of a heat-stable chitin deacetylase from Aspergillus nidulans and its role in cell wall degradation, Curr. Microbiol. 30 (1) (1995) 49 54. Available from: doi:10.1007/BF00294524. [32] P. Nahar, V. Ghormade, M.V. Deshpande, The extracellular constitutive production of chitin deacetylase in Metarhizium anisopliae: possible edge to entomopathogenic fungi in the biological control of insect pests, J. Invertebr. Pathol. 85 (2) (2004) 80 88. [33] N. Jeraj, B. Kunic, H. Lenasi, K. Breskvar, Purification and molecular characterization of chitin deacetylase from Rhizopus nigricans, Enzyme Microb. Technol. 39 (6) (2006) 1294 1299. [34] J. Cai, J. Yang, Y. Du, L. Fan, Y. Qiu, J. Li, et al., Purification and characterization of chitin deacetylase from Scopulariopsis brevicaulis, Carbohydr. Polym. 65 (2) (2006) 211 217. Available from: doi:10.1016/j.carbpol.2006.01.003. [35] S.J. Moon, J.W. Lee, Current views on insect feed and its future, Entomol. Res. 45 (6) (2015) 283 285. Available from: doi:10.1111/1748-5967.12138. [36] C.L.R. Payne, D. Dobermann, A. Forkes, J. House, J. Josephs, A. McBride, et al., Insects as food and feed: European perspectives on recent research and future priorities, J. Insects as Food Feed 2 (4) (2016) 269 276. Available from: doi:10.3920/ JIFF2016.0011. [37] J. Berg, K. Wendin, M. Langton, A. Josell, F. Davidsson, State of the art report: insects as food and feed, Ann. Exp. Biol. 5 (2) (2017) 37 46. [38] J. Ramos-Elorduy Bla´squez, J.M. Pino Moreno, V.H. Martinez Camacho, Could grasshoppers be a nutritive meal? Food Nutr. Sci. 3 (2) (2012) 164 175. Available from: doi:10.4236/fns.2012.32025. [39] Y. Feng, X.M. Chen, M. Zhao, Z. He, L. Sun, C.Y. Wang, et al., Edible insects in China: utilization and prospects, Insect Sci. 25 (2) (2018) 184 198. Available from: doi:10.1111/1744-7917.12449. [40] A. Mol, M. Kaya, M. Mujtaba, B. Akyuz, Extraction of high thermally stable and nanofibrous chitin from Cicada (Cicadoidea), Entomol. Res. 48 (6) (2018) 480 489. Available from: doi:10.1111/1748-5967.12299. [41] A. Caligiani, A. Marsegli, G. Leni, S. Baldassarre, L. Maistrello, A. Dossena, et al., Composition of black soldier fly prepupae and systematic approaches for extraction and fractionation of proteins, lipids and chitin, Food Res. Int. 105 (2018) 812 820. Available from: doi:10.1016/j.foodres.2017.12.012. [42] M.W. Kim, Y.S. Song, Y.S. Han, Y.H. Jo, M.H. Choi, Y.K. Park, Production of chitin and chitosan from the exoskeleton of adult two-spotted field crickets (Gryllus bimaculatus), Entomol. Res. 47 (5) (2017) 279 285. Available from: doi:10.1111/17485967.12239. [43] M.D. Finke, Estimate of chitin in raw whole insects, Zoo Biol. 26 (2) (2007) 105 115. Available from: doi:10.1002/zoo.20123. [44] G. Tan, M. Kaya, A. Tevlek, I. Sargin, T. Baran, Antitumor activity of chitosan from mayfly with comparison to commercially available low, medium and high molecular weight chitosans, In Vitro Cell. Dev-An. 54 (5) (2018) 366 374. Available from: doi:10.1007/s11626-018-0244-8. [45] Y.S. Song, M.W. Kim, C. Moon, D.J. Seo, Y.S. Han, Y.H. Jo, et al., Extraction of chitin and chitosan from larval exuvium and whole body of edible mealworm, Tenebrio molitor, Entomol. Res. 48 (3) (2018) 227 233. Available from: doi:10.1111/17485967.12304. [46] Q. Luo, Y. Wang, Q. Han, L. Ji, H. Zhang, Z. Fei, et al., Comparison of the physicochemical, rheological, and morphologic properties of chitosan from four insects, Carbohydr. Polym. 209 (2019) 266 275. Available from: doi:10.1016/j.carbpol.2019.01.030. [47] Gran View Research, Chitosan market estimates and trend analysis by application (water treatment, pharmaceutical & biomedical, cosmetics, food & beverage), by

Handbook of Chitin and Chitosan

244

[48]

[49]

[50]

[51]

[52]

[53]

[54]

[55]

[56]

[57]

[58]

[59]

[60]

8. Alternative methods for chitin and chitosan preparation, characterization, and application

region (North America, Europe, Asia Pacific, RoW), by country, and segment forecasts, 2018 2025. ,https://www.grandviewresearch.com/industry-analysis/globalchitosan-market., 2017 (accessed 08.03.19). Report buyer, chitin and chitosan derivatives: technologies, applications and global markets. ,https://www.reportbuyer.com/product/4806925/chitin-and-chitosan-derivatives-technologies-applications-and-globalmarkets.html., 2017 (Accessed 25.03.19). K. Pulidindi, H. Pandey, Chitosan market size by source (shrimps, prawns, crabs, lobsters), by end-user (water treatment, cosmetic & toiletries, food & beverage, healthcare/medical, agrochemical, biotechnology), Industry Analysis Report, Regional Outlook (U.S., Canada, Germany, UK, France, Spain, Italy, China, India, Japan, Australia, Indonesia, Malaysia, Brazil, Mexico, South Africa, Saudi Arabia, UAE), Application Growth Potential, Price Trends, Competitive Market Share & Forecast, 2018 2024. ,https://www.gminsights.com/industry-analysis/chitosanmarket., 2018 (accessed 08.03.19). S. Kaur, G.S. Dhillon, Recent trends in biological extraction of chitin from marine shell wastes: a review, Crit. Rev. Biotechnol. 35 (1) (2013) 44 61. Available from: doi:10.3109/07388551.2013.798256. L.A. Cira, S. Huerta, G.M. Hall, K. Shirai, Pilot scale lactic acid fermentation of shrimp wastes for chitin recovery, Process Biochem. 37 (12) (2002) 1359 1366. doi:10.1016/S0032-9592(02)00008-0. J.M. Plascencia, M.A. Olvera, J.L.A. Arredondo, K. Shirai, Feasibility of fishmeal replacement by shrimp head silage protein hydrolysates in Nile Tilapia (Oreochromis niloticus L) diets, J. Sci. Food Agri. 82 (7) (2002) 753 759. M. Gimeno, J.Y. Ramı´rez-Herna´ndez, C. Ma´rtinez-Ibarra, N. Pacheco, R. Garcı´aArrazola, E. Ba´rzana, et al., One-solvent extraction of astaxanthin from lactic acid fermented shrimp wastes, J. Agri. Food Chem. 55 (25) (2007) 10345 10350. B. Flores-Albino, L. Arias, J. Go´mez, A. Castillo, M. Gimeno, K. Shirai, Chitin and L (1)-lactic acid production from crab (Callinectes bellicosus) wastes by fermentation of Lactobacillus sp. B2 using sugar cane molasses as carbon source, Bioproc. Biosyst. Eng. 35 (7) (2012) 1193 1200. Available from: doi:10.1007/s00449-012-0706-4. K. Shirai, I. Guerrero, S. Huerta, G. Saucedo, A. Castillo, R.O. Gonza´lez, et al., Effect of initial glucose concentration and inoculation level of lactic acid bacteria in shrimp waste ensilation, Enzyme Microb. Technol. 28 (4) (2001) 446 452. N. Pacheco, M. Ga´rnica-Gonza´lez, J.Y. Ramı´rez-Herna´ndez, B. Flores-Albino, M. Gimeno, E. Ba´rzana, et al., Effect of temperature on chitin and astaxanthin recoveries from shrimp waste using lactic acid bacteria, Bioresour. Technol. 100 (11) (2009) 2849 2854. M. Bajaj, A. Freiberg, J. Winter, Y. Xu, C. Gallert, Pilot-scale chitin extraction from shrimp shell waste by deproteination and decalcification with bacterial enrichment cultures, Appl. Microbiol. Biotechnol. 99 (22) (2015) 9835 9846. Available from: doi:10.1007/s00253-015-6841-5. S.H.O. Bahasan, S. Satheesh, M.A. Ba-akdah, Extraction of chitin from the shell wastes of two shrimp species Fenneropenaeus semisulcatus and Fenneropenaeus indicus using microorganisms, J. Aqua. Food Prod. Technol. 26 (4) (2017) 390 405. Available from: doi:10.1080/10498850.2016.1188191. R. Aranday-Garcia, A. Roma´n Guerrero, K. Shirai, S. Ifuku, Successive inoculation of Lactobacillus brevis and Rhizopus oligosporus on shrimp wastes for recovery of chitin and added-value products, Process Biochem. 58 (2017) 17 24. Available from: doi:10.1016/j.procbio.2017.04.036. R. Aranday-Garcı´a, H. Saimoto, K. Shirai, S. Ifuku, Chitin biological extraction from shrimp wastes and its fibrillation for elastic nanofiber sheets preparation, Carbohydr. Polym. 213 (2019) 112 120. Available from: doi:10.1016/j.carbpol.2019.02.083.

Handbook of Chitin and Chitosan

References

245

[61] A. Haddar, N. Hmidet, O. Ghorbel-Bellaaj, N. Fakhfakh-Zouari, A. Sellami-Kamoun, M. Nasri, Alkaline proteases produced by Bacillus licheniformis RP1 grown on shrimp wastes: application in chitin extraction, chicken feather-degradation and as a dehairing agent, Biotechnol. Bioproc. Eng. 16 (2011) 669 678. Available from: doi:10.1007/ s12257-010-0410-7. [62] O. Ghorbel-Bellaaj, I. Younes, H. Maaˆlej, S. Hajji, M. Nasri, Chitin extraction from shrimp shell waste using Bacillus bacteria, Int. J. Biol. Macromol. 51 (5) (2012) 1196 1201. Available from: doi:10.1016/j.ijbiomac.2012.08.034. [63] J. Chakravarty, C.L. Yang, J. Palmer, C.J. Brigham, Chitin extraction from lobster shell waste using microbial culture-based methods, Appl. Food Biotechnol. 5 (3) (2018) 141 154. Available from: doi:10.22037/afb.v5i3.20787. [64] C. Lopes, L.T. Antelo, A. Franco-Urı´a, A.A. Alonso, R. Pe´rez-Martı´n, Chitin production from crustacean biomass: sustainability assessment of chemical and enzymatic processes, J. Clea. Prod. 172 (2018) 4140 4151. Available from: doi:10.1016/j. jclepro.2017.01.082. [65] O. Ghorbel-Bellaaj, K. Jellouli, I. Younes, L. Manni, M. Ouled Salem, M. Nasri, A solvent-stable metalloprotease produced by Pseudomonas aeruginosa A2 grown on shrimp shell waste and its application in chitin extraction, Appl. Biochem. Biotechnol. 164 (4) (2011) 410 425. Available from: doi:10.1007/s12010-010-9144-4. [66] L. Manni, O. Ghorbel-Bellaaj, K. Jellouli, I. Younes, M. Nasri, Extraction and characterization of chitin, chitosan, and protein hydrolysates prepared from shrimp waste by treatment with crude protease from Bacillus cereus SV1, Appl. Biochem. Biotechnol. 162 (2) (2009) 345 357. Available from: doi:10.1007/s12010-0098846-y. [67] I. Younes, S. Hajji, V. Frachet, M. Rinaudo, K. Jellouli, M. Nasri, Chitin extraction from shrimp shell using enzymatic treatment. Antitumor, antioxidant and antimicrobial activities of chitosan, Int. J. Biol. Macromol. 69 (2014) 489 498. Available from: doi:10.1016/j.ijbiomac.2014.06.013. [68] S. Mhamdi, I. Bkhairia, R. Nasri, T. Mechichi, M. Nasri, A.S. Kamoun, Evaluation of the biotechnological potential of a novel purified protease BS1 from Bacillus safensis S406 on the chitin extraction and detergent formulation, Int. J. Biol. Macromol. 104 (2017) 739 747. Available from: doi:10.1016/j.ijbiomac.2017.06.062. [69] N.Y. Giyose, N.T. Mazomba, L.V. Mabinya, Evaluation of proteases produced by Erwinia chrysanthemi for the deproteinization of crustacean waste in a chitin production process, Afr. J. Biotechnol. 9 (5) (2010) 707 711. Available from: doi:10.5897/ AJB09.1262. [70] N. Nwe, W.F. Stevens, Production of fungal chitosan by solid substrate fermentation followed by enzymatic extraction, Biotechnol. Lett. 24 (2) (2002) 131 134. Available from: doi:10.1023/A:1013850621734. [71] A. Ramos-Puebla, C. De Santiago, S. Trombotto, L. David, C.P. Larralde-Corona, K. Shirai, Addition of abscisic acid increases the production of chitin deacetylase by Colletotrichum gloeosporioides in submerged culture, Process Biochem. 51 (8) (2016) 959 966. Available from: doi:10.1016/j.procbio.2016.05.003. [72] W. Arbia, L. Arbia, L. Adour, A. Amrane, Chitin extraction from crustacean shells using biological methods, Food Technol. Biotechnol. 51 (1) (2013) 12 25. [73] Y. Dun, Y. Li, J. Xu, Y. Hu, C. Zhang, Y. Liang, et al., Simultaneous fermentation and hydrolysis to extract chitin from crayfish shell waste, Int. J. Biol. Macromol. 123 (2019) 420 426. Available from: doi:10.1016/j.ijbiomac.2018.11.088. [74] A. Espindola-Cortes, R. Moreno-Tovar, L. Bucio, M. Gimeno, J.L. Ruvalcaba-Sil, K. Shirai, Hydroxyapatite crystallization in shrimp cephalothorax wastes during subcritical water treatment for chitin extraction, Carbohydr. Polym. 172 (2017) 332 341. Available from: doi:10.1016/j.carbpol.2017.05.055.

Handbook of Chitin and Chitosan

246

8. Alternative methods for chitin and chitosan preparation, characterization, and application

[75] B. Duan, Y. Huang, A. Lu, L. Zhang, Recent advances in chitin based materials constructed via physical methods, Prog. Polymer Sci. 82 (2018) 1 33. Available from: doi:10.1016/j.progpolymsci.2018.04.001. [76] T. Uto, S. Idenoue, K. Yamamoto, J. Kadokawa, Understanding dissolution process of chitin crystal in ionic liquids: theoretical study, Phys. Chem. Chem. Phys. 20 (31) (2018) 20669 20677. Available from: doi:10.1039/c8cp02749h. [77] P. Zhu, Z. Gu, S. Hong, H. Lian, One-pot production of chitin with high purity from lobster shells using choline chloride malonic acid deep eutectic solvent, Carbohydr. Polym. 177 (2017) 217 223. Available from: doi:10.1016/j.carbpol.2017.09.001. [78] R. Devi, R. Dhamodharan, Pretreatment in hot glycerol for facile and green separation of chitin from prawn shell waste, ACS Sustain. Chem. Eng. 6 (1) (2018) 846 853 (2018). Available from: doi:10.1021/acssuschemeng.7b03195. [79] M.C. Gortari, R.A. Hours, Biotechnological processes for chitin recovery out of crustacean waste: a mini-review, Electron. J. Biotechnol. 16 (3) (2013) 14. [80] G.M. Hall, S. Da Silva, Lactic acid fermentation of shrimp (Penaeus monodon) waste for chitin recovery, in: C.J. Brine, P.A. Sandford, J.P. Zikakis (Eds.), Advances in Chitin and Chitosan, Elsevier Applied Sciences, London, 1992, pp. 633 638. [81] Z. Zakaria, G.M. Hall, G. Shama, Lactic acid fermentation of scampi waste in a rotating horizontal reactor for chitin recovery, Process Biochem. 33 (1) (1998) 1 6.

Handbook of Chitin and Chitosan

C H A P T E R

9 Current research on the blends of chitosan as new biomaterials A. Rajeswari, Sreerag Gopi, E. Jackcina Stobel Christy, K. Jayaraj and Anitha Pius Department of Chemistry, The Gandhigram Rural Institute—Deemed to be University, Dindigul, India O U T L I N E 9.1 Introduction

248

9.2 Chitosan biomaterial 9.2.1 Preparation of chitosan 9.2.2 Properties of chitosan 9.2.3 Applications of chitosan

249 249 253 255

9.3 Modification of chitosan 9.3.1 Polymer blending technique 9.3.2 Chitosan blends

256 256 257

9.4 Natural polymers blends with chitosan 9.4.1 Chitosan/starch 9.4.2 Chitosan/collagen 9.4.3 Chitosan/proteins 9.4.4 Chitosan/natural rubber latex

258 258 260 261 263

9.5 Chitosan blends with synthetic polymers 9.5.1 Chitosan/hydrophilic polymers 9.5.2 Chitosan/nylon 9.5.3 Chitosan/polyacrylamide 9.5.4 Chitosan/poly(lactic acid)

264 265 268 271 273

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00009-2

247

© 2020 Elsevier Inc. All rights reserved.

248

9. Current research on the blends of chitosan as new biomaterials

9.6 Chitosan-based hydrogels

274

9.7 Conclusions

275

Acknowledgment

275

References

275

9.1 Introduction With the evolution of human civilization, the field of biomaterial evolved different materials at multiple length scales from the macro- to micro- to nanolevel with a simple focus to extend human life and improve the quality of life. A sustainable biobased product is derived from renewable resources with recyclability and biodegradability with commercial viability and environmental acceptability (Fig. 9.1). Biodegradation was defined by Albertsson and Karlsson as an event which takes place through the action of enzymes and/or chemical decomposition associated with living organisms (bacteria, fungi, etc.) or their secretion products [1]. However, it is necessary to consider abiotic reactions (e.g., photodegradation, oxidation, and hydrolysis) that may also alter the polymer before, during, or instead of biodegradation because of environmental factors. Albertsson and Karlsson have also recognized that nature can be used as a model in the design of degradable polymeric materials, because it can combine polymeric materials with different degradation times into a hierarchical system that

FIGURE 9.1 Scheme for sustainable environment.

Handbook of Chitin and Chitosan

9.2 Chitosan biomaterial

249

optimizes both energy and material properties [2]. Naturally occurring biopolymers are susceptible to environmental degradation factors and their breakdown may be caused by a combination of these; for example, wood is both oxidized and hydrolyzed. Tailoring the properties of polymers to a wide range of uses and developing a predetermined service life for the materials have become increasingly important and Albertsson and Karlsson suggest four different strategies: • the use of cheap, synthetic, bulk polymers with the addition of a biodegradable or photooxidizable component; • chemical modification of the main polymer chain of synthetic polymers by the introduction of hydrolyzable or oxidizable groups; • the use of biodegradable polymers and their derivatives, with poly (hydroxyl alkanoate)s (PHAs) being the most studied; and • tailormake new hydrolyzable structures, for example, polyesters, polyanhydrides, and polycarbonates.

9.2 Chitosan biomaterial In the search for new materials with improved physicochemical and mechanical properties, the blending of polymers has received considerable attention of researchers in the past several decades. Chitosan is a modified natural polymer prepared by the deacetylation of chitin, which is a water-insoluble polymer. It is a natural polycation, nontoxic, biocompatible, and biodegradable material. Chitosan (ß-(1,4)-2-amino-2-deoxyD-glucopyranose) is the second most abundant biopolymer in nature next to cellulose. Compared with other polysaccharides, chitosan has several important advantages, including biocompatibility, biodegradability, nontoxicity, good film-forming characteristics, excellent chemical resistance, and electrolytic properties. So, chitosan has been widely studied for use in clinics, drug delivery systems, solid polyelectrolytes, surfactants, and membranes for ultrafiltration, reverse osmosis, and evaporation. However, its mechanical properties and other physical/chemical properties are not good enough to meet this wide range of applications.

9.2.1 Preparation of chitosan Chitosan (CS) is the deacetylated derivative of chitin [3,4]. Among numerous polysaccharides, chitin occupies a special position due to its versatility, facile modification, and unique properties. Chitin is usually distributed in marine invertebrates, the most important sources of chitin are crustaceans [5
8], such as shrimps, squids, and crabs. The different sources are shown in Fig. 9.2. From a practical viewpoint, the shells of marine

Handbook of Chitin and Chitosan

250

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.2 Sources of chitosan.

crustaceans such as crabs and shrimps are convenient; they are available as waste from the seafood processing industry and used for commercial production of chitin. The exoskeleton of crustacean samples consists of 30%
40% proteins, 30%
50% calcium carbonate, and 20%
30% chitin [9]. Pigments and other metal salts are only minor components. β-Chitin is less common than α-chitin, but this allomorph is obtained in a certain amount from squid pens. β-Chitin also occurs in Aphrodite chaetae, lorica of sessile ciliates, pogonophore tubes, and diatom spines. It is attractive as another form of chitin with characteristics that are noticeably different from those of α-chitin and understanding of the chemistry is steadily progressing [10]. Other possible sources of chitin production include krill, clams, oysters, insects, and fungi. Table 9.1 lists the rough estimates of chitin contents from different sources. Cell walls of some fungi (Zygomycetes) contain chitosan and will be potential sources of chitosan. The chemical structure of acetylated and deacetylated structure is given in Fig. 9.3.

Handbook of Chitin and Chitosan

251

9.2 Chitosan biomaterial

TABLE 9.1

Sources and contents of chitin.

Source

Chitin (%)

Shrimp cuticle

30
40

Squid pen

20
40

Krill cuticle

20
30

Crab cuticle

15
30

Fungi cell wall

10
25

Insect cuticle

5
25

Clam/oyster shell

3
6

FIGURE 9.3 Chemical structure of (A) chitin and (B) chitosan.

Chitin exists in tightly bound complexes with other substances in the cuticles of crabs and shrimps and some portions of polypeptides are suggested to be linked covalently to the amino groups. Chitin is not soluble in ordinary solvents and this makes the solvent extraction method inapplicable for the isolation of chitin. However, chitin is fairly stable under mild acidic as well as basic conditions and thus is obtained as the residue remaining after decomposition of the other components with acid and alkali [11,12]. Chitin and chitosan are both prepared using the common process illustrated and described in Fig. 9.4 which shows the extraction of

Handbook of Chitin and Chitosan

252

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.4 Extraction of chitosan from crustaceans wastes.

chitosan from crustacean waste. Chemical extraction of chitin from the crustacean exoskeleton consists of three basic steps: 1. demineralization, 2. deproteinization, and 3. deacetylation. These three steps are the standard procedure for chitin production [13,14]. The deacetylation of chitin is generally achieved by treatment with concentrated sodium hydroxide solution (40%
50%) at 90 C
100 C or higher temperature in order to remove some or all of the acetyl groups from the chitin [15,16]. Chitosan is commercially available. The majority of its commercial samples are available with degree of deacetylation (DD) ranging between 70% and 90% and always less than 95% [17]. Tolaimate et al. [18] have reported that chitosan with DD higher than 95% may be obtained via further deacetylation steps. However, this may result in partial depolymerization as well as increase the cost of the preparation. Chitosan dissolves readily in dilute solutions of organic acids, including malic, acetic, tartaric, glycolic, citric, and ascorbic acid solution [19]. It consists of an amino group which exhibits protonation under acidic conditions. The structure of chitosan is given in Fig. 9.3. Chitosan is also found in some microorganisms like yeast and fungi. The primary unit in the chitin polymer is 2-deoxy-2-(acetylamino) glucose. These units are combined by β-(1,4)-glucosidic linkages forming a long-chain linear polymer. The degree of hydrolysis (deacetylation) has a significant effect on the solubility and rheological properties of the polymer. When

Handbook of Chitin and Chitosan

9.2 Chitosan biomaterial

253

FIGURE 9.5 Repeating unit of chitin.

dissolved in an acidic solution chitosan gives a viscous solution. The viscosity determines the extent of penetration of chitosan into fabric structure. The structure of chitosan is essential for the synthetic chemistry in site selective modification due to the different reactivities of the amino group at the C2 position and the primary and secondary hydroxyl groups at the C3 and C6 positions, as shown in Fig. 9.5 [20].

9.2.2 Properties of chitosan Chitosan has become the subject of research due to its unique properties and ease of modification [21
23]. The physical, chemical, and biological properties [24
26] of chitin and chitosan depend on two parameters: the degree of deacetylation (DD) and molecular weight distribution [27], which are dictated by the chitin sources and the method of preparation [28]. In fact, the degree of deacetylation of chitosan influences not only its physicochemical characteristics, but also its biodegradability [29,30]. In addition, the degree of deacetylation is also reported to have an impact on the performance of chitosan in many of its applications [31,32]. 9.2.2.1 Physical properties Chitosan exists in a form of white and yellowish flakes, which can be converted to bead or powders. The degree of deacetylation also plays a vital role in the molecular weight of chitosan. The lower the deacetylation, the higher the molecular weight, which provides higher chemical stability and mechanical strength. The average molecular weight of chitosan is approximately 1.2 3 105 g/mol [33]. Chitin and chitosan are amorphous solids and almost insoluble in water. This is mostly due to intermolecular hydrogen bonding, which can form between the neutral molecules of chitosan. The solubility of chitosan in water depends upon

Handbook of Chitin and Chitosan

254

9. Current research on the blends of chitosan as new biomaterials

the balance between the electrostatic repulsion resulting from the protonated amine functions and the hydrogen bonding due to the free amino groups [34]. Chitosan can be dissolved in aqueous organic acid solutions, such as formic acid and acetic acid at a pH below 6, and becomes a cationic polymer due to the protonation of the amino groups available in its molecular structure. However, chitosan hardly dissolves in pure acetic acid. In fact, concentrated acetic acid solutions at high temperatures may cause the depolymerization of chitosan. The solubility of chitosan in dilute acids depends on the degree of deacetylation in the polymer chain. Consequently, this property can be used to distinguish between chitin and chitosan [35]. The degree of deacetylation has to be at least 85% complete so that the desired solubility of chitosan can be achieved. Furthermore, the properties of chitosan solutions depend not only on its average degree of deacetylation but also on the distribution of the acetyl groups along the chain [36,37], the acid concentration, and the type of acid [38]. 9.2.2.2 Chemical properties Similar to most natural polymers, chitosan has an amphiphilic character, which can influence its physical properties in solutions and solid states. This is attributed to the presence of the hydrophilic amino groups together with the hydrophobic acetyl groups in its molecular structure. The presence of a large number of amino groups also confers upon chitosan a strong positive charge unlike most polysaccharides. Chitosan has a cationic nature due to the presence of amino and hydroxyl groups, which make it modifiable by various means including complexation, grafting, cross-linking, and blending [39]. Chitosan is a rigid polymer due to the presence of hydrogen bonding in its molecular structure. Consequently, it can be easily transformed into films with a high mechanical strength. Chitosan is a weak polyelectrolyte that may be regarded as a very poor anion-exchanger. Therefore it is likely to form films on negatively charged surfaces in addition to the ability to chemically bind with negatively charged fats, cholesterol, proteins, and macromolecules [40,41]. 9.2.2.3 Biological properties Chitin and chitosan are nontoxic, biocompatible, and biodegradable. They are amino polysaccharides and show interesting biological, physiological, and pharmacological properties. Their notable bioactivities include the promotion of wound healing, hemostatic activity, immune enhancement, hypolipidemic activity, mucoadhesion, eliciting biological responses and antimicrobial activity. Chitosan is also promising as a supporting polymer for gene delivery, cell culture, and tissue engineering. Certain chitin and chitosan oligomers have physiological functions, including the induction of phytoalexins, antimicrobial activity, and

Handbook of Chitin and Chitosan

9.2 Chitosan biomaterial

255

immune-enhancing activity. Chitin and chitosan as dietary fibers exhibit hypolipidemic activity, as confirmed by the reduced cholesterol and triglyceride levels in blood plasma (serum) and liver of rats [42]. Chitosan was much more effective than chitin [43]. Chitosan and its derivatives may thus be of medicinal importance [44].

9.2.3 Applications of chitosan Chitosan is considered as a potential polysaccharide because of its free amino groups that contribute to polycationic, chelating, and dispersion forming properties along with ready solubility in dilute acetic acid. Chitosan possesses exceptional chemical and biological qualities that can be used in a wide variety of applications, ranging from biomedical, pharmaceutical, and cosmetic products to water treatment and plant protection. In different applications, different properties of chitosan are required. These properties change with, for example, degree of deacetylation and molecular weight. 9.2.3.1 In wastewater treatment Among other polysaccharides chitosan is a positively charged molecule, so it can be used as a flocculating agent and can also act as a chelating agent and heavy metal trapper. That is why it finds potential applications in water treatment [45]. In terms of utilization, crawfish chitosan as a coagulant for recovery of organic compounds in wastewater was demonstrated to be equivalent or superior to the commercial chitosans from shrimp and crab waste shell and synthetic polyelectrolytes in turbidity reduction [46]. The wastewater released from food processing plants, typically in the seafood, dairy, or meat processing industries, contains an appreciable amount of protein which can be recovered with the use of chitosan; this protein, after drying and sterilization, makes a great source of feed additives for farm animals [47]. 9.2.3.2 In food industry Chitosan is already used as a food ingredient in Japan, in Europe, and in the United States as a lipid trap, an important dietetic breakthrough. Since chitosan is not digested by the human body, it acts as a fiber, a crucial diet component. It has the unique property of being able to bind lipids arriving in the intestine, thereby reducing by 20%
30% the amount of cholesterol absorbed by the human body. In solutions, chitosan has thickening and stabilizing properties, both essential to the preparation of sauces and other culinary dishes that hold their consistency well. Finally, as a flocculating agent, it is used to clarify beverages. Because of its phytosanitary properties, it can be sprayed in dilute form

Handbook of Chitin and Chitosan

256

9. Current research on the blends of chitosan as new biomaterials

on foods such as fruits and vegetables, creating a protective, antibacterial, fungistatic film. The principal commercial applications of chitosan include preservatives, food stabilizers, animal feed additives, and anticholesterol additives [48]. 9.2.3.3 In agriculture Chitosan offers a natural alternative to the use of chemical products that are sometimes harmful to humans and their environment. Chitosan triggers the defensive mechanisms in plants (acting much like a vaccine in humans), stimulates growth, and induces certain enzymes (synthesis of phytoalexins, chitinases, pectinases, glucanases, and lignin). This new organic control approach offers promise as a biocontrol tool. In addition to the growth-stimulation properties and antifungal activity, chitosans are used for seedcoating, frost protection, bloom and fruit-setting stimulation, timed release of product into the soil (fertilizers, organic control agents, nutrients), and as protective coatings for fruits and vegetables [48].

9.3 Modification of chitosan Chitosan has a number of outstanding properties; some properties specially desired for a particular application can be incorporated by further modification of the chitosan backbone. The presence of certain functionalities like 2 NH2 and 2 OH groups in the chitosan molecules provides the basis for interaction with other polymers and biological molecules. A number of methods have been reported for the modification of chitosan [49]. The most commonly used methods for the modification of chitosan are blending, graft copolymerization and curing. Blending is the technique used to get a polymer with desired properties by the physical mixing of two or more polymers. However, graft copolymerization is the method in which one polymer is covalently bonded to the other polymer chain. There is no time limitation for the graft copolymerization as it can take place in a few seconds, minutes, or even in hours. On the other hand, curing is a very fast and rapid process and takes place in a fraction of seconds.

9.3.1 Polymer blending technique The new polymer modification process based on a simple mechanical mixture of two polymers generated a new polymer class called polymer blends [50]. Polymer blends are physical mixtures of structurally different polymers or copolymers, which interact through secondary forces with no covalent bonding, and which are miscible at the molecular level. The basis of polymer
polymer miscibility may arise from any

Handbook of Chitin and Chitosan

9.3 Modification of chitosan

257

specific interaction, such as hydrogen bonding, dipole
dipole forces, and charge transfer, complexes for homopolymer mixtures [51
53]. There have been various techniques used to study the miscibility of the polymer blends [54
60]. Some of these techniques are complicated, costly, and time-consuming. Hence, it is desirable to identify simple, low-cost, and rapid techniques to study the miscibility of polymer blends. The modification of polymers has received much attention in relation to their potential application [61].

9.3.2 Chitosan blends Generally, there are two main methods that are commonly used in the blending of chitosan: (1) dissolving in a solvent followed by evaporation (solution blending) [62,63] and (2) mixing under fusion conditions (melt blending) [64]. However, according to the literature, solution blending is the most applied method for preparing chitosan blends due to its simplicity and suitability for producing various forms of blends (beads, microspheres, films, and fibers). In solution blending, chitosan is dissolved in an appropriate solvent (usually diluted acetic acid) with continuous stirring at room temperature. This is followed by mixing a desired amount of another polymer after being dissolved in a solvent under continuous stirring conditions. The blend solution of chitosan is often cross-linked by addition of a cross-linking agent to improve mechanical properties. Subsequently, the blend solution is filtered and then cast on a glass plate or a petri dish and left to dry under room or oven temperature [65,66]. Eventually, the blend is washed with NaOH solution to remove the excess acetic acid. Like other polymer blends, the properties of chitosan blends depend upon the miscibility of its components at the molecular scale, which is attributed to specific interactions between polymeric components. The most common interactions in the chitosan blends are hydrogen bonding, ionic and dipole, π-electrons and charge-transfer complexes. Various techniques, such as thermal analysis [67], electron microscopy [68], viscometric measurements [69], and dynamic mechanical studies [70], have been used to investigate the polymer
polymer miscibility in solution or in solid state. The purposes of chitosan blending vary depending upon the application demands. This includes the following: (1) to enhance hydrophilicity [71,72], (2) to enhance mechanical properties [73
77], (3) to improve blood compatibility [78
80], and (4) to enhance antibacterial properties [81]. In blood contact applications, chitosan promotes surface-induced thrombosis and embolization. The selection of the polymers to be blended with the chitosan depends on the property to be conferred or boosted. For example, the hydrophilic property of chitosan

Handbook of Chitin and Chitosan

258

9. Current research on the blends of chitosan as new biomaterials

is modified by blending with polymers such as PEG and PVA. Chitosan was also blended with several polymers, such as polyamides, poly (acrylic acid), gelatin, silk fibroin, and cellulose, to enhance mechanical properties. To enhance antibacterial properties, chitosan is blended with cellulose [71
81].

9.4 Natural polymers blends with chitosan Blending of two polymers is an approach to develop new biomaterials exhibiting combinations of properties that could not be obtained by individual polymers [82]. Blends made of synthetic and natural polymers can be endowed with a wide range of physicochemical properties and processing techniques of synthetic polymers as well as the biocompatibility and biological interactions of natural polymers. Natural polymers as biotechnological or biomedical resources have been investigated because of their unique properties including nontoxicity, degradability, and biological compatibility. Blending of natural polymers with chitosan is a very interesting methodology by which to modify the properties of natural polymers and to develop novel composite materials based on natural polymers.

9.4.1 Chitosan/starch Starch is composed of amylose and amylopectin with relative amounts of each component varying according to its source. As an example, corn starch has about 28 wt.% amylose compared with cassava starch with 17 wt.% [83
85]. Rice starch and its major components, amylose and amylopectin, are attractive raw materials for use in various applications. They have been used to produce biodegradable films to partially or entirely replace plastic polymers because of the low cost and renewability, as well as possession of distinct mechanical properties [86]. However, the wide application of starch film is limited by its mechanical properties and efficient barrier against low polarity compounds [87]. This constraint has led to the development of the improved properties of natural source-based films by modifying the starch properties and/or incorporating other materials. Jagannath et al. [88] blended starch with different proteins to decrease the water vapor permeability of the films and to increase their tensile strength. However, these films still did not perform well compared to synthetic polymer-based films. Starch-based films, however, are brittle and hydrophilic, therefore limiting their processing and application. In order to overcome these drawbacks, starch can be mixed with various synthetic and natural polymers. These approaches are multilayer structures with aliphatic polyesters, blends with natural rubber or zein, and composites with

Handbook of Chitin and Chitosan

9.4 Natural polymers blends with chitosan

259

fibers. Another widely used approach to improve the mechanical properties and processability of starch films is the addition of chitosan. Thawien Bourtoom et al. have developed [89] biodegradable blend films from rice starch
chitosan by casting a film solution on leveled trays. The properties of rice starch
chitosan biodegradable blend film and selected biopolymer and synthetic polymer films were compared; the results demonstrated that rice starch
chitosan biodegradable blend film had mechanical properties similar to the other chitosan films. However, the water vapor permeability of rice starch
chitosan biodegradable blend film was characterized (shown in Figs. 9.6 and 9.7) by relatively lower water vapor permeability than chitosan films but higher than polyolefin.

FIGURE 9.6 Effects of chitosan ratios on (A) tensile strength (TS) and (B) elongation at break (E) of the biodegradable blend films. (a
c) Means with the different letters represent significantly different value at P , .05 using Duncan’s Multiple Range Test. Source: T. Bourtoom, M.S. Chinnan, Preparation and properties of rice starch—chitosan blend biodegradable film, LWT—Food Sci. Technol. 41 (2008) 1633
1641. Reproduced with permission from Copyright r 2007 Elsevier Ltd.

Handbook of Chitin and Chitosan

260

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.7 Effects of chitosan ratios on (A) water vapor permeability (WVP) and (B) film solubility (FS) of the biodegradable blend films. (a
c) Means with the different letters represent significantly different value at P , .05 using Duncan’s Multiple Range Test. Source: T. Bourtoom, M.S. Chinnan, Preparation and properties of rice starch—chitosan blend biodegradable film, LWT—Food Sci. Technol. 41 (2008) 1633
1641, Reproduced with permission from Copyright r 2007 Elsevier Ltd.

9.4.2 Chitosan/collagen Collagen, which is the most abundant protein in mammals, makes up 25%
35% of the body’s protein content and represents the major constituent of animal cartilage and connective tissue. It is widely used as the scaffold during bone regeneration due to its outstanding cell adhesion, proliferation, and differentiation properties. Collagen has been reported to promote MC3T3-E1 cell proliferation and enhance stem cell differentiation into osteoblasts [90
92]. However, it is difficult to use alone as a

Handbook of Chitin and Chitosan

9.4 Natural polymers blends with chitosan

261

biomaterial because of its low mechanic properties and easy degradation during bone tissue engineering. Chitosan
collagen composite made using different methods can be used as a tissue engineering material. Blended films with different chitosan and collagen mass ratios significantly affected the cell morphology, adhesive force, and Young’s modulus. Stem cells could be cultured on chitosan
collagen formed sponges [93]. The process of bone marrow mesenchymal stem cell development, proliferation, and osteogenic differentiation was enhanced after culturing on chitosan hydrogel and collagen [94]. Many studies have shown that chitosan and collagen can be used in bone tissue engineering to influence osteoblast proliferation, differentiation and mineralization. While previous studies focused on the overall cell performance, little is known about the molecular mechanism of the cells when cultured on these biomaterials. Therefore, it is important to characterize the molecular mechanism of the cell behavior when the cells are cultured on chitosan and collagen composites to achieve better use in bone tissue engineering. Martı´nez et al. have demonstrated that chitosan/collagen (Chit/Col) blends great potential for use in tissue engineering (TE) applications. The essential properties of the scaffolds such as morphology, mechanical stiffness, swelling, degradation and cytotoxicity were studied [95]. Fig. 9.8 shows the stress
strain curves for hydrated nonchemically cross-linked and cross-linked scaffolds such as 80Chit/20Col; (B) 50Chit/50Col (C) 20Chit/80Col and (D) Typical profile of three repeat tests of the same scaffold specimen (n 5 5) tested under compression at 5 mm/min at room temperature.

9.4.3 Chitosan/proteins Biopolymers, including proteins and chitosan, have been the focal point of an expanding number of studies reporting their potential use in new materials, such as edible film, drug delivery, and membranes. He Huang et al. have made protein
chitosan films by casting a solution of proteins and chitosan on pyrolytic graphite electrodes. Myoglobin (Mb), hemoglobin (Hb), and horseradish peroxidase (HRP) incorporated in chitosan films gave a pair of stable, well-defined and quasireversible cyclic voltammetric peaks at about
0.33 V versus saturated calomel electrode in pH 7 buffers, respectively, while catalase (Ct) in chitosan films showed a peak pair at about
0.46 V which was not stable [96]. Many polymeric membranes have been investigated for the purpose of wound covering on account of its importance in the treatment of burns, prevention of postsurgical adhesions, and cosmetic surgery. These materials include synthetic polymers like polyurethane, polyethylene, polylactides, polyglycolides, and polyacrylonitrile. However

Handbook of Chitin and Chitosan

262

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.8 Stress
strain curves for hydrated nonchemically cross-linked scaffolds (A) 80Chit/20Col, (B) 50Chit/50Col, and (C) 20Chit/80Col. (D) Typical profile of three repeat tests of the same scaffold specimen. Source: A. Martı´neza, M.D. Blanco, N. Davidenko, R.E. Cameron, Tailoring chitosan/collagen scaffolds for tissue engineering: Effect of composition and different crosslinking agents on scaffold properties, Carbohydr. Polym. 132 (2015) 606
619. Reproduced with permission from Copyright r 2015 Elsevier Ltd.

some of these polymers have disadvantages in such applications, that is, poor biocompatibility and the release of acidic degradation products [97,98]. One alternative approach involves the use of biodegradable polymers from renewable resources, including starch, collagen, gelatin, chitosan, and proteins (soy protein, casein, silk fibroin and wheat), since these polymers are widely available in nature, and are biodegradable and nontoxic [99
104]. Among these renewable polymers, soy protein, the major component of the soybean, has the advantages of being economically competitive and presents good water resistance as well as storage stability. The combination of these properties with a similarity to tissue constituents and a reduced susceptibility to thermal degradation makes soy an ideal template to be used as a biomaterial for skin tissue engineering. Abugoch et al. [105] have prepared quinoa protein/chitosan films by solution casting of blends of quinoa protein extract (PE) and chitosan.

Handbook of Chitin and Chitosan

9.4 Natural polymers blends with chitosan

263

FIGURE 9.9 (A) FTIR and (B) XRD spectra of PE, chitosan, and PE/chitosan blend film. Source: L.E. Abugoch, C. Tapia, M.C. Villama´n, M. Yazdani-Pedram, M. Dı´az-Dosque, Characterization of quinoa protein
chitosan blend edible films, Food Hydrocoll. 25 (2011) 879
886. Reproduced with permission from Copyright r 2015 Elsevier Ltd.

Films from a PE/chitosan blend were characterized by FTIR, X-ray diffraction, thermal analysis, and SEM. Fig. 9.9A shows the FTIR spectrum of PE powder, chitosan powder, and PE/chitosan blend film. A significant shift of the broad absorption band due to the O-H vibration of PE at 3402
3435/cm is seen for the PE/chitosan blend film. The tensile mechanical, barrier, and sorption properties of the films were also evaluated. Fig. 9.9B shows the XRD peaks of chitosan powder with two main diffraction peaks at 2θ 10.1 and 20.1 degrees which agree with previously published results [106,107]. New diffraction peaks at 2θ 29.7, 31.2, and 36.1 degrees suggest the existence of intermolecular interactions between quinoa PE and chitosan.

9.4.4 Chitosan/natural rubber latex The emergence of thermoplastic elastomers is one of the important developments in the field of polymer science and technology in recent years. Thermoplastic elastomers are a new class of materials which combine the properties of rubber with the ease of processability of thermoplastics. One type of fast-growing thermoplastic elastomer which is easier to process is made by blending rubber and plastic in definite proportions. Characteristically, this is a family of materials consisting of a rubber soft segment which gives rise to elastomeric properties and a crystalline hard segment which acts as a cross-link and filler. Ismail et al. have prepared chitosan-filled rubber compounds using a laboratory-sized two-roll mill. The effect of chitosan loading on three

Handbook of Chitin and Chitosan

264

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.10

(A) SEM micrographs of unfilled compounds of STR 5 L taken at magnification of 300 3 . (B) SEM micrographs of unfilled compounds ENR-25 taken at magnification of 300 3 . (C) SEM micrographs of unfilled compounds of SBR taken at magnification of 300 3 . Source: H. Ismail, S.M. Shaari, N. Othman, The effect of chitosan loading on the curing characteristics, mechanical and morphological properties of chitosan-filled natural rubber (NR), epoxidised natural rubber (ENR) and styrene-butadiene rubber (SBR) compounds, Poly. Test. 30 (2011) 784
790. Reproduced with permission from Copyright r 2011 Elsevier Ltd.

different types of rubber (STR 5 L, ENR-25, and SBR) was investigated. Morphological studies (Fig. 9.10) of the tensile fractured surface of the vulcanisates using scanning electron microscopy (SEM) indicated that chitosan interacts less well with SBR than with STR 5 L and ENR-25 [108].

9.5 Chitosan blends with synthetic polymers Blending of chitosan with synthetic polymers is of great interest due to their significant characterizations. The modification of synthetic polymers can enhance their properties such as flexibility, conductivity, good water retention, enhanced mechanical properties, etc., while

Handbook of Chitin and Chitosan

9.5 Chitosan blends with synthetic polymers

265

maintaining biodegradability and sorption performance [109]. Recently, one of the natural biopolymers playing a great role in this field has been chitosan. Because of the numerous applications of chitosan in various fields, blends with synthetic polymers with a wide range of physicochemical properties have been prepared on various occasions with solution blending by many researchers. The advantages of this method compared to the conventional techniques of polysaccharide mixture production have been shown [110]. Chitosan contains high-polarity groups that can form hydrogen bonds with the corresponding functional groups of synthetic polymers that, in turn, lead to their enhanced compatibility to each other; the synthetic polymers capable of hydrogen bond formation include polyamides, polyesters, and many vinyl polymers. However, the tendency of these components to self-associate often hinders the interaction and compatibility of polysaccharides with a second polymer component. Several properties of chitosan are, however, undesirable from the processing and manufacturing point of view. For example, wellcharacterized, purified chitosan is expensive. Chitosan is also not an easy polymer to work with, in regard to solubility, film processing, and film quality. We have thus chosen to work with chitosan blends with other hydrophilic polymers; this study evaluates the efficiency for the absorption enhancement of a model drug using blends, compared with the homopolymers, as well as presents some relevant material properties of the blends.

9.5.1 Chitosan/hydrophilic polymers In nature hydrophilic products like protein, keratin, or wool are responsible for water vapor permeability. The easiest way to obtain water vapor permeability for a chemist should be the use of a hydrophilic polymer. There are a lot of existing polymeric materials which are hydrophilic, for example, polyethylene glycol ethers, polyurethanes with polyethylene glycol, ether soft segments, ethoxylated graft polymers, polyvinyl alcohol (PVA), polyvinyl pyrrolidone, and polyethylene oxide (Fig. 9.11). As a hydrophilic material for pervaporation membranes, chitosan has drawn a great deal of attention in the past decade [111
115]. In addition to its high hydrophilicity, the good film-forming property, high mechanical strength, and chemical resistance make chitosan a very promising material. PVA is used as the first material for pervaporation membranes for the separation of water
ethanol mixtures [116
118]. The composite ˝ Trenntechnik). membrane was produced by GFT (Gesellschaft fur The PVA membranes also have been widely used in various pervaporation dehydration applications in industries, such as dehydration of

Handbook of Chitin and Chitosan

266

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.11 Various types of hydrophilic polymers.

alcohols, esters, acids, and volatile organic compounds [119]. PVA is an aliphatic polymer containing hydroxyl groups. PVA can be cross-linked by glutaraldehyde [120] and multicomponent carboxylic acids. Structures of PVA are illustrated in Fig. 9.12. Svang-Ariyaskul et al. [121] have prepared homogeneous membranes by casting the solution of blended chitosan and PVA on a glass plate. This membrane showed very excellent performance with good mechanical strength. It is promising to develop this membrane for industrial uses Fig. 9.13A and B show the effects of the membrane composition on the pervaporation performance—flux and water content in permeate— at 60 C for the separation of isopropanol (90
95 wt.%). At 90 and 95 wt. % isopropanol in the feed, total flux increased gradually with the chitosan content in the membrane. It confirmed the explanation previously about the effects of chitosan composition that the higher chitosan in the membrane, the higher the total permeation flux. According to Fig. 9.14A, the swelling degree of the membrane increased when the isopropanol content in the solution decreased. This is due to the fact that all membranes are highly hydrophilic and this characteristic results in the increase of solution adsorbed in the membrane when water concentration in solution increases. Fig. 9.14B confirms that all membranes are highly hydrophilic because in both 35 and 75 wt.% isopropanol aqueous solutions, water adsorbed in the membranes was higher than 80%.

Handbook of Chitin and Chitosan

9.5 Chitosan blends with synthetic polymers

267

FIGURE 9.12 Effects of concentration on (A) total flux (B) water content in the permeate at 60 C. Source: A. Svang-Ariyaskul, R.Y. Huang, P.L. Douglas, R. Pal, X. Feng, P. Chen, et al., Blended chitosan and polyvinyl alcohol membranes for the pervaporation dehydration of isopropanol, J. Membr. Sci. 280 (2006) 815
823. Reproduced with permission from Copyright r 2006 Elsevier Ltd.

Rajeswari et al. [122] have investigated the removal of phosphate ions using polyethylene glycol/chitosan and PVA/chitosan composites. The polyethylene glycol/chitosan and PVA/chitosan composites were found to be effective for the removal of phosphate anions [123]. SEM pictures taken before and after adsorption of phosphate onto PEG/chitosan and PVA/chitosan composites are shown in Fig. 9.14A and B, respectively. The effect of pH was determined at five different pH levels, viz 3, 5, 7, 9, and 11, on the adsorption of phosphate on both PEG/chitosan and PVA/chitosan. The results indicate that the pH of the solution has an influence on the adsorption in the case of phosphate on to PEG/

Handbook of Chitin and Chitosan

268

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.13 Effects of chitosan composition in membranes on (A) swelling degree and (B) amount of water adsorbed in membranes. Source: A. Svang-Ariyaskul, R.Y. Huang, P.L. Douglas, R. Pal, X. Feng, P. Chen, et al., Blended chitosan and polyvinyl alcohol membranes for the pervaporation dehydration of isopropanol, J. Membr. Sci. 280 (2006) 815
823 Reproduced with permission from Copyright r 2006 Elsevier Ltd.

chitosan and PVA/chitosan, that is adsorption efficiency was high at pH below 6. The high adsorption capacity at low pH is mainly due to the strong electrostatic interaction between the positively charged sites of the adsorbent and the phosphate anion Fig. 9.15.

9.5.2 Chitosan/nylon Nylons are also called polyamides because of the characteristic amide groups in the backbone chain. These amide groups are moderately polar, and can hydrogen bond with one another. Nylons are often

Handbook of Chitin and Chitosan

9.5 Chitosan blends with synthetic polymers

269

FIGURE 9.14 SEM images of (A) PEG/chitosan, (B) PVA/chitosan, (C) phosphateadsorbed PEG/chitosan, and (D) phosphate-adsorbed PVA/chitosan. Source: A. Rajeswari, A. Amalraj, A. Pius, Removal of phosphate using chitosan-polymer composites, J. Environ. Chem. Engine. 3 (2015) 2331
2341. Reproduced with permission from Copyright r 2006 Elsevier Ltd.

crystalline due to the polar groups present in the molecule, leading to the formation of hydrogen bonds, which thereby enables tighter packing. Nylon 66 is usually synthesized by reacting adipic acid with hexamethylene diamine [124,125]. Smitha et al. [126] carried out the pervaporation separation of 1,4-dioxane/water mixtures using cross-linked blend membranes of chitosan and nylon 66 (NYL). These membranes were characterized by FTIR, TGA, XRD, and tensile strength to assess the intermolecular interactions, thermal stability, crystallinity, and mechanical strength, respectively. Sorption studies were carried out in pure liquids and binary mixtures of different

Handbook of Chitin and Chitosan

270

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.15 Mechanism for the effect of pH on (A) PEG/chitosan and (B) PVA/chitosan composite. Source: A. Rajeswari, A. Amalraj, A. Pius, Removal of phosphate using chitosan-polymer composites, J. Environ. Chem. Engine. 3 (2015) 2331
2341. Reproduced with permission from Copyright r 2006 Elsevier Ltd.

compositions to evaluate polymer
liquid interactions. The membrane performance was also investigated for the separation of various feed compositions of 1,4-dioxane
water mixtures and permeate pressures. The azeotrope formed at 82-wt.% dioxane was easily broken with a selectivity of 865 and water flux of 0.089 kg/m2 h (Fig. 9.16) [127].

Handbook of Chitin and Chitosan

9.5 Chitosan blends with synthetic polymers

271

FIGURE 9.16 Schematic representation of microstructures of (A) CS-NYL blend, (B) CS-NYL blend cross-linked with sulfuric acid, and (C) interaction of cross-linked polymer matrix with water (A lines, chitosan chains; B line, nylon chain). Source: B. Smitha, G. Dhanuja, S. Sridhar, Dehydration of 1,4-dioxane by pervaporation using modified blend membranes of chitosan and nylon 66, Carbohydr. Polym. 66 (2006) 463
472. Reproduced with permission from Copyright r 2006 Elsevier Ltd.

9.5.3 Chitosan/polyacrylamide Poly(acrylamide) (PAA), a synthetic polymer, is widely used as a flocculent, a paper strengthening agent, and can also be applied to enhance the recovery of oil [128]. Moreover, PAA is a water-soluble polymer of biomedical and pharmaceutical interest widely studied as a hydrogel for blood compatible material [129]. PAA can interact with some other functional groups, such as
COOH,
NH2, and
C 5 O groups, due to its backbone chain having several primary amido groups. With the purpose of producing new polymeric materials that possess both good mechanical properties and biocompatibility, PAA as a candidate blended with chitosan was therefore selected. Poly(acrylamide) is a hydrophilic, highmolecular-weight synthetic polymer which like chitosan has
NH2 groups on its side chain and can form hydrogen bonds with other polymers. PAA has already proved to be a useful material in water purification as a flocculent to bind heavy metal ions by forming coordination bonds [130] and cationic PAA has also been used for antimicrobial applications [131]. However, a superhigh molecular weight is necessary for high flocculating efficiency. Obviously, grafting PAA onto chitosan would improve the properties and increase the potential application of both materials. PAA has been widely used for wastewater treatment. Keyur Desai et al. [132] reported the formation of nonwoven fibers without bead defects by electrospinning blend solutions of chitosan and polyacrylamide (PAAm) with blend ratios varying from 75 to 90 wt.% chitosan using a modified electrospinning unit, wherein polymer solutions can be spun at temperatures greater than ambient up to 100 C. Electrospinning at elevated temperature leads to further expansion of the processing window, by producing fibers with fewer defects at higher chitosan weight

Handbook of Chitin and Chitosan

272

9. Current research on the blends of chitosan as new biomaterials

FIGURE 9.17 SEM images of 1.4 wt.% HMW chitosan/PAAm blends at different blend ratios and hot air blown at 25 ft3/h at different temperatures; (A)
(C) are HMW Chitosan/ PAAm 95:05 blend ratio; (D)
(F) are HMW chitosan/PAAm 90:10 blend ratio; and (G)
(I) are HMW chitosan/PAAm 75:25 blend ratio fibers. Source: K. Desai, K. Kit, Effect of spinning temperature and blend ratios on electrospun chitosan/poly(acrylamide) blends fibers, Polymer 49 (2008) 4046
4050. Reproduced with permission from Copyright r 2008 Elsevier Ltd.

percentage in the blends. Effects of varying blend ratios, spinning temperatures, and molecular weights on fiber formation were studied and optimum conditions for the formation of uniform nonwoven fiber mats with potential applications for air and water filtration were obtained. Uniform beadless fiber mats with fiber diameter as low as 307 6 67 nm were formed by spinning 90% chitosan in blend solutions at 70 C. Fig. 9.17 summarizes the effect of blend ratios and spinning solution temperature on fiber formation. It can be seen that with an increase in temperature the fiber diameter increases slightly and the bead density decreases. SEM images of electrospun solutions containing 95% chitosan (Fig. 9.17A) show poor fiber formation at room temperature, and very few fibers are collected on the target. As the temperature is increased (Fig. 9.17B and C) fiber formation is improved with beadless fibers formed at 70 C. When chitosan content was reduced to 90% (Fig. 17D
F) with increasing spinning temperature, we see the transformation from beaded fibers to uniform beadless fibers.

Handbook of Chitin and Chitosan

9.5 Chitosan blends with synthetic polymers

273

Further reduction to 75% chitosan in the blend leads to the formation of beadless fibers at room temperature (Fig. 9.17G). As spinning temperature is increased (Fig. 9.17H and I), an increase in fiber diameter is seen.

9.5.4 Chitosan/poly(lactic acid) Poly(lactic acid) (PLA) belongs to the family of aliphatic polyester commonly made from lactic acid, which can be produced from renewable resources such as starch via fermentation processes [133]. It is a thermoplastic, high-strength, high-modulus polymer and is considered biodegradable and compostable [134]. Moreover, it is possible to use it for food contact, that is, it is classified as GRAS (generally recognized as safe) [135]. Thanks to its relatively hydrophobic nature the use of this polyester could reduce the hydrophobilic nature of chitosan-based films and consequently improve their moisture barrier properties and decrease overall the water/matrix interactions. Fimbeau Se´bastien et al. [136] have prepared composite films from chitosan and PLA by solution mixing and a film casting procedure. They studied the elaboration and the characterization of chitosan/PLA based biopackaging for potential food applications and the study of antifungal activity of coatings and films on three mycotoxinogen fungal strains, Fusarium proliferatum, Fusarium moniliforme, and Aspergillus ochraceus (Fig. 9.18).

FIGURE 9.18 Ratio of different peak intensities evaluated by FTIR measurements in function of the PLA content. Source: F. Se´bastien, G. Ste´phane, A. Copinet, V. Coma, Novel biodegradable films made from chitosan and poly (lactic acid) with antifungal properties against mycotoxinogen strains, Carbohydr. Polym. 65 (2006) 185
193. Reproduced with permission from Copyright r 2006 Elsevier Ltd.

Handbook of Chitin and Chitosan

274

9. Current research on the blends of chitosan as new biomaterials

The infrared spectra of several composite film samples with 10, 20, and 30% (w/w) of PLA were carried out (data not shown). This would explain the inhibitory properties of composite films further observed. These results were in accordance with those of Peesan, Supaphol, and Rujiravanit (2005) obtained from a FTIR study. These results indicated that no significant interaction between chemically modified chitosan (hexanoyl chitosan) and PLA molecule was observed, even if the same solvent was used.

9.6 Chitosan-based hydrogels Polymers that form hydrogels may have hydrophilic or hydrophobic functional groups. Hydrophilic functional groups, such as hydroxyl (
OH), amine (NH2), and amide (
CONH-CONH2) enable the hydrogel to absorb water leading to hydrogel expansion, which is known as swelling. During swelling, the cross-linked structure of hydrogels prevents complete destruction of the hydrogel cross-links and dissolution. Hydrogels with hydrophobic chains such as PLA have lower water swelling capacity than hydrophilic lattices [137]. Hydrogels can be prepared from synthetic or natural polymers, involving a wide range of chemical compositions and with different mechanical, physical, and chemical properties. Natural polymers such as alginate, chitosan, hyaluronic acid, cellulose, starch, ulvan, gelatin, and pullulan are used to prepare hydrogel.[138
148]. Hydrogels, the three-dimensional networks composed of a polymer backbone, water, and a cross-linking agent to produce a complex network of high molecular weight are gaining much importance in a wide variety of applications in the medical, pharmaceutical, and related fields, such as for wound dressings, contact lenses, artificial organs, and drug delivery systems [149
152]. In recent years, particular interest has been devoted to gels exhibiting phase transitions (i.e., volume change) in response to changes in external conditions, such as pH, ionic strength, temperature, and electric currents, known as the “intelligent” gels [153]. A number of studies on the volume phase transition of synthetic polyelectrolyte amphoteric gels have been reviewed [154
157]. Various hydrogels and microspheres have been employed as injectable scaffolds for a variety of biomedical applications [158
163]. Injectable, biodegradable hydrogels could be utilized as delivery systems, cell carriers, and scaffolds for tissue engineering, which allow easy and homogenous drug or cell distribution within any defect size or shape. Some hydrogels have antibacterial and antifungal activities and these properties could be useful for wound dressings and accelerating the wound healing process.

Handbook of Chitin and Chitosan

References

275

9.7 Conclusions This chapter describes the current research on the blends of chitosan as new biomaterials. Chitosan is a natural polymer that has been proposed for a wide variety of applications ranging from food packaging to as a drug carrier. This chapter is an attempt to better understand chitosan and its blends in various aspects, including its physical, chemical, and biological properties, as well as the preparation and characterization of its blends with other materials and their applications. The number of papers published in the area of polymeric blends may suggest that in the near future a huge interest in the production of new products should arise based on the blends of natural and synthetic polymers.

Acknowledgment The authors would like to thank the University Grants Commission, Government of India for the financial support (F.No.25
1/2014-15/ (BSR)/7
225/2008/(BSR)) and also thank the authorities of Gandhigram Rural Institute-DU for the encouragement.

References [1] A.C. Albertsson, S. Karlsson, Degradable polymers for the future, chemistry and technology of biodegradable polymers, Acta Polym. 46 (1994) 114
123. [2] A.C. Albertsson, S. Karlsson, Macromolecular architecture-nature as a model for degradable, Polym. J. Macromol. Sci. A 33 (1996) 1565
1570. [3] R.A.A. Muzzarelli, Carboxymethylated chitins and chitosans, Carbohydr. Polym. 8 (1988) 1
21. [4] J.P. Jikakis (Ed.), Chitin, Chitosan and Related Enzymes, Academic, Orlando, FL, 1984. [5] Y. Sawayanagi, N. Nambu, T. Nagai, Dissolution properties and bioavailability of phenytoin from ground mixtures with chitin or chitosan, Chem. Pharm. Bull. 31 (1983) 2064
2068. [6] T.C. Yang, R.R. Zull, Absorption of metals by natural polymers generated from seafood processing wastes, Ind. Eng. Chem. Prod. Res. Dev. 23 (1984) 168
172. [7] J. Yanga, I.I. Shih, Y. Tzengc, S. Wang, Production and purification of protease from a Bacillus subtilis that can deproteinize crustacean wastes, Enzyme Microb. Technol. 26 (2000) 406
413. [8] T.A. Khan, K.K. Peh, H.S. Ch’ng, Reporting degree of deacetylation values of chitosan: the influence of analytical methods, J. Pharm. Pharmaceut. Sci. 5 (2002) 205
212. [9] D. Knorr, Use of chitinous polymers in food-a challenge for food research and development, Food Technol-Chicago 38 (1984) 85
97. [10] K. Kurita, β-chitin and the reactivity characteristics, in: M.F.A. Goosen (Ed.), Applications of Chitin and Chitosan, Technomic Publishing, Lancaster, PA, 1997, pp. 79
87. [11] M.F. Bade, Structure and isolation of native animal chitins, in: M.F.A. Goosen (Ed.), Applications of Chitin and Chitosan, Technomic Publishing, Lancaster, PA, 1997, pp. 255
277.

Handbook of Chitin and Chitosan

276

9. Current research on the blends of chitosan as new biomaterials

[12] H.K. No, S.P. Meyers, Preparation of chitin and chitosan, in: R.A.A. Muzzarelli, M.G. Peter (Eds.), Chitin Handbook, Atec Edizioni) (, Grottammare, 1997, pp. 475
489. [13] H.K. No, S.P. Meyers, Preparation and characterization of chitin and chitosan—a review, J. Aqua. Food Prod. Technol. 4 (1995) 27
52. [14] G. Galed, E. Diaz, A. Heras, Conditions of N-deacetylation on chitosan production from alpha chitin, Nat. Prod. Commun. 3 (2008) 543
550. [15] E.S. Abdou, K.S.A. Nagy, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from local sources, Bioresour. Technol. 99 (2008) 1359
1367. [16] F.A. Al Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from marine sources in Arabian Gulf, Carbohydr. Polym. 77 (2009) 410
419. [17] E. Guibal, Interactions of metal ions with chitosan-based sorbents: a review, Sep. Purif. Technol. 38 (2004) 43
74. [18] A. Tolaimate, J. Desbrieres, M. Rhazi, A. Alagui, Contribution to the preparation of chitins and chitosans with controlled physico-chemical properties, Polymer 44 (2003) 7939
7952. [19] V. Singh, K. Kumari, Some physicochemical measurements of chitosan/starch polymers in acetic acid-water mixtures, Macromol. Symp. 320 (2012) 81
86. [20] V.K. Mourya, N.N. Inamdara, A. Tiwari, Carboxymethyl chitosan and its applications, Adv. Mat. Lett. 1 (2010) 11
33. [21] S. Mima, M. Miya, R. Iwamoto, S. Yoshikawa, Highly deacetylated chitosan and its properties, J. Appl. Polym. Sci. 28 (1983) 1909
1917. [22] T. Seo, H. Ohtake, T. Kanbara, K. Yonetake, T. Iijima, Preparation and permeability properties of chitosan membranes having hydrophobic groups, Die Makromol. Chem 192 (1991) 2447
2461. [23] M. Mucha, Rheological characteristics of semi-dilute chitosan solutions, Macromol. Chem. Phys. 198 (1997) 471
484. [24] H. Miyoshi, K. Shimahara, K. Watanabe, K. Onodera, Characterization of some fugal chitosan, Biosci. Biotech. Biochem 56 (1992) 1901
1905. [25] K. Nishimura, S. Nishimura, N. Nishi, I. Saiki, S. Tokura, I. Azuma, Immunological activity of chitin and its derivatives, Vaccine 93 (1984) 93
99. [26] K. Nishimura, S. Nishimura, N. Nishi, F. Numata, Y. Tone, S. Tokura, et al., Adjuvant activity of chitin derivatives in mice and guinea-pigs, Vaccine 3 (1985) 379
384. [27] E.I. Rabea, M.E.T. Badawy, C.V. Stevens, G. Smagghe, W. Steurbaut, Chitosan as antimicrobial agent: applications and mode of action, Biomacromolecules 4 (2003) 1457
1465. [28] C.J. Brine, P.R. Austin, Chitin variability with species and method of preparation, Comp. Biochem. Physiol-B Biochem. Mol. Biol 69 (1981) 283
286. [29] N. Hutadilok, T. Mochimasu, H. Hisamori, K. Hayashi, H. Tachibana, S. Ishii, et al., The effect of N-substitution on the hydrolysis of chitosan by an endo-chitosanase, Carbohydr. Res. 268 (1995) 143
149. [30] R.J. Nordtveit, K.M. Varum, O. Smidsrod, Degradation of partially N-acetylated chitosans with hen egg white and human lysozyme, Carbohydr. Polym. 29 (1996) 163
167. [31] R.A.A. Muzzarelli, Chitin, Pergamon Press Ltd, Hungary, 1977, pp. 83
143. [32] S.E. Lower, Polymers from the sea chitin and chitosan, Manuf. Chem. 55 (1984) 1
17. [33] K. Nagasawa, Y. Tohira, Y. Inoue, N. Tanoura, Reaction between carbohydrates and sulfuric acid: part I. Depolymerization and sulfation of polysaccharides by sulfuric acid, Carbohydr. Res. 18 (1971) 95
102. [34] A. Domard, Physico-chemical and structural basis for applicability of chitin and chitosan, in: The proceedings of the Second Asia Pacific Symposium of chitin and Chitosan, Bangkok, Thailand, vol. 1, 1996, pp. 8
12.

Handbook of Chitin and Chitosan

References

277

[35] M.G. Peter, Applications and environmental aspects of chitin and chitosan, J. Macromol. Sci. Pure Appl. Chem. 32 (1995) 629
640. [36] S. Aiba, Studies on chitosan: 3. Evidence for the presence of random and block copolymer structures in partially N-acetylated chitosans, Int. J. Biol. Macromol 13 (1991) 40
44. [37] N. Kubota, Y. Eguchi, Facile preparation of water-soluble N-acetylated chitosan and molecular weight dependence of its water-solubility, Polym. J. 29 (1997) 123
127. [38] K. Kurita, Controlled functionalization of the polysaccharide chitin, Prog. Polym. Sci. 26 (2001) 1921
1971. [39] A. Denuziere, D. Ferrier, O. Damour, A. Domard, Chitosan-chondroitin sulfate and chitosan
hyaluronate polyelectrolyte complexes: biological properties, Biomaterials 19 (1998) 1275
1285. [40] Q. Li, E.T. Dunn, E.W. Grandmaison, M.F. Goosen, Applications and properties of chitosan, J. Bioact. Compat. Pol. 7 (1992) 1
9. [41] P.A. Sandford, Front C. High purity chitosan and alginate: preparation, analysis, and applications, Carbohydr. Res. 2 (1992) 250
269. [42] J.J. Nagyvary, J.D. Falk, M.L. Hill, M.L. Schmidt, A.K. Wilkins, E.L. Bradbury, The hypolipidemic activity of chitosan and other polysaccharides in rats, Nutr. Rep. Int. 20 (1979) 677
684. [43] M. Sugano, S. Watanabe, A. Kishi, M. Izume, A. Ohtakara, Hypocholesterolemic action of chitosans with different viscosity in rats, Lipids 23 (1988) 187
191. [44] R.A.A. Muzzarelli, M.G. Peter (Eds.), Chitin Handbook, Atec Edizioni, Grottammare, 1997. [45] P.K. Dutta, M.N.V. Ravi Kumar, J. Dutta, Chitin and chitosan: chemistry, properties and applications, JMS—Polym. Rev 42 (2002) 20
31. [46] Y.I. Cho, H.K. No, S.P. Meyers, Physicochemical characteristics and functional properties of various commercial chitin and chitosan products, J. Agric. Food Chem. 46 (1998) 3839
3843. [47] S.K. Rout, Physicochemical, Functional and Spectroscopic Analysis of Crawfish Chitin and Chitosan as Affected by Process Modification (LSU Historical Dissertations and Theses), Dissertation, 2001. [48] http://www.plasticstrends.net. [49] J. Zhang, W. Xia, P. Liu, Q. Cheng, T. Tahirou, W. Gu, et al., Chitosan modification and pharmaceutical/biomedical applications, Mar. Drugs 8 (2010) 1962
1987. [50] G.R. Strobl (Ed.), The Physics of Polymer: Concept for Understanding their Structure and Behavior, Springer-Verlag, New York, 1996. [51] A.R. Sarasam, R.K. Krishnaswamy, S.V. Madihally, Blending chitosan with polycaprolactone: Effects on physicochemical and antibacterial properties, Biomacromolecules 7 (2006) 1131
1138. [52] Y. Wan, H. Wu, A.X. Yu, D.J. Wen, Biodegradable polylactide/chitosan blend membranes, Biomacromolecules 7 (2006) 1362
1372. [53] D.F. Varenell, J.P. Runt, M.M. Coleman, FT ir and thermal analysis studies of blends of poly (ε-caprolactone) with homo-and copolymers of poly (vinylidene chloride), Polymer 24 (1983) 37
42. [54] D.F. Varenell, M.M. Coleman, FT i.r. studies of polymer blends: V. Further observations on polyester-poly(vinyl chloride) blends, Polymer 22 (1981) 1324
1328. [55] E.M. Woo, J.W. Barlow, D.R. Paul, Phase behavior of blends of aliphatic polyesters with a vinylidene chloride/vinyl chloride copolymer, J. Appl. Polym. Sci. 32 (1986) 3889
3897. [56] Y. Liu, M.C. Messmer, Surface structures and segregation of polystyrene/poly (methyl methacrylate) blends studied by sum-frequency (sf) spectroscopy, J. Phys. Chem. B 107 (2003) 9774
9779.

Handbook of Chitin and Chitosan

278

9. Current research on the blends of chitosan as new biomaterials

[57] X. Zhang, D.M. Kale, S.A. Jenekhe, Electroluminescence of multicomponent conjugated polymers. 2. Photophysics and enhancement of electroluminescence from blends of polyquinolines, Macromolecules 35 (2002) 382
393. [58] K.K. Chee, Determination of polymer-polymer miscibility by viscometry, Eur. Polym. Mater 26 (1990) 423
426. [59] R. Paladhi, R.P. Singh, Ultrasonic and rheological investigations on interacting blend solutions of poly(acrylic acid) with poly(vinyl pyrrolidone) or poly(vinyl alcohol), Eur. Polym. Mater 30 (1994) 251
257. [60] U.S. Toti, T.M. Aminabhavi, Different viscosity grade sodium alginate and modified sodium alginate membranes in pervaporation separation of water 1 acetic acid and water 1 isopropanol mixtures, J. Membr. Sci. 228 (2004) 199
208. [61] M.N.V. Ravikumar, R.A.A. Muzzarelli, C. Muzzarelli, H. Sashiwa, A.J. Domb, Chitosan chemistry and pharmaceutical perspectives, Chem. Rev. 104 (2004) 6017
6084. [62] D.K. Singh, A.R. Ray, Radiation-induced grafting of N,N0 -dimethylamino ethylmethacrylate onto chitosan films, J. Appl. Polym. Sci. 66 (1997) 869
877. [63] S. Yoshikawa, T. Takeshi, N. Tsubokawa, Grafting reaction of living polymer cations with amino groups on chitosan powder, J. Appl. Polym. Sci. 68 (1998) 1883
1889. [64] V.M. Correlo, L.F. Boesel, M. Bhattacharya, J.F. Mano, N.M. Neves, R.L. Reis, Grafting reaction of living polymer cations with amino groups on chitosan powder, Mater. Sci. Eng. A 403 (2005) 57
68. [65] N.F.M. Nasir, N.M. Zain, M.G. Raha, N.A. Kadri, Characterization of chitosan-poly (ethylene oxide) blends as haemodialysis membrane, Am. J. Eng. Appl. Sci 2 (2005) 1578
1583. [66] Qing-lei Qi, Qiang Li, Jian-wei Lu, Zhao-xia Guo, Jian Yu, Preparation and characterization of soluble eggshell membrane protein/chitosan blend films, Chinese J. Polym. Sci. 27 (2009) 387
392. [67] J.C. Cabanelas, B. Serrano, J. Baselga, Development of cocontinuous morphologies in initially heterogeneous thermosets blended with poly (methyl methacrylate), Macromolecules 38 (2005) 961
970. [68] O. Olabisi, L.M. Robeson, M.T. Shaw, Polymer
Polymer Miscibility, Academic Press, New York, 1979, pp. 164
169. [69] W.H. Jiang, S. Han, An improved criterion of polymer
polymer miscibility determined by viscometry, Europ. Polym. J. 34 (1998) 1579
1584. [70] K. Lewandowska, The miscibility of poly(vinyl alcohol)/poly(N-vinylpyrrolidone) blends investigated in dilute solutions and solids, Europ. Polym. J. 41 (2005) 55
64. [71] D.K. Kweon, D.W. Kang, Drug-release behavior of chitosan-g-poly(vinyl alcohol) copolymer matrix, J. Appl. Polym. Sci. 74 (1999) 458
464. [72] M. Zhang, X.H. Li, Y.D. Gong, N.M. Zhao, X.F. Zhang, Properties and biocompatibility of chitosan films modified by blending with PEG, Biomaterials 23 (2002) 2641
2648. [73] I. Arvanitoyannis, I. Nakayama, S. Aiba, Chitosan and gelatin based edible films: state diagrams, mechanical and permeation properties, Carbohydr. Polym. 37 (1998) 371
382. [74] A. Isogai, R.H. Atalla, Preparation of cellulose-chitosan polymer blends, Carbohydr. Polym. 25 (1992) 25
28. [75] J.W. Lee, S.Y. Kim, S.S. Kim, Y.M. Lee, K.H. Lee, S.J. Kim, Synthesis and characteristics of interpenetrating polymer network hydrogel composed of chitosan and poly (acrylic acid), J. Appl. Polym. Sci. 73 (1999) 113
120. [76] S.R. Park, K.Y. Lee, W.S. Ha, S.Y. Park, Structural changes and their effect on mechanical properties of silk fibroin/chitosan blends, J. Appl. Polym. Sci. 74 (1999) 2571
2575.

Handbook of Chitin and Chitosan

References

279

[77] J.A. Ratto, C.C. Chen, R.B. Blumstein, Phase behavior study of chitosan/polyamide blends, J. Appl. Polym. Sci. 59 (1996) 1451
1461. [78] M.M. Amiji, Pyrene fluorescence study of chitosan self-association in aqueous solution, Carbohydr. Polym. 26 (1995) 211
213. [79] T. Chandy, C.P. Sharma, Prostaglandin E1-immobilized poly (vinyl alcohol)-blended chitosan membranes: blood compatibility and permeability properties, J. Appl. Polym. Sci. 44 (1992) 2145
2156. [80] M. Ishihara, K. Nakanishi, K. Ono, M. Sato, M. Kikuchi, Y. Saito, et al., Photocrosslinkable chitosan as a dressing for wound occlusion and accelerator in healing process, Biomaterials 23 (2002) 833
840. [81] Y.B. Wu, S.H. Yu, F.L. Mi, C.W. Wu, S.S. Shyu, C.K. Peng, Preparation and characterization on mechanical and antibacterial properties of chitsoan/cellulose blends, Carbohydr. Polym. 57 (2004) 435
440. [82] M. Ratajska, S. Boryniec, Physical and chemical aspects of biodegradation of natural polymers, Reactive Funct. Polym. 38 (1998) 35
49. [83] P. Forssell, R. Lahtinen, M. Lahelin, P. Mylla¨rinen, Oxygen permeabilityof amylose and amylopectin films, Carbohydr. Polym 47 (2002) 125
129. [84] J.M. Raquez, Y. Nabar, M. Srinivasan, B.Y. Shin, R. Narayan, P. Dubois, Maleated thermoplastic starch by reactive extrusion, Carbohydr. Polym. 74 (2008) 159
169. [85] A. Rindlava, S.H.D. Hulleman, P. Gatenholma, Formation of starch filmswith varying crystallinity, Carbohydr. Polym. 34 (1997) 25
30. [86] X.Y. Xu, K.M. Kim, M.A. Hanna, D. Nag, Chitosan-starch composite film: preparation and characterization, Ind. Crops Prod. 21 (2005) 185
192. [87] J.H. Jagannath, C. Nanjappa, D.K. Das Gupta, A.S. Bawa, Mechanical and barrier properties of edible starch-protein-based films, J. Appl. Polym. Sci. 88 (2003) 64
68. [88] A. Jangchud, M.S. Chinnan, Properties of peanut protein film: sorption isotherm and plasticizer effect, LWT—Food Sci. Technol 2 (1999) 89
94. [89] T. Bourtoom, M.S. Chinnan, Preparation and properties of rice starch—chitosan blend biodegradable film, LWT—Food Sci. Technol 41 (2008) 1633
1641. [90] S. Cao, H. Li, K. Li, J. Lu, L. Zhang, In vitro mineralization of MC3T3-E1 osteoblastlike cells on collagen/nano-hydroxyapatite scaffolds coated carbon/carbon composites, J. Biomed. Mater. Res. A 104 (2016) 533
543. [91] J. Liu, et al., The effect of different molecular weight collagen peptides on MC3T3-E1 cells differentiation, Biomed. Mater. Eng. 26 (Suppl. 1) (2015) 2041
2047. [92] R.M. Salasznyk, W.A. Williams, A. Boskey, A. Batorsky, G.E. Plopper, Adhesion to vitronectin and collagen I promotes osteogenic differentiation of human mesenchymal stem cells, J. Biomed. Biotechnol. 2004 (2004) 24
34. [93] I.Y. Kim, Chitosan and its derivatives for tissue engineering applications, Biotechnol. Adv. 26 (2008) 1
21. [94] L. Wang, J.P. Stegemann, Thermogelling chitosan and collagen composite hydrogels initiated with beta-glycerophosphate for bone tissue engineering, Biomaterials 31 (2010) 3976
3985. [95] A. Martı´neza, M.D. Blanco, N. Davidenko, R.E. Cameron, Tailoring chitosan/ collagen scaffolds for tissue engineering: effect ofcomposition and different crosslinking agents on scaffold properties, Carbohydr. Polym. 132 (2015) 606
619. [96] H. Huang, N. Hu, Y. Zeng, G. Zhou, Electrochemistry and electrocatalysis with heme proteins in chitosan biopolymer films, Anal. Biochem. 308 (2002) 141
151. [97] F.L. Mi, Y.B. Wu, S.S. Shyu, A.C. Chao, J.Y. Lai, C.C. Su, Asymmetric chitosan membranes prepared by dry/wet phase separation: a new type of wound dressing for controlled antibacterial release, J. Membr. Sci. 212 (2003) 237
254. [98] P.A. Gunatilake, R. Adhikari, Biodegradable synthetic polymers for tissue engineering, Eur. Cells Mat 5 (2003) 1
16.

Handbook of Chitin and Chitosan

280

9. Current research on the blends of chitosan as new biomaterials

[99] T.A. Khan, K.K. Peh, Pharm, a preliminary investigation of chitosan film as dressing for punch biopsy wounds in rats, Pharm. Sci 6 (2003) 20
26. [100] D. Bakos, J. Koller, Polymer based systems on tissue engineering, replacement and regeneration, in: R.L. Reis, D. Cohn (Eds.), NATO Science Series, Kluwer Academic Publishers, 2001, p. 55. [101] D. Demirgoz, C. Elvira, J.F. Mano, A.M. Cunha, E. Piskin, R.L. Reis, Chemical modification of starch based biodegradable polymeric blends: effects on water uptake, degradation behaviour and mechanical properties, Polym. Degrad. Stabil. 70 (2000) 161
170. [102] F.L. Mi, S.S. Shyu, Y.B. Wu, S.T. Lee, J.Y. Shyong, R.N. Huang, Fabrication and characterization of a sponge-like asymmetric chitosan membrane as a wound dressing, Biomaterials 22 (2001) 165
173. [103] C.M. Vaz, M. Fossen, R.F. Van Tuil, L.A. De Graaf, R.L. Reis, A.M. Cunha, Casein and soybean protein-based thermoplastics and composites as alternative biodegradable polymers for biomedical applications, J. Biom. Mater. Res. 65A (2003) 60
70. [104] R.M. Silva, G.A. Silva, O.P. Coutinho, J.F. Mano, R.L. Reis, Physical properties and biocompatibility of chitosan/soy blended membranes, J. Mater. Sci. Mater. Med. (2003) 1
14. [105] L.E. Abugoch, C. Tapia, M.C. Villama´n, M. Yazdani-Pedram, M. Dı´az-Dosque, Characterization of quinoa protein
chitosan blend edible films, Food Hydrocoll. 25 (2011) 879
886. [106] G. Ritthidej, T. Phaechamud, T. Koizumi, Moist heat treatment on physicochemical change of chitosan salt films, Int. J. Pharm. 232 (2002) 11
22. [107] B. Wang, K. Wang, W. Dan, T. Zhang, Y. Ye, Konjac glucomannan collagen-chitosan blend films, J. Biomed. Sci. Eng. 23 (2006) 102
106. [108] H. Ismail, S.M. Shaari, N. Othman, The effect of chitosan loading on the curing characteristics, mechanical and morphological properties of chitosan-filled natural rubber (NR), epoxidised natural rubber (ENR) and styrene-butadiene rubber (SBR) compounds, Poly. Testing 30 (2011) 784
790. [109] I. Engelberg, J. Kohn, Physico-mechanical properties of degradable polymers used in medical applications: a comparative study, Biomaterials 12 (1991) 292
304. [110] S.Z. Rogovina, G.A. Vikhoreva, Polysaccharide-based polymer blends: methods of their production, Glycoconjug. J. 23 (2006) 611
618. [111] Y.M. Lee, Modified chitosan membranes for pervaporation, Desalination 90 (1993) 277
290. [112] L.G. Wu, C.L. Zhu, M. Liu, Study of a new pervaporation membrane. Part 1. Preparation and characteristics of the new membrane, J. Membr. Sci 90 (1994) 199
216. [113] T. Uragami, T. Masuda, T. Miyata, Structure of chemically modified chitosan membranes and their characteristics of permeation and separation of aqueous ethanol solutions, J. Membr. Sci. 88 (1994) 243
249. [114] X. Feng, R.Y.M. Huang, Pervaporation with chitosan membranes. I. Separation of water from ethylene glycol by a chitosan/polysulfone composite membrane, J. Membr. Sci. 116 (1996) 67
76. [115] J. Shieh, R.Y.M. Huang, Pervaporation with chitosan membranes II. Blended membranes of chitosan and polyacrylic acid and comparison of homogenous and compostie membrane based on polyelectrolyte complexes of chitosan and polyacrylic acid for the separation of ethanol
water mixtures, J. Membr. Sci. 127 (1997) 185
202. [116] J. Neel, Introduction to pervaporation, in: R.Y.M. Huang (Ed.), Pervaporation Membrane Separation Processes, Elsevier, Amsterdam, 1991. [117] N.P. Wynn, Pervaporation, Kirk-Othmer Encyclopedia of Chemical Technology, John Wiley & Sons, 2003.

Handbook of Chitin and Chitosan

References

281

[118] H.L. Fleming, C.S. Slater, Part III pervaporation, in: W.S.W. Ho, K.K. Sirkar (Eds.), Membrane Handbook, Van Nostrand Reinhold, New York, 1992. [119] A. Jonquieres, R. Clement, P. Lochon, J. Neel, M. Dresch, B. Chretien, Industrial state-of-the-art of pervaporation and vapour permeation in the western countries, J. Membr. Sci. 206 (2002) 87
94. [120] J. Yu, C.H. Lee, W.H. Hong, Performances of crosslinked asymmetric poly(vinyl alcohol) membranes for isopropanol dehydration by pervaporation, Chem. Eng. Process. 41 (2002) 693
706. [121] A. Svang-Ariyaskul, R.Y. Huang, P.L. Douglas, R. Pal, X. Feng, P. Chen, et al., Blended chitosan and polyvinyl alcohol membranes for the pervaporation dehydration of isopropanol, J. Membr. Sci. 280 (2006) 815
823. [122] A. Rajeswari, A. Amalraj, A. Pius, Removal of phosphate using chitosan-polymer composites, J. Environ. Chem. Engine 3 (2015) 2331
2341. [123] J. Brandrup, E.H. Immergut, E.A. Grulke, Polymer handbook, fourth ed., Wiley, New York, 1999. Deanin, R. D. Polymer structure, properties and application (1972). [124] V.R. Gowariker, N.V. Vishwanathan, J. Sridhar. Polymer Science. New Age International Ltd, New Delhi, 1999. [125] P. Meares, Polymer Structure and Bulk Properties, Van Nostrand, London, 1965. [126] B. Smitha, G. Dhanuja, S. Sridhar, Dehydration of 1, 4-dioxane by pervaporation using modified blend membranes of chitosan and nylon 66, Carbohydr. Polym. 66 (2006) 463
472. [127] F. Yan, C. Zheng, X. Zhai, D. Zhao, Preparation and characterization of polyacrylamide in cationic microemulsion, J. Appl. Polym. Sci. 67 (1998) 747
754. [128] N.A. Peppas, Chapter 2—Classes of materials used in medicine, Biomater. Sci 1 (1996) 37
130. [129] T. Siyam, Development of acrylamide polymers for the treatment of waste water, Des. Monomers Polym. 4 (2001) 107
168. [130] B.J. Gao, S.X. He, J.F. Guo, R.X. Wang, Antibacterial property and mechanism of copolymer of acrylamide and quaternary salt of 4-vinyl pyridine, J. Appl. Polym. Sci. 2 (2006) 1531
1537. [131] C. Xiao, L. Weng, Y. Lu, L. Zhang, Blend films from chitosan and polyacrylamide solutions, J. Macromol. Sci. Pure Appl. Chem. 8 (2001) 761
771. [132] K. Desai, K. Kit, Effect of spinning temperature and blend ratios on electrospun chitosan/poly(acrylamide) blends fibers, Polymer 49 (2008) 4046
4050. [133] D. Garlotta, A literature review of poly (lactic acid), J. Polym. Environ 9 (2001) 63
84. [134] A. Jarerat, Y. Tokiwa, Degradation of poly (L-lactide) by a fungus, Macromol. Biosci. 1 (2001) 136
140. [135] R.E. Conn, J.J. Kolstad, J.F. Borzelleca, D.S. Dixler, L.J. Filer, B.N. LaDu, Safety assessment of polylactide (PLA) for use as a food-contact polymer, Food Chem. Toxicol 33 (1995) 273
283. [136] F. Se´bastien, G. Ste´phane, A. Copinet, V. Coma, Novel biodegradable films made from chitosan and poly (lactic acid) with antifungal properties against mycotoxinogen strains, Carbohydr. Polym. 65 (2006) 185
193. [137] F. Ahmadi, Z. Oveisi, S. Mohammadi Samani, Z. Amoozgar, Chitosan based hydrogels: characteristics and pharmaceutical applications, Res. Pharma. Sci 10 (2015) 1
16. [138] K. Murakami, H. Aoki, S. Nakamura, S. Nakamura, M. Takikawa, M. Hanzawa, et al., Hydrogel blends of chitin/chitosan, fucoidan and alginate as healingimpaired wound dressings, Biomaterials 31 (2010) 83
90. [139] B. Balakrishnan, M. Mohanty, P.R. Umashankar, A. Jayakrishnan, Evaluation of an in situ forming hydrogel wound dressing based on oxidized alginate and gelatin, Biomaterials 26 (2005) 6335
6342.

Handbook of Chitin and Chitosan

282

9. Current research on the blends of chitosan as new biomaterials

[140] J.O. Kim, J.K. Park, J.H. Kim, S.G. Jin, C.S. Yong, D.X. Li, et al., Development of polyvinyl alcohol
sodium alginate gel-matrix-based wound dressing system containing nitrofurazone, Int. J. Pharma 359 (2008) 79
86. [141] E.A. Kamoun, E.S. Kenawy, T.M. Tamer, M.A. El-Meligy, M.S. Mohy Eldin, Poly (vinyl alcohol)- alginate physically crosslinked hydrogel membranes for wound dressing applications: characterization and bio-evaluation, Arabian J. Chem. 8 (2015) 38
47. [142] N.B. Milosavljevi´c, L.M. Kljajevi´c, I.G. Popovi´c, J.M. Filipovi´c, M.T. Kalagasidis Kruˇsi´c, Chitosan, itaconic acid and poly(vinyl alcohol) hybrid polymer networks of high degree of swelling and good mechanical strength, Polym. Int. 59 (2010) 686
694. [143] L. Racine, I. Texier, R. Auze´ly-Velty, Chitosan-based hydrogels: recent design concepts to tailor properties and functions, Polym. Int. 66 (2017) 981
998. [144] L.P. Silva, T.C. Santos, D.B. Rodrigues, R.P. Pirraco, M.T. Cerqueira, R. Reis, et al., Stem cell-containing hyaluronic acid-based spongy hydrogels for integrated diabetic wound healing, J. Investig. Dermatology 137 (2017) 1541
1551. [145] Y. Luo, K.R. Kirker, G.D. Prestwich, Cross-linked hyaluronic acid hydrogel films: new biomaterials for drug delivery, J. Control. Release 69 (2000) 169
184. [146] T. Segura, B.C. Anderson, P.H. Chung, R.E. Webber, K.R. Shull, L.D. Shea, Crosslinked hyaluronic acid hydrogels: a strategy to functionalize and pattern, Biomaterials 26 (2005) 359
371. [147] C. Chang, B. Duan, L. Zhang, Fabrication and characterization of novel macroporous cellulose
alginate hydrogels, Polymer 50 (2009) 5467
5473. [148] T. Maneerung, S. Tokura, R. Rujiravanit, Impregnation of silver nanoparticles into bacterial cellulose for antimicrobial wound dressing, Carbohydr. Polym. 72 (2008) 43
51. [149] N.A. Peppas, Hydrogels in Medicine and Pharmacy, CRC Press, Boca Raton, FL, 1985. [150] G.V.N.D. Rthna, D.V. Mohan Rao, P.R. Chatterji, Hydrogels of gelatin-sodium carboxy methyl cellulose: synthesis and swelling kinetics, J. Mater. Sci. Pure Appl. Chem 33 (1996) 1199
1207. [151] N.J. Einerson, K.R. Stevens, W.J. Kao, Synthesis and physicochemical analysis of gelatin-based hydrogels for drug carrier matrices, Biomaterials 24 (2002) 509
523. [152] N.A. Peppas, K.B. Keys, M. Torres-Lugo, A.M. Lowman, Poly(ethylene glyco)-containing hydrogels in drug delivery, J. Control Rel. 62 (1999) 81
87. [153] T. Tanaka, D. Fillmor, S.T. Sun, I. Nishio, Phase transitions in ionic gel, Phys. Rev. Lett. 45 (1980) 1636
1639. [154] E.K. Sarkyt, B.S. Vladimir, Swelling, shrinking, deformation and oscillation of polyampholyte gels based on vinyl 2-aminoethyl ether and sodium acrylate, Langmuir 15 (1999) 4230
4235. [155] H.H. Hooper, J.P. Baker, H.W. Blanch, J.M. Prausnitz, Swelling equilibria for positively ionized polyacrylamide hydrogels, Macromolecules 23 (1990) 1096
1104. [156] J.P. Baker, D.R. Stephens, H.W. Blanch, J.M. Prausnitz, Swelling equilibria for acrylamide-based polyampholyte hydrogels, Macromolecules 25 (1992) 1955
1998. [157] S. Katayama, A. Myoga, Y. Akahori, Swelling behavior of amphoteric gel and the volume phase transition, J. Phys. Chem. 96 (1992) 4698
4701. [158] J.S. Tememoff, A.G. Mikos, Injectable biodegradable materials for orthopedic tissue engineering, Biomaterials 21 (2000) 2405
2412. [159] J.L. Drury, D.J. Mooney, Hydrogels for tissue engineering: scaffold design variables and applications, Biomaterials 24 (2003) 4337
4351. [160] Q.P. Hou, P.A. De Bank, K.M. Shakesheff, Injectable scaffolds for tissue regeneration, J. Mater. Chem. 14 (2004) 1915
1923.

Handbook of Chitin and Chitosan

References

283

[161] A.B. Pratt, F.E. Weber, H.G. Schmoekel, R. Muller, J.A. Hubbell, Synthetic extracellular matrices for in situ tissue engineering, Biotechnol. Bioeng. 86 (2004) 27
36. [162] L. Lao, H. Tan, Y. Wang, C. Gao, Chitosan modified poly(L-lactide) microspheres as cell microcarriers for cartilage tissue engineering, Colloids Surf.- B Biointerf. 66 (2008) 218
225. [163] J.B. McGlohorn, L.W. Grimes, S.S. Webster, K.J.L. Burg, Characterization of cellular carriers for use in injectable tissue-engineering composites, J. Biomed. Mater. Res. 66A (2003) 441
449.

Handbook of Chitin and Chitosan

C H A P T E R

10 Chitin and chitosan-based aerogels E. Jackcina Stobel Christy, A. Rajeswari, Sreerag Gopi and Anitha Pius Department of Chemistry, The Gandhigram Rural Institute—Deemed to be University, Dindigul, India O U T L I N E 10.1 Introduction 10.1.1 Classification of aerogels 10.1.2 Chitin and chitosan: sources and properties

286 288 290

10.2 Chitin and chitosan-based aerogels: preparation process

292

10.3 Characterization of chitin and chitosan-based aerogels 10.3.1 Morphological analysis/microscopic analysis 10.3.2 Porosity 10.3.3 Thermal properties 10.3.4 Raman spectra 10.3.5 Tensile property of aerogels/mechanical property 10.3.6 Fourier transform infrared spectroscopy 10.3.7 X-ray diffraction 10.3.8 X-ray photoelectron spectroscopy 10.3.9 Nuclear magnetic resonance spectroscopy 10.3.10 Water contact angle

296 296 304 309 312 314 317 319 319 322 323

10.4 Future aspects of aerogel

324

10.5 Conclusions

327

Acknowledgments

328

References

328

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00010-9

285

© 2020 Elsevier Inc. All rights reserved.

286

10. Chitin and chitosan-based aerogels

10.1 Introduction Research on existing materials evolves and develops to create new and innovative materials that appear to be socially and economically useful. New materials can show superior functionality and performance to existing materials and are finally subjected to practical uses in which the safety and reliability of the materials plays an important role. In this context materials research has emerged as one of the key technologies for cutting-edge industries and an exciting area of research crossing the fields of science, engineering, and technology. It is influenced by various economic factors and conditions, as well as by national and social needs. The new generation equipment being developed must be lighter, stronger, tougher, consume less energy, and be adaptable to the current environmental situation. The fundamental constituents of this advancement are perhaps the development of nanomaterials, thin films, quantum dots, aerogels, membranes etc. for technologically important applications such as water treatments, photocatalystss, supercapacitor electrodes, optoelectronics, storage devices, gas sensors, detectors, hygienic products, agriculture, drug delivery systems, sealing, coal dewatering, food additives, pharmaceuticals, biomedical applications like tissue engineering and regenerative medicines, diagnostics, wound dressing, separation of biomolecules or cells, barrier materials to regulate biological adhesions, and also biosensors (Fig. 10.1). Among the other trending material research, aerogels have tremendous potential in a wide range of applications in which high pore

FIGURE 10.1 Overview diagram for application of material research.

Handbook of Chitin and Chitosan

10.1 Introduction

287

volume and high surface area play major roles. Aerogels are the lightest solid material known and are produced nowadays at an industrial scale [1]. Aerogels are low-density nanoporous solids with a fine, open-pore structure resulting in low densities (0.003 0.15 kg/m3), high porosity, and large surface areas (500 1000 m2/g) [2]. This combination of properties makes them unique in many ways, opening possibilities to a huge number of applications, some of which have been commercialized already [3]. Aerogels were first introduced by Dr. S. Kistler in the 1930s when he extracted the pore-filing liquid of wet gels using a supercritical drying approach [4] to obtain an air-filled solid material with nearly the same dimension as the original wet gel [5]. He defined areogels as “gels in which the liquid has been replaced by air, with very moderate shrinkage of the solid network.” The definition of aerogels has been discussed a lot; according to Smirnova and Gurikov an aerogel is an open colloidal or polymeric network consisting of loosely packed, bonded particles or fibers that is expanded throughout its volume by gas and therefore exhibits very low density [1] (Fig. 10.2). After the 1930s more and more interest was given to aerogel preparation due to its applications like thermal insulation, electrochemistry (supercapacitors), carrier of catalysts and other active agents, filling materials, materials for tissue engineering, etc. The aerogels were prepared by varying the precursor materials, for example, organic-based

FIGURE 10.2

Dr. S. Kistler, inventor of aerogels.

Handbook of Chitin and Chitosan

288

10. Chitin and chitosan-based aerogels

aerogels, natural compounds-based aerogels, bioaerogels, polymerbased aerogels. This was a new trend in the aerogel field, although the idea was proposed by Dr. Kistler who prepared protein (ovalbumin and collagen)-based aerogels. In general, aerogels are composed of a network of clustered nanoparticles. The materials usually have unique properties including high strength to density and high surface area to volume ratios. They are manufactured by subjecting a wet gel precursor to critical point drying in order to remove the liquid through supercritical drying, without disturbing the network.

10.1.1 Classification of aerogels Generally, aerogels can be classified based on their material application, porous structure, and by their composition as explained in Fig. 10.3. • Based on their application as materials, aerogels are classified as monolith, powder, and film forms. • Aerogels are classified based on specific properties, such as photoluminescence (quantum dots and metals chalcogenide aerogels), high catalytic property (semiconductor aerogels), good conductivity (metal aerogels), high surface area, supercapacitors (carbon and metal-based aerogels), oil sorption (hydrophobic silica aerogels and polymeric aerogels), high elasticity (graphene-coated aerogels). This field of aerogels is improving to attain good mechanical properties. Organic aerogels are stronger, and made up of organic polymers, but they have limitations for use in situations that need robustness. However, based on their composition, advances in aerogel synthesis and their drying technologies, aerogels are classified as • inorganic (such as SiO2 [10], TiO2, Al2O3, ZrO2, Cu NW aerogel [11]); • organic [i.e., resorcinol-formaldehyde (RF), poly(vinyl alcohol) and melamine-formaldehyde aerogels [12] polyurethane, polyimide, polystyrene, etc.] [9,10]; or • carbon (i.e., carbon, carbon nanotubes, graphene) [11 14]. Additionally, besides the single-component aerogels mentioned above the compositing of one of those aerogels with a specific component has often conferred an additional functionality, such as mechanical strength (glass bead; GB) and nanosilica aerogel (NSA), hydrophobicity, removal and catalytic features to pristine materials [polydopamine onto the surface of CNFs, which were cross-linked with polyethylenimine (PEI) to form the aerogels], and has enlarged the above classification for some high-performance applications [5,13 16].

Handbook of Chitin and Chitosan

10.1 Introduction

289

FIGURE 10.3 Classification of aerogels. Source: Reprinted with permission from X.D. Gao, Y. Di Huang, T.T. Zhang, Y.Q. Wu, X.M. Li, Amphiphilic SiO2 hybrid aerogel: an effective absorbent for emulsified wastewater, J. Mater. Chem. A. 5 (2017) 12856 12862. doi:10.1039/ c7ta02196h [6] Royal Society of Chemistry, Copyright r 2017; S. Naskar, J.F. Miethe, S. Sa´nchezParadinas, N. Schmidt, K. Kanthasamy, P. Behrens, et al., Photoluminescent aerogels from quantum wells, Chem. Mater. 28 (2016) 2089 2099. doi:10.1021/acs.chemmater.5b04872 [7] American Chemical Society, Copyright r 2016; A.M. Joseph, B. Nagendra, P. Shaiju, K.P. Surendran, E.B. Gowd, Aerogels of hierarchically porous syndiotactic polystyrene with a dielectric constant near to air, J. Mater. Chem. C. 6 (2018) 360 368. doi:10.1039/c7tc05102f [8] Royal Society of Chemistry, Copyright r 2017; and Y. Tang, K.L. Yeo, Y. Chen, L.W. Yap, W. Xiong, W. Cheng, Ultralowdensity copper nanowire aerogel monoliths with tunable mechanical and electrical properties, J. Mater. Chem. A. 1 (2013) 6723 6726. doi:10.1039/c3ta10969k [9] Royal Society of Chemistry, Copyright r 2013.

Apart from the conventional aerogels, sustainable environment materials like biopolymers are driving researchers to develop novel aerogels with biocompatibility, abundant resources, low environmental loads, and flexibility for functionalization [17]. Biopolymers are a class of polymers produced by living organisms which represent the most abundant organic compounds on Earth with suitable properties for sustainable chemistry today. They are generally classified as polysaccharides, polypeptides, polynucleotides, polyisoprenoids, and polyphenols, in which cellulose, starch, chitin and chitosan, proteins and peptides, lignin alginate, gelatine, etc. are biopolymers (i.e., polysaccharides) with characteristic polymeric properties,

Handbook of Chitin and Chitosan

290

10. Chitin and chitosan-based aerogels

structural features, functional groups, nontoxic aquatic nature, etc. [18 21]. The intrinsic advantages of biopolymers with the unique threedimensional structures of aerogels have opened up attractive new potential applications in flexible thermal superinsulation, biomedical scaffolds, energy storage devices, drug carriers, catalysts, flame retardants, oil and water separations, and food processing [19,20,22 25]. Recently, chitin and chitosan-based aerogels have attracted a lot of attention from researchers. The importance of these biopolymers is increasing and is highlighted in the form of a high number of publications. Chitin (Ch), the second abundant biopolymer present as a highly crystalline nanofibrils on Earth, has many intrinsic properties, such as biocompatibility, biodegradability, renewability, and nontoxicity similar to cellulose, resulting in a wide variety application in biomedical science [26 28].

10.1.2 Chitin and chitosan: sources and properties Chitin is available in large quantities in the form of the exoskeletons of arthropods (crabs, lobsters, and shrimps) and the flexible internal backbone of cephalopods (internal shells of squids), worms, webs of spiders, cell walls of fungi and yeasts. It is the least utilized biopolymer as it has bulk structure [29]. It consists of 2-acetamido-2-deoxy-β-D-glucose with β-(1-4) linkage and as discussed, it is often considered to be similar to cellulose. In cellulose, the C2 position is occupied by a hydroxyl group, whereas in chitin it is replaced by an acetamido group ( NHCOCH3) [30 33]. Similar to cellulose, chitin also occurs in nature in a microfibrillated form having both amorphous and crystalline regions and these fibrils are typically embedded in a protein matrix, depending on the origin of the source [34]. Chitin has many useful features, such as hydrophilicity, biocompatibility, antibacterial properties, an affinity for proteins, and easy recyclability, which have made it to be a green solution for renewable, biodegradable polymers in large-scale applications. Chitosan comprises glucosamine and N-acetylglucosamine residues derived from the partial deacetylation of chitin [35,36], as shown in Fig. 10.4, in which about 50% of the acetyl groups will be removed from the chitin by a hydration process or enzyme hydrolysis. The following four steps are the process to produce chitosan from chitin sources: (1) deproteinization, (2) demineralization, (3) decoloration, and (4) deacetylation [32]. The material obtained is a linear polymer α-(1-4)-linked 2-amino-2deoxy-β-D-glucopyranose. Due to N-deacetylation, chitosan is found to be a copolymer of N-acetyl glucosamine and glucosamine [32,37].

Handbook of Chitin and Chitosan

10.1 Introduction

FIGURE 10.4

291

Sources of chitin and preparation of chitosan from chitin.

Compared to all other naturally occurring polysaccharides, like cellulose, pectin, agar, dextran etc., it possesses many important properties, including low toxicity, adsorption capacities, film-forming ability, bacteriostatic action, chelating activities, biocompatibility, and biodegradability [38]. Chitosan is highly basic in nature, whereas the other previously mentioned polysaccharides are either neutral or acidic. Hence chitosan has been widely used in broad applications like drug delivery systems, solid polyelectrolytes, surfactants and membranes in ultrafiltration, reverse osmosis and evaporation, food processing and agricultural packaging, oil water separations, pharmaceuticals, and wastewater treatments [33,39 43]. New approaches and advanced technologies to modernize the chitin and chitosan-based material for the betterment of humans and the environment have been adopted. Aerogel has been prepared from chitin and chitosan-based biopolymers and employed in advanced applications such as wound dressing, drug delivery, bone repair, biosensors, food packaging, water treatment, etc., Recently Suenaga et al. specifically functionalized and fabricated β-chitin nanofiber aerogels by lyophilization that showed higher strength and ideal pore structure [28]. Highly porous and mechanically strong chitin aerogels were reported by Ding et al. [44]; COF (covalent organic framework)-based poroplastic chitosan aerogels was built and used in highly efficient continuous flow-through microreactors for chlorobenzene (CB) dechlorination in water at room temperature [45]. Photoacoustic studies on solid monoliths of Au(III) chitosan silica aerogels were carried out by Kuthirummal et al. for UV-induced reduction of Au(III) to Au(0) [46].

Handbook of Chitin and Chitosan

292

10. Chitin and chitosan-based aerogels

Chau et al. have prepared barium peroxotitanate/chitin aerogel composites which are novel semiconducting photocatalytics used for gassensing for optoelectronic device fabrications [47]. These are some of the chitin and chitosan-based papers reported with specific application. The following section gives detailed information about chitin and chitosanbased aerogels preparation and their applications. This review paper aims at a comprehensive and timely overview of the recent advances in the development of aerogels with a particular emphasis on the synthesis, processing, and applications of almost all aerogels and their composites from different environmental remediation aspects.

10.2 Chitin and chitosan-based aerogels: preparation process Aerogels from polysaccharides such as chitin and chitosan have been the subject of much attention due to their renewable starting materials. Chitin and chitosan-based aerogels have wide applications based on modifications in their functional group and surface morphology. In general, aerogel preparation follows inexpensive and flexible production of the desired morphology by varying the processing step. So the aerogels obtained have different applications, for example, from drug delivery to water treatment, thermal insulators, etc. Common process involved in almost all type of aerogels preparation as shown in Fig. 10.5. Preparation procedure of aerogels involves three important steps (1) sol gel process, (2) aging, and (3) drying (i.e., supercritical drying, freeze-drying, and ambient temperature drying) [3,48]. Among the mentioned steps, drying of the gels is the key step which determines many properties of the aerogels. Because drying of formed gels without damaging the original porous structure is an important step during aerogel preparation process, several conventional drying methods are used to obtain wet gels such as supercritical drying (using i.e., alcohol, acetone, or CO2), ambient pressure drying, and freezedrying, as shown in Table 10.1 and Fig. 10.5. Specific features of these processes with respect to drying steps are summarized in Table 10.1, as reported by Irina Smirnova and Pavel Gurikov [1]. Supercritical drying is the most appropriate and efficient way to dry the wet gels by removing the pores’ liquid under supercritical conditions. From a practical point of view, during supercritical drying, the wet gel is placed inside a closed pressure vessel, so that the pressure and temperature of vessel passes the critical point (TC, PC) of the solvent entrapped in the pores of wet gels. The drying of these gel-based hydrocolloids with supercritical CO2 has been found to prevent the commonly observed collapse of the pores normally attributed to

Handbook of Chitin and Chitosan

10.2 Chitin and chitosan-based aerogels: preparation process

293

FIGURE 10.5 Common preparation scheme of aerogels. Source: Reprinted with permission from C.A. Garcı´a-Gonza´lez, L. Dura˜es, P. del Gaudio, H. Maleki, M. Mahmoudi, A. Portugal, Synthesis and biomedical applications of aerogels: possibilities and challenges, Adv. Colloid Interf. Sci. 236 (2016) 1 27. doi:10.1016/j.cis.2016.05.011 [48]. Elsevier B.V., Copyright r 2016.

the capillary stress exerted on the network upon solvent removal [49]. Highly porous and nanostructured polysaccharides-based aerogels were synthesized via dissolution-solvent exchange-drying with supercritical CO2. The resulting aerogels were mesoporous, with the highest density (0.12 0.15 g/cm3) and highest specific surface area (around 600 m2/g). The results obtained provide the guidelines for making aerogel matrices with fully controlled morphology and properties [50] (Fig. 10.6). Ambient pressure drying is a promising, simple, and safe technique to dry the gels under the ambient temperate conditions and maybe a suitable way for mass industrial production. However, in this method,

Handbook of Chitin and Chitosan

294

Drying technologies for aerogel production.

TABLE 10.1

Preparation steps prior to drying

Main energy costs/risks

Ambient drying

Ambient pressure, room or slightly elevated temperature

Chemical hydrophobization of the matrix is often needed

Low energy costs; well established process; extensive use of hydrophobization agents

Freeze-drying

Vacuum (P , 100 mbar); 270 , T , 2 20 C

Addition of modifier (e.g., tertbutanol) to avoid structure compaction

High energy costs to maintain low temperature; batch process

Direct supercritical drying (high temperature)

Over critical point of organic solvent used for gel formation T . 100 C P . 30 bar

No solvent exchange needed; direct conversion of the solvent to critical conditions

Moderate energy costs to reach critical conditions (heating); batch process, High explosion and toxicity hazards

Supercritical drying by CO2

T . 31 C P . 74 bar Typically 40 C, 100 150 bar

Solvent should be reasonably mixable with CO2 at process conditions; For hydrogels solvent exchange needed

Significant energy costs due to CO2 Compression (may be improved by process optimization); so far bath process; lower explosion risk due to low temperature and CO2 nature

Supercritical drying

Reprinted with permission from I. Smirnova, P. Gurikov, Aerogel production: current status, research directions, and future opportunities, J. Supercrit. Fluid. 134 (2018) 228 233. Elsevier B.V., Copyright r 2018.

10. Chitin and chitosan-based aerogels

Handbook of Chitin and Chitosan

Conditions

10.2 Chitin and chitosan-based aerogels: preparation process

295

FIGURE 10.6 Schematic presentation of two main methods of structure formation, from solution to aerogel. Source: Reprinted with permission from S. Groult, T. Budtova, Tuning structure and properties of pectin aerogels, Eur. Polym. J. 108 (2018) 250 261. doi:10.1016/j.eurpolymj.2018.08.048. Elsevier B.V., Copyright r 2016.

the surface of the pore walls inside the gel must be chemically treated with some nonpolar groups in order to disrupt further condensations in neighboring surface functionality after being compressed by capillary stresses. A chitosan silica hybrid aerogel was produced whose primary precursors (chitosan and silica) were extracted from agricultural waste using the sol gel method at ambient pressure drying technique [51]. Freeze-drying is a simple, economical, and environment-friendly drying approach which is employed to prepare aerogels with reasonably porous structures. It seems to be an alternative, if hydrogels can be directly dried without the severe loss of the pore volume and surface area, as is, for instance, the case for high-density gels. Moreover, freezedrying is usually a long process (tens of hours), leaving room for process intensification [1]. Three-dimensional (3D) network interpenetrated KGM-SiO2 aerogel with thermal insulation performance and high mechanical properties prepared by a facile freeze-drying process, as shown in Fig. 10.7 [52]. The low temperature of freeze-drying favors the development of aerogels used for medical studies as high temperature may damage the medicine [5,48,53]. It is obvious that each drying process has its own advantages and disadvantages. However, supercritical drying by CO2 and freeze-drying were employed in the most general preparation of aerogels, due to being safe and environment-friendly ways of drying, suitable for virtually all kinds of gels. Still, a lot of efforts are needed to reduce the drying time and the energy costs associated with this process.

Handbook of Chitin and Chitosan

296

10. Chitin and chitosan-based aerogels

FIGURE 10.7 Influence of KGM concentration on KGM aerogel morphology and physical properties at Na2CO3-to-KGM mass ratio of 0.12:1 (A) morphological changes in the preparation of KGM aerogels; (B) image of KGM aerogel on the top of a leaf. Source: Reprinted with permission from J. Zhu, J. Hu, C. Jiang, S. Liu, Y. Li, Ultralight, hydrophobic, monolithic konjac glucomannan-silica composite aerogel with thermal insulation and mechanical properties, Carbohydr. Polym. 207 (2019) 246 255. doi:10.1016/j.carbpol.2018.11.073 [52]. Elsevier B.V., Copyright r 2018.

10.3 Characterization of chitin and chitosan-based aerogels 10.3.1 Morphological analysis/microscopic analysis Understanding the texture of material requires the synergy of several techniques. Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) are techniques that are able to provide quantitative information on the shape and the size of the secondary structures and porous nature of polysaccharide aerogels. This information is useful in choosing the proper model for the interpretation of data [54]. 10.3.1.1 Scanning electron microscope SEM is a tool used for seeing the invisible worlds at the micro- and nanoscale. Since the wavelength of electrons is much smaller than the wavelength of light, the resolution of SEM is superior to that of a light microscope. However, the aerogels melt under the electron beam of SEM within a short period of time, causing their mesoporous structures to disappear gradually. Considering that the aerogels were degassed at 100 C for 10 h, the high specific surface area indicated that the mesoporous structures were stable at 100 C. The key factor to obtaining stable aerogels with a high specific surface area is the full replacement of the solvent with absolute ethanol. If the solvent is not fully replaced by absolute ethanol, only a broken wet-gel will be produced. In this context, Chang et al. prepared cross-linked chitosan-based aerogels by freeze-drying and atmospheric drying, but only dry gels without porous structures were achieved and it is well-demonstrated in the SEM images in Fig. 10.8. It explains the mesoporous structures formation of these

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

297

FIGURE 10.8 (A) Photographs of the wet gel (A1) and aerogel (A2) and SEM image (A3) of chitosan glutaraldehyde, (B) chitosan glyoxal, and (C) chitosan formaldehyde. Source: Reprinted with permission from X. Chang, D. Chen, X. Jiap, Chitosan-based aerogels with high adsorption performance, J. Phys. Chem. B. 112 (2008) 7721 7725. doi:10.1021/jp8011359. American Chemical Society, Copyright r 2016.

prepared (A3) chitosan glutaraldehyde, (B) chitosan glyoxal, and (C) chitosan formaldehyde materials, and their pore sizes [55]. Kadib et al. investigated the morphology of the prepared inorganic oxide microspheres incorporated polymeric network using SEM. The images indicate that the size of the void between the fibers is in a range larger than 50 nm, which is characteristic of materials featuring a dual meso- and macroporous network [56] (Fig. 10.9). Ma et al. prepared chitosan-based aerogels. Fig. 10.10 shows the photographs and SEM micrographs of the silica aerogel, chitosan aerogel, and chitosan silica composite aerogel (20-CS). The silica aerogel exhibited a porous structure after the hydrophobic treatment. Chitosan aerogel with glutaraldehyde as a cross-linker had a micron-sized porous structure. In addition, chitosan silica composite aerogel was full of a large amount of nanopores and also exhibited a looser microstructure after hydrophobic treatment [57]. Hao et al. fabricated NiCo2S4 nanotube array/carbon aerogel and NiCo2O4 nanoneedle array/carbon aerogel hybrid supercapacitor electrode materials, by assembling a Ni-Co precursor needle array on the surface of channel walls of hierarchical porous carbon aerogels derived from chitosan [58] (Fig. 10.11). Structure morphology of the hybrid nanostructures was characterized by SEM (Fig. 10.12). From the observation, the morphology of NCOC is similar to that of the Ni-Co precursor/CA hybrid nanostructure; there is

Handbook of Chitin and Chitosan

298

10. Chitin and chitosan-based aerogels

FIGURE 10.9 SEM analysis of (A) M1, (B) M2, (C) M3, (D) M1 (high magnification), and (E) native chitosan (M0). Photographic images of titanium dioxide microspheres (material M2). Source: Reprinted with permission from A. El Kadib, K. Molvinger, T. Cacciaguerra, M. Bousmina, D. Brunel, Chitosan templated synthesis of porous metal oxide microspheres with filamentary nanostructures, Microporous Mesoporous Mater. 142 (2011) 301 307. doi:10.1016/j.micromeso.2010.12.012. Elsevier B.V., Copyright r 2010.

FIGURE 10.10 Photographs of the aerogels: (A0) silica aerogel (0-CS), (B0) chitosan aerogel (100-CS) and (C0) chitosan silica composite aerogel (20-CS) and SEM micrographs of these aerogels: (A1) unmodified silica aerogel, (A2) HMDS-modified silica aerogel, (B1, B2) chitosan aerogel with glutaraldehyde as the cross-linker, (C1) unmodified composite aerogel, and (C2) HMDS-modified composite aerogel. Source: Reprinted with permission from Q. Ma, Y. Liu, Z. Dong, J. Wang, X. Hou, Hydrophobic and nanoporous chitosan-silica composite aerogels for oil absorption, J. Appl. Polym. Sci. 132 (2015) 1 11. doi:10.1002/app.41770 [57]. Wiley Periodical, Inc., Copyright r 2014. Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

299

FIGURE 10.11

Schematic illustration of the formation process of hybrid nanostructures of NCOC and NCSC. Source: Reprinted with permission from P. Hao, J. Tian, Y. Sang, C.C. Tuan, G. Cui, X. Shi, et al., 1D Ni-Co oxide and sulfide nanoarray/carbon aerogel hybrid nanostructures for asymmetric supercapacitors with high energy density and excellent cycling stability, Nanoscale. 8 (2016) 16292 16301. doi:10.1039/c6nr05385h. Royal Society of Chemistry, Copyright r 2016.

FIGURE 10.12 SEM images of (A C) NCOC and (D F) NCSC hybrid nanostructures. Source: Reprinted with permission from P. Hao, J. Tian, Y. Sang, C.C. Tuan, G. Cui, X. Shi, et al., 1D Ni-Co oxide and sulfide nanoarray/carbon aerogel hybrid nanostructures for asymmetric supercapacitors with high energy density and excellent cycling stability, Nanoscale. 8 (2016) 16292 16301. doi:10.1039/c6nr05385h. Royal Society of Chemistry, Copyright r 2016.

no abscission between the 1D Ni-Co oxide and CA in the oxidation process (Fig. 10.11A). And as shown in Fig. 10.11B and C, NiCo2O4 nanoneedles with a diameter of about 60 70 nm and a length of 3 4 μm

Handbook of Chitin and Chitosan

300

10. Chitin and chitosan-based aerogels

FIGURE 10.13 SEM images of (A) CS-GA, (D) β-CD-CS@HMTA, (G) β-CDCS@HMTA-Cr; EDX pattern of (B, C) CS-GA, (E, F) β-CD-CS@HMTA, (H, J) β-CDCS@HMTA-Cr (insets show the photos of CS-GA, β-CD-CS@HMTA, β-CD-CS@HMTA-Cr.) Source: Reprinted with permission from X.L. Wang, D.M. Guo, Q. Da An, Z.Y. Xiao, S.R. Zhai, High-efficacy adsorption of Cr(VI) and anionic dyes onto β-cyclodextrin/chitosan/hexamethylenetetramine aerogel beads with task-specific, integrated components, Int. J. Biol. Macromol. 128 (2019) 268 278. doi:10.1016/j.ijbiomac.2019.01.139. Elsevier B.V., Copyright r 2019.

covered the whole surface of the channel of CA uniformly. Fig. 10.11D F shows the morphology of the sample of NCSC; it can be seen that the as-prepared sample is different from the precursors in morphology. Wang et al. have prepared β-cyclodextrin/chitosan/hexamethylenetetramine aerogel beads and SEM images of CS-GA, β-CD-CS@HMTA and β-CD-CS@HMTA-Cr are shown in Fig. 10.13A, D, and G. A three-dimensional network structure is observed, revealing that aerogel beads are composited with mesopores and the pore sizes are heterogeneous, which probably could be attributed to the nonuniform reaction. Compared with neutral CS, after modifying CS with β-cyclodextrin and hexamethylenetetramine, it is realized that the CS fiber is thicker and tighter. The element type and content of aerogel beads were shown in EDX mapping (Fig. 10.13C, F, and J) in which the cross profile of β-CD-

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

301

CS@HMTA is covered by the Cr element. This suggested that the Cr(VI) is adsorbed on the aerogel beads successfully. Fig. 10.13E and H expressed the EDX spectra of β-CD-CS@HMTA before and after adsorption which further proved that the Cr(VI) ions had been adsorbed on modified CS beads [59]. A new kind of poly(ethylenimine)/chitosan (PEI/CS) aerogel beads was successfully synthesized using controllable sol gel and freezedrying methods. Optical images of the CS GA, CS GA PEI, and CS/ PEI GA beads are shown in Fig. 10.14A, C, and E. The microscopic morphologies of these three adsorbents were characterized by using SEM and the results are shown in Fig. 10.14B, D, and F. Among these, CS GA PEI exhibited an ideal microscopic three-dimensional network

FIGURE 10.14 Optical microscopy images of (A) CS GA, (C) CS GA PEI, (E) CS/ PEI GA beads, and the inset images are the equivalent hydrogel beads before freezedrying. (B), (D), and (F) are the FE-SEM images of three different beads, respectively. Source: Reprinted with permission from R. Li, Q. Da An, Z.Y. Xiao, B. Zhai, S.R. Zhai, Z. Shi, Preparation of PEI/CS aerogel beads with a high density of reactive sites for efficient Cr(VI) sorption: Batch and column studies, RSC Adv. 7 (2017) 40227 40236. doi:10.1039/c7ra06914f. Royal Society of Chemistry, Copyright r 2017.

Handbook of Chitin and Chitosan

302

10. Chitin and chitosan-based aerogels

structure compared to CS GA and CS/PEI GA. Clearly, this obvious network structure might be helpful for increasing the adsorption capacity for heavy metal ions from aqueous solutions [60]. 10.3.1.2 Transmission electron microscopy TEM is the easiest and most widely used technique for structural and compositional characterization of aerogel in bright-field imaging [61] and it is a very powerful tool for material science. Basic information about the typical mesoparticle size, confirmation of the porous nature, the distribution of incorporated particles and morphological homogeneity of aerogel images were obtained, and have been used to develop new aerogel-based materials. The full range of advanced TEM analysis, not just bright-field imaging, is possible, despite the low thermal and electrical conductivity of most aerogels [61,62]. Singh et al. have prepared highly soluble chitosan-L-glutamic acid (CL-GA) aerogel derivative. The shape of the CL-GA derivative was measured in the solid-state using TEM (Fig. 10.15). The chitosan derivative formed regularly packed aggregates of spherical particles. The dark lines represent the cross-section of the chitosan derivative layer and the gray area to the matrix. The TEM study is simply indicative of the helical conformation of the chitosan acid derivative [63]. Yu et al. has fabricated graphene oxide (GO) chitosan (CS) aerogel by the lyophilization technique. The structure of GO CS aerogel surface was characterized under TEM. From Fig. 10.16, a series of folding and stacking that has occurred in GO CS can be seen. In contrast, bare GO sheets only showed small wrinkles and were generally flat, as shown in Fig. 10.16C [64].

FIGURE 10.15 TEM image of CL-GA derivative. Source: Reprinted with permission from J. Singh, P.K. Dutta, J. Dutta, A.J. Hunt, D.J. Macquarrie, J.H. Clark, Preparation and properties of highly soluble chitosan-L-glutamic acid aerogel derivative, Carbohydr. Polym. 76 (2009) 188 195. doi:10.1016/j.carbpol.2008.10.011 [63]. Elsevier B.V., Copyright r 2018.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

303

FIGURE 10.16 TEM images of GO CS (A) and (B) and free GO (C). Source: Reprinted with permission from B. Yu, J. Xu, J.H. Liu, S.T. Yang, J. Luo, Q. Zhou, et al., Adsorption behavior of copper ions on graphene oxide-chitosan aerogel, J. Environ. Chem. Eng. (2013). doi:10.1016/j. jece.2013.08.017. Elsevier B.V., Copyright r 2013.

FIGURE 10.17 TEM of (A) M1, (B) M2, (C) M3, and (D) M1 (high magnification) Source: Reprinted with permission from A. El Kadib, K. Molvinger, T. Cacciaguerra, M. Bousmina, D. Brunel, Chitosan templated synthesis of porous metal oxide microspheres with filamentary nanostructures, Microporous Mesoporous Mater. 142 (2011) 301 307. doi:10.1016/j. micromeso.2010.12.012. Elsevier Inc., Copyright r 2010.

Inorganic oxide (ZrO2, Al2O3 and SnO2)-coated chitosan microspheres were synthesized by Kadib et al. and networks of the materials show several branched filaments (Fig. 10.17). These filaments consist of small particles. The initially formed inorganic oligomers seem to selfassemble or entangle in the fibrous network of the polysaccharides. This

Handbook of Chitin and Chitosan

304

10. Chitin and chitosan-based aerogels

FIGURE 10.18 TEM images of (A) ESXS aerogels, (B) ESXS aerogels with particle size 5 20 nm AuNPs formed inside, (C) ESXS aerogels with particle size 2 45 nm AgNPs formed inside, (D) size distribution histograms of AuNPs, and (E) AgNPs. Source: Reprinted with permission from S. Zhao, H. Xu, L. Wang, P. Zhu, W.M. Risen, J. William Suggs, Synthesis of novel chitaline-silica aerogels with spontaneous Au and Ag nanoparticles formation in aerogels matrix, Microporous Mesoporous Mater. 171 (2013) 147 155. doi:10.1016/j.micromeso.2012.12.038. Elsevier B.V., Copyright r 2013.

resulting structure is a consequence of the petrification of the chitosan fibers by inorganic oxide particles during the growth of the precursors inside the polymeric scaffold. At higher magnification, the inorganic network seems to resemble the continuous filaments rather than the aggregation of small nanoparticles [56]. Zhao et al. prepared Au(0) and Ag(0) nanoparticles doped in the chitaline silica aerogels. TEM was undertaken and the representative images are shown in Fig. 10.18. The average sizes and size distribution of these particles were determined by analysis of several different images, shown in Fig. 10.3D and E. It turned out that the AuNPs and AgNPs have a relatively wide size distribution, AuNPs range from 5 to 20 nm, while the AgNPs range from 2 to 45 nm [65].

10.3.2 Porosity Specific surface area (SBET) and pore size distribution are the main parameters characterizing aerogel texture. As for classical aerogels, specific surface area of bioaerogels is determined using nitrogen adsorption technique and Brunauer Emmett Teller (BET) theory. The standard

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

305

FIGURE 10.19 Nitrogen adsorption desorption isotherms and pore size distribution curves (inset) of CSGA, CS-GA-PEI and CS/PEI-GA. Source: Reprinted with permission from R. Li, Q. Da An, Z.Y. Xiao, B. Zhai, S.R. Zhai, Z. Shi, Preparation of PEI/CS aerogel beads with a high density of reactive sites for efficient Cr(VI) sorption: Batch and column studies, RSC Adv. 7 (2017) 40227 40236. doi:10.1039/c7ra06914f. Royal Society of Chemistry, Copyright r 2017.

method for measuring pore volume and size distribution uses the Barrett Joyner Halenda (BJH) approach [66]. Bioaerogels possess macro- and mesopores and are often have large macropores (several hundreds of nanometers up to several microns). The nature of the functional groups of the polysaccharide significantly influences the adsorption of N2 on the surface of the aerogel. The net enthalpy of adsorption increases with the polarity of the surface groups of the biopolymers, in the order chitin , agar # chitosan , carrageenan , alginic acid, alginate. The surface area and the mesopore distribution of the aerogels depend both on the dispersion of the parent hydrogel as well as the behavior of each polymer in the drying method. The surface area of the aerogel can bring reliable information about the size of the secondary units which form the polymer network, providing further details about the shape of the aggregates and the density of the polymer, which can be reasonably assessed by independent techniques. Good retention of the structure of the hydrogel in the supercritically dried aerogel will be discussed below [54]. Fig. 10.19 shows N2 adsorption desorption isotherms of CS GA, CS GA PEI, and CS/PEI GA beads. These three types of adsorption materials show type IV curves with hysteresis loops, clearly proving that these samples are porous materials. The surface areas of CS GA,

Handbook of Chitin and Chitosan

306

10. Chitin and chitosan-based aerogels

CS GA PEI, and CS/PEI GA were 59.4, 13.7, and 23.0 m2/g, respectively. The average pore diameters of these three different adsorbents were approximately 19.5, 22.7, and 3.8 nm, respectively. Taking these together, the results of CS GA PEI show that it can be considered as a potential adsorbent with favorable textural characteristics [60]. Tsioptsias et al. have prepared chitin aerogels successfully via supercritical drying. The aerogels exhibited high porosity, high surface area, and low density compared to chitin. N2 adsorption was measured at 77K and a representative isotherm is illustrated in Fig. 10.20. The aerogel exhibits the typical form of type II or IV isotherms corresponding to physical sorption with mesoporous structured material (pore sizes between 2 and 50 nm) [27]. Tsutsumi et al. have synthesized chitin nanofibrous hydrogel and aerogel by a freeze-drying method. The specific surface area of the prepared aerogel was found to be 289 m2/g (Fig. 10.21) and there was excellent cycling stability toward the Knoevenagel reaction for the exposed C2-amines [29]. Takeshita et al. prepared hexanal modified chitosan aerogel [67]. The nitrogen adsorption desorption isotherm of aerogel was analyzed and it exhibited type IV curves with a hysteresis, indicating a mesoporous structure. The specific surface areas are 672 and 581 m2/g for unmodified and hexanal-modified samples, respectively. From BJH plots, the hexanal modified sample has a sharper and slightly larger pore distribution than that of the unmodified sample (Fig. 10.22).

FIGURE 10.20 Representative N2 adsorption isotherm at 77K of chitin aerogels. Source: Reprinted with permission from C. Tsioptsias, C. Michailof, G. Stauropoulos, C. Panayiotou, Chitin and carbon aerogels from chitin alcogels, Carbohydr. Polym. 76 (2009) 535 540. doi:10.1016/j.carbpol.2008.11.018. Elsevier Ltd., Copyright r 2008.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

307

FIGURE 10.21 Chitin nanofibrous hydrogel (A) and aerogel (B), SEM of aerogel (C) and the base catalysis Knoevenagel reaction. (D) Nitrogen adsorption isotherm of chitin aerogel. Source: Reprinted with permission from Y. Tsutsumi, H. Koga, Z.D. Qi, T. Saito, A. Isogai, Nanofibrillar chitin aerogels as renewable base catalysts, Biomacromolecules. 15 (2014) 4314 4319. doi:10.1021/bm501320b. American Chemical Society, Copyright r 2014.

FIGURE 10.22 Nitrogen adsorption isotherms at 77K of (left) unmodified and (right) hexanal-modified (13.3 vol.%) chitosan aerogels. Source: Reprinted with permission from American S. Takeshita, A. Konishi, Y. Takebayashi, S. Yoda, K. Otake, Aldehyde approach to hydrophobic modification of chitosan aerogels, Biomacromolecules. 18 (2017) 2172 2178. doi:10.1021/acs.biomac.7b00562. Chemical Society, Copyright r 2017.

Chitosan clay hybrid materials have been widely applied in different fields of application. Ennajih et al. reported the development of a MMT chitosan hybrid and analyzed the pore nature of materials using adsorption desorption isotherms. From Fig. 10.23, the continuous network of pores of sizes ranging from the mesoporous to macroporous domains can be seen [49]. To examine the porosity of prepared chitosan SH, COF-IL@chitosan, and COF-IL, N2 adsorption desorption were performed at 77K [68]. As

Handbook of Chitin and Chitosan

308

10. Chitin and chitosan-based aerogels

FIGURE 10.23 Nitrogen sorption isotherms and SEM micrographs of MMT chitosan hybrid microspheres (scale bar 1 lm). (A) M:30 70. (B) M:50 50. (C) M:70 30. Source: Reprinted with permission from H. Ennajih, R. Bouhfid, E.M. Essassi, M. Bousmina, A. El Kadib, Chitosan-montmorillonite bio-based aerogel hybrid microspheres, Microporous Mesoporous Mater. 152 (2012) 208 213. doi:10.1016/j.micromeso.2011.11.032. Elsevier Inc., Copyright r 2011.

FIGURE 10.24 Brunauer Emmett Teller (BET) isotherm for chitosan SH, COFIL@chitosan, and COF-IL. Source: Reprinted with permission from L.G. Ding, B.J. Yao, F. Li, S. C. Shi, N. Huang, H.B. Yin, et al., Ionic liquid-decorated COF and its covalent composite aerogel for selective CO2 adsorption and catalytic conversion, J. Mater. Chem. A. 7 (2019) 4689 4698. doi:10.1039/c8ta12046c. Royal Society of Chemistry, Copyright r 2019.

shown in Fig. 10.24, the N2 adsorption desorption curve of COF-IL showed a type I isotherm (microporous structure). The N2 uptake amount is 197 cm3/g, and its BET specific surface area was 291 m2/g.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

309

As the ice-templating generated macropores did not significantly contribute to the surface area, so the COFIL@ chitosan showed a decreased N2 adsorption capacity (162.6 cm3/g) and a lower BET specific surface area (103.3 m2/g) in comparison with the COF-IL. On the other hand, the involved COF crystals significantly enhanced the gas capacity and BET specific surface area of the pure chitosan SH aerogel (3.05 cm3/g and 4.67 m2/g) [69]. Nitrogen adsorption desorption isotherms and pore size distribution curves of CS-O, CS-F, and CS-PDA were analyzed by Guo et al. BET areas of CS-F and CS-PDA were found to be 66.2 m2/g and 77.3 m2/g, respectively, which were much higher than that of CS-O (4.3 m2/g), suggesting that the freeze-drying method could effectively enlarge a specific surface area of chitosan beads. CS-PDA aerogels possessed a pore size of 24.9 nm, larger than the pore size of CS-F. Additionally, the adsorption desorption isotherms of these adsorbents were type IV, revealing the existence of a mesoporous structure which contributed to the mass transfer of metal [70]. Suenaga et al. have prepared β-ChNF aerogels and analyzed nitrogen adsorption desorption isotherms. The relative surface area of the β-ChNF [acid] aerogel was two times greater than that of the β-ChNF [neutral] aerogel (Fig. 10.25). The relative surface area of the aerogel prepared by the quick-freeze method was three times greater than that of the respective former aerogels [28].

10.3.3 Thermal properties 10.3.3.1 Thermogravimetric analysis Thermogravimetric analysis (TGA) was used to determine the thermal stability of the aerogel blankets in an air atmosphere. Chitosanbased aerogel membrane was prepared and TGA was analyzed [71]. The results (Fig. 10.26) of the blend (Agr-CS) as well as genipin crosslinked blend (Agr-CS-G) prepared in the presence of cross-linking agent showed the high thermal stability in comparison to the pristine constituents. Minimum residual mass of 20.53%, 25.05%, and B31.21% was obtained for Agr, CS, and CS-Agr blends, respectively. However, genipin cross-linked blend (Agr-CS-G) retained as high as 41.08% residual mass at 599.5 C. Therefore, it directly implies that the rigid network is a result of genipin cross-linking in aerogel membranes. The TGA results show that the CTS GO aerogels are more thermally stable than the neat chitosan aerogel [72]. TGA provides information on the proximate analysis (moisture, volatile matter, fixed carbon, and ash contents) of aerogels by Takeshita et al. Graphene oxide/chitosan aerogel (GOCA) was prepared by a

Handbook of Chitin and Chitosan

310

10. Chitin and chitosan-based aerogels

FIGURE 10.25 (A) Nitrogen adsorption and desorption isotherms and (B) relative surface area of the aerogels. Source: Reprinted with permission from S. Suenaga, M. Osada, Preparation of β-chitin nanofiber aerogels by lyophilization, Int. J. Biol. Macromol. 126 (2019) 1145 1149. doi:10.1016/j.ijbiomac.2019.01.006. Elsevier B.V., Copyright r 2019.

facile ice-templating technique. This study revealed that the incorporation of GO in chitosan improved the thermal stability of chitosan and the results are shown in Table 10.2 [73]. 10.3.3.2 Differential scanning calorimetry DSC can be used to study cross-linking as well as to determine substrate uniformity. Pressure DSC can determine the oxidative stability of a material and analyze pressure-sensitive reactions. Singh et al. have prepared CL-GA in chitosan and analyzed the DSC thermogram. The results indicate that the structure of chitosan chains has been changed due to the introduction of glutamic acid and the reduced ability of crystallization. The lack of the exothermic decomposition in CL-GA (present

Handbook of Chitin and Chitosan

311

10.3 Characterization of chitin and chitosan-based aerogels

FIGURE 10.26

Thermal behavior of neat chitosan at room temperature and the studied aerogels after thermal treatment up to 100 C. Source: Reprinted with permission from A.A. Alhwaige, T. Agag, H. Ishida, S. Qutubuddin, Biobased chitosan hybrid aerogels with superior adsorption: role of graphene oxide in CO2 capture, RSC Adv. 3 (2013) 16011 16020. doi:10.1039/ c3ra42022a. Elsevier Inc., Copyright r 2013.

TABLE 10.2

Thermal stability of GOCA.

Thermodynamics

ΔG (kJ) 





ΔH (kJ/mol)

ΔS (kJ/mol K)



Co (mg/L)

30 C

40 C

50 C

60 C

20

226.98

227.08

227.66

227.93

216.50

0.0343

40

227.73

228.17

228.12

227.30

232.11

20.0134

60

227.74

227.72

227.00

225.08

254.57

20.0870

80

226.18

225.79

226.17

224.33

242.09

20.0518

100

225.33

224.64

224.81

223.61

240.45

20.0498

200

221.44

220.87

221.84

220.79

224.33

20.0098

300

219.76

219.80

220.17

219.43

221.71

20.0060

400

218.69

218.68

219.47

217.91

223.61

20.0155

500

218.20

218.68

218.24

217.53

225.95

20.0245

600

217.45

217.97

217.34

217.25

221.41

20.0123

Reprinted with permission from K.C. Lai, B.Y.Z. Hiew, L.Y. Lee, S. Gan, S. Thangalazhy-Gopakumar, W.S. Chiu, et al., Ice-templated graphene oxide/chitosan aerogel as an effective adsorbent for sequestration of metanil yellow dye, Bioresour. Technol. 274 (2019) 134 144. doi:10.1016/j.biortech.2018.11.048. Elsevier Ltd., Copyright r 2018.

Handbook of Chitin and Chitosan

312

10. Chitin and chitosan-based aerogels

at 376.5 C in chitosan) provides good evidence toward the successful modification of this material [63].

10.3.4 Raman spectra Raman spectroscopy is an effective tool monitoring the evolution of the aerogel systems. It is used to determine physisorbed as well as chemisorbed water in aerogels (e.g., in aerogel window systems) as well as to detect the disorder, space expansion, and dislocation of materials. Spatially resolved Raman spectroscopy can be used to investigate the mechanism of water diffusion in the aerogel network. Alhwaige et al. investigated the effect of calcination on the stability of GO in the hybrid aerogels by Raman spectroscopy. The Raman spectrum of CTS-GO-10% shows amorphous structure of CTS-GO hybrids. There is no signature for graphene, as observed in Fig. 10.27. The ratio of D and G bands reveals that GO remains in the matrix after treatment up to 400 C [72]. Zhai et al. studied GO/CS aerogel and rGO/CS aerogel structural changes through Raman spectra. Fig. 10.28 clearly shows that in the presence of chitosan, the ID/IG ratio trend does not show a clear increment despite no characteristic peak appearing in the Raman spectra of pristine CS and CS with 1 h annealing at 200 C [74]. Similarly, Frindy et al. studied raman spectrum for CS-GO at different ratios. They confirmed the intactness of graphene oxide within the

FIGURE 10.27 Raman spectra of (A) graphite, (B) GO at 60 C, and (C) CTS-GO-10% at 400 C. Source: Reprinted with permission from A.A. Alhwaige, T. Agag, H. Ishida, S. Qutubuddin, Biobased chitosan hybrid aerogels with superior adsorption: Role of graphene oxide in CO2 capture, RSC Adv. 3 (2013) 16011 16020. doi:10.1039/c3ra42022a. Elsevier Inc., Copyright r 2013.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

313

FIGURE 10.28

Raman spectra of (A) GO before thermal annealing, rGO powder annealed at 200 C for 12 min and 1 h, (B) GO/CS aerogel before thermal annealing, rGO/CS aerogel annealed at 200 C for 12 min and 1 h, (C) CS powder before thermal annealing and annealed at 200 C for 1 h. Source: Reprinted with permission from T. Zhai, L. Verdolotti, S. Kaciulis, P. Cerruti, G. Gentile, H. Xia, et al., High piezo-resistive performances of an anisotropic composite realized by embedding rgo-based chitosan aerogel in open cell polyurethane foams, Nanoscale 11 (2019) 8835 8844. doi:10.1039/c9nr00157c. Royal Society of Chemistry, Copyright r 2019.

FIGURE 10.29 Raman of GO pristine material and the two hybrids before (CS-GO-20%-m) and after (CS-rGO-20%-h) reduction. Source: Reprinted with permission from S. Frindy, A. Primo, H. Ennajih, A. el kacem Qaiss, R. Bouhfid, M. Lahcini, et al., Chitosan graphene oxide films and CO2-dried porous aerogel microspheres: interfacial interplay and stability, Carbohydr. Polym. 167 (2017) 297 305. doi:10.1016/j.carbpol.2017.03.034. Elsevier Ltd., Copyright r 2017.

porous network of CS-GO-20%-m, as shown in Fig. 10.29. Upon hydrazine treatment, a substantial increase in the ID/IG intensity is observed. It is evidence for the successful reduction of graphene oxide within the porous network of the polymer [75].

Handbook of Chitin and Chitosan

314

10. Chitin and chitosan-based aerogels

10.3.5 Tensile property of aerogels/mechanical property Mechanical properties are useful measurements for industrial-based applications such as packaging and construction but are also important for medical applications, such as tissue culture. For the last several years, mechanical properties of the polysaccharides-based aerogels have been improved by different methods. For instance, there have been many reinforcing agents incorporated with polysaccharide aerogels to improve the mechanical properties. To determine the mechanical property different loading studies are carried out, which include compression, bending, torsion, tension, and multiaxial stress states under quasistatic, dynamic, and fatigue loading conditions. The mechanical behavior of chitosan aerogels have been studied by Takeshita et al.; the aerogels also showed flexibility and higher mechanical toughness than conventional silica aerogels. The characteristic

FIGURE 10.30 Mechanical properties of chitosan aerogels. (A) Compression stress strain curves. Inset: expanded curves in the small stress region. (B) Changes with density in elastic modulus, stress at 50% strain, and yield stress. The yield stress of C4F7 is not shown because of an ambiguous yield point. (C) Photographs of a 0.6-mm-thick sample of C4F7. Source: Reprinted with permission from S. Takeshita, S. Yoda, Chitosan aerogels: transparent, flexible thermal insulators, Chem. Mater. 27 (2015) 7569 7572. doi:10.1021/acs.chemmater.5b03610. American Chemical Society, Copyright r 2015.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

315

mechanical properties associated with a low thermal conductivity enable us to realize a new category of thermal insulators: environmentfriendly, transparent, and flexible [76] (Fig. 10.30). Superhydrophobic and superoleophilic chitosan sponge (MCTCS) was successfully prepared via a facile freeze-drying method by Su et al. [77]. The cyclic stress strain curves showed that the MCTCS sponges could be compressed to large strains (60%) at relatively low stress (0.023 MPa), owing to the high porosity and elasticity of the sponge. The reproducible cyclic curves in Fig. 10.31B and C demonstrated the good recovery of elasticity in the process of repeated compressions. It indicates that the MCTCS sponges possessed ideal elasticity and compressive durability, which was very important for the practical application of the sponges. The mechanical properties of hemicelluloses citrate chitosan foam and hemicellulose chitosan foam were investigated with dynamic mechanical analysis (DMA). From Fig. 10.32 it is observed that the storage modulus of hemicelluloses citrate chitosan foam was significantly higher than that of hemicellulose chitosan foam and the tan delta lower, reflecting increased cross-linking of the hemicellulose citrate to

FIGURE 10.31 Photographs of reversible compression of the MCTCS sponge (A). Cyclic stress strain curves of the MCTCS sponge subjected to a compressive strain of 20% (B) and 60% (C). Source: Reprinted with permission from C. Su, H. Yang, H. Zhao, Y. Liu, R. Chen, Recyclable and biodegradable superhydrophobic and superoleophilic chitosan sponge for the effective removal of oily pollutants from water, Chem. Eng. J. (2017). doi:10.1016/j.cej.2017.07.157. Elsevier Ltd., Copyright r 2017.

Handbook of Chitin and Chitosan

316

10. Chitin and chitosan-based aerogels

FIGURE 10.32 DMA of (H C) hemicellulose chitosan and (HC C) hemicelluloses citrate chitosan foam. Reaction conditions: 1:1 of HC:C, 3 h, solid to liquid ratio 1:100, pH 3.5, 110 C. Source: Reprinted with permission from A. Salam, R.A. Venditti, J.J. Pawlak, K. El-Tahlawy, Crosslinked hemicellulose citrate-chitosan aerogel foams, Carbohydr. Polym. (2011). doi:10.1016/j.carbpol.2011.01.008. Elsevier Ltd., Copyright r 2011.

FIGURE 10.33 Compression stress strain curves of P/CA along the axial direction (A) and the radial one (B). Photos along the axial direction (C) and the radial one (D). Source: Reprinted with permission from S. Zhang, J. Feng, J. Feng, Y. Jiang, L. Li, Ultra-low shrinkage chitosan aerogels trussed with polyvinyl alcohol, Mater. Des. 156 (2018) 398 406. doi:10.1016/j.matdes.2018.07.004. Elsevier Ltd., Copyright r 2018.

chitosan relative to the hemicellulose to chitosan and a more elastic-like behavior of the material [78]. The mechanical characteristics including toughness and robustness need to be soundly matched to the requisites in a practical application. The compression stress strain curves (Fig. 10.33) of P/CA, along the

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

317

FIGURE 10.34 Photographs of the aerogels obtained after lyophilization Fig. 10.6. Stress strain curves for the aerogels. Source: Reprinted with permission from S. Suenaga, M. Osada, Preparation of β-chitin nanofiber aerogels by lyophilization, Int. J. Biol. Macromol. 126 (2019) 1145 1149. doi:10.1016/j.ijbiomac.2019.01.006. Elsevier B.V., Copyright r 2019.

axial and radial directions (Fig. 10.33C and D), are parallel to those of nonbrittle porous polymer foams. The addition of a linear polymer is more beneficial to render the prepared aerogel with linear elasticity at a low strain range. On further expanding the strain to 70%, a significant stress increase is monitored due to the densification of pore skeletons. Accordingly, the mechanical property of P/CA is fully satisfactory to the routine usage of thermal insulators at unevenly varied ambient temperature [79]. Fig. 10.34 shows the stress strain curves of the aerogels. The compressive modulus and the energy absorption of the β-ChNF [acid] aerogels were 30 and three times greater than those of the β-ChNF [neutral] aerogels, respectively, probably because of the aerogel uniformity. The elastic deformation region was clearly observed, and the compressive modulus was high. The β-ChNF [acid] aerogels are likely suitable for scaffolding in terms of mechanical strength [28]. Other characterization techniques of aerogels, including Fourier transform infrared spectroscopy (FTIR), X-ray diffraction (XRD), Nuclear magnetic resonance spectroscopy (NMR), X-ray photoelectron spectroscopy (XPS), and water contact angle, are discussed below.

10.3.6 Fourier transform infrared spectroscopy FTIR analysis has been used to complement water uptake or contact angle studies. It gives direct structural information about the chemical bonds present in an aerogel sample. The new porous cross-linked aerogel materials were structurally characterized by FTIR. Fig. 10.35 confirms that the aerogels obtained with different cross-linkers have no further free cross-linker molecules [80].

Handbook of Chitin and Chitosan

318

10. Chitin and chitosan-based aerogels

FIGURE 10.35 FTIR spectrum of different cross-linker added aerogels. Source: Reprinted with permission from R. Valentin, B. Bonelli, E. Garrone, F. Di Renzo, F. Quidnard, Accessibility of the functional group of chitosan aerogel probed by FT-IR-monitored deuteration, Biomacromolecules 8 (2007) 3646 3650. doi:10.1021/bm070391a. American Chemical Society Copyright r 2017.

FIGURE 10.36 FTIR spectra of no-GEL and GEL CS aerogels cross-linked with ΦGA 5 (A) 5 and (B) 15 wt.% after thermal treatment. Source: Reprinted with permission from M. Salzano de Luna, C. Ascione, C. Santillo, L. Verdolotti, M. Lavorgna, G.G. Buonocore, et al., Optimization of dye adsorption capacity and mechanical strength of chitosan aerogels through crosslinking strategy and graphene oxide addition, Carbohydr. Polym. 211 (2019) 195 203. doi:10.1016/j.carbpol.2019.02.002. Elsevier Ltd., Copyright r 2019.

Accessibility of the functional group of chitosan aerogel probed by FT-IR-monitored deuteration. The FTIR spectrum of no-GEL and GEL aerogels at the end of the thermal treatment was shown in Fig. 10.36. The possible effects of the different cross-linking strategy on the reaction path was elaborated using FTIR analysis. Further, the results of FTIR confirm the occurrence of the cross-linking reactions irrespective of preparation procedure and amount of GA [81]. Fig. 10.37 shows an FT-IR spectrum of pure chitosan (A), chitosan aerogel (B), and chitosan aerogels modified with Au nanoparticles

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

319

FIGURE 10.37

FT-IR spectrum of (A) pure chitosan; (B) CSAG-85; (C) CSAG-85Au; (D) CSAG90Au; and (E) CSAG95Au. Source: Reprinted with permission from M. Pia˛tkowski, J. Radwan-Pragłowska, Ł. Janus, D. Bogdał, D. Matysek, V. Cablik, Microwave-assisted synthesis and characterization of chitosan aerogels doped with Au-NPs for skin regeneration, Polym. Test. 73 (2019) 366 376. doi:10.1016/j.polymertesting.2018.11.024. Elsevier Ltd., Copyright r 2018.

(C E). The results show chitosan structural modification throughout Au nanoparticles addition [82].

10.3.7 X-ray diffraction To elucidate interactions between the molecules and its arrangement XRD was performed. The XRD patterns of the chitosan, silica aerogel, and composite aerogel with 20 wt.% chitosan are shown in Fig. 10.38. The coexistence of the chitosan and silica diffraction peaks indicate the chitosan silica composite aerogel was prepared successfully [57] The prepared COF-IL showed good crystallinity, which was revealed by the measured powder XRD (PXRD) pattern (Fig. 10.39). The structural modeling was thus conducted using the software of Materials Studio (Version 7.0) for more precise structural information [69].

10.3.8 X-ray photoelectron spectroscopy XPS spectra are usually employed to detect the existence of a certain element and determine the species of the same element, because the electron configuration of the atom would be changed by the change of chemical bonds.

Handbook of Chitin and Chitosan

320

10. Chitin and chitosan-based aerogels

FIGURE 10.38 XRD patterns of the (A) chitosan, (B) silica aerogel (0-CS), and (C) composite aerogel (20-CS). Source: Reprinted with permission from Q. Ma, Y. Liu, Z. Dong, J. Wang, X. Hou, Hydrophobic and nanoporous chitosan-silica composite aerogels for oil absorption, J. Appl. Polym. Sci. 132 (2015) 1 11. doi:10.1002/app.41770. Wiley Periodical, Inc., Copyright r 2014.

FIGURE 10.39 (A) The simulated and measured PXRD patterns of COF-IL. (B) Crystal packing pattern viewed down the crystallographic c and b axes based on the PXRD data obtained by using Materials Studio. Source: Reprinted with permission from L.G. Ding, B.J. Yao, F. Li, S.C. Shi, N. Huang, H.B. Yin, et al., Ionic liquid-decorated COF and its covalent composite aerogel for selective CO2 adsorption and catalytic conversion, J. Mater. Chem. A. 7 (2019) 4689 4698. doi:10.1039/c8ta12046c. Royal Society of Chemistry, Copyright r 2019.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

321

FIGURE 10.40 XPS spectra of P/CA for C 1 s (A), N 1 s (B), and O 1 s (C). (D) FTIR spectra of the control, CTS and P/CA the development of biomass aerogels with controllable low-shrinkage feature from final wet gel to aerogel, used for thermal insulator in energy-saving buildings. Source: Reprinted with permission from S. Zhang, J. Feng, J. Feng, Y. Jiang, L. Li, Ultra-low shrinkage chitosan aerogels trussed with polyvinyl alcohol, Mater. Des. 156 (2018) 398 406. doi:10.1016/j.matdes.2018.07.004. Elsevier Ltd., Copyright r 2018.

FIGURE 10.41 XPS spectra of (A D) NCOC [(A) C1s, (B) Ni2p, (C) Co2p, (D) O1s] and (E H) NCSC [(E) C1s, (F) Ni2p, (G) Co2p, (H) S2p]. Source: Reprinted with permission from P. Hao, J. Tian, Y. Sang, C.C. Tuan, G. Cui, X. Shi, et al., 1D Ni-Co oxide and sulfide nanoarray/carbon aerogel hybrid nanostructures for asymmetric supercapacitors with high energy density and excellent cycling stability, Nanoscale. 8 (2016) 16292 16301. doi:10.1039/ c6nr05385h. Royal Society of Chemistry, Copyright r 2016.

Handbook of Chitin and Chitosan

322

10. Chitin and chitosan-based aerogels

FIGURE 10.42 1H NMR spectra of (A) Na-CMCS and (B) NaO-CMCS. Source: Reprinted with permission from B. Doshi, E. Repo, J.P. Heiskanen, J.A. Sirvio¨, M. Sillanpa¨a¨, Sodium salt of oleoyl carboxymethyl chitosan: A sustainable adsorbent in the oil spill treatment, J. Clean. Prod. 170 (2018) 339 350. doi:10.1016/j.jclepro.2017.09.163. Elsevier Ltd., Copyright r 2017.

A new type of chitosan aerogel with ultralow shrinkage from its final wet gel to aerogel was prepared successfully from CTS and linear PVA polymer by using supramolecular interaction and covalent crosslinking. XPS clearly shows the interface bonding and chemical composition of CTS aerogels [79] (Fig. 10.40). The XPS spectra of NCOC and NCSC are shown in Fig. 10.41. Results conclude that the surfaces of NCOC and NCSC have the chemical composition of substituted spinel-type NiCo2O4 and NiCo2S4, respectively.

10.3.9 Nuclear magnetic resonance spectroscopy Information about the structure of a sample can be deduced through analysis of its NMR spectrum. There are a wide variety of NMR experiments and techniques, including multidimensional NMR spectroscopy, which can be employed to obtain detailed structural information about samples. NMR spectroscopy of solid-state samples, such as aerogels, is more complicated than for nonviscous solutions. The rapid tumbling of molecules in solution and spinning of sample tubes averages out the anisotropic effects. But this tumbling does not occur in solid samples, so the observed shifts contain an anisotropic component. Grinding the sample to small, uniform particle size and spinning it at the magic angle reduces these effects somewhat. Amphiphilic sodium salt of oleoyl carboxymethyl chitosan (NaOCMCS) was synthesized and characterized by NMR [33]. The 1H NMR spectra of Na-CMCS and NaO-CMCS are presented in Fig. 10.42A and B, respectively. The successful attachment of oleoyl substituent to NaCMCS was confirmed.

Handbook of Chitin and Chitosan

10.3 Characterization of chitin and chitosan-based aerogels

323

FIGURE 10.43 1H NMR spectra of hexanal reagent, unmodified and hexanal-modified (13.3 vol.%) chitosan aerogels. Source: Reprinted with permission from S. Takeshita, A. Konishi, Y. Takebayashi, S. Yoda, K. Otake, Aldehyde approach to hydrophobic modification of chitosan aerogels, Biomacromolecules 18 (2017) 2172 2178. doi:10.1021/acs.biomac.7b00562. American Chemical Society, Copyright r 2017.

Hydrophobic biobased nanofibrous aerogels using the combination of chitosan and alkyl aldehydes by were examined [67]. From the NMR spectrum (Fig. 10.43) the broadening of peaks after hexanal-modified modification was noted, which indicates that their movement is restricted by being connected to chitosan chains.

10.3.10 Water contact angle Hydrophobic aerogel was obtained from modifying hydrophilic groups on chitosan. The modification does not damage its mechanical properties, because no obvious changes were found in the structure of the modified sample T-CS-OCA. As shown in Fig. 10.44B and E, both hydrophilic and oleophilic properties were found for the unmodified

Handbook of Chitin and Chitosan

324

10. Chitin and chitosan-based aerogels

FIGURE 10.44

Water contact angle images of CS-OCA (A) and T-CS-OCA (D). Water and oil droplets on CS-OCA (B) and T-CS-OCA (E). SEM images of CS-OCA (C) and T-CSOCA (F). Source: Reprinted with permission from Z. Li, L. Shao, W. Hu, T. Zheng, L. Lu, Y. Cao, et al., Excellent reusable chitosan/cellulose aerogel as an oil and organic solvent absorbent, Carbohydr. Polym. 191 (2018) 183 190. doi:10.1016/j.carbpol.2018.03.027. Elsevier Ltd., Copyright r 2018.

sample CS-OCA, while the modified sample T-CS-OCA was converted to hydrophobic and remained lipophilic. The water droplet was absorbed instantly by CS-OCA (as shown in Fig. 10.44A) when it came in contact with the surface of CS-OCA, while the water contact angle of T-CS-OCA was up to 152.8 degrees (as shown in Fig. 10.44D). The water absorption capacity of T-CS-OCA was only 0.16 g/g, which decreased 99.15% and showed a hydrophobic ability. While the oil absorption capacity of T-CSOCA was 14.76 g/g, and the lipophilicity was almost unchanged. Hence, after modification, TCS-OCA exhibited high oil/ water selectivity [83].

10.4 Future aspects of aerogel The aerogel field will continue its rapid development in the coming years. They are promising materials for a variety of high-performance applications because of their unique and adjustable physical properties, such as low density, extremely high specific surface area, and porosity. These remarkable properties, combined with readily modifiable surface chemistry can lead to the development of new types of aerogels to produce target product properties, this is a result of the versatility of the synthesis approach. The major application of aerogels is in thermal insulation (aerospace and building sectors), but they are also promising

Handbook of Chitin and Chitosan

10.4 Future aspects of aerogel

325

candidates for many other applications, such as adsorption and environmental cleanup, chemical sensors, biomedical, and pharmaceutical, due to their tunable surface chemistry and excellent textural properties. However, there are still some issues regarding the preparation and modification of chitin and chitosan-based aerogels. The main issues of directly using the biopolymers are the structural strength and stability performance (such as thermal stability and capability for repeated adsorption) of chitin and chitosan aerogels which can be improved by physical mixing or chemical modifications. In this regard, the superior applications of aerogels are relied on because they act as an effective material for capturing the CO2 and toxic volatile organic compounds from atmospheric and industrial release. In the outlook of water treatment, aerogels have been considered as interesting material for an effective abatement of various toxic organic solvents as well as dyes and heavy metal ions that are discharged from industrial and municipal wastes to water sources and they also can be employed for oil removal. A few developing and developed material applications are also discussed below. As a principal product of hemoglobin metabolism, bilirubin is normally conjugated with albumin and transported to the liver for excretion. When excess bilirubin accumulates in human blood, it will result in hyperbilirubinemia, which may bring about a yellow discoloration of the skin or other tissues, and lead to hepatic coma and even to death. Therefore the removal of excess bilirubin from the blood of patients who suffer from liver failure is vital to obtain enough time for liver transplantation or recovery of the damaged liver. Song et al. fabricated

FIGURE 10.45 Graphical representation of ch/GO aerogel beads and its adsorption of bilirubin. Source: Reprinted with permission from X. Song, X. Huang, Z. Li, Z. Li, K. Wu, Y. Jiao, et al., Construction of blood compatible chitin/graphene oxide composite aerogel beads for the adsorption of bilirubin, Carbohydr. Polym. 207 (2019) 704 712. doi:10.1016/j.carbpol.2018.12.005. Elsevier Ltd., Copyright r 2018.

Handbook of Chitin and Chitosan

326

10. Chitin and chitosan-based aerogels

FIGURE 10.46 Digital photographs (A F) of removal of oil from the water surface by MHPCA and an external magnetic field. Source: Reprinted with permission from Z. Xu, X. Jiang, H. Zhou, J. Li, Preparation of magnetic hydrophobic polyvinyl alcohol (PVA) cellulose nanofiber (CNF) aerogels as effective oil absorbents, Cellulose. 25 (2018) 1217 1227. doi:10.1007/ s10570-017-1619-9. Springer Nature Copyright r 2017.

a novel Ch/GO (chitin/graphene oxide) aerogel beads which showed higher bilirubin adsorption capacity [26] (Fig. 10.45). High selectivity, high oil absorption capacity, fast absorption rate, and easy collection showed the effective oil absorption of MHPCA; the MHPCA can be magnetically driven to the polluted region and quickly absorbs oil while the sample repels the oil. After absorption, the sample loaded with oil remains floating on the water surface [84] (Fig. 10.46). Chitosan aerogels loaded with a bioactive compound (vancomycin) were evaluated for their potential use in chronic wounds healing. The loading of vancomycin in the aerogels provides a fast local administration of the antibiotic at the wound site to prevent infections shortly after wound debridement without compromising cell viability. Vancomycinloaded chitosan aerogels arise as a promising formulation to be incorporated in dressings for the management of chronic wounds [85] (Fig. 10.47). Selective CO2 adsorption using COF-IL@chitosan aerogel [69]. GOCA was prepared by a facile ice-templating technique without using any cross-linking reagent for metanil yellow dye sequestration [73]. RSM yielded an adsorption capacity of 430.99 mg/g under these optimum conditions. Polydopamine-modified chitosan (CS-PDA) aerogels were synthesized through dopamine self-polymerization and glutaraldehyde

Handbook of Chitin and Chitosan

10.5 Conclusions

327

FIGURE 10.47

Overall application of vancomycin-loaded chitosan aerogel. Source: Reprinted with permission from C. Lo´pez-Iglesias, J. Barros, I. Ardao, F.J. Monteiro, C. AlvarezLorenzo, J.L. Go´mez-Amoza, et al., Vancomycin-loaded chitosan aerogel particles for chronic wound applications, Carbohydr. Polym. 204 (2019) 223 231. doi:10.1016/j.carbpol.2018.10.012. Elsevier Ltd., Copyright r 2018.

cross-linking reactions to enhance the adsorption capacity and acid resistance of chitosan. CS-PDA exhibited superior adsorption performances in the removal of Cr(VI), Pb(II), and organic dyes. The maximum adsorption capacities of CS-PDA for Cr(VI) and Pb(II) were 374.4 and 441.2 mg/g, respectively. These superiorities make CS-PDA a promising multifunctional adsorbent for the purification of metal ions and dyes [70]. The chitosan aerogels showed flexibility and higher mechanical toughness than conventional aerogels. The characteristic mechanical properties associated with a low thermal conductivity enable us to realize a new category of thermal insulators: environmentfriendly, transparent, and flexible [76].

10.5 Conclusions This chapter describes the bioaerogels, especially chitin and chitosanbased aerogels, their classification, and characterization studies using various techniques. The process of preparation of aerogels comprised the formation of physical gels and drying techniques. Mechanical strength and porosity are essential for determining the application of aerogels in practical fields. The reported aerogels have specific surface areas over 350 m2/g and most of the modified aerogels are considered to be mesoporous material. The internal structure of the obtained aerogels appears as porous aggregated networks in microscopy images observed by using SEM. The aerogels maintained the chemical identity

Handbook of Chitin and Chitosan

328

10. Chitin and chitosan-based aerogels

of a chitosan or chitin backbone when modified according to FTIR analysis, indicating the presence of their characteristic functional groups. Further NMR spectra were used to confirm structural modifications and functional groups of aerogels. TEM is useful for the smaller scale up to nanolevels and shows mesoparticle size, confirmation of the porous nature, the distribution of incorporated particles, morphological homogeneity of aerogel images, and also shows the exfoliated structure. XPS can be used to investigate a wide range of surface modifications on aerogel. However, the application of aerogels depends on the nature of the materials (environment-friendly, renewability, biocompatibility, and biodegradability), but also some other properties such as low density, high porosity, and a high specific surface area play important roles. Chitin and chitosan-based aerogels are particularly well-suited for applications in the areas of adsorption and the separation of biomedical and thermal insulation materials, as well as many other fields. Rapid growth in the research areas of life science, environmental applications, and electrochemistry is expected, which can be achieved by the further development of organic and hybrid aerogels and the optimization of their manufacturing processes at pilot and production scale.

Acknowledgments The authors would like to thank the authorities of Gandhigram Rural Institute, DU for the encouragement.

References [1] I. Smirnova, P. Gurikov, Aerogel production: current status, research directions, and future opportunities, J. Supercrit. Fluids 134 (2018) 228 233. Available from: https:// doi.org/10.1016/j.supflu.2017.12.037. [2] X. Yang, E.D. Cranston, Chemically cross-linked cellulose nanocrystal aerogels with shape recovery and superabsorbent properties, Chem. Mater. 26 (2014) 6016 6025. Available from: https://doi.org/10.1021/cm502873c. [3] A. Du, B. Zhou, Z. Zhang, J. Shen, A special material or a new state of matter: a review and reconsideration of the aerogel, Mater. (Basel) 6 (2013) 941 968. Available from: https://doi.org/10.3390/ma6030941. [4] S.S. Kistler, Coherent expanded aerogels and jellies, Nature 127 (1931) 741. [5] H. Maleki, Recent advances in aerogels for environmental remediation applications: a review, Chem. Eng. J. 300 (2016) 98 118. Available from: https://doi.org/10.1016/j. cej.2016.04.098. [6] X.D. Gao, Y. Di Huang, T.T. Zhang, Y.Q. Wu, X.M. Li, Amphiphilic SiO2 hybrid aerogel: an effective absorbent for emulsified wastewater, J. Mater. Chem. A 5 (2017) 12856 12862. Available from: https://doi.org/10.1039/c7ta02196h. [7] S. Naskar, J.F. Miethe, S. Sa´nchez-Paradinas, N. Schmidt, K. Kanthasamy, P. Behrens, et al., Photoluminescent aerogels from quantum wells, Chem. Mater. 28 (2016) 2089 2099. Available from: https://doi.org/10.1021/acs.chemmater.5b04872.

Handbook of Chitin and Chitosan

References

329

[8] A.M. Joseph, B. Nagendra, P. Shaiju, K.P. Surendran, E.B. Gowd, Aerogels of hierarchically porous syndiotactic polystyrene with a dielectric constant near to air, J. Mater. Chem. C. 6 (2018) 360 368. Available from: https://doi.org/10.1039/ c7tc05102f. [9] Y. Tang, K.L. Yeo, Y. Chen, L.W. Yap, W. Xiong, W. Cheng, Ultralow-density copper nanowire aerogel monoliths with tunable mechanical and electrical properties, J. Mater. Chem. A 1 (2013) 6723 6726. Available from: https://doi.org/10.1039/ c3ta10969k. [10] T. Xia, H. Yang, J. Li, C. Sun, C. Lei, Z. Hu, et al., Synthesis and physicochemical characterization of silica aerogels by rapid seed growth method, Ceram. Int. 45 (2019) 7071 7076. Available from: https://doi.org/10.1016/j.ceramint.2018.12.209. [11] X. Xu, R. Wang, P. Nie, Y. Cheng, X. Lu, L. Shi, et al., Copper nanowire-based aerogel with tunable pore structure and its application as flexible pressure sensor, ACS Appl. Mater. Interfaces 9 (2017) 14273 14280. Available from: https://doi.org/10.1021/ acsami.7b02087. [12] K. Shang, J.C. Yang, Z.J. Cao, W. Liao, Y.Z. Wang, D.A. Schiraldi, Novel polymer aerogel toward high dimensional stability, mechanical property, and fire safety, ACS Appl. Mater. Interfaces 9 (2017) 22985 22993. Available from: https://doi.org/ 10.1021/acsami.7b06096. [13] T. Shimizu, K. Kanamori, K. Nakanishi, Silicone-Based Organic Inorganic Hybrid Aerogels and Xerogels (2017) 1 13. Available from: https://doi.org/10.1002/ chem.201603680. [14] B.G. Hasegawa, T. Shimizu, K. Kanamori, A. Maeno, H. Kaji, Highly flexible hybrid polymer aerogels and xerogels based on resorcinol-formaldehyde with enhanced elastic stiffness and recoverability: insights into the origin of their mechanical properties (n.d.). [15] Q. Zeng, T. Mao, H. Li, Y. Peng, Thermally insulating lightweight cement-based composites incorporating glass beads and nano-silica aerogels for sustainably energysaving buildings, Energy Build. 174 (2018) 97 110. Available from: https://doi.org/ 10.1016/j.enbuild.2018.06.031. [16] J. Tang, Y. Song, F. Zhao, S. Spinney, J. da Silva Bernardes, K.C. Tam, Compressible cellulose nanofibril (CNF) based aerogels produced via a bio-inspired strategy for heavy metal ion and dye removal, Carbohydr. Polym. 208 (2019) 404 412. Available from: https://doi.org/10.1016/j.carbpol.2018.12.079. [17] G. Crini, Recent developments in polysaccharide-based materials used as adsorbents in wastewater treatment, Prog. Polym. Sci. 30 (2005) 38 70. Available from: https:// doi.org/10.1016/j.progpolymsci.2004.11.002. [18] C.A. Garcı´a-Gonza´lez, M. Alnaief, I. Smirnova, Polysaccharide-based aerogels— promising biodegradable carriers for drug delivery systems, Carbohydr. Polym. 86 (2011) 1425 1438. Available from: https://doi.org/10.1016/j.carbpol.2011.06.066. [19] L. Baldino, S. Cardea, M. Scognamiglio, E. Reverchon, A new tool to produce alginatebased aerogels for medical applications, by supercritical gel drying, J. Supercrit. Fluids 146 (2019) 152 158. Available from: https://doi.org/10.1016/j.supflu.2019.01.016. [20] K.S. Mikkonen, K. Parikka, A. Ghafar, M. Tenkanen, Prospects of polysaccharide aerogels as modern advanced food materials, Trends Food Sci. Technol. 34 (2013) 124 136. Available from: https://doi.org/10.1016/j.tifs.2013.10.003. [21] F. De Cicco, P. Russo, E. Reverchon, C.A. Garcı´a-Gonza´lez, R.P. Aquino, P. Del Gaudio, Prilling and supercritical drying: a successful duo to produce core-shell polysaccharide aerogel beads for wound healing, Carbohydr. Polym. 147 (2016) 482 489. Available from: https://doi.org/10.1016/j.carbpol.2016.04.031. [22] S. Takeshita, A. Sadeghpour, W.J. Malfait, A. Konishi, K. Otake, S. Yoda, Formation of nanofibrous structure in biopolymer aerogel during supercritical CO2 processing:

Handbook of Chitin and Chitosan

330

[23]

[24] [25]

[26]

[27]

[28]

[29]

[30]

[31]

[32] [33]

[34]

[35]

[36]

[37]

10. Chitin and chitosan-based aerogels

the case of chitosan aerogel, Biomacromolecules. (2019). Available from: https://doi. org/10.1021/acs.biomac.9b00246. acs.biomac.9b00246. J. Jiang, Q. Zhang, X. Zhan, F. Chen, Renewable, Biomass-Derived, Honeycomblike Aerogel a Robust. Oil Absorbent Two-Way Reusability (2017) 10307 10316. Available from: https://doi.org/10.1021/acssuschemeng.7b02333. Highly efficient flame retardant polyurethane foam with alginate/clay aerogel coating (2016). doi:10.1021/acsami.6b11659. B. Duan, Y. Huang, A. Lu, L. Zhang, Recent advances in chitin based materials constructed via physical methods, Prog. Polym. Sci. 82 (2018) 1 33. Available from: https://doi.org/10.1016/j.progpolymsci.2018.04.001. X. Song, X. Huang, Z. Li, Z. Li, K. Wu, Y. Jiao, et al., Construction of blood compatible chitin/graphene oxide composite aerogel beads for the adsorption of bilirubin, Carbohydr. Polym. 207 (2019) 704 712. Available from: https://doi.org/10.1016/j. carbpol.2018.12.005. C. Tsioptsias, C. Michailof, G. Stauropoulos, C. Panayiotou, Chitin and carbon aerogels from chitin alcogels, Carbohydr. Polym. 76 (2009) 535 540. Available from: https://doi.org/10.1016/j.carbpol.2008.11.018. S. Suenaga, M. Osada, Preparation of β-chitin nanofiber aerogels by lyophilization, Int. J. Biol. Macromol. 126 (2019) 1145 1149. Available from: https://doi.org/ 10.1016/j.ijbiomac.2019.01.006. Y. Tsutsumi, H. Koga, Z.D. Qi, T. Saito, A. Isogai, Nanofibrillar chitin aerogels as renewable base catalysts, Biomacromolecules. 15 (2014) 4314 4319. Available from: https://doi.org/10.1021/bm501320b. L. Panariello, M.B. Coltelli, M. Buchignani, A. Lazzeri, Chitosan and nanostructured chitin for biobased anti-microbial treatments onto cellulose based materials, Eur. Polym. J. 113 (2019) 328 339. Available from: https://doi.org/10.1016/j. eurpolymj.2019.02.004. S. Gopi, P. Balakrishnan, A. Pius, S. Thomas, Chitin nanowhisker (ChNW)-functionalized electrospun PVDF membrane for enhanced removal of Indigo carmine, Carbohydr. Polym. 165 (2017) 115 122. Available from: https://doi.org/10.1016/j. carbpol.2017.02.046. M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (2006) 603 632. Available from: https://doi.org/10.1016/j.progpolymsci.2006.06.001. B. Doshi, E. Repo, J.P. Heiskanen, J.A. Sirvio¨, M. Sillanpa¨a¨, Sodium salt of oleoyl carboxymethyl chitosan: a sustainable adsorbent in the oil spill treatment, J. Clean. Prod. 170 (2018) 339 350. Available from: https://doi.org/10.1016/j.jclepro.2017.09.163. S. Gopi, P. Balakrishnan, C. Divya, S. Valic, E. Govorcin Bajsic, A. Pius, et al., Facile synthesis of chitin nanocrystals decorated on 3D cellulose aerogels as a new multifunctional material for waste water treatment with enhanced anti-bacterial and antioxidant properties, N. J. Chem. 41 (2017) 12746 12755. Available from: https://doi. org/10.1039/c7nj02392h. Y. Zhao, L. Ma, R. Zeng, M. Tu, J. Zhao, Preparation, characterization and protein sorption of photo-crosslinked cell membrane-mimicking chitosan-based hydrogels, Carbohydr. Polym. 151 (2016) 237 244. Available from: https://doi.org/10.1016/j. carbpol.2016.05.067. L. Pereira, M. Alves, Dyes-environmental impact and remediation, Environmental Protection Strategies for Sustainable Development (2012) 111 162. Available from: https://doi.org/10.1007/978-94-007-1591-2_4. M. Yadav, P. Goswami, K. Paritosh, M. Kumar, N. Pareek, V. Vivekanand, Seafood waste: a source for preparation of commercially employable chitin/chitosan materials, Bioresour, Bioprocess 6 (2019). Available from: https://doi.org/10.1186/s40643019-0243-y.

Handbook of Chitin and Chitosan

References

331

[38] S.S. Silva, J.F. Mano, R.L. Reis, Ionic liquids in the processing and chemical modification of chitin and chitosan for biomedical applications, Green. Chem. 19 (2017) 1208 1220. Available from: https://doi.org/10.1039/c6gc02827f. [39] M.S. Rao, S.R. Kanatt, S.P. Chawla, A. Sharma, Chitosan and guar gum composite films: preparation, physical, mechanical and antimicrobial properties, Carbohydr. Polym. 82 (2010) 1243 1247. Available from: https://doi.org/ 10.1016/j.carbpol.2010.06.058. [40] K.M. Reddy, V.R. Babu, M. Sairam, M.C.S. Subha, N.N. Mallikarjuna, P.V. Kulkarni, et al., Development of chitosan-guar gum semi-interpenetrating polymer network microspheres for controlled release of cefadroxil, Des. Monomers Polym. 9 (2006) 491 501. Available from: https://doi.org/10.1163/156855506778538047. [41] T. Dai, M. Tanaka, Y.Y. Huang, M.R. Hamblin, Chitosan preparations for wounds and burns: antimicrobial and wound-healing effects, Expert. Rev. Anti. Infect. Ther. 9 (2011) 857 879. Available from: https://doi.org/10.1586/eri.11.59. [42] G.A. Morris, S.M. Ko¨k, S.E. Harding, G.G. Adams, Polysaccharide drug delivery systems based on pectin and chitosan, Biotechnol. Genet. Eng. Rev. 27 (2010) 257 284. Available from: https://doi.org/10.1080/02648725.2010.10648153. [43] P.R. Sivashankari, M. Prabaharan, Prospects of chitosan-based scaffolds for growth factor release in tissue engineering, Int. J. Biol. Macromol. 93 (2016) 1382 1389. Available from: https://doi.org/10.1016/j.ijbiomac.2016.02.043. [44] B. Ding, J. Cai, J. Huang, L. Zhang, Y. Chen, X. Shi, et al., Facile preparation of robust and biocompatible chitin aerogels, J. Mater. Chem. 22 (2012) 5801 5809. Available from: https://doi.org/10.1039/c2jm16032c. [45] F. Li, L. Ding, B. Yao, N. Huang, J. Li, Q. Fu, et al., Pd loaded and covalent-organic framework involved chitosan aerogels and their application for continuous flowthrough aqueous CB decontamination, J. Mater. Chem. A Mater. Energy Sustain. 00 (2018) 1 7. Available from: https://doi.org/10.1039/C8TA03275K. [46] N. Kuthirummal, A. Dean, C. Yao, W. Risen, Photo-formation of gold nanoparticles: photoacoustic studies on solid monoliths of Au(III)-chitosan-silica aerogels, Spectrochim. Acta—Part A Mol. Biomol. Spectrosc. 70 (2008) 700 703. Available from: https://doi.org/10.1016/j.saa.2007.09.011. [47] T.T.L. Chau, D.Q.T. Le, H.T. Le, C.D. Nguyen, L.V. Nguyen, T.D. Nguyen, Chitin liquid-crystal-templated oxide semiconductor aerogels, ACS Appl. Mater. Interfaces 9 (2017) 30812 30820. Available from: https://doi.org/10.1021/acsami.7b07680. [48] C.A. Garcı´a-Gonza´lez, L. Dura˜es, P. del Gaudio, H. Maleki, M. Mahmoudi, A. Portugal, Synthesis and biomedical applications of aerogels: possibilities and challenges, Adv. Colloid Interf. Sci. 236 (2016) 1 27. Available from: https://doi. org/10.1016/j.cis.2016.05.011. [49] H. Ennajih, R. Bouhfid, E.M. Essassi, M. Bousmina, A. El Kadib, Chitosanmontmorillonite bio-based aerogel hybrid microspheres, Microporous Mesoporous Mater. 152 (2012) 208 213. Available from: https://doi.org/10.1016/j. micromeso.2011.11.032. [50] S. Groult, T. Budtova, Tuning structure and properties of pectin aerogels, Eur. Polym. J. 108 (2018) 250 261. Available from: https://doi.org/10.1016/j. eurpolymj.2018.08.048. [51] H.B. Chen, P. Shen, M.J. Chen, H.B. Zhao, D.A. Schiraldi, Highly efficient flame retardant polyurethane foam with alginate/clay aerogel coating, ACS Appl. Mater. Interfaces 8 (2016) 32557 32564. Available from: https://doi.org/10.1021/acsami.6b11659. [52] J. Zhu, J. Hu, C. Jiang, S. Liu, Y. Li, Ultralight, hydrophobic, monolithic konjac glucomannan-silica composite aerogel with thermal insulation and mechanical properties, Carbohydr. Polym. 207 (2019) 246 255. doi:10.1016/j.carbpol.2018.11.073.

Handbook of Chitin and Chitosan

332

10. Chitin and chitosan-based aerogels

[53] D.R. Chen, X.H. Chang, X.L. Jiao, Aerogels in the Environment Protection, Elsevier B. V, 2014. Available from: https://doi.org/10.1016/B978-0-444-63283-8.00022-3. [54] M. Robitzer, A. Tourrette, R. Horga, R. Valentin, M. Boissie`re, J.M. Devoisselle, et al., Nitrogen sorption as a tool for the characterisation of polysaccharide aerogels, Carbohydr. Polym. 85 (2011) 44 53. Available from: https://doi.org/10.1016/j. carbpol.2011.01.040. [55] X. Chang, D. Chen, X. Jiap, Chitosan-based aerogels with high adsorption performance, J. Phys. Chem. B 112 (2008) 7721 7725. Available from: https://doi.org/ 10.1021/jp8011359. [56] A. El Kadib, K. Molvinger, T. Cacciaguerra, M. Bousmina, D. Brunel, Chitosan templated synthesis of porous metal oxide microspheres with filamentary nanostructures, Microporous Mesoporous Mater. 142 (2011) 301 307. Available from: https:// doi.org/10.1016/j.micromeso.2010.12.012. [57] Q. Ma, Y. Liu, Z. Dong, J. Wang, X. Hou, Hydrophobic and nanoporous chitosansilica composite aerogels for oil absorption, J. Appl. Polym. Sci. 132 (2015) 1 11. Available from: https://doi.org/10.1002/app.41770. [58] P. Hao, J. Tian, Y. Sang, C.C. Tuan, G. Cui, X. Shi, et al., 1D Ni-Co oxide and sulfide nanoarray/carbon aerogel hybrid nanostructures for asymmetric supercapacitors with high energy density and excellent cycling stability, Nanoscale 8 (2016) 16292 16301. Available from: https://doi.org/10.1039/c6nr05385h. [59] X.L. Wang, D.M. Guo, Q. Da, An, Z.Y. Xiao, S.R. Zhai, High-efficacy adsorption of Cr (VI) and anionic dyes onto β-cyclodextrin/chitosan/hexamethylenetetramine aerogel beads with task-specific, integrated components, Int. J. Biol. Macromol. 128 (2019) 268 278. Available from: https://doi.org/10.1016/j.ijbiomac.2019.01.139. [60] R. Li, Q. Da An, Z.Y. Xiao, B. Zhai, S.R. Zhai, Z. Shi, Preparation of PEI/CS aerogel beads with a high density of reactive sites for efficient Cr(VI) sorption: batch and column studies, RSC Adv. 7 (2017) 40227 40236. Available from: https://doi.org/ 10.1039/c7ra06914f. [61] R.M. Stroud, J.W. Long, J.J. Pietron, D.R. Rolison, A practical guide to transmission electron microscopy of aerogels, J. Non. Cryst. Solids (2004) 277 284. Available from: https://doi.org/10.1016/j.jnoncrysol.2004.05.013. [62] A.A. Michael, L. Nicholas, M.M. Koebel, Resorcinol-Formaldehyde Aerogels (2011). Available from: https://doi.org/10.1007/978-1-4614-1957-0. [63] J. Singh, P.K. Dutta, J. Dutta, A.J. Hunt, D.J. Macquarrie, J.H. Clark, Preparation and properties of highly soluble chitosan-L-glutamic acid aerogel derivative, Carbohydr. Polym. 76 (2009) 188 195. Available from: https://doi.org/10.1016/j. carbpol.2008.10.011. [64] B. Yu, J. Xu, J.H. Liu, S.T. Yang, J. Luo, Q. Zhou, et al., Adsorption behavior of copper ions on graphene oxide-chitosan aerogel, J. Environ. Chem. Eng. (2013). Available from: https://doi.org/10.1016/j.jece.2013.08.017. [65] S. Zhao, H. Xu, L. Wang, P. Zhu, W.M. Risen, J. William Suggs, Synthesis of novel chitaline-silica aerogels with spontaneous Au and Ag nanoparticles formation in aerogels matrix, Microporous Mesoporous Mater. 171 (2013) 147 155. Available from: https://doi.org/10.1016/j.micromeso.2012.12.038. [66] T. Budtova, Cellulose II Aerogels: A Review, Springer, Netherlands, 2019. Available from: https://doi.org/10.1007/s10570-018-2189-1. [67] S. Takeshita, A. Konishi, Y. Takebayashi, S. Yoda, K. Otake, Aldehyde approach to hydrophobic modification of chitosan aerogels, Biomacromolecules 18 (2017) 2172 2178. Available from: https://doi.org/10.1021/acs.biomac.7b00562. [68] D.K. Singh, V. Kumar, V.K. Singh, S.H. Hasan, R.N. Moussawi, D. Patra, et al., Modeling of adsorption behavior of the amine-rich GOPEI aerogel for the removal of

Handbook of Chitin and Chitosan

References

[69]

[70]

[71]

[72]

[73]

[74]

[75]

[76]

[77]

[78]

[79]

[80]

[81]

[82]

333

As(iii) and As(v) from aqueous media, RSC Adv. 6 (2016) 56684 56697. Available from: https://doi.org/10.1039/C6RA10518A. L.G. Ding, B.J. Yao, F. Li, S.C. Shi, N. Huang, H.B. Yin, et al., Ionic liquid-decorated COF and its covalent composite aerogel for selective CO2 adsorption and catalytic conversion, J. Mater. Chem. A 7 (2019) 4689 4698. Available from: https://doi.org/ 10.1039/c8ta12046c. D.-M. Guo, Q.-D. An, Z.-Y. Xiao, S.-R. Zhai, D.-J. Yang, Efficient removal of Pb(II), Cr(VI) and organic dyes by polydopamine modified chitosan aerogels, Carbohydr. Polym. 202 (2018) 306 314. Available from: https://doi.org/10.1016/j.carbpol.2018.08.140. J.P. Chaudhary, N. Vadodariya, S.K. Nataraj, R. Meena, Chitosan-based aerogel membrane for robust oil-in-water emulsion separation, ACS Appl. Mater. Interfaces 7 (2015) 24957 24962. Available from: https://doi.org/10.1021/acsami.5b08705. A.A. Alhwaige, T. Agag, H. Ishida, S. Qutubuddin, Biobased chitosan hybrid aerogels with superior adsorption: role of graphene oxide in CO2 capture, RSC Adv. 3 (2013) 16011 16020. Available from: https://doi.org/10.1039/c3ra42022a. K.C. Lai, B.Y.Z. Hiew, L.Y. Lee, S. Gan, S. Thangalazhy-Gopakumar, W.S. Chiu, et al., Ice-templated graphene oxide/chitosan aerogel as an effective adsorbent for sequestration of metanil yellow dye, Bioresour. Technol. 274 (2019) 134 144. Available from: https://doi.org/10.1016/j.biortech.2018.11.048. T. Zhai, L. Verdolotti, S. Kaciulis, P. Cerruti, G. Gentile, H. Xia, et al., High piezoresistive performances of an anisotropic composite realized by embedding rGObased chitosan aerogel in open cell polyurethane foams, Nanoscale 11 (2019) 8835 8844. Available from: https://doi.org/10.1039/c9nr00157c. S. Frindy, A. Primo, H. Ennajih, A. el kacem Qaiss, R. Bouhfid, M. Lahcini, et al., Chitosan graphene oxide films and CO2-dried porous aerogel microspheres: interfacial interplay and stability, Carbohydr. Polym. 167 (2017) 297 305. Available from: https://doi.org/10.1016/j.carbpol.2017.03.034. S. Takeshita, S. Yoda, Chitosan aerogels: transparent, flexible thermal insulators, Chem. Mater. 27 (2015) 7569 7572. Available from: https://doi.org/10.1021/acs. chemmater.5b03610. C. Su, H. Yang, H. Zhao, Y. Liu, R. Chen, Recyclable and biodegradable superhydrophobic and superoleophilic chitosan sponge for the effective removal of oily pollutants from water, Chem. Eng. J. (2017). Available from: https://doi.org/10.1016/j. cej.2017.07.157. A. Salam, R.A. Venditti, J.J. Pawlak, K. El-Tahlawy, Crosslinked hemicellulose citrate-chitosan aerogel foams, Carbohydr. Polym. (2011). Available from: https:// doi.org/10.1016/j.carbpol.2011.01.008. S. Zhang, J. Feng, J. Feng, Y. Jiang, L. Li, Ultra-low shrinkage chitosan aerogels trussed with polyvinyl alcohol, Mater. Des. 156 (2018) 398 406. Available from: https://doi.org/10.1016/j.matdes.2018.07.004. R. Valentin, B. Bonelli, E. Garrone, F. Di Renzo, F. Quidnard, Accessibility of the functional group of chitosan aerogel probed by FT-IR-monitored deuteration, Biomacromolecules 8 (2007) 3646 3650. Available from: https://doi.org/10.1021/ bm070391a. M. Salzano de Luna, C. Ascione, C. Santillo, L. Verdolotti, M. Lavorgna, G.G. Buonocore, et al., Optimization of dye adsorption capacity and mechanical strength of chitosan aerogels through crosslinking strategy and graphene oxide addition, Carbohydr. Polym. 211 (2019) 195 203. Available from: https://doi.org/10.1016/j. carbpol.2019.02.002. M. Pia˛tkowski, J. Radwan-Pragłowska, Ł. Janus, D. Bogdał, D. Matysek, V. Cablik, Microwave-assisted synthesis and characterization of chitosan aerogels doped with

Handbook of Chitin and Chitosan

334

10. Chitin and chitosan-based aerogels

Au-NPs for skin regeneration, Polym. Test. 73 (2019) 366 376. Available from: https://doi.org/10.1016/j.polymertesting.2018.11.024. [83] Z. Li, L. Shao, W. Hu, T. Zheng, L. Lu, Y. Cao, et al., Excellent reusable chitosan/cellulose aerogel as an oil and organic solvent absorbent, Carbohydr. Polym. 191 (2018) 183 190. Available from: https://doi.org/10.1016/j.carbpol.2018.03.027. [84] Z. Xu, X. Jiang, H. Zhou, J. Li, Preparation of magnetic hydrophobic polyvinyl alcohol (PVA) cellulose nanofiber (CNF) aerogels as effective oil absorbents, Cellulose 25 (2018) 1217 1227. Available from: https://doi.org/10.1007/s10570-017-1619-9. [85] C. Lo´pez-Iglesias, J. Barros, I. Ardao, F.J. Monteiro, C. Alvarez-Lorenzo, J.L. Go´mezAmoza, et al., Vancomycin-loaded chitosan aerogel particles for chronic wound applications, Carbohydr. Polym. 204 (2019) 223 231. Available from: https://doi. org/10.1016/j.carbpol.2018.10.012.

Handbook of Chitin and Chitosan

C H A P T E R

11 Chitin, chitosan, marine to market G.M. Oyatogun1, T.A. Esan2, E.I. Akpan3, S.O. Adeosun4, A.P.I. Popoola5, B.I. Imasogie1, W.O. Soboyejo6, A.A. Afonja1, S.A. Ibitoye1, V.D. Abere7, A.O. Oyatogun1, K.M. Oluwasegun1, I.E. Akinwole1 and K.J. Akinluwade8 1

Department of Materials Science and Engineering, Obafemi Awolowo University, Ile-Ife, Nigeria, 2Department of Restorative Dentistry, Obafemi Awolowo University, Ile-Ife, Nigeria, 3Institute for Composite Materials, Technical University, Kaiserslautern, Germany, 4Department of Metallurgical and Materials Engineering, University of Lagos, Lagos, Nigeria, 5Deparment of Chemical, Metallurgical and Materials Engineering, Tshwane University of Technology, Pretoria, South Africa, 6Faculty of Engineering, Wisconsin Polytechnic Institute, Menomonie, WI, United States, 7Department of Mineral Processing, National Metallurgical Development Centre, Jos, Nigeria, 8Department of Research and Development, Prototype Engineering Development Institute (National Agency for Science and Engineering Infrastructure, NASENI), Ilesa, Nigeria O U T L I N E 11.1 Introduction

336

11.2 Origin and sources of chitin and chitosan

337

11.3 Synthesis of chitin and chitosan

339

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00011-0

335

© 2020 Elsevier Inc. All rights reserved.

336

11. Chitin, chitosan, marine to market

11.3.1 Synthesis of chitin 11.3.2 Synthesis of chitosan 11.3.3 Synthesis of derivatives of chitin and chitosan

339 344 346

11.4 Properties of chitin and chitosan 11.4.1 Physiochemical properties of chitin and chitosan 11.4.2 Biological properties of chitosan

348 349 351

11.5 Potential applications of chitin and chitosan 11.5.1 Biomedical application of chitosan 11.5.2 Industrial applications of chitosan

354 354 357

11.6 Economic potential of chitin and chitosan

361

11.7 Conclusions

364

References

364

Further reading

376

11.1 Introduction Chitin is the second most abundant natural amino polysaccharide on Earth, after cellulose [1 3]. It is a nitrogen-modified, high-molecularweight, linear homopolysaccharide that consists of repeated units of Nacetyl-D-glucosamine, bound together in β-(1 4)-N-acetyl-D-glucosamine bonds [2]. Chitin has a chemical formula of (C8H13O5N)n, hence it is considered to be a complex polysaccharide, whose structure resembles that of cellulose but with one hydroxyl group on each monomer replaced by an acetyl amine group, as shown in Fig. 11.1A. There are three different allomorphs of chitin. These are α-chitin, β-chitin, and γ- chitin. The main differences between the three are the degree of hydration, the size of the unit cell, and the number of chitin chains per unit cell [3,4]. The most abundant polymorph is the α-chitin, which has a tightly compacted orthorhombic cell, formed by alternating

FIGURE 11.1 Structure of (A) chitin and (B) chitosan.

Handbook of Chitin and Chitosan

11.2 Origin and sources of chitin and chitosan

337

sheets of antiparallel chains [3]. This polymorph occurs in fungal and yeast cell walls, in krill, lobster and crab tendons, crustacean shells, as well as in insect cuticles. The isomorph β-chitin has a monoclinic unit cell, with polysaccharide chains attached in a parallel manner and is found in association with proteins in squid pens [4]. The γ-chitin is more of a combination of both α-chitin and β-chitin rather than a third polymorph [3]. The compact structure of chitin limits its reactivity and solubility in most solvents, which consequently limits its application. This had led to several chemical modifications of chitin in an attempt to produce more soluble and reactive derivatives, one of which is chitosan [2 9]. Chitosan is a natural carbohydrate polymer obtained from deacetylation of chitin under alkaline condition [1,3,5]. It has a linear chain chemical structure that consists of β-(1,4)-linked 2-acetamino-2-deoxyβ-D-glucopyranose with 2-amino-2-deoxy-β-D-glucopyranose (see Fig. 11.1B). Chitosan is characterized by the presence of three functional groups; an amino group and primary and secondary hydroxyl groups situated at the C-2, C-3, and C-6 positions, respectively [2,4,7]. Chitosan has been found to be more reactive than chitin. It is biocompatible, biodegradable, nontoxic, and possesses good antimicrobial characteristics, good film-forming ability, good chelation, and good absorption properties. Consequently, chitosan has found diverse applications in fields such as environmental remediation, biomedical engineering, water engineering, biotechnology, food processing, cosmetics industry, textile industry, paper industry, agriculture, and photography [1,2,8 20]. Chitin and chitosan are therefore natural and abundant polymers with extensive structural possibilities for chemical and physical modifications [7 9]. These often culminate in novel properties that meet diverse functional requirements, consequently facilitating the versatile applications of chitin and chitosan [1,2,8,9,12 16]. Shrimp, crab, squid, lobster, insect cuticle, fungi, and yeast are the best naturally occurring sources of chitin. This chapter discusses the isolation of chitin from various marine-based sources. It also reviewed the synthesis of chitosan and the diverse applications of these versatile biopolymers.

11.2 Origin and sources of chitin and chitosan Chitin is a polysaccharide found in the exoskeleton of crustaceans, such as shrimp, crab, lobster, crawfish shells, and other marine zooplankton species such as cnidarian, foraminifera, and mollusks [16,17,20 29]. Similarly, marine gastropods such as sea shells, cone snails, coral and cowry shells, have been reported to be the other main sources of chitin [3,25 27,29 34]. Insect’s wing and fungi’s cell wall are

Handbook of Chitin and Chitosan

338

11. Chitin, chitosan, marine to market

FIGURE 11.2 Some marine-based sources of chitin.

also reported to contain chitin [5,21]. Fig. 11.2 shows some of the marine-based sources of chitin and chitosan. The chitinous solid waste fraction of the average Indian landing of shellfish was observed to range from 60,000 to 80,000 tonnes [23]. It has also been reported that dry prawn waste contained 23% chitin while dried squilla contained 15% chitin [5,23,28,29]. Similarly, Asford et al. reported that chitin represents 14% 27% and 13% 15% of the dry weight of shrimp and crab processing wastes, respectively [28]. Consequently, commercial chitins are usually isolated from marine crustaceans, because a large amount of waste is available as a by-product during food processing. According to Ibrahim et al., Shrimp consists of about 45% of the raw material used by the seafood processing industry while about 30% 40% by weight of this, that is, the exoskeleton (shells) is discarded as waste [28]. If not properly disposed, this may pose a threat to the environment. The conversion of this waste to useful and economically viable materials such as chitin and chitosan will maximize the economic potential of this waste and minimize its threat as a potential source of environmental pollution [29]. Dutta et al. affirmed that chitin and chitosan, which are waste products of the crabbing and shrimp canning industry are natural resources waiting for a market [2]. They also reported that in 1973 over 150,000 Mt of chitin was produced from waste from shellfish, krill, clams, oysters, squid, and fungi in the United States alone [2]. The innovative use of this renewable product to produce economically valuable and sustainable materials will therefore facilitate the conversion of waste to wealth, which could open up an untapped market.

Handbook of Chitin and Chitosan

11.3 Synthesis of chitin and chitosan

339

11.3 Synthesis of chitin and chitosan The exoskeleton of arthropods, crustaceans, marine zooplankton, and marine gastropods are composed of chitin that is found as a constituent of a complex network of proteins and calcium carbonate deposits that form the rigid shell [35]. Consequently, chitin isolation from these sources require the removal of two major constituents of the shell, that is, proteins by deproteinization and inorganic calcium carbonate by demineralization [3,35]. Most often, this is followed by decolorization to remove some of the residual pigments [16,17]. Many methods of chitin isolation have been articulately documented, although no standard method has been adopted. Both deproteinization and demineralization could be carried out using chemical or enzymatic treatments in which the order of the two steps may be reversed [3]. Microbial fermentation has also been employed to isolate chitin; in this case however deproteinization and demineralization steps are carried out simultaneously [3]. The chitin residue is then converted to chitosan by the standard deacetylation process. This section presents an in-depth review of the techniques used for chitin isolation and chitosan production, with an emphasis on chitin from marine-based sources.

11.3.1 Synthesis of chitin The biosynthesis of chitin is catalyzed by the enzyme chitin synthase, the pivotal enzyme in the chitin synthesis pathway that exists in every chitin-synthesizing organism [22]. The enzyme utilizes uridine-diphosphate-N-acetylglucosamine (UDP-GlcNAc), as the activated sugar donor to form the chitin polymer and remains bound to the growing polymer chains as it undergoes addition polymerization, that is, sequential addition of single GlcNAc units to the nonreducing end of the extending chain [21,22,30]. The linear polymer chain spontaneously assembles into microfibrils of varying diameter and length [22,30]. After polymerization, these are transported to the extracellular space where they are embedded in a matrix of protein, calcium carbonate, and phosphate [21]. Consequently, chitin is widely found naturally in marine invertebrates, insects, fungi, yeast, and other chitin-synthesizing organisms. Chitin was first synthesized by Prof. Henri Braconnot in 1811, who isolated it from mushrooms and named it fungine [31]. Subsequently, in 1823 Antoine Odier synthesized chitin from beetle cuticles and named it chitin, after the Greek word chiton, meaning a coat of mail [31]. Since then, global attention has been focused on the isolation of chitin and extensive work on chitin isolation from various chitin-synthesizing organisms has been articulately documented. In this chapter, attention

Handbook of Chitin and Chitosan

340

11. Chitin, chitosan, marine to market

is focused on the isolation of chitin from marine chitin-synthesizing organisms, since these have been reported as the most abundant sources of chitin [29,30,33]. 11.3.1.1 Synthesis of chitin by chemical method Synthesis of chitin by a chemical procedure is the most common method used to isolate chitin. Several studies have been documented on the successful isolation of chitin from crustacean shells by chemical process [33,34,36 47]. This process comprises deproteinization, demineralization, and decolorization. Demineralization involves the removal of inorganic matter, such as calcium carbonate, by acid treatment, while deproteinization entails the extraction of protein in an alkaline medium, and decolorization consists of the removal of pigments by chemical reagents to achieve a colorless product. Fig. 11.3 is a schematic flow chart of the process of isolation of chitin from marine-based exoskeletons. 11.3.1.1.1 Deproteinization

Deproteinization entails the removal of the protein content of the shell. According to Younes and Rinaudo, a wide range of alkaline solutions, such as sodium hydroxide (NaOH), sodium carbonate (Na2CO3), sodium hydrogen carbonate (NaHCO3), potassium hydroxide (KOH), potassium carbonate (K2CO3), calcium hydroxide (Ca(OH)2), sodium sulfite (Na2SO3), sodium hydrogen sulfite (NaHSO3), calcium hydrogen

FIGURE 11.3 Process of chemical isolation of chitin.

Handbook of Chitin and Chitosan

11.3 Synthesis of chitin and chitosan

341

sulfite (Ca(HSO3)2), trisodium phosphate (Na3PO4), and sodium sulfide (Na2S), have been studied as deproteinization reagents under different reaction conditions [3]. These authors, however, reported that NaOH was found to be the preferred reagent when applied at concentrations ranging from 0.125 to 5.0 M, at varying temperature (up to 160 C) and duration, from a few minutes up to a few days [3]. Subsequently, chemical deproteinization is often accomplished by reacting the shells with the aqueous alkaline solution of NaOH [30]. In some of the reported work, deproteinization was carried out after demineralization, hence the order in which any of these two processes is carried out during chitin isolation is not fixed [16,17,30]. 11.3.1.1.2 Demineralization

Demineralization entails the removal of minerals, specifically calcium carbonate (CaCO3), by acid treatment. It is usually carried out using any of the following acids: hydrochloric acid (HCl), nitric acid (HNO3), sulfuric acid (H2SO4), or acetic acid (CH3COOH). The conventional demineralization process is usually carried out using dilute hydrochloric acid [3]. During demineralization, decomposition of calcium carbonate into the water-soluble calcium salts with the release of carbon dioxide, as shown in Eq. (11.1), occurs. Simultaneously, other mineral matter in chitinbearing shells are also removed by this reaction to give soluble salts, 2HCl 1 CaCO3 -CaCl2 1 H2 O 1 CO2 m

(11.1)

which are then separated by filtration to obtain chitin residue. Extensive washing of the residue is carried out using deionized water. This is followed by acidimetric titration: to ensure that the solid residue is chitin [3]. According to Younes and Rinaudo, the demineralization process is often empirical and found to vary with the mineralization degree of shell, extraction time, temperature, particle size, acid concentration, and solute to solvent ratio [3]. These authors reported that, using HCl, demineralization could be achieved in 2 3 h under stirring. The reaction time was however found to be dependent on preparation methods [3]. Longer demineralization time was found to result in a slight drop in ash content along with polymer degradation [3]. In addition, Truong et al. reported that carrying out demineralization at high temperature accelerates the demineralization reaction [48]. Furthermore, to have a complete reaction and ensure total removal of all the minerals, it was suggested that acid intake should be greater than the stoichiometric amount of minerals, or that acid with higher concentration be utilized [37,38]. Marquis-Duval noted that the decisive factor in demineralization is the contact area between the chitin matrix and the solvent [49]. It was therefore established that high temperatures, longer incubations,

Handbook of Chitin and Chitosan

342

11. Chitin, chitosan, marine to market

high acid concentrations, and particle size affect the final physicochemical properties of the resulting chitin [3]. Consequently, the development of different demineralization processes evolved with varying demineralization process parameters to enhance the efficiency of the process [34,39,40]. For example, Horowitz et al. and Synowiecki et al. carried out demineralization at room temperature using 90% formic acid and 22% HCl, respectively [42,43]. Foster et al., however, stated that these drastic treatments may result in polymer degradation as the molecular weight (Mw) and acetylation degree was lowered [44]. Other researchers therefore considered the use of mild acid to prevent this [45,46]. For example, Austin et al. carried out demineralization using ethylene-diamine-tetracetic acid (EDTA) [45]. Similarly, Brine and Austin made use of acetic acid in their work [46], while Peniston and Johnson studied the sulfurous acid process [47]. 11.3.1.1.3 Decolorization

Decolorization is carried out to remove natural pigment existing in chitin. This is often accomplished by the addition of acetone to chitin residue under reflux conditions for a period of time [30,50]. However, Teli and Sheikh accomplished this using potassium permanganate (KMnO4) and oxalic acid [16]. Although the most documented method for isolating chitin from crustacean shells is the chemical method, the use of enzymatic hydrolysis for deproteinization [51] and microorganisms for both demineralization and deproteinization has been also reported [51,52]. The chemical extraction method was however reported to have higher yields and to produce higher purity chitin when compared to biological extraction [52]. The next section will give a brief review of chitin isolation by biological methods. 11.3.1.2 Synthesis of chitin by biological method Several works on synthesis of chitin by biological methods have been reported [3,53 58]. Bahasan et al. studied the isolation of chitin from the shrimp shell waste using microbes isolated from fermented milk and bread [53]. These authors concluded that the application of microorganisms for the extraction of chitin from the shrimp shell waste could be an alternative to the traditional chemical methods. According to Kaur and Dhillon, the chemical methods of chitin extraction utilize harsh chemicals at elevated temperatures for a prolonged time, which often affect the physicochemical properties of the isolated chitin and constitute serious environmental hazards [58]. These authors declared that green extraction methods are increasingly gaining popularity due to their environment-friendly nature, although not currently exploited to their maximum potential at the commercial level [58]. They therefore

Handbook of Chitin and Chitosan

11.3 Synthesis of chitin and chitosan

343

advocated for this novel method of chitin synthesis, stating that microorganisms-mediated fermentation processes are easy to handle, simple, fast, and can be optimized by controlling the process parameters, while the use of ambient temperature and negligible solvent consumption would result in reduced adverse environmental impact and costs [58]. Adour et al. extracted chitin from the teguments of white shrimp, Parapenaeus longirostris, by means of Lactobacillus helveticus grown on different media for comparison. The media comprised date juice waste and glucose. The authors extracted the chitin by inoculating the shrimp shells with a suspension of Lactobacillus helveticus strain, Milano, in a fermentor [54]. It was observed that within the pH range of 8.5 9.0 and a temperature of 30 C maximum deproteinization and demineralization obtained were 76% and 53%, for the 80 and 300 g/L glucose, respectively. Increasing the processing temperature from 30 C to 35 C was noted to result in a 60% increase in demineralization level. The use of date juice, as an alternative to the use of glucose, however, resulted in low demineralization, with the highest demineralization obtained being 44%. Similarly, increasing temperature and total sugar content were found to enhance the deproteinization process [54]. According to Healy et al. the conventional harsh, chemical method of chitin isolation is extremely hazardous, energy-consuming, and not ecofriendly because it employs high concentrations of mineral acid and alkali [55]. They therefore investigated the potential of the isolation of chitin from prawn shell by a fermentation process. This was claimed to provide a multiproduct process that will eliminate waste and generate revenue. The authors carried out anaerobic fermentations of prawn shell waste in a benchtop stirred-tank bioreactor from which they obtained various components of the fermentation products. They reported that the unique microbial mixture used in the fermentation of prawn shell waste resulted in the production of a purified chitin with calcium removal as high as 93.8%. They also reported the production of byproducts, which consisted of a pigmented liquor containing peptides and amino acids [55]. In a similar study, biological treatment of prawn waste for chitin production was investigated by Aytekin and Elibol [56]. These authors fermented prawn shell waste using two different microorganisms, Lactococcus lactis and Teredinobacter turnirae, in order to extract chitin from prawn waste in the presence of different glucose concentrations [56]. They applied different strategies in the course of the study. In one set of experiments both the bacteria were inoculated individually, while in cocultivation experiments three different strategies were applied. The first consisted of simultaneous inoculation of the bacteria, the second entailed starting the fermentation with protease producer and inoculating Teredinobacter turnirae at the end of the fourth day

Handbook of Chitin and Chitosan

344

11. Chitin, chitosan, marine to market

of cultivation, while the third strategy was the reverse of the second, that is, first Lactococcus lactis and then Teredinobacter turnirae inoculations were carried out. They reported that the individually cultured approach resulted in efficient removal of inorganic mineral matter when cultured with Lactococcus lactis while the Teredinobacter turnirae culture gave better deproteinization. The authors also reported that cofermentation of both bacteria using three different protocols resulted in the highest yield, 95.5%, when Teredinobacter turnirae was first inoculated [56]. They however affirmed that the extraction of chitin by biological treatment, though environment-friendly, was incomplete when compared to that of the chemical method. Furthermore, Cremades et al. obtained carotenoproteins and chitin from crawfish by a combined process that was based on flotation (sedimentation) and in situ lactic acid production [57]. The chitin isolated by this process was reported to be of high quality, comparable to that available commercially for medical and nutritional uses. In conclusion, isolation of chitin by biological methods, though not as efficient as the chemical method, resulted in the production of high-quality chitin, under environment-friendly conditions. The biological treatment of shell waste for chitin extraction may therefore be considered as an alternative to the chemical method.

11.3.2 Synthesis of chitosan Chitosan is mostly obtained by deacetylation of chitin using alkaline hydrolysis or an enzymatic method [30,57 68]. The use of certain bacteria and fungi has been reported to result in enzymatic deacetylation of chitin [61]. These deacetylases have been isolated from various types of fungi, namely Mucor rouxii, Aspergillus nidulans, and Colletotrichum lindemuthianium. The activity of these deacetylases is, however, reported to be severely limited by the insolubility of the chitin substrate [61]. According to Younes and Rinaudo, chemical methods are used extensively for the commercial synthesis of chitosan due to its low production cost and the ability to adapt it for mass production [3]. Furthermore, it was suggested that either acids or alkalis may be used to deacetylate chitin; however, the use of acid was reported to result in the destruction of the chain because of the susceptibility of the glycosidic bonds to acid, hence the alkali deacetylation process is often used to obtain chitosan from chitin [32]. Generally, the different alkaline hydrolysis methods may be broadly classified into two main categories: (1) the heterogeneous deacetylation of solid chitin and (2) the homogeneous deacetylation of preswollen chitin under reduced pressure in an aqueous medium [60]. In both

Handbook of Chitin and Chitosan

11.3 Synthesis of chitin and chitosan

345

processes the deacetylation reaction involves the use of concentrated alkali solutions and long processing times, which may vary from 1 to 80 h [60,61]. The preferred industrial method is however the heterogeneous deacetylation, which involves preferential reaction in the amorphous regions of the polymer while leaving, almost intact, the intractable crystalline native regions in the parent chitin [60]. Homogeneous deacetylation, on the other hand entails the hydrolysis of preswollen chitin using moderately concentrated alkali (13% w/w) at 25 C 40 C for 12 24 h [60]. The quality of chitosan obtained was found to be dependent on the source of chitin and the isolation process [67]. It has been reported that the deacetylation of chitin results in the degradation of the polymeric chain [68]. Harsh reaction conditions have also been established to damage the crystallinity of chitosan [69]. Consequently, the reaction conditions must be controlled when preparing chitosan [68,70]. The use of heterogeneous conditions with NaOH 75% (w/v) and a temperature of 110 C have been established to result in the deacetylation of chitin with minimal degradation [70]. The degree of deacetylation (DD) in both processes was found to be dependent on the concentration of the alkali, previous treatment, particle size, and density of chitin [68]. Several studies on chitosan synthesis with variation in processing parameters have been carried out and articulately documented [13,17,33,50]. Toan investigated the effects of varying alkaline concentration on the deacetylation of chitin obtained from shrimp shell using four different concentrations of sodium hydroxide, NaOH (30%, 40%, 50%, 60%) at 65 C with a solid to solvent ratio 1:10 (w/v) for 20 h [62]. He reported that although 60% NaOH treatment yielded the highest deacetylated chitosan with maximum solubility, 50% NaOH treatment could be used to get high-quality chitosan of 79.57% DD and 97.02% solubility [62]. Al-Sagheer et al. investigated the effects of deacetylation time on the Mw and crystallinity of synthesized chitosan [30]. They converted chitin to chitosan using two methods of deacetylation. The first was the standard deacetylation method, which involved treating chitin with 45% concentration of NaOH in 1 g:15 mL solid to solvent ratio at 110 C. The second method was the microwave method. This consisted of a mixture of chitin in 45% concentration of NaOH in a conical flask covered tightly with cotton under microwave radiation. They reported that the microwave technique was able to reduce deacetylation time from 8 h to approximately 15 min and that it resulted in the synthesis of chitosan with higher Mw and crystallinity [30]. Based on the aforementioned, several alternative processing methods have been developed to reduce processing times and to also minimalize quantities and concentration of alkali needed for deacetylation. Some documented processes include the use of successive alkali treatments using thiophenol in DMSO [63], a thermomechanical process using a

Handbook of Chitin and Chitosan

346

11. Chitin, chitosan, marine to market

cascade reactor operated under low alkali concentration [64], flash treatment under saturated steam [65], and the use of microwave dielectric heating [66]. The commercial production of chitin and chitosan have already commenced in some parts of the world, for example, the United States, Japan, Poland, India, Australia, and Norway. Global attention has also been focused on chitin and chitosan to tailor and impart the required functionalities in an attempt to maximize their effectiveness and expand their scope of applications [1,2,12 16,32,33,35,43 45,50,51,60].

11.3.3 Synthesis of derivatives of chitin and chitosan The reactivity of the primary amino group and the primary and secondary hydroxyl side groups result in the possibilities of chemical modifications and the formation of varieties of derivatives, with different structure and properties, which in turn result in versatile applications [71 74]. For example, the introduction of alkyl or carboxymethyl groups to the chitin or chitosan structure was reported to drastically increase the solubility of chitin and chitosan at neutral and alkaline pH, while substitution with carboxyl groups has been reported to yield polymers with polyampholytic properties [73,74]. Furthermore, derivatives of chitin and chitosan have shown promise for metal ions adsorption. They have also found diverse applications in drug delivery, tissue engineering, and wound healing. Chitin, chitosan, and their derivatives are widely researched as antimicrobial agents and as components in cosmetics and food [71 73]. According to Kim et al., derivatives of chitin may be classified into two categories. These authors reported that in each case the N-acetyl groups were removed while the exposed amino function reacts either with acyl chlorides, or anhydrides to give the group NHCOR, or is modified by reductive amination to NHCH2COOH [72]. According to the authors, derivatives of both types are formed by reaction with bi- or polyfunctional reagents that result in the formation of highly reactive derivatives with great potential [72]. Research attention has therefore been focused on the chemical modification of chitin and chitosan because of the great potential that is yet to be fully exploited. Some of the useful derivatives of chitin and chitosan are produced by cross-linking, graft copolymerization, complexation, chemical modifications, and blending [71 77]. Glycol chitin, a partially o-hydroxyethylated chitin, was the first derivative of chitin of practical importance synthesized [75]. Some of the recent works on chitin and chitosan derivatives are discussed below.

Handbook of Chitin and Chitosan

11.3 Synthesis of chitin and chitosan

347

Kurita [18] was among those who investigated the effects of chemical modifications of chitosan on its solubility. The author treated fully deacetylated chitosan with phthalic anhydride in dimethylformamide to give N-phthaloyl-chitosan. It was reported that the new derivative was readily soluble in polar organic solvents [18]. In the same vein Sashiwa et al. [76] successfully synthesized dendronized chitosan sialic acid hybrids using the convergent grafting of preassembled dendrons built on gallic acid and tri(ethylene glycol) backbone. The new derivatives were found to be soluble in water. It was also confirmed that the water solubility of the novel derivatives was further enhanced by N-succinylation of the remaining amine functionality [76]. Baba et al. [73] synthesized methylthiocarbamoyl and phenylthiocarbamoyl chitosan derivatives and examined their selectivity toward metal ions from aqueous ammonium nitrate solution. The authors reported that the adsorption of copper(II) on the synthesized derivative is dependent on the pH, in a low pH region. The authors suggested that the metals adsorption occurred by chelate formation that was accompanied by the release of hydrogen ions. They also noted that selective adsorption of copper(II) by the derivative is greater than that of iron(III), indicating that the new derivative will be an effective selective adsorbent of copper ions [73]. Similarly, Zhang et al. fabricated chitosan microparticles with 251.22 mg/g of adsorption capacity for Cd(II), higher than most of the counterparts, by chemical coprecipitation, spray drying, and Michael addition reaction, without any cross-linker participation [77]. The authors suggested that this might serve as a promising adsorbent for the recycling of contaminated water. The synthesis of chitosan hydrogels by direct grafting of D, L-lactic and/or glycolic acid onto chitosan in the absence of catalysts was carried out by Qu et al. [78]. They demonstrated that a stronger interaction existed between water and chitosan chains after grafting lactic and/or glycolic acid. According to the authors, the side chains aggregated to form physical cross-linking, which resulted in pH-sensitive chitosan hydrogels that are considered potentially useful for biomedical applications [78]. Recent work has also reported the potential of gadolinium neutron capture therapy by chitosan nanoparticles [79,80]. The authors demonstrated that the novel gadolinium-loaded nanoparticles are potentially suitable for intratumoral injection into solid tumor [80]. Research focused on chitin and its derivatives has therefore resulted in the production of polymers with extensive structural possibilities that culminated in novel properties and meet diverse functional requirements, thus expanding its scope of applications [1,2,4,6,12 16,18 20,72 117]. Several reports demonstrated that the development and applications of these functional derivatives facilitated

Handbook of Chitin and Chitosan

348

11. Chitin, chitosan, marine to market

the development of various products such as hydrogels [81], membranes [82,83], nanofibers [84 86], beads [87], micro/nanoparticles [79,80,88,91], scaffolds [81,89 93], and sponges [94,95] with broad applications. According to Kumar, the potential and traditional applications of chitin, chitosan, and their derivatives are estimated to be above 200 [19]. It can therefore be affirmed that chitin and its derivatives are indeed an expanded market in waiting.

11.4 Properties of chitin and chitosan Chitin contains a high content of GlcNAc units, hence it is found to be insoluble in water and most organic solvents [7]. However, when the degree of N-acetylation, which is defined as the average number of Nacetyl-D-glucosamine units per 100 monomers, is less than 50%, chitin becomes soluble in aqueous acidic solutions (pH , 6.0) and is called chitosan [113]. Chitosan therefore comprises a group of fully and partially deacetylated chitin [19]. The effects of degree of acetylation (DA) on the solubility, conformation, and dimensions of chitosan chains in aqueous media have been extensively studied [118 122]. The general laws of chitosan behavior in aqueous solutions were proposed by Schatz et al. [120]. According to these authors, chitosan exhibits the highest structural charge density and displays polyelectrolyte behavior related to long-distance intra- and intermolecular electrostatic interactions responsible for chain expansion, high solubility, and ionic condensation at DA below 20%. They also affirmed that hydrophilic and hydrophobic interactions are progressively counterbalanced when the values of DA are in the range of 20% 50%. DA above 50% was however found to result in electrostatic interactions that are essentially short-distance interactions, rather than hydrophobic interactions, due to the increase in the acetyl group content [120]. Similarly, Sandford [123] and Aranaz et al. [60] affirmed that chitosan with DA values from 0% to 30% has the optimum biological properties, hence is the most useful for biomedical applications. Currently, research attention is focused on chitosan and its derivatives because of their extensive array of properties that have made them versatile [1 3,8 21]. According to Aranaz et al. [60] most of the characteristic properties of chitosan are strictly related to its average Mw and the high content of glucosamine residues containing primary amino groups. The presence of the amino ( NH2) groups had been found to enhance the polymer’s reactivity and made it considerably more versatile than cellulose. It has also facilitated the production of functional derivatives of chitosan by chemical modifications, graft reactions, and

Handbook of Chitin and Chitosan

11.4 Properties of chitin and chitosan

349

ionic interactions [98,104,111]. This has further enhanced the properties of the polymer and expanded its range of application. This section discusses extensively some of the physiochemical and biological properties of chitin, chitosan, and its derivatives.

11.4.1 Physiochemical properties of chitin and chitosan Chitin is a polymorphic substance that occurs in three different crystalline structure, α-, β-, and γ-chitin [1,51]. The most abundant polymorph of the three is the α-chitin that occurs in the exoskeleton of crustaceans such as lobsters, crabs, and other marine-based invertebrate [3,108]. Fungal and yeast cell walls and arthropods are the other main sources of α-chitin while the β-chitin is found in association with proteins in squid pens [4,108]. These show that different polymorphic forms, with different physiochemical properties, are associated with different chitin sources. According to Khor [108] and Jang et al. [109], the differences between the three polymorphs depend on the arrangement of chains in the crystalline regions. In the α form, all chains exhibit an antiparallel orientation while in the β form the chains are arranged in a parallel manner, see Fig. 11.4A and B. In the γ form sets of two parallel strands alternate with single antiparallel strands (Fig. 11.4C). This shows that γ-chitin is composed of both α and β chitin [108]. Several authors, including Muzzarelli et al. [110], reported that the crystal structure of β-chitin lacks hydrogen bonds along the b axis [3,4,110]. The susceptibility of β-chitin to intracrystalline swelling, acid hydrolysis, and loss of scarcely crystalline fractions was ascribed to this lack of intrasheet hydrogen bonds [3,4,110]. β-Chitin is therefore more reactive and shows a higher affinity for solvents than α-chitin. Synowiecki and Al-Khateeb [111] reported that the DD may range from 30% to 95%, depending on the source and processing technique. The crystallinity of the semicrystalline polymer has been found to be dependent on the DD [114,115]. Rinaudo [12] affirmed that the origin of

FIGURE 11.4 γ-chitin [60].

Three polymorphic configurations of (A) α-chitin, (B) β-chitin, and (C)

Handbook of Chitin and Chitosan

350

11. Chitin, chitosan, marine to market

chitin influences not only its crystallinity and purity but also its polymer chain arrangement and consequently its properties. Consequently, the influence of the chitin source on the structure and subsequently properties of chitin and chitosan cannot be overemphasized. Due to these differences in properties of chitin polymorphs, chitin from different sources possesses different properties [3,109,110]. Knowledge of the source will therefore facilitate a good understanding of the crystalline structure, which is crucial to comprehending the structure, properties, and applicational relationships of chitin and its derivatives. Apart from the sources of chitin, the processes and conditions under which they are prepared affect the physiochemical properties of chitosan [12,14,17,18,30,46,60,111,112]. Extensive studies have shown that the Mw and subsequently DA are dependent on both processing methods and conditions [60,100,102,105 108]. Generally, commercial chitins are prepared by a chemical method. It was however noted that when deproteinization preceded demineralization a collapsed chitin, which results in the loss of its native structure was isolated while when demineralization occurred before deproteinization, compacted chitin in which the native chain and fibrous structures are intact and stabilized was obtained [96,97]. Furthermore, the use of harsh chemicals for the decolorization of chitin was also observed to result in distortion of the chitin structure [96,97]. Research focused on varying processing parameters to facilitate the production of chitin and subsequently chitosan with desirable properties has been extensively documented [46,60 76,102,105,107]. The physicochemical properties of chitin and chitosan have been found to be influenced by the variation in Mw [106]. The variation in the Mw of chitin/chitosan is extensive and was reported to range from several to more than thousands of kDa [102 104]. Based on this, the polymer was categorized into three categories. These are: 1. low-molecular-weight chitosan (LMWC); 2. medium-molecular-weight chitosan (MMWC); and 3. high-molecular-weight chitosan (HMWC). Several works have been carried out to investigate the effects of increasing Mw on the properties of chitin and chitosan [104 107]. Khan and Peh [107] reported that the density of chitosan and its solution increased with decreasing Mw. Similarly, solubility in water, permeation into cell nucleus, antioxidant activity, and biodegradability were reported to increase with decreasing Mw [60,100]. Conversely, the viscosity of chitosan solution and adsorption to fat droplets were reported to be enhanced by increasing Mw [60,107]. However, the effect of Mw on pH, surface tension, and conductivity of chitosan solutions was found to be unpredictable [107].

Handbook of Chitin and Chitosan

11.4 Properties of chitin and chitosan

351

Apart from the influence of averaged Mw on the physiochemical properties of chitin and chitosan, characteristic properties of chitosan were also observed to be related to the high content of glucosamine residues containing primary amino groups. Its cationic properties such as solubility, biodegradability, and absorption of its substrates have been ascribed to the amount of protonated amino groups in the polymer chain and thus on its DD [114 116]. The cationic nature of chitosan was also observed to facilitate electrostatic interactions with anionic glycosaminoglycan and proteoglycans [103]. The high density of positive charges of chitosan has been claimed to be responsible for the versatile nature of chitosan, considering that it facilitates the formation of different derivatives, which promotes the utilization of chitosan in diverse applications [107,114,116,117,124 127]. Moreover, the presence of the hydroxyls and amino groups, which act as electron donors, has been established to enhance chelating ability for many transitional metals [8,34]. Metal binding by chitosan takes place under acidic or near neutral pH. The amino groups of chitosan and hydroxyl groups are protonated and interact with metal anions by electrostatic attraction under acidic conditions and adsorption at pH close to neutral. The chelating process depends on the pH, ion type, composition of the solution, and the chitosan’s DD and Mw [116]. Jung and Zhao studied the chelating ability on ferrous ions of chitosan with a wide range of Mw and observed the highest values for 4 5 kDa chitosan, while 280 300 kDa chitosan did not exhibit this property.

11.4.2 Biological properties of chitosan In addition to the various physiochemical properties, chitin, chitosan, and their derivatives have been shown to possess diverse biological properties that are directly related to their physicochemical properties [3,60,101 108,116,125 127]. Indeed, the reactive amino and hydroxyl groups confer on chitosan important biological properties, such as antitumor, antimicrobial, antiinflammatory, bone regeneration, human hemostasis, antioxidant, hypocholesterolemic, antihypertensive, prebiotic, and ion binding activities. Most of these biological properties are dependent on the extent of acetylation, Mw of the chitosan, its polydispersity, crystallinity, and the distribution of GlcNAc and GlcN units along the polymeric chain [107,116,118 123,127 134]. Extensive studies investigating the effects of Mw on the antimicrobial properties of chitosan and its derivatives have been documented [3,101 103,125 128]. Sudarshan et al. [125] stated that low Mw chitosan could easily penetrate the cell wall of bacteria, interact with DNA, and

Handbook of Chitin and Chitosan

352

11. Chitin, chitosan, marine to market

as a result inhibit synthesis of mRNA and DNA transcription. It had also been shown that HMWC could interact with the cell surface and consequently alter cell permeability [126]. Lim and Hudson [101] ascribed the antimicrobial activity of chitosan to its polycationic nature that resulted in its interactions with predominantly anionic components with the subsequent change in permeability. These interactions have been associated with the death of cells by induced leakage of intracellular components [20,101,135]. Similarly, chitosan has been reported to adsorb electronegative substrate in the cell of microbe proteins, thereby disrupting the physiological activities of the microorganism, subsequently leading to the death of cells [102]. It has been demonstrated that the DD greatly influences the positive charge density of chitosan [103]. Higher DD was found to result in higher positive charge density when compared to that of moderate, or low DD. Subsequently, chitosan with higher DD may result in stronger antibacterial activity when compared to that of lower DD [103]. Similarly, the pH of the system has been found to influence its antimicrobial activities [3]. Younes and Rinaudo [3] reported that absorption of chitosan by bacteria occurs more at lower pH, as this results in an increase in chitosan’s positive ionic charge. The authors therefore affirmed that low DA, low Mw, and low pH offered higher antibacterial efficiency [3]. The effects of Mw on antioxidant activities have been extensively studied [129 132]. Anraku et al. [129] demonstrated that LMWC had higher antioxidant activity when compared to the HMWC. In the same vein, Chang et al. [130] established that chitosan with a low Mw significantly increased the antioxidant scavenging activity against 2,2-diphenyl-1-picrylhydrazyl (DPPH) radicals. Chien et al. [131] produced LMWC (12 kDa) that exhibited stronger scavenging activity toward DPPH radicals and higher ferrous ion chelating activity when compared to medium- and HMWC (95 and 318 kDa, respectively). Tomida et al. [132] studied the relationship between antioxidant properties and Mw of chitosan. The authors also affirmed that lower-molecularweight chitosan has impressive antioxidant properties, which was ascribed to the ability of the polymer to scavenge hydroxyl radicals and reduce cupric acid ions. Furthermore, Anraku et al. [134] investigated the effect of HMWC supplements in normal volunteers and reported that treatment with chitosan for 8 weeks produced a decrease in oxidized to reduced albumin ratio and an increase in total plasma antioxidant activity. These authors also reported that a significant decrease in total cholesterol levels and atherogenic index, along with increase in levels of high-density lipoprotein (HDL) were observed. With these findings they demonstrated the antioxidative potential of HMWC in the systemic circulation in human and suggested that

Handbook of Chitin and Chitosan

11.4 Properties of chitin and chitosan

353

HMWC significantly reduced the levels of prooxidants such as cholesterol and uremic toxins in the gastrointestinal tract, thus inhibiting the subsequent development of oxidative stress in the systemic circulation in humans [134]. Indeed, a vast amount of work has been documented on the study of the antifungal nature of chitin and chitosan [136 143]. Allan and Hadwiger [136] confirmed the ability of chitosan to induce the in vitro growth of a number of fungi, excluding Zygomycytes that has chitosan as a component of its cell walls. According to Chen et al. [137] direct applications of chitosan film to the colony-forming Rhodotorula rubra and Penicillium notatum inhibit the growth of the organisms. Jung et al. [138] studied the effects of chitosan graft copolymers on Candida albicans, Trichophyton rubrum, and Trichophyton violaceum. The authors observed that the number and type of grafted chains, as well as the pH, substantially influenced the activities of these fungi. In the same vein, extensive studies investigating the antifungal activity of chitosan derivatives were articulately documented by Rabea et al. [139]. They carried out a radial hyphal growth bioassay of B. cinerea and P. grisea to assess the fungicidal activity of 24 new derivatives of chitosan (i.e., N-alkyl, N-benzylchitosans) and reported that all the derivatives are better fungicides than native chitosan. These authors further established that the N-dodecylchitosan, N-(p-isopropylbenzyl) chitosan, and N-(2,6-dichlorobenzyl) chitosan were the most active against B. cinerea while the N-(m-nitrobenzyl) chitosan was the most against P. grisea. A similar work carried out by Zhong et al. [140] using 12 types of new hydroxylbenzenesulfonilanide derivatives of chitosan, carboxymethyl chitosan, and chitosan sulfate reported that all the derivatives displayed stronger antifungal properties than the original materials. Nutritional studies on chitosan show that it is able to reduce serum cholesterol, have excellent lipid-binding properties and exercise a hypocholesterolemic effect on animals [141 143]. Chitosan is not digested in the gastrointestinal tract by the digestive enzymes, hence it acts as a dietary fiber in humans and animals [141,142]. According to Lian et al. [143] LMWC is a good dietary fiber that acts as a fat blocker though it increases the intestinal excretion of essential fatty acids and decreases absorption of fat-soluble vitamins and minerals. HMWC (150 600 kDa) was however considered a more effective dietary fiber with none of the side effects associated with LMWC [143]. Some other biological properties that facilitate its versatile applications include biocompatibility, biodegradability, immunogenicity, wound healing activity, low-toxicity, bioactivity, and its mucoadhesive properties [33,114,116,144,145]. The positive surface charge of this biomaterial and its biocompatibility enable it to effectively support cell growth, while the hydrophilic surface facilitates adhesion, proliferation,

Handbook of Chitin and Chitosan

354

11. Chitin, chitosan, marine to market

and differentiation of cells [145]. According to Kumirska et al. [116] the properties of chitin/chitosan are interrelated and many times the polymer could elicit conflicting biological response such as obtained when wound healing properties and permeation enhancement properties are inevitably linked to others such as mucoadhesion and cytotoxicity [116]. The authors suggested that research efforts toward investigating these conflicting biological responses be intensified to facilitate the production of safe and efficient chitin and chitosan-based biomedical products [116].

11.5 Potential applications of chitin and chitosan Chitin and its deacetylated form chitosan are established marine biopolymers that have found diverse industrial and biomedical applications [12 21]. The potential applications of chitin and chitosan depend on their physicochemical and biological properties [116]. These outstanding properties have resulted in the versatile applications of the biopolymer in diverse areas such as in pharmaceutics, gene delivery, tissue engineering, ophthalmology, cosmetics, water treatment, textile industry, and paper industry [1 3,12 16,19,32,51,75,85,89 93,102]. This section discussed some of the diverse applications of chitin/chitosan and their derivatives and also identifies opportunities to develop valueadded products from this marine-based biopolymer.

11.5.1 Biomedical application of chitosan The marine biopolymer and its derivatives have attracted considerable research interest for biomedical applications based on their outstanding biological attributes, such as biocompatibility, biodegradability, biosafety, and nontoxicity. Chitin/chitosan has also been found to be hemostatic, fungistatic, bacteriostatic, spermicidal, anticholestermic, and anticarcinogenic, which had further expanded its range of biomedical applications [12,13,19,33,114,116,144 180]. Additionally, the polycationic surface of the biopolymer has resulted in its capability to form inter- and intramolecular hydrogen bonding, which has also facilitated its use in the development of novel biomedical products [3,12,13,19,99]. 11.5.1.1 Wound dressing/wound healing Wound dressings are used to prevent the infection of wounded skin. Purified chitin-based wound dressings have been found to prevent bacteria infiltration, enhance dermal regeneration, and accelerate wound healing [87,107]. Other attributes such as good strength,

Handbook of Chitin and Chitosan

11.5 Potential applications of chitin and chitosan

355

flexibility, and bioabsorbable properties facilitated fibers made of chitin/chitosan to be utilized as absorbable sutures and wound dressing materials [117,148,149]. Unlike other absorbable sutures chitin/chitosan-based sutures have been found to resist attack in bile, urine, and pancreatic juice [148]. These chitin/chitosan-based materials have also been reported to accelerate wound healing when incorporated in spray, gel, and gauze [107,150,151]. Le et al. confirmed that wound dressings made of chitin and chitosan fibers accelerate the healing of wounds by about 75% [148]. Moreover, chitin/chitosan has been used as a coating on normal biomedical materials to enhance the wound healing process. Sutures made from materials such as standard silk and catgut when coated with regenerated chitin/chitosan were reported to show enhanced wound healing activities [148]. 11.5.1.2 Burn treatment/artificial skin graft Artificial skin graft has been identified as another likely application of chitosan in the medical and cosmetic industries [2,8,107]. According to Dutta et al. [2] chitin/chitosan is a promising candidate for burn treatment. The authors ascribed this to the fact that chitosan can form tough, water-absorbent, biocompatible films directly on the burn. They noted that the use of chitosan for burn treatment allows excellent oxygen permeability, which is important to prevent oxygendeprivation of injured tissues. They further stated that chitosan films have the ability to absorb water and are naturally degraded by body enzymes, subsequently there is no need for removal of the wound dressing [2]. The use of a nonantigenic membrane that may serve as a biodegradable template for the synthesis of neodermal tissue has been the focus of research in the design of artificial skin applicable for long-term chronic use [2,178]. Due to similar structural characteristics to glycosamino glycans, the suitability of chitosan for the development of this substratum for skin replacement, such as may be required in brain-scalp damage, plastic skin surgery, etc., has been investigated [178,179]. 11.5.1.3 Tissue engineering Tissue engineering is another thriving area of biomedical application of chitin/chitosan and their derivatives. The development and manipulation of laboratory-grown cells, tissues, or organs that would replace or support the function of defective or injured parts of the body would facilitate the revolution of current technology in the repair, support, or replacement of organs or tissues that have lost their function [2]. The current methods used include the use of transplants, implants, or surgical reconstruction, all of which are fraught with diverse limitations. Most

Handbook of Chitin and Chitosan

356

11. Chitin, chitosan, marine to market

implant materials, although biocompatible, may lack the ability to meet the long-term mechanical, geometrical, and functional requirements of the body, hence the development of artificial tissues that can mimic the natural ones by combining with modulated cells with different types of scaffolding materials offers a better option [2,173]. The use of chitin/chitosan as scaffolds has been reported to have an acceleratory effect on the tissue engineering processes owing to their polycationic nature that enhances the adsorption of cells on the polymer. Consequently, research efforts have been focused on the use of chitin/chitosan as scaffolding material in tissue engineering. It has been demonstrated that chitosan can be easily processed into porous scaffolds, films and beads [174], while Kast et al. [175] affirmed that chitosan thioglycolic acid (chitosan-TGA) conjugate is a promising candidate as a scaffolding material in tissue engineering. Similarly, Zhang and Zhang [176] synthesized and characterized microporous chitosan/calcium phosphate composite scaffolds for tissue engineering application. The authors showed that the chitosan was able to provide a scaffold form while the bioactivity of the calcium phosphates encouraged osteoblast attachment and strengthened the scaffold, resulting in the development of stronger, bioactive and biodegradable scaffold [176]. In the same vein, Madihally and Matthew attempted to develop procedures for synthesizing many porous chitosan scaffolds [177]. They were able to prepare bulk, planar, and tubular scaffolds that could be used to develop different types of engineered tissue that may be applied in tissue engineering of nerves and blood vessels as a template for cells. Furthermore, Prasitsilp et al. [178] investigated the effects of DD of chitosan on its in vitro cellular responses and reported that cells are more readily attached to more highly deacetylated chitosan. 11.5.1.4 Drug delivery Drug delivery is another important area in which chitin/chitosan is finding a broad range of applications. Development of drug delivery systems such as nanoparticles, hydrogels, microspheres, films, and tablets using this important marine biopolymer has been extensively researched and documented [106,125,126,146,147,156 159]. Its pharmaceutical applications include nasal, ocular, oral, parenteral, and transdermal drug delivery. Chitosan has also been shown to be a suitable material for efficient nonviral gene therapy [156,160]. In addition, chitin/chitosan has been found to inhibit tumor cell growth [161 166] and studies on targeted delivery of drugs to tumor tissues have demonstrated their potential for applications in cancer therapy and diagnosis [167,168].

Handbook of Chitin and Chitosan

11.5 Potential applications of chitin and chitosan

357

11.5.1.5 Ophthalmology Characteristics required of an ideal contact lens, such optical clarity, mechanical stability, sufficient optical correction, gas permeability, wettability, and immunological compatibility, are possessed by chitin/ chitosan, thus making it an ideal candidate for ophthalmological applications [2,169,180]. Contact lenses made from partially depolymerized and purified squid pen chitosan that are clear, tough, and possess other required properties, such as modulus, tensile strength, elongation, and oxygen permeability have been reported [2]. Chitin/chitosan’s antimicrobial and wound healing properties, coupled with excellent film-forming capability were also claimed to be responsible for the suitability of the marine biopolymer for the development of ocular bandage lens [169,180].

11.5.2 Industrial applications of chitosan Chitin and chitosan possess very interesting physiochemical and biological properties which facilitate their utilization in the manufacturing of a vast array of widely different products. Some of the diverse array of industrial applications of chitin/chitosan and their derivatives include water engineering, biotechnology, food processing, cosmetics industries, textile industry, paper industry, agriculture, photography, and electronics [1 3,12 16,21,27 30]. Different arrays of properties are required by different industrial applications, most of which are met by the diverse array of properties of this important marine-based biopolymer [2]. This section gives an overview of some of the important industrial applications of chitin/chitosan and their derivatives. 11.5.2.1 Water engineering Water pollution caused by chemical contamination of water from a wide range of toxic products, such as metals, aromatic molecules, and dyes, constitutes a serious environmental problem [60]. Research towards proffering solutions to this hazard is therefore critical to forestall the potential toxicity that may result from this pollution. The use of chitin and chitosan to remove water pollutants has been extensively researched and documented [20,60,181 190]. Due to their polycationic nature, chitin/chitosan can be used as flocculating agents, they can also act as a chelating agent and a heavy metals trapper [2]. Guibal et al. [183,184] examined the effect of chitosan properties on the adsorption of metals, dyes, and organic compounds. Similarly, the use of chitin/chitosan-based material for the removal of anionic dyes was extensively reviewed and documented by Crini and Badot [181]. The coagulation flocculation process and the adsorption process were

Handbook of Chitin and Chitosan

358

11. Chitin, chitosan, marine to market

reported to depend on the DD. Saha et al. affirmed that chitosan with higher DD has a higher efficiency for adsorption of an azo dye [190]. When compared to chitin, chitosan was found to be more efficient in the removal of metal ions [183,187], polychlorinated biphenyls (PCBs) [188], and anionic dyes [182]. Conversely, chitin was found to be more efficient than chitosan in the removal of polycyclic aromatic hydrocarbons from petrochemical wastewater [189]. Chitosan has been used as adsorbent, coagulant, and bactericide in the treatment of aquaculture wastewater [191 193]. Weltroswki et al. [192] used chitosan N-benzyl sulfonate derivatives as sorbents for the removal of metal ions in an acidic medium. Considerable amounts of world production of chitin and chitosan and derivatives were reported to be used in wastewater treatment [194]. 11.5.2.2 Agriculture Threat posed by antimicrobial-resistant pathogenic organisms to human and animal health led to the need to investigate the development of natural products that exhibit unique, superior, cost-effective, and safe microbicides [169]. It has been articulately reported that chitin/chitosan possess antimicrobial activity against a number of Gramnegative and Gram-positive bacteria [72,125,152,170]. Cuero [154] demonstrated that chitin and chitosan from crustacean sources exhibited antifungal activity against a large number of human pathogenic fungi. The author however noted that the biopolymer acts more quickly on fungi than on bacteria. Furthermore, chitosan was reported to have a higher antifungal activity than chitin, although it is found to be less effective against fungi that have chitin or chitosan components in their cell walls [136]. These natural antibacterial and antifungal characteristics have been widely explored in the development of commercial disinfectant products [135 140,152,153] and has also facilitated its agrochemical applications [155]. Extensive work has been carried out on the use of chitosan coating as a protective barrier to extend the shelf life of many fruits and vegetables [195 200]. Chien et al. [198] demonstrated that coating with low Mw chitosan retarded ripening, water loss, and decay of sliced red pitayas. The authors affirm that the coated samples exhibited greater antifungal resistance than thiabendazole [198]. In addition to this the utilization of chitin/chitosan as foliage spray to induce disease resistance and increase quality and production of plants has been reported [2,12,60]. The successful applications of the marine biopolymer in crops such as rice, palm, corn, cassava, and many other tropical fruits to inhibit infection has been reported [2,60]. Similarly, several works have been carried out on the application of chitosan to induce resistance to disease in animals and for the

Handbook of Chitin and Chitosan

11.5 Potential applications of chitin and chitosan

359

enhancement of the shelf life of animal products [2,12,60]. Jeon et al. [199] compared the preservative efficacy of different viscosity chitosan in coated herring and Atlantic cod and demonstrated the potential of chitosan as a preservative coating in reducing or preventing moisture loss, lipid oxidation, and microbial growth for the fish. It was reported that the higher viscosity (360 cP) chitosan exerted a better preservative effect in both fish model systems [199]. In a similar study, pr-soaking of fish fillets with a high DD chitosan solution was found to extend shelf life from 5 to 9 days [200]. Investigation into enhancing the shelf life of eggs has shown that chitosan offers a protective barrier against the transfer of carbon dioxide and moisture through the eggshell [201]. Kim and No [201] also reported that this barrier was effective in keeping a high Haugh unit and yolk index because it was able to prevent diffusion of water from the albumen. Furthermore, chitin/chitosan has been incorporated into animal feed for fish and shrimps, as feed coating, and also used as a supplement in the drinking water of poultry, cattle, and porcine to prevent infection [2,12,60]. The use of chitosan as a fertilizer and pesticide has been documented [202]. Hadwiger reported that when absorbed by a plant, chitosan activates and improves its natural defense mechanisms by influencing the biochemistry of plant cells. He further demonstrated that the application of this chitosan-based solution enhances seed germination, plant growth, and yields. It may therefore be concluded that the applications of chitin/chitosan in agriculture are versatile and crucial to meet the food demands of the ever-increasing population. 11.5.2.3 Food processing Due to its outstanding attributes chitin/chitosan and their derivatives have found a wide range of unique applications in the food industry [20]. According to Aranaz et al. [60] microcrystalline chitin (MCC), which shows good emulsifying properties, superior thickening, and gelling activity, has found application in stabilizing food [60]. Due to the antimicrobial action against food spoilage microorganisms and antioxidant properties, the use of chitin/chitosan in the protection of food from microbial deterioration has been articulately documented [60,195 200]. Edible, semipermeable chitosan films and coatings have been reported to not only retard ripening, water loss, and reduce decay [195 200], but to also create a controlled atmosphere, such as used in storage, at a lower cost. This also results in the preserved food’s flavor and color being maintained and not altered by the preservatives as is usually the case with other edible coatings [2,60]. It has also been reported that chitin/chitosan exhibits anticholesterolemic properties, that is, cholesterol-binding capacity, which facilitate application as dietary agents in multiple nutritional supplement products [2,60,203 205].

Handbook of Chitin and Chitosan

360

11. Chitin, chitosan, marine to market

Dutta et al. [2] suggested that chitin/chitosan can be used as a nonabsorbable carrier for highly concentrated food ingredients such as food dyes and nutrients. 11.5.2.4 Cosmetics industries Chitin/chitosan and their derivatives have found use in three main areas of cosmetics: hair care, skin care, and oral care [2]. Owing to the fact that chitin/chitosan and hair carry opposite electrical charges—chitosan positive and hair negative—they readily complement each other. According to Dutta et al. [2] a clear solution that contains chitosan forms a clear, elastic film on hair, that enhance the softness, smoothness, and mechanical strength of hair. Therefore, chitin/chitosan are found in hair products such as shampoos, rinses, permanent wave agents, hair colorants, styling lotions, hair sprays, and hair tonics [2]. The physiochemical and biological attributes of chitosan and its derivatives facilitate its use in the development of skin care products. The Mw of most chitosan products is so high that they cannot penetrate the skin, hence chitosan has found use in the production of skin moisturizer and will be a costeffective replacement for hyaluronic acid that has been hitherto used as skin moisturizer [2,206,207]. Consequently, chitin/chitosan and their derivatives are found in cosmetics products such as creams, lotions, antiaging cosmetics, nail enamel, nail lacquers, foundation, eye shadow, lipstick, and cleansing materials [2,206,207]. The antimicrobial and antifungal attributes of chitin/chitosan and their derivatives facilitate application in the production of oral care products, such as toothpaste, mouthwashes, and chewing gum; their components in the products result in fresh breath and prevent the formation of plaque and tooth decay [2]. 11.5.2.5 Photography Chitin/chitosan has found important applications in photography due to the resistance to abrasion, optical characteristics, and filmforming ability [8]. According to Muzzarelli chitosan can easily release silver complexes, used in photography, so that it can easily penetrate from one film layer to another by the diffusion transfer reversal process [8]. Dutta et al. [2] reported the use of chitosan as a fixing agent for the acid dyes in gelatin and that it also acts as an aid to improve diffusion in color photography. 11.5.2.6 Chitosan gel for light-emitting device The use of dyes containing chitosan gels as a potential component in lasers and other light-emitting devices (LEDs) has been reported [1,2]. According to Dutta et al. [2] the doping process utilizes dyes such as porphyrin compounds that resemble the heme groups in blood.

Handbook of Chitin and Chitosan

11.6 Economic potential of chitin and chitosan

361

According to the authors, research on porphyrins and other dyes, such as fluorinated coumarin and rhodamine B for transparent thin films are ongoing [2]. 11.5.2.7 Textile industry Chitin/chitosan has also found application in the textile industry [2]. Incorporation of chitin/chitosan fibers into both woven and nonwoven fabric has been reported to result in fabrics that can control odor and prevent microbial growth [2]. According to Kulkarni et al. [208] “The fusion of conventional structural textile materials with advanced properties given by ‘smart’ functional finishing technology offers a wide range of high added-value product options.” Consequently, the authors were able to devise an innovative strategy for functional finishing of textile materials that is based on the incorporation of a thin layer of surface-modifying systems (SMS), in the form of stimuli-sensitive microgels or hydrogels, to produce textiles with smart functional finishing that is responsive to both temperature and pH [208]. Campos et al. [209] investigated the effect of pretreatment of cotton with chitosan in natural dyeing. The authors reported that fabrics pretreated with a natural mordant have better color strength than fabrics which have not been pretreated. Thus it can be concluded that chitin/chitosan and their derivatives are vital materials that are required in all the phases of production in the textile industry. 11.5.2.8 Paper industry Cellulose constitutes the main raw materials for paper manufacturing. Due to the structural similarities between cellulose and chitosan, it is currently being used in the manufacturing of paper in place of cellulose [1,2]. According to Dutta et al. [1] chitosan utilization in paper making also saves chemical additives, increases output and results in the production of paper with better quality [1]. The use of the marine-based biopolymer in paper recycling results in the production of strengthened recycled paper, while at the same time reducing environmental degradation. Chitosan has therefore found application in the paper industry in the manufacturing of products such as toilet paper, wrapping paper, and cardboard used for packaging.

11.6 Economic potential of chitin and chitosan The seafood processing industry produces hundreds of thousands of tonnes of shell waste that are discarded on a regular basis. Thailand is one of the major producers and exporters of shrimp and other marine products worldwide that deals with over 600,000 metric tonnes of

Handbook of Chitin and Chitosan

362

11. Chitin, chitosan, marine to market

marine products containing chitinous materials, such as shrimp, crab, and squid, yearly [128,210]. Production of this vast amount of seafood will result in enormous waste generation. Disposal of this waste is not only costly but may result in serious environmental threats as a result of toxins seeping into the environment. Conversion of this waste to wealth, that is, the production of useful products such as chitin, chitosan, and other derivatives, which will result in value additions. Furthermore, the versatile nature of chitin/chitosan and their derivatives has resulted in the expanded scope of applications of the polymer. Consequently, processing of this seafood waste will culminate in socioeconomic advancement. In addition, the environmental challenges related to the use of petroleum-based products, such as synthetic plastics, has resulted in an intensive research drive toward the use of materials derived from renewable natural resources, such as chitin and chitosan, that are biodegradable. This will enhance the maintenance of global environmental sustainability by reducing the reliance on fossil fuels. The sustainable and cost-effective production of the hitherto petroleum-based products from chitin will also improve the economy. According to Hayes, the first patent on chitosan production was introduced in the 1920s and today there are several hundred patents on the production of chitin and its derivatives along with their applications [211]. Large-scale production of the biopolymer was confirmed to have started approximately two decades ago but its growth was hindered due to several factors, such as the unavailability of a large and reliable supply of raw material, inconsistent quality, the presence of pollutants such as heavy metals, ash, and other foreign materials, and finally the high production cost [211]. Presently, marine-based sources of raw material for chitin and chitosan production, that is, by-products of the seafood processing industry, are readily available. These include marine zooplankton, the exoskeleton of crustaceans such as crab shell, lobster shell, shrimps, and prawns. [3,27 30,210]. Other untapped sources of chitin are fungal mycelia [69]. The seafood industries however constitute the main sources of raw materials for the commercial production of chitin/chitosan, and their derivatives [19,28,30]. Since these wastes are relatively inexpensive, the production of chitosan on a large scale from this renewable bioresource is reported to be economically feasible [128]. The extraction of chitin from crustacean shells has been commercialized in the last couple of decades in different parts of the world. These include Europe, Latin America, North America, Asia, Middle East, and Africa [128,210]. An improved production process has also facilitated the production of high-purity chitin and chitosan and consequently expanding the scope of biomedical applications [1 3].

Handbook of Chitin and Chitosan

11.6 Economic potential of chitin and chitosan

363

Recent advances in chitin and chitosan research have dramatically increased their applications in various fields and spurred the development of new products, with more potential applications either in the incubation phase, or yet to be discovered [1 3,108,124,128]. The applications of chitin/chitosan in agriculture, water treatment, textile industry, paper industry, cosmetics, and biomedicine have been articulately documented, some of which have been extensively discussed in the previous sections [1 3,108,124,128]. The emerging applications of chitin and chitosan have resulted in an increasing demand for chitin and its derivatives by these end-use industries. Furthermore, with considerable research efforts being directed toward developing safe and efficient chitin/chitosan-based products, the expansion of their scope of applications and subsequently demand will be increased. Industries such as medicine, water treatment plant, food industry, beverage industry, pharmaceutical companies, bioplastics, cosmetics industry, textile industry, paper manufacturing, and agriculture constitute some of the end users that have also fueled the market globally. This has made the global market of chitin, chitosan, and their derivatives extremely attractive. Based on the Global Industry Analysts, Inc. report, the global demand for chitin in 2015 was above 60,000 tonnes, while its global production was around 28,000 tonnes [212]. As at 2015 the global chitosan market was reported to be US$1205 million [212]. In 2016 the global chitin and chitosan market was reported to be US$2 billion and was forecasted to reach US$4.2 billion by 2021, at a compound annual growth rate (CAGR) of 15.4%. The report estimated that the global market for chitin derivatives, including chitosan, should reach US$63 billion by 2024 [212]. The forecasted increase in CAGR within this period is ascribed to its growing applications in various end-use industries. The Asia-Pacific region was reported to hold the highest share of the global market, that is, a two-thirds share, while the European market is less developed [212]. Of the different grades of chitin/chitosan, the industrial grade constitutes the most widely used grade. Some of these diverse industrial applications include water treatment, food and beverage processing, agrochemicals, cosmetics industry, biomedical, and pharmaceuticals industry. The increasing demand for chitosan in these diverse areas has resulted in the growth of the industrial grade segment of the chitosan market and may be responsible for the estimated increase in CAGR. Of the industrial segment, the water treatment segment is projected to have the largest share of the chitosan market. This was attributed to the ongoing rapid industrialization, the increasing inflow of wastewater from various industries, and the increase in the scarcity of freshwater, which has led to an increase in the global demand for treated recycled water [213 215].

Handbook of Chitin and Chitosan

364

11. Chitin, chitosan, marine to market

The Asia-Pacific region has been projected to further fuel the growth of the industrial grade chitosan market due to the ongoing rapid industrialization in China and India [215]. According to Prasad, China’s chitosan market is projected to grow at the highest CAGR, during the forecast period, followed by Vietnam and India [204]. Prasad stated that countries such as Thailand, Indonesia, and Malaysia, that have emerged as manufacturing hubs for different industries, might also fuel the growth of the Asia-Pacific chitosan market [215]. Meanwhile, due to the strong pharmaceutical sector in North America, the region is projected to witness gains close to 18% in the coming years [213]. These extremely attractive global chitosan markets may further be enhanced by extensive research and development initiatives to ascertain other future novel applications of the marine biopolymer [213]. According to Pulidindi and Pandey, chitosan market size for the cosmetics and toiletries segment is projected to witness gains closer to 18.5% in the coming years [213]. This was attributed the use of chitin/ chitosan and their derivatives in the manufacturing of antiaging creams, shampoos, styling gels, hair sprays, and hair colorants. Due to the antiplaque and antidecay properties, they have also found application in the manufacture of toothpastes, mouthwashes, and chewing gums, which will further expand the chitosan market size for the cosmetics segment [213].

11.7 Conclusions Indeed, the journey of chitin/chitosan from the marine environment to the end-user industry and finally the consumer can be described as a viable and highly lucrative emerging market.

References [1] P.K. Dutta, M.N.V. Ravikumar, J. Dutta, Chitin and chitosan for versatile applications, JMS Polym. Rev. C42 (2002) (2002) 307. [2] P.K. Dutta, J. Dutta, V.S. Tripathi, Chitin and chitosan: chemistry, properties and applications, J. Sci. Ind. Res. 63 (2004) 20 31. [3] S. Younes, M. Rinaudo, Chitin and chitosan preparation from marine sources. Structure, properties and applications, Mar. Drugs 13 (2015) 1133 1174. [4] S.S. Kim, Y.M. Lee, C.S. Cho, Synthesis and properties of semi-interpenetrating polymer networks composed of β-chitin and poly(ethylene glycol) macromer, Polymer 36 (1995) 4497. [5] D. Wanule, J. Balkhande, P. Ratnakar, A.N. Kulkarni, C.S. Bhowate, Extraction and FTIR analysis of chitosan from American cockroach, periplaneta, Americana, Int. J. Eng. Sci. Innov. Technol. (IJEIT) 3 (2014) 299 304. [6] M.N.V. Ravikumar, Chitin and chitosan fibres: a review, Bull. Mater. Sci., Ind. Acad. Sci 22 (5) (1999) 905 915.

Handbook of Chitin and Chitosan

References

365

[7] K. Kurita, Controlled functionalization of the polysaccharide chitin, Progr. Polym. Sci. 26 (2001) 1921 1971. [8] R.A.A. Muzzarelli, Some modified chitosans and their niche applications, chitin handbook, in: R.A.A. Muzzarelli, M.G. Peter (Eds.), Chitin Handbook, European Chitin Society, Pergamon Press, Oxford, 1997, pp. 47 52. [9] K.D. Yao, F. Zhao, F. Li, Y.J. Yin, in: M. Schwartz (Ed.), Chitosan-based Gels, Encyclopedia of Smart Materials, vol. 1, John Wiley, New York, 2002, pp. 182 190. [10] D.V. Luyen, V. Rossbach, Mixed esters of chitin, J. Appl. Polym. Sci. 55 (1995) 679. [11] N. Kubota, Permeability properties of chitosan-transition metal complex membranes, J. Appl. Polym. Sci. 64 (1997) 819. [12] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (603) (2006) 32. [13] M. Rinaudo, Physical properties of chitosan and derivatives in sol and gel states, in: B. Sarmento, J. das Neves (Eds.), in: Chitosan-based Systems for Biopharmaceuticals: Delivery, Targeting and Polymer Therapeutics, John Wiley & Sons, Chichester, 2012, pp. 23 44. [14] M. Rinaudo, Main properties and current applications of some polysaccharides as biomaterials, Polym. Int. 57 (2008) 397 430. [15] M. Rinaudo, Materials based on chitin and chitosan, in: S. Kabasci (Ed.), in: Biobased Plastics: Materials and Applications, John Wiley and Sons, Chichester, 2014, pp. 63 80. [16] M.D. Teli, J. Sheikh, Extraction of chitosan from shrimp shells waste and application in antibacterial finishing of bamboo rayon, Int. J. Biol. Macromol. 50 (2012) 1195 1200. [17] M.M. Islam, S.M. Masum, M.A.I. Molla, M.M. Rahman, A.A. Shaikh, S.K. Roy, Preparation of chitosan from shrimp shell and investigation of its properties, Int. J. Basic. Appl. Sci. 11 (1) (2011) 116 130. [18] K. Kurita, Chemistry and application of chitin and chitosan, Polym. Degrad. Stab. (59)(1998) 117. [19] M.N.V.R. Kumar, A review of chitin and chitosan applications, React. Funct. Polym. 46 (1) (2000) 1 27. [20] F. Shahidi, R. Abuzaytoun, Chitin, chitosan, and co-products: chemistry, production, applications, and health effects, Adv. Food Nutr. Res. (49)(2005) 93 135. [21] R.N. Tharanathan, F.S. Kittur, Chitin-the undisputed biomolecule of great potential, Crit. Rev. Food Sci. Nutr. (43)(2003) 61 87. [22] H. Merzendorfer, L. Zimoch, Chitin metabolism in insects: structure, function and regulation of chitin synthases and chitinases, J. Exp. Biol 206 (Pt 24) (2003) 4393 4412 [PubMed: 14610026].. [23] P. Madhavan, K.G.R. Nair, Proceedings of the First International Conference on Chitin and Chitosan, MIT, Cambridge, 1978, pp. 88 102. [24] F. Nessa, Md Shah, S.M. Masum, M. Asaduzzaman, S.K. Roy, M.M. Hossain, et al., A process for the preparation of chitin and chitosan from prawn shell waste, Bangladesh J. Sci. Ind. Res. 45 (4) (2010) 323 330. [25] I.E. Akinwole, G. Alebiowu, G.M. Oyatogun, D.V. Abere, K.M. Oluwasegun, A.O. Oyatogun, et al., Synthesis and characterization of cowry and crab shells based chitosan for drug delivery, Bioceram. Dev. Appl. (8)(2018) 107. Available from: https:// doi.org/10.4172/2090-5025.1000107. [26] K.J. Akinluwade, G.M. Oyatogun, G. Alebiowu, A.P.I. Popoola, Morphological analysis of chitosan-acacia gum nanoparticles loaded and unloaded with cisplatin, Eur. J. Pharm. Med. Res. 5 (6) (2018) 162 166. [27] N.A. Ashford, D. Hattis, A.E. Murray, Industrial prospects for chitin and protein from shellfish wastes, MIT Sea Grant Report MISG, MIT, Cambridge, MA, 1977. 77-3.

Handbook of Chitin and Chitosan

366

11. Chitin, chitosan, marine to market

[28] H.M. Ibrahim, M.F. Salama, H.A. El-Banna, Shrimp’s waste: chemical composition nutritional value and utilization, Nahrung (43)(1991) 418 423. [29] S. Subasinghe, The Development of Crustacean and Mollusk Industries, vol. 27, Ampnag Press, Malaysia, 1995. [30] F.A. Al-Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from marine sources in Arabian Gulf, Carbohydr. Polym. 77 (1) (2009) 410 419. [31] R.A.A. Muzzarelli, R. Jeniaux, G.W. Gooday, Chitin in Nature and Technology, Plenum Publishing Corporation, New York, 1986. [32] I. Aranaz, D. Elieh-Ali-Komi, M.R. Hamblin, Chitin and chitosan: production and application of versatile biomedical nanomaterials, Int. J. Adv. Res. (Indore) 4 (3) (2016) 411 427. [33] D. Zvezdova, Synthesis and characterization of chitosan from marine sources in black sea, Scientific Papers of the University of Russia, (49) 9, 2010, 65 69. [34] R.A.A. Muzzarelli, F. Tanfani, M. Emanuelli, S. Gentile, The chelation of cupric ions by chitosan membranes [Callinectes sapidus, blue crab shell], J. Appl. Biochem. (2) (1980) 380 389. [35] S.A. Anicuta, L. Dobre, M. Stroescu, L. Jipa, Fourier transformer infrared (FTIR) spectroscopy for characterization of antimicrobal films containing chitosan, Analele ´ Universit˘aNii din Oradea Fascicula: Ecotoxicologie, Zootehnie s¸ i Tehnologii de Industrie Alimentar˘a (2010) 1234 1240. [36] N. Acosta, C. Jimenez, V. Borau, A. Heras, Extraction and characterization of chitin from crustaceans, Biomass Bioenergy. (5)(1995) 145 153. [37] E.L. Johnson, Q.P. Peniston, Utilization of shellfish waste for chitin and chitosan production, in: R.E. Martin, G.J. Flick, C.E. Hebard, D.R. Ward (Eds.), Chemistry and Biochemistry of Marine Food Products, AVI Publishing Co, Westport, CT, 1982, p. 415. Chapter 19. [38] F. Shahidi, J. Synowiecki, Isolation and characterization of nutrients and value-added products from snow crab (Chifroeceles opilio) and shrimp (Panda- 111sb orealis) processing discards, J. Agric. Food Chem. (39)(1991) 1527 1532. [39] R.H. Hackman, Studies on chitin. I. Enzymatic degradation of chitin and chitin esters, Aust. J. Biol. Sci. (7)(1954) 168 178. [40] R.H. Hakman, M. Goldberg, Light-scattering and infrared-spectrophotometric studies of chitin and chitin derivatives, Carbohydr. Res. (38)(1974) 35 45. [41] G.G. Anderson, N. de Pablo, C. Romo, Antartic krill (Euphausia superba) as a source of chitin and chitosan, in: R.A.A. Muzzarelli, E.R. Priser (Eds.), Proceedings of First International Conference on Chitin and Chitosan, 1978, Mit Sea Grant Program, Cambridge, MA, 1978, pp. 54 63. [42] S.T. Horowitz, S. Roseman, H.J. Blumental, Preparation of glucosamine oligosaccharides. 1. Separation, J. Am. Chem. Soc. (79)(1957) 5046 5049. [43] J. Synowiecki, Z.E. Sikorski, M. Naczk, Immobilization of invertase on krill chitin, Biotechnol. Bioeng. (23)(1981) 231 233. [44] A.B. Foster, J.M. Webber, Chitin, Adv. Carbohydr. Chem. 15 (1960) 371 393. [45] P.R. Austin, C.J. Brine, J.E. Castle, J.P. Zikakis, Chitin: facets of research, Science (212)(1981) 749 753. [46] C.J. Brine, P.R. Austin, Chitin variability with species and method of preparation, Comp. Biochem. Physiol. (69B)(1981) 283 286. [47] Q.P. Peniston, E.L. Lohnson, Process for demineralization of crustacea shells. U.S. Patent 4,066,735, January 3, 1978. [48] T. Truong, R. Hausler, F. Monette, P. Niquette, Fishery industrial waste valorization for the transformation of chitosan by hydrothermo-chemical method, Rev. Sci. Eau (20)(2007) 253 262.

Handbook of Chitin and Chitosan

References

367

[49] F.O. Marquis-Duval, Isolation et valorisation des constituants de la carapace de la crevette nordique (PhD dissertation), Laval University, Quebec, Canada. [50] H. Mirzadeh, N. Yaghobi, S. Amanpour, H. Ahmadi, M.A. Mohagheghi, F. Hormozi, Preparation of chitosan derived from shrimp’s shell Persian Gulf as Blood Hemostasis Agent, Iran. Polym. J. 11 (1) (2002) 63 68. [51] N.M. Alves, J.F. Mano, Chitosan derivatives obtained by chemical modifications for biomedical and environmental applications, Int. J. Biol. Macromol. (43)(2008) 401 414. ISSN 0141-8130.. [52] K. Shirai, D. Palella, Y. Castro, I. Guerrero-Legarreta, G. Saucedo-Castaneda, S. Huerta-Ochoa, et al., Characterization of chitins from lactic acid fermentation of prawn wastes, in: R.H. Chen, H.C. Chen (Eds.), Advances in Chitin Science, vol. III, Elsevier, Taiwan, 1998, pp. 103 110. [53] S.H.O. Bahasan, S. Sathianeson Satheesh, M.A. Mohammad Abdulaziz Ba-akdah, Extraction of chitin from the shell wastes of two shrimp species Fenneropenaeus semisulcatus and Fenneropenaeus indicus using microorganisms, J. Aquat. Food Product. Technol. 26 (4) (2017) 390 405. [54] L. Adour, W. Arbia, A. Amrane, N. Mameri, Combined use of waste materials— recovery of chitin from shrimp shells by lactic acid fermentation supplemented with date juice waste or glucose, J. Chem. Technol. Biotechnol. (83)(2008) 1664 1669. [55] M. Healy, A. Green, A. Healy, Bioprocessing of marine crustacean shell waste, J. Eng. Life Sci. 23 (2 3) (2003) 151 160. [56] O. Aytekin, M. Elibol, Cocultivation of Lactococcus lactis and Teredinobacter turnirae for biological chitin extraction from prawn waste, Bioprocess. Biosyst. Eng (33)(2010) 393 399. [57] O. Cremades, E. Ponce, R. Corpas, J.F. Gutie´rrez, M. Jover, M.C. Alvarez-Ossorio, et al., Processing of crawfish (Procambarus clarkii) for the preparation of carotenoproteins and chitin, J. Agric. Food Chem. 49 (11) (2001) 5468 5470. [58] S. Kaur, G.S. Dhillon, Recent trends in biological extraction of chitin from marine shell wastes: a review, Crit. Rev. Biotechnol. 35 (1) (2015) 44 61. [59] K.L. Morley, G. Chauve, R. Kazlauskas, C. Dupont, F. Shareck, R.H. Marchessault, Acetyl xylan esterase-catalyzed deacetylation of chitin and chitosan, Carbohydr. Polym. (63)(2006) 310 315. [60] I. Aranaz, M. Mengı´bar, R. Harris, I. Pan˜os, B. Miralles, N. Acosta, et al., Functional characterization of chitin and chitosan, Curr. Chem. Biol. (3)(2009) 203 230. [61] G.W. Gooday, J.I. Prosser, K. Hillman, M.G. Cross, Mineralization of chitin in an estuarine sediment: the importance of the chitosan pathway, Biochem. Syst. Ecol. 19 (5) (1991) 395 400. [62] N.V. Toan, Production of chitin and chitosan from partially autolyzed shrimp shell materials, Open. Biomater. J. (1)(2009) 21 24. [63] A. Domard, M. Rinadudo, Preparation and characterization of fully deacetylated chitosan, I. J. Biol. Macromol. (5)(1983) 49 52. [64] A. Pelletier, I. Lemire, J. Sygusch, E. Chornet, R.P. Overend, Chitin/chitosan transformation by thermo-mechano-chemical treatment including characterization by enzymatic depolymerization, Biotechnol. Bioeng. (36)(1990) 310 315. [65] B. Focher, P.L. Beltrane, A. Naggi, G. Torri, Alkaline N-deacetylation of chitin enhanced by flash treatments. Reaction kinetics and structure modifications, Carbohydr. Polym. (12)(1990) 405 418. [66] F.M. Goycoolea, I. Higuera-Ciapara, J.L.J. Hernandez, K.D. Garcı´a, Preparation of chitosan from squid (Loligo spp) pen by a microwave-accelerated thermochemical process, in: A. Domard, G.A.F. Roberts, K.M. Va˚rum (Eds.), Advances in Chitin Science, Jacques Andre´, Lyon, 1998, pp. 78 83.

Handbook of Chitin and Chitosan

368

11. Chitin, chitosan, marine to market

[67] G. Galed, B. Miralles, I. Pan˜os, A. Santiago, A.N. Heras, Deacetylation and depolymerization reactions of chitin/chitosan: influence of the source of chitin, Carbohydr. Polym. 62 (4) (2005) 316 320. [68] P.R. Rege, L.H. Block, Chitosan processing: influence of process parameters during acidic and alkaline hydrolysis and effect of the processing sequence on the resultant chitosan’s properties, Carbohyd. Res. 321 (3 4) (1999) 235 245. [69] M. Sukwattanasinitt, H. Zhu, H. Sashiwa, S. Aiba, Utilization of commercial non-chitinase enzymes from fungi for preparation of 2-acetamido-2-deoxy—glucose from [beta]-chitin, Carbohydr. Res. 337 (2) (2002) 133 137. [70] G. Galed, E. Diaz, A. Heras, Conditions of N-deacetylation on chitosan production from alpha chitin, Nat. Prod. Commun. 3 (4) (2008) 543 550. [71] V. Zargar, M. Asghari, A. Dashti, A review on chitin and chitosan polymers: structure, chemistry, solubility, derivatives, applications, Chem. Bio. Eng. Rev. 2 (3) (2015) 204 226. [72] C.H. Kim, J.W. Choi, H.J. Chun, K.S. Choi, Synthesis of chitosan derivatives with quaternary ammonium salt and their antibacterial activity, Polym. Bull. (38)(1997) 387. [73] Y. Baba, H. Noma, R. Nakayama, Y. Matsushita, Preparation of chitosan derivatives containing methylthiocarbamoyl and phenylthiocarbamoyl groups and their selective adsorption of copper (II) over iron (III), Anal. Sci. 18 (3) (2002) 359 361. [74] M. Sugimoto, M. Morimoto, H. Sashiwa, H. Saimoto, Y. Shigemasa, Preparation and characterization of water-soluble chitin and chitosan derivatives, Carbohydr. Polym. 36 (1) (1998) 49 59. [75] Q. Li, E.T. Dunn, E.W. Grandmaison, M.F.A. Goosen, in: M.F.A. Goosen (Ed.), Applications and Properties of Chitosan, Applications of Chitin and Chitosan, Technomic Publishing Company, Lancaster, 1997, pp. 3 29. [76] H. Sashiwa, Y. Shigemasa, R. Roy, Chemical modification of chitosan: synthesis of dendronized chitosan-sialic acid hybrid using convergent grafting of preassembled dendrons built on gallic acid and tri(ethylene glycol) backbone, Macromolocules 2001 (2001) 3905. [77] H. Zhang, Q. Dang, C. Liu, D. Yub, Y. Wang, X. Pu, et al., Fabrication of methyl acrylate and tetraethylenepentamine grafted magnetic chitosan microparticles for capture of Cd(II) from aqueous solutions, J. Hazard. Mater. (366)(2019) 346 357. [78] X. Qu, A. Wirsen, A.C. Albertsson, Effect of lactic/glycolic acid side chains on the thermal degradation kinetics of chitosan derivatives, Polymer (41)(2001) 4841. [79] H. Tokumitsu, J. Hiratsuka, Y. Sakurai, T. Kobayashi, H. Ichikawa, Y. Fukumori, Gadolinium neutron-capture therapy using novel gadopentetic acid-chitosan complex nanoparticles: in vivo growth suppression of experimental melanoma solid tumor, Cancer Lett. (150)(2000) 177. [80] H. Tokumitsu, H. Ichikawa, Y. Fukumori, Chitosangadopentetic acid complex nanoparticles for gadolinium neutron-capture therapy of cancer: preparation by novel emulsiondroplet coalescence technique and characterization, Pharm. Res. (16)(1999) 1830. [81] H. Tamura, T. Furuike, S.V. Nair, R. Jayakumar, Biomedical applications of chitin hydrogel membranes and scaffolds, Carbohydr. Polym. (84)(2011) 820 824. [82] H. Nagahama, T. Kashiki, N. Nwe, R. Jayakumar, T. Furuike, H. Tamura, Preparation of biodegradable chitin/gelatin membranes with GlcNAc for tissue engineering applications, Carbohydr. Polym. (73)(2008) 456 463. [83] H. Nagahama, N. Nwe, R. Jayakumar, S. Koiwa, T. Furuike, H. Tamura, Novel biodegradable chitin membranes for tissue engineering applications, Carbohydr. Polym. (73)(2008) 295 302. [84] K. Azuma, S. Ifuku, T. Osaki, Y. Okamoto, S. Minami, Preparation and biomedical applications of chitin and chitosan nanofibers, J. Biomed. Nanotechnol. (10)(2014) 2891 2920.

Handbook of Chitin and Chitosan

References

369

[85] Y. Fan, T. Saito, A. Isogai, Preparation of chitin nanofibers from squid pen betachitin by simple mechanical treatment under acid conditions, Biomacromolecules (9)(2008) 1919 1923. [86] S. Ifuku, M. Nogi, K. Abe, M. Yoshioka, M. Morimoto, H. Saimoto, et al., Preparation of chitin nanofibers with a uniform width as alpha-chitin from crab shells, Biomacromolecules (10)(2009) 1584 1588. [87] N.L.B.M. Yusof, L.Y. Lim, E. Khor, Preparation and characterization of chitin beads as a wound dressing precursor flexible chitin films as potential wound-dressing materials: wound model studies, J. Biomed. Mater. Res. (54)(2001) 59 68. 23. [88] A. Anitha, N. Deepa, K.P. Chennazhi, S.V. Nair, H. Tamura, R. Jayakumar, Development of mucoadhesive thiolated chitosan nanoparticles for biomedical applications, Carbohydr. Polym. (83)(2010) 66 73. [89] M. Peter, Sudheesh, P.T. Kumar, N.S. Binulal, S.V. Nair, H. Tamura, et al., Development of novel chitin/nano bioactive glass ceramic nanocomposite scaffolds for tissue engineering applications, Carbohydr. Polym. (78)(2009) 926 931. [90] M. Peter, N.S. Binulol, S. Soumya, S.V. Nair, H. Tamura, R. Jayakumar, Nanocomposite scaffolds of bioactive glass ceramic nanoparticles disseminated chitosan matrix for tissue engineering applications, Carbohydr. Polym. (79)(2010) 284 289. [91] C.A. Ibekwe, G.M. Oyatogun, T.A. Esan, K.M. Oluwasegun, Synthesis and characterization of chitosan/gum arabic nanoparticles for bone regeneration, Am. J. Mater. Sci. Eng. 5 (1) (2017) 28 36. [92] M. Prabaharan, R. Jayakumar, Chitosan-graft-β-cyclodextrin scaffolds with controlled drug release capability for tissue engineering applications, Int. J. Biol. Macromol. (44)(2009) 320 325. [93] Y. Maeda, R. Jayakumar, H. Nagahama, T. Furuike, H. Tamura, Synthesis, characterization and bioactivity studies of novel β-chitin scaffolds for tissue-engineering applications, Int. J. Biol. Macromol. (42)(2008) 463 467. [94] K. Muramatsu, S. Masuda, Y. Yoshihara, Fujisawa, In vitro degradation behavior of freeze-dried carboxymethyl-chitin sponges processed by vacuum-heating and gamma irradiation, Polym. Degradat. Stabil. (81)(2003) 327 332. [95] A. Portero, D. Teijeiro-Osorio, M.J. Alonso, C. Remunan-Lopez, Development of chitosan sponges for buccal administration of insulin, Carbohydr. Polym. (68)(2007) 617 625. [96] M.L. Bade, Structure and isolation of native animal chitins, in: M.F.A. Goosen (Ed.), Applications of Chitin and Chitosan, Technomic Publishing, Lancaster, 1997, pp. 253 277. [97] K. Kurita, T. Sannan, Y. Iwakura, Studies on chitin, evidence for formation of block and random copolymers of N-acetyl-D-glucosamine and D-glucosamine by heteroand homogeneous hydrolyses, Makromol. Chem. (178)(1997) 3197 3202. [98] B. Klaykruayat, K. Siralertmukul, K. Srikulkit, Chemical modification of chitosan with cationic hyperbranched dendritic polyamidoamine and its antimicrobial activity on cotton fabric, Carbohydr. Polym. 80 (1) (2010) 197 207. [99] K. Tomihata, Y. Ikada, In vitro and in vivo degradation of films of chitin and its deacetylated derivatives, Biomaterials 18 (7) (1997) 567 575. [100] H. Zhang, S.H. Neau, In vitro degradation of chitosan by a commercial enzyme preparation: effect of molecular weight and degree of deacetylation, Biomaterials 22 (12) (2001). 1653 1658. [101] S.H. Lim, S.M. Hudson, Synthesis and antimicrobial activity of a water-soluble chitosan derivative with a fiber-reactive group, Carbohydr. Res. 339 (2) (2004) 313 319. [102] L.Y. Zheng, J.F. Zhu, Study on antimicrobial activity of chitosan with different molecular weights, Carbohydr. Polym. 54 (4) (2003) 527 530.

Handbook of Chitin and Chitosan

370

11. Chitin, chitosan, marine to market

[103] M. Kong, X.G. Chen, K. Xing, H.J. Park, Antimicrobial properties of chitosan and mode of action: a state of the art review, Int. J. Food Microbiol. 144 (1) (2010) 51 63. [104] K.V. Harish-Prashanth, R.N. Tharanathan, Chitin/chitosan: modifications and their unlimited application potential—an overview, Trends Food Sci. Technol. (18)(2007) 117 131. pISSN 0924-2244. [105] S.B. Lin, Y.C. Lin, H.H. Chen, Low molecular weight chitosan prepared with the aid of cellulase, lysozyme and chitinase: characterisation and antibacterial activity, Food Chem. (116)(2009) 47 53. ISSN 0308-8146. [106] Y. Sun, F. Cui, K. Shi, J. Wang, M. Niu, R. Ma, The effect of chitosan molecular weight on the characteristics of spray-dried methotrexate-loaded chitosan microspheres for nasal administration, Drug. Dev. Ind. Pharm. (35)(2009) 379 386. ISSN 0363-9045. [107] T.A. Khan, K.K. Peh, Influence of chitosan molecular weight on its physical properties, Int. Med. J. 2 (1) (2003). ISSN 1823-4631. [108] E. Khor, Chitin: fulfilling a biomaterials promise, Biomaterials 23 (18) (2001) 3913 3915. [109] M.K. Jang, B.G. Kong, Y.I. Jeong, C.H. Lee, J.W. Nah, Physicochemical characterization of α-chitin, β-chitin and γ-chitin separated from natural resources, J. Polym. Sci. A1 (42) (2004) 3423 3432. [110] R.A.A. Muzzarelli, M. El-Mehtedi, M. Mattioli-Belmonte, Emerging biomedical applications of nano-chitins and nano-chitosans obtained via advanced eco-friendly technologies from marine resources, Mar. Drugs (12)(2014) 5468 5502. [111] J. Synowiecki, N.A. Al-Khateeb, Production, properties and some new applications of chitin and its derivatives, Crit. Rev. Food Sci. Nutr. (43)(2003). 145-171. [112] I.F.M. Rumengan, E. Suryanto, R. Modaso, S. Wullur, T.E. Tallei, D. Limbong, Structural characteristics of chitin and chitosan isolated from the biomass of cultivated Rtifer, Brachionus rotundiformis, Int. J. Fish. Aquat. Sci. 3 (1) (2014) 12 18. [113] C.K.S. Pillai, W. Paul, C.P. Sharma, Chitin and chitosan polymers: chemistry, solubility and fiber formation, Prog. Polym. Sci. (34)(2009) 641 678. [114] A. Costa-Pinto, R. Reis, N.M. Neves, Scaffolds based bone tissue engineering: the role of chitosan, Tissue Eng. Part B 17 (5) (2011) 1 18. [115] M. Rodriguez-vazquez, B. Vega-Ruiz, R. Ramos-Zuniga, D.A. Saldana-Koppel, L.F. Quinones-Olvera, Chitosan and its potential use as a scaffold for tissue engineering in regenerative medicine, Biomed. Res. Int. 2015 (2015) 1 15. [116] J. Kumirska, M.X. Weinhold, J. Tho¨ming, P. Stepnowski, Biomedical activity of chitin/chitosan based materials-influence of physicochemical properties apart from molecular weight and degree of N-acetylation, Polymers (3)(2011) 1875 1901. [117] S.M. Hudson, C. Smith, Polysaccharide: chitin and chitosan: chemistry and technology of their use as structural materials, in: D.L. Kaplan (Ed.), Biopolymers from Renewable Resources, Springer-Verlag, New York, 1998, pp. 96 118. [118] G. Berth, H. Dautzenberg, The degree of acetylation of chitosans and its effect on the chain conformation in aqueous solution, Carbohydr. Polym. (47)(2002) 39 51. ISSN 0144- 8617. [119] G. Berth, H. Dautzenberg, M.G. Peter, Physico-chemical characterization of chitosans varying in degree of acetylation, Carbohydr. Polym. (36)(1998) 205 216. ISSN 0144-8617. [120] C. Schatz, C. Viton, T. Delair, C. Pichot, A. Domard, Typical physicochemical behaviors of chitosan in aqueous solution, Biomacromolecules (4)(2003) 641 648. ISSN 1525-7797. [121] P. Sorlier, A. Denuziere, C. Viton, A. Domard, Relation between the degree of acetylation and the electrostatic properties of chitin and chitosan, Biomacromolecules (2) (2001) 765 772. ISSN 1525-7797.

Handbook of Chitin and Chitosan

References

371

[122] P. Sorlier, C. Viton, A. Domard, Relation between solution properties and degree of acetylation of chitosan: role of aging, Biomacromolecules (3)(2002) 1336 1342. pISSN 1525-7797. [123] P. Sandford, Chitosan: commercial uses and potential applications, in: E. SkjakBraek, T. Anthonsen, P. Standorf (Eds.), Chitin and Chitosan: Sources Chemistry, Biochemistry, Physical Properties and Applications, Elsevier Applied Science, London, 1989, pp. 51 69. [124] G. Gomez d’Ayala, M. Malinconico, P. Laurienzo, Marine derived polysaccharides for biomedical applications: chemical modification approaches, Molecules 2008 (13) (2008) 2069 2106. [125] N.R. Sudarshan, D.G. Hoover, D. Knorr, Antibacterial action of chitosan, Food Biotechnol. (6)(1992) 257 272. [126] S. Leuba, P. Stossel, Chitosan and other polyamines: antifungal activity and interaction with biological membranes, in: R.A.A. Muzzarelli, C. Jeuniaux, C. Gooday (Eds.), Chitin in Nature and Technology, Plenum Press, New York, 1985, p. 217. [127] I. Younes, S. Sellimi, M. Rinaudo, K. Jellouli, M. Nasri, Influence of acetylation degree and molecular weight of homogeneous chitosans on antibacterial and antifungal activities, Int. J. Food Microbiol. (185)(2014) 57 63. [128] E.I. Rabea, M.E.T. Badawy, C.V. Stevens, G. Smagghe, W. Steurbaut, Chitosan as antimicrobial agent: applications and mode of action, Biomacromolecules (4)(2003) 1457 1465. [129] M. Anraku, A. Michihara, T. Yasufuku, K. Akasaki, D. Tsuchiya, H. Nishio, et al., The antioxidative and antilipidemic effects of different molecular weight chitosans in metabolic syndrome model rats, Biol. Pharmaceut. Bull. 33 (12) (2010) 1994 1998. [130] S.H. Chang, C.H. Wu, G.J. Tsai, Effects of chitosan molecular weight on its antioxidant and antimutagenic properties, Carbohydr. Polym. (181)(2018) 1026 1032. [131] P.J. Chien, F. Sheu, W.T. Huang, M.S. Su, Effect of molecular weight of chitosans on their antioxidative activities in apple juice, Food Chem. (102)(2007) 1192 1198. [132] W. Xie, P. Xu, Q. Liu, Antioxidant activity of watersoluble chitosan derivatives, Bioorg. Med. Chem. Lett. (11)(2001) 1699 1701. [133] H. Tomida, T. Fujii, N. Furutani, A. Michihara, T. Yasufuku, K. Akasaki, et al., Antioxidant properties of some different molecular weight chitosans, Carbohydr. Res. (344)(2009) 1690 1696. [134] M. Anraku, T. Fujii, Y. Kondo, E. Kojima, T. Hata, N. Tabuchi, et al., Antioxidant properties of high molecular weight dietary chitosan in vitro and in vivo, Carbohydr. Polym. (83)(2011) 501 505. [135] I.M. Helander, E.L. Nurmiaho-Lassila, R. Ahvenainen, J. Rhoades, S. Roller, Chitosan disrupts the barrier properties of the outer membrane of gram-negative bacteria, Int. J. Food Microbiol. (71)(2001) 235 244. ISSN 0168-1605. [136] C. Allan, L.A. Hadwiger, The fungicidal effect of chitosan on fungi of varying cell wall composition, Exp. Mycol. (3)(1979) 285 287. ISSN 0147-5975. [137] M.C. Chen, G.H.C. Yeh, B.H. Chiang, Antimicrobial and physicochemical properties of methylcellulose and chitosan films containing a preservative, J. Food Process. Preserv. (20)(1996) 379 390. ISSN 01458892. [138] B.O. Jung, C.H. Kim, K.S. Choi, Y.M. Lee, J.J. Kim, Preparation of amphiphilic chitosan and their antimicrobial activities, J. Appl. Polym. Sci. (72)(1999) 1713 1719. ISSN 1097-4628. [139] E.I. Rabea, M.T. El Badawy, T.M. Rogge, C.V. Stevens, M. Ho¨fte, W. Steurbaut, et al., Insecticidal and fungicidal activity of new synthesized chitosan derivatives, Pest Manag. Sci. (61)(2005) 951 960. pISSN 1526-498X.. [140] Z. Zhong, R. Chen, R. Xing, X. Chen, S. Liu, Z. Guo, et al., Synthesis and antifungal properties of sulfanilamide derivatives of chitosan, Carbohydr. Res. (342)(2007) 2390 2395. ISSN 0008-6215.

Handbook of Chitin and Chitosan

372

11. Chitin, chitosan, marine to market

[141] I. Ikeda, Y. Tomari, M. Sugano, Interrelated effects of dietary fiber and fat on lymphatic cholesterol and triglyceride absorption in rats, J. Nutr. (119)(1989) 1383 1387. [142] C.M. Gallaher, J. Munion, R. Hesslink, J. Wise, D. Gallaher, Cholesterol reduction by glucomannan and chitosan is mediated by changes in cholesterol absorption and bile acid and fat excretion in rats, J. Nutr. (130)(2000) 2753 2759. [143] C.M. Lian, K.L. Chen, K.C. Chan, Effect of high molecular weight chitosan on nutrient digestibility, blood lipids and fecal contents in rats, in: R.H. Chen, H. Chen (Eds.), Advances in Chitosan Science, vol. 3, National Taiwan Ocean University, Taiwan, 1999, pp. 299 306. [144] P. He, S.S. Davis, L. Illum, In vitro evaluation of the mucoadhesive properties of chitosan microspheres, Int. J. Pharm. (166)(1998) 75 88. pISSN 0378-5173. [145] J. Mota, N. Yu, S.G. Caridade, G.M. Luz, E.M. Gomes, R.L. Reis, et al., Chitosan/ bioactive glass nanoparticle composite membranes for periodontal regeneration, Acta Biomater. (8)(2012) 4173 4180. [146] C. Saikia, P. Gogoi, T.K. Maji, Chitosan: a promising biopolymer in drug delivery applications, J. Mol. Genet. Med. (S4)(2015) 006. Available from: https://doi.org/ 10.4172/1747-0862.S4-006. [147] M.N.V. RaviKumar, R.A.A. Muzzarelli, C. Muzzarelli, H.A. Sashiwa, J. Domb, Chitosan chemistry and pharmaceutical perspectives, Chem. Rev. (104)(2004) 6017 6084. [148] Y. Le, S.C. Anand, A.R. Horrocks, Recent developments in fibres and materials for wound management, Indian J. Fibre Text. (22)(1997) 337. [149] K. Murakami, H. Aoki, S. Nakamura, S. Nakamura, M. Takikawa, M. Hanzawa, et al., Hydrogel blends of chitin/chitosan, fucoidan and alginate as healingimpaired wound dressings, Biomaterials (31)(2010) 83 90. [150] N.L. Yusof, A. Wee, L.Y. Lim, E. Khor, Flexible chitin films as potential wound-dressing materials: wound model studies, J. Biomed. Mater. Res. Part A (66A)(2003) 224 232. [151] P. Wongpanit, N. Sanchavanakit, P. Pavasant, P. Supaphol, S. Tokura, R. Rujiravanit, Preparation and characterization of microwave-treated carboxymethylchitin and carboxymethylchitosan films for potential use in wound care application, Macromol. Biosci. 5 (2005) 1001 1012. [152] Y.C. Chung, H.L. Wang, Y.M. Chen, S.L. Li, Effect of abiotic factors on the antibacterial activity of chitosan against waterborne pathogens, Biores. Technol. (88)(2003) 179 184. [153] Z. Limam, S. Selmi, S. Sadok, A. El Abed, Extraction and characterization of chitin and chitosan from crustacean by-products: biological and physicochemical properties, Afr. J. Biotechnol. 10 (4) (2011) 640 647. [154] R.G. Cuero, Antimicrobial action of exogenous chitosan, EXS (87)(1999) 315 333. [155] S.L. Wang, J.R. Huang, Microbial reclamation of shellfish wastes for the production of chitinases, Enzyme Microb. Tech. (28)(2001) 376 382. [156] R. Jayakumar, R.L. Reis, J.F. Mano, Phosphorous containing chitosan beads for controlled oral drug delivery, J. Bioact. Compat. Polym. (21)(2006) 327 340. [157] N. Acosta, I. Aranaz, C. Peniche, A. Heras, Tramadol release from a delivery system based on alginate-chitosan microcapsules, Macromol. Biosci. (3)(2003) 546 551. [158] C. Peniche, W. Arguelles-Monal, H. Peniche, N. Acosta, Chitosan: an attractive biocompatible polymer for microencapsulation, Macromol. Biosci. (3)(2003) 511 520. [159] L. Illum, N.F. Farraj, S.S. Davis, Chitosan as novel nasal delivery system for peptides drugs, Pharm. Res. (11)(1994) 1186 1189. [160] S. Mao, W. Sun, T. Kissel, Chitosan-based formulations for delivery of DNA and siRNA, Adv. Drug. Deliv. Rev. (62)(2010) 12 27. [161] A. Tokoro, N. Tatewaki, K. Suzuki, T. Mikami, S. Suzuki, M. Suzuki, Growth inhibitory effect of hexa-N-acetylchitohexaose and chitohexaos and Meth-A solid tumor, Chem. Pharm. Bull. (Tokyo) (36)(1998) 784 790.

Handbook of Chitin and Chitosan

References

373

[162] J. Murata, I. Saiki, S. Nishimura, N. Nishi, S. Tokura, I. Azuma, Inhibitory effect of chitin heparinoids on the lung metastasis of B16-BL6 melanoma, Jpn. J. Cancer Res. (80)(1986) 866 872. [163] M. Hasegawa, K. Yagi, S. Iwakawa, M. Hirai, Chitosan induces apoptosis via caspase-3 activation in bladder tumor cells, Jpn. J. Cancer Res (92)(2001) 459 466. [164] L. Qi, Z. Xu, M. Chen, In vitro and in vivo suppression of hepatocellular carcinoma growth by chitosan nanoparticles, Eur. J. Cancer (43)(2007) 184 193. [165] M. Guminska, J. Ignacak, E. Wojcik, In vitro inhibitory effect of chitosan and its degradation products on energy metabolism in Ehrlich ascites tumour cells (EAT), Pol. J. Pharmacol. (48)(1996) 495 501. [166] S.Y. Lin, H.Y. Chan, F.H. Shen, M.H. Chen, Y.J. Wang, C.K. Yu, Chitosan prevents the development of AOM-induced aberrant crypt foci in mice and suppressed the proliferation of AGS cells by inhibiting DNA synthesis, J. Cell Biochem. (100)(2007) 1573 1580. [167] C.R. Dass, P.F. Choong, The use of chitosan formulations in cancer therapy, J. Microencapsul (25)(2008) 275 279. [168] W.R. Chen, R.L. Adams, R. Carubelli, R.E. Nordquist, Laser-photosensitizer assisted immunotherapy: a novel modality for cancer treatment, Cancer Lett. (115)(1997) 25 30. [169] M.L. Markey, L.M. Bowman, V.M. Bergamini, Contact lenses made of chitosan, in: G. Skjak-Braek, T. Anthonsen, P. Sandford (Eds.), Chitin and Chitosan: Source, Chemistry, Biochemistry, Physical Properties and Applications, Elsevier Applied Science, London and New York, 1989, pp. 713 717. [170] M. Anand, M. Maruthupandy, S. Prasanna kumar, S. Ravikumar, N. Jeyaraj, R. Kumaran, Antimicrobial potential of chitosan from marine biowastes—a wealth from waste approach, J. Chitin Chitosan Sci. 3 (2) (2015) 1 4. [171] Z.S. Jia, D.F. Shen, W.L. Xu, Synthesis and antibacterial activities of quaternary ammonium salt of chitosan, Carbohydr. Res. 333 (1) (2001). [172] H. Liu, Y.M. Du, J.H. Yang, H.Y. Zhu, Structural characterization and antimicrobial activity of chitosan/betaine derivative complex, Carbohydr. Polym. (55)(2004) 291 297. [173] J. Dutta, P.K. Dutta, Tissue engineering: an emergent process for development of bio-products, Industrial Products Finder, Business Press, Mumbai, India, 2003, pp. 246 253. [174] C. Jarry, C. Chaput, A. Chenite, M.A. Renaud, M. Buschmann, J.C. Leroux, Effects of steam sterilization on thermogelling chitosan-based gels, J. Biomed. Mater. Res. (58)(2001) 127 132. [175] C.E. Kast, W. Frick, U. Losert, A.B. Schnurch, Chitosanthioglycolic acid conjugate: a new scaffold material for tissue engineering, Int. J. Pharm. (256)(2003) 183 189. [176] Y. Zhang, M. Zhang, Synthesis and characterization of macroporous chitosan/ calcium phosphate composite scaffolds for tissue engineering, J. Biomed. Mater. Res. (55)(2001) 304 307. [177] S.V. Madihally, H.W.T. Matthew, Porous chitosan scaffolds for tissue engineering, Biomaterials (20)(1999) 1133 1337. [178] M. Prasitslip, R. Jenwithisuk, K. Kongsuwan, N. Damrongchai, P. Watts, Cellular responses to chitosan in vitro: the importance of deacetylation, J. Mater. Sci: Mater. Med. (11)(2000) 773. [179] M. Mucha, Rheological characteristics of semi-dilute chitosan solutions, Macromol. Chem. Phys. (198)(1997) 471 475. [180] H. Jing, W. Su, S. Caracci, T.G. Bunning, T. Cooper, W. Adams, Optical waveguiding and morphology of chitosan thin films, J. Appl. Polym. Sci. (61)(1996) 1163 1169.

Handbook of Chitin and Chitosan

374

11. Chitin, chitosan, marine to market

[181] G. Crini, Recent developments in polysaccharide-based materials used as adsorbents in wastewater treatment, Prog. Polym. Sci. 30 (1) (2005) 38 70. [182] H.K. No, S.P. Meyers, Application of chitosan for treatment of wastewaters, Rev. Environ. Contam. Toxicol. (163)(2000) 1 28. [183] E. Guibal, Interactions of metal ions with chitosan-based sorbents: a review, Sep. Purif. Technol. 38 (1) (2004) 43 74. [184] E. Guibal, M. Van-Vooren, B.A. Dempsey, J.A. Roussy, Review of the use of chitosan for the removal of particulate and dissolved contaminants, Sep. Sci. Technol. 41 (11) (2006) 2487 2514. [185] J. Roussy, M. Van-Vooren, B.A. Dempsey, E. Guibal, Influence of chitosan characteristics on the coagulation and the flocculation of bentonite suspensions, Water Res. 39 (14) (2005) 3247 3258. [186] G. Crini, P. Badot, Application of chitosan, a natural aminopolysaccharide, for dye removal from aqueous solutions by adsorption processes using batch studies: a review of recent literature, Prog. Polym. Sci. (33)(2008) 399 447. [187] R. Muzzarelli, O. Tuberni, Chitin and chitosan as chromatographic supports and adsorbents for collection of metal ions from organic and aqueous solutions and sea-water, Talanta (71)(1969) 1571 1577. [188] J.P. Thome´, Y. Van-Daele, Adsorption of polychlorinated biphenyls (PCB) on chitosan and application to decontaminated of polluted stream waters, in: R.A.A. Muzzarelli, C. Jeuniaux, G.W. Gooday (Eds.), Proceedings of the Third International Conference on Chitin and Chitosan. Senigallia, 1986, pp. 551 554. [189] R. Crisafully, M.A.L. Milhome, R.M. Cavalcante, E.R. Silveira, D. De Keufeleire, R.F. Nascimiento, Removal of some polycyclic aromatic hydrocarbons from petrochemical wastewater using low-cost adsorbents of natural origin, Biores Technol. (99) (2008) 4515 4519. [190] T.K. Saha, S. Karmaker, H. Ichikawa, Y. Fukumori, Mechanisms and kinetics of trisodium 2-hydroxy-1,1-azonaphthalene-3,4,6-trisulfonate adsorption onto chitosan, J. Colloid Interf. Sci. (286)(2005) 433 439. [191] Y.C. Chung, Y.H. Li, C.C. Chen, Pollutant removal from aquaculture wastewater using the biopolymer chitosan at different molecular weights, Environ. Sci. Health A (40)(2005) 1775 1790. [192] M. Weltrowski, B. Martel, M. Morcellet, Chitosan N-benzyl sulfonate derivatives as sorbents for removal of metal ions in an acidic medium, J. Appl. Polym. Sci. (59) (1996) 647 654. [193] K.D. Bhavani, P.K. Dutta, Physico-chemical adsorption properties on chitosan for dyehouse effluent, Am. Dyest. Rep. (88)(1999) 53 58. [194] T.R. Sridhari, P.K. Dutta, Synthesis and characterization of maleilated chitosan for dye house effluent, Indian. J. Chem. Tech. (7)(2000) 198 201. [195] A. El Ghaouth, J. Arul, C. Wilson, N. Benhamou, Biochemical and cytochemical aspects of the interactions of chitosan and Botrytis cinerea in bell pepper fruit, Postharvest Biol. Tec. (12)(1997) 183 194. [196] P. Herna´ndez-Mun˜oz, E. Almenar, M. Ocio, R. Gavara, Effect of calcium dips and chitosan coatings on postharvest life of strawberries (Fragaria 3 ananassa), Postharvest Biol. Tec. 39 (3) (2006) 247 253. [197] C. Ribeiro, A. Vicente, J. Teixeira, C. Miranda, Optimization of edible coating composition to retard strawberry fruit senescence, Postharvest Biol. Tec. 44 (1) (2007) 63 70. [198] P.J. Chien, F. Sheu, H.R. Lin, Quality assessment of low molecular weight chitosan coating on sliced red pitayas, J. Food Eng. 79 (2) (2007) 736 740.

Handbook of Chitin and Chitosan

References

375

[199] Y. Jeon, J. Kamil, F. Shahidi, Chitosan as an edible invisible film for quality preservation of herring and Atlantic cod, J. Agric. Food Chem. 50 (18) (2002) 5167 5178. [200] G. Tsai, W. Su, H. Chen, C. Pan, Antimicrobial activity of shrimp chitin and chitosan from different treatments and applications of fish preservation, Fish. Sci. (68)(2002) 170 177. [201] S. Kim, H. No, W. Prinyawiwatkul, Effect of molecular weight, type of chitosan, and chitosan solution pH on the shelf-life and quality of coated eggs, J. Food Sci. 72 (1) (2007) S044 S048. [202] L.A. Hadwiger, Multiple effects of chitosan on plant systems: solid science or hype, Plant Sci. 208 (2013) 42 49. [203] J. Liu, J. Zhang, W. Xia, Hypocholesterolaemic effects of different chitosan samples in vitro and in vivo, Food Chem. 107 (1) (2008) 419 425. [204] L. Zeng, C. Qin, W. Wang, W. Chi, W. Li, Absorption and distribution of chitosan in mice after oral administration, Carbohydr. Polym. 71 (3) (2008) 435 440. [205] M. Sumiyoshi, Y. Kimura, Low molecular weight chitosan inhibits obesity induced by feeding a high-fat diet long-term in mice, J. Pharm. Pharmacol. (58)(2006) 201 207. [206] P. Morgantim, Chitin nanofibrils and their derivatives as cosmeceuticals, in: S.K. Kim (Ed.), Chitin, Chitosan, Oligosaccharides and their Derivatives: Biological Activities and Application, CRC Press, New York, 2010, pp. 531 542. [207] P. Morganti, P. Palombo, M. Palombo, G. Fabrizi, A. Cardillo, F. Svolacchia, et al., A phosphatidylcholine hyaluronic acid chitin nanofibrils complex for a fast skin remodeling and a rejuvenating look, Clin. Cosmet. Investig. Dermatol. (5)(2012) 213 220. [208] A. Kulkarni, A. Tourrette, M.M.C.G. Warmoeskerken, D. Jocic, Microgel-based surface modifying system for stimuli-responsive functional finishing of cotton, Carbohydr. Polym. 82 (4) (2010) 1306 1314. ISSN 0144-8617. [209] J. Campos, P. Dı´az-Garcı´a, I. Montava, M. Bonet-Aracil, E. Bou-Belda, Chitosan pretreatment for cotton dyeing with black tea, IOP Conf. Ser. Mater. Sci. Eng. 254 (2017) 112001. Available from: https://doi.org/10.1088/1757-899X/254/11/112001. [210] A.K. Singla, M. Chawla, Chitosan: some pharmaceutical and biological aspects—an update, J. Pharm. Pharmacol. (53)(2001) 1047 1067. [211] M. Hayes, Chitin, chitosan and their derivatives from marine rest raw materials: potential food and pharmaceutical applications, in: M. Hayes (Ed.), Marine Bioactive Compounds, Springer, Boston, MA, 2012, pp. 115 128. [212] Global Industry Analysts Inc., Chitin and chitosan derivatives market report—2015. Available from: ,https://www.alliedmarketresearch.com/chitosan-market.. [213] K. Pulidindi, H. Pandey, Potential, price trends, competitive market share and forecast, 2018 2024, global industry size report, chitosan market analysis 2018 2024. Available from: ,https://www.gminsights.com/industry-analysis/chitosan-market.. [214] Future market insights report, chitin market: global industry analysis (2012 2016) and opportunity assessment (2017 2027). Available from: ,https://www.futuremarketinsights.com/reports/chitin-market.. [215] E. Prasad, Chitosan market by grade (industrial, food, and pharmaceutical), application (water treatment, food and beverages, cosmetics, medical and pharmaceuticals, and agrochemicals), and region (Asia Pacific, North America, Europe, Row)—Global Forecast to 2022. Available from: ,https://www.reportsnreports. com/reports/1467340-chitosan-market-by-grade-industrial-food-and-pharmaceutical-application-water-treatment-food..

Handbook of Chitin and Chitosan

376

11. Chitin, chitosan, marine to market

Further reading M.N. Horst, A.N. Walker, E. Klar, The pathway of crustacean chitin synthesis, in: M.N. Horst, J.A. Freeman (Eds.), The Crustacean Integument: Morphology and Biochemistry, CRC, Boca Raton, FL, 1993, pp. 113 149. M.S. Hossain, A. Iqbal, Production and characterization of chitosan from shrimp waste, J. Bangladesh Agril. Univ. 12 (1) (2014) 153 160. Z. Sheikh, S. Najeeb, Z. Khurshid, V. Verma, H. Rashid, M. Glogauer, Biodegradable materials for bone repair and tissue engineering applications, Materials (8) (2015) 2953 2993. Available from: ,www.mdpi.com/journal/Materials..

Handbook of Chitin and Chitosan

C H A P T E R

12 Miscibility, properties, and biodegradability of chitin and chitosan Muhammad Arshad, Muhammad Zubair and Aman Ullah Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, AB, Canada O U T L I N E 12.1 Introduction

378

12.2 Physicochemical properties of chitin and chitosan 12.2.1 Miscibility/solubility of chitin and chitosan 12.2.2 Dissolution mechanism

378 379 383

12.3 Biological properties of chitin and chitosan 12.3.1 Antioxidant property 12.3.2 Anticancer/antitumor property 12.3.3 Antimicrobial property 12.3.4 Antiinflammatory property 12.3.5 Neuroprotective property

385 386 386 387 388 388

12.4 Biodegradability of chitin and chitosan 12.4.1 factors affecting the biodegradability

389 391

12.5 Concluding remarks

392

References

392

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00012-2

377

© 2020 Elsevier Inc. All rights reserved.

378

12. Miscibility, properties, and biodegradability of chitin and chitosan

12.1 Introduction Chitin is the second most abundant natural polysaccharide after cellulose and is composed of N-acetyl-D-glucosamine units having β-(1,4) linkage [1]. Chitin is mainly produced from crustaceans, mollusks, insects, algae, fungi, and similar kinds of related organisms [2,3]. Being biodegradable, renewable, biocompatible, biofunctional, and environmentfriendly, chitin has found uses in drug delivery systems, water treatment, membranes, biodegradable adhesive tape, etc. [4]. These characteristic properties of the biomaterial gained higher attention. However, the poor solubility of chitin makes its use limited by various industries, especially in the pharmaceutical sector. Strong intra- and interhydrogen bonding in chitin molecules makes it completely insoluble in common organic and dilute aqueous solvents. Chitosan is a modified form of polysaccharide polymer, which is derived or obtained by the deacetylation and depolymerization of chitin in the presence of alkaline conditions or enzymes named as chitin deacetylase [1,5], and it is completely soluble in an acid medium. After deacetylation, the released amino groups make chitosan a versatile polymer and can be modified into a variety of materials with suitable properties for distinct applications. Both chitin and chitosan have numerous application as a sorbent for the removal of undesired metal ions [6] and dyes [7] during wastewater treatment, and in membranes for purification processes [8], in the food industry [9
11], agriculture [12,13], cosmetics [14], pulp and paper industry [15,16], wound healing [17], tissue engineering [18,19], and as carriers for gene [20,21] and drug delivery [17,18]. Chitin and chitosan can be easily modified and transformed into gels [22], beads [23], nanofibers [24,25], scaffold [26
28], sponges [29], microparticles [30], and nanoparticles [31].

12.2 Physicochemical properties of chitin and chitosan Chitin polymer is mainly composed of poly-(2-acetamido-2-deoxy-Dglucose) units having a degree of acetylation (DA) more than 50% (acetyl contents 70%
80%, amino contents 20%
30%) and a molecular weight in the range of 100
1000 kDa. This structural composition of chitin makes it insoluble in most solvents at any pH. However, its deacetylated derivative, chitosan, has a reduced degree of deacetylation, which contains 20%
30% acetyl contents, 70%
80% amino contents, and molecular weight in the range of 3.8
20 kDa. These characteristics of chitosan make it soluble in most of the solvents at pH , 6.3 [32
37].

Handbook of Chitin and Chitosan

12.2 Physicochemical properties of chitin and chitosan

379

12.2.1 Miscibility/solubility of chitin and chitosan Although chitin and chitosan’s molecular structures are almost similar, their physical properties and chemical reactions are often different due to the less crystalline structure of chitosan and the presence of a different percentage of reactive functional groups (amines, hydroxyls) on their linear chains. The variation in their solubility is because of their structural differences [38]. In most of the organic solvents, chitin is insoluble, while chitosan is dissolved easily in dilute acidic solutions with pH below 6, which is due to the presence of free amino groups making it a strong base with pKa value of 6.3 [39]. In an acidic medium, these amine groups become positively charged after getting protonated and make chitosan a watersoluble polyelectrolyte.

Among the organic acids, lactic acid and formic acid can dissolve chitosan [40,41]. Out of these two, formic acid (0.2%
100% in aqueous systems) is the best solvent to dissolve chitosan [42]. Chitosan also showed good solubility in dilute nitric acid and 1% hydrochloric acid, while in phosphoric and sulfuric acid it is insoluble. However, hightemperature conditions using concentrated acetic acid solution results in the depolymerization of chitosan [41,43]. Chitosan displayed no solubility in any of the organic solvents, including dimethyl sulfoxide and dimethylformamide. There are various factors affecting the solubility of chitosan such as temperature, time, concentration of alkali, pretreatments to chitin, ratio of chitin to alkali concentration used, and particle size [44]. Molecular weight and distribution of acetyl groups along the main chain also plays a critical role in chitosan solubility [45
48]. Recently, Geying Ru et al. reported the effect of aqueous alkali (KOH, NaOH, and LiOH) on the solubility of deacetylated chitin. This study shows that the solubility of chitin depends on the DA of the chitin chain, i.e., KOH . NaOH .. LiOH for chitin (DA 5 0.94
0.74), KOH  LiOH  NaOH for chitosan I (DA 5 0.53
0.25), while it is inverse LiOH .. KOH . NaOH for chitosan II (DA , 0.25) [49].

Handbook of Chitin and Chitosan

380

12. Miscibility, properties, and biodegradability of chitin and chitosan

In addition to acids and alkalis, glycerol-2-phosphate and polyethyleneglycol (PEG) are reported to aid the preparation of water-soluble chitosan at neutral pH [50
53]. 12.2.1.1 Dissolution by inorganic reagents Several attempts have been made and reported for the dissolution of chitin using sodium hydroxide as a strong inorganic base and inorganic salts [54
56]. Kunike reported the solubility of deacetylated chitin in acetic acid obtained by treating it with 5% caustic soda for 14 days at 60 C [57]. Dissolution of chitin by inorganic salts, such as CaCl2, CaBr2, LiSNS, CaI2, and Ca(CNS)2, has been reported by Weimarn [56]. Use of additives with NaOH like urea and thiourea enhance the solubility of chitin. Use of urea (4 wt.%) with NaOH (8 wt.%) at
20 C increases the solubility of chitin as reported by Kennedy and coworkers [1]. Seorv and coworkers reported the complete solubility of chitin in an aqueous solution of NaOH (10%), when 5% urea and 5% thioureas were used as additives [58]. 12.2.1.2 Dissolution by polar solvents and strong acids Dissolution of chitin is reported by strong polar protic solvents like trichloroacetic acid (TCA), dichloroacetic acid (DCA), etc. and using a combination of solvent systems (formic acid, diisopropyl ether, and DCA) [59,60]. As TCA and DCA are corrosive in nature, they can break the polymer into a lower molecular weight which can affect the strength of fibers. Highly polar solvents such as methane sulfonic acid, hexafluoroisopropyl alcohol, and hexafluoracetone sesquihydrate have also been reported for the solubilization of chitin [38]. 12.2.1.3 Solubility in ionic liquids Ionic liquids (ILs) act as a solvent with low melting points and have been found to dissolve natural polysaccharides [61
65]. Various studies have been reported on the dissolution of chitin using appropriate ILs to fabricate new chitin-based functional materials [62
68]. An IL [1-allyl-3-methylimidazolium bromide (AMIMBr)] has been found to dissolve α-chitin up to 5 wt.% by a simple process [69,70] (Fig. 12.1). A series of 1-allyl-3-methylimidazolium acetate (AMIMOAc), alkylimidazolium chloride, and dimethyl phosphate have been studied to explore their role for the dissolution of chitin; only former ILs were found to dissolve chitin up to 5 wt.% [64]. The dissolution behavior of chitin in ILs varies depending upon its crystallinity, degree of deacetylation, and molecular weight. Moreover, 1-butyl- and 1-ethyl-3methylimidazolium acetates (BMIMOAc and EMIMOAc, respectively) have also shown their capability to dissolve a certain percentage of

Handbook of Chitin and Chitosan

12.2 Physicochemical properties of chitin and chitosan

381

FIGURE 12.1 Representative ionic liquids for the dissolution of chitin. Source: Reproduced with permission from Elsevier.

chitin [64,71,72]. In a recent study, tetrabutylphosphonium amino acid salts and 1-ethyl-3-methylimidazolium alkanoates have also shown potential to dissolve chitin (Fig. 12.1) [73]. Furthermore, a novel process for the dissolution of chitin and chitosan using an IL 1-butyl-3-methylimidazolium chloride (BMIMCl) has been developed and reported by Xie et al. [74]. The dissolution behavior of chitin on treating with AMIMBr was reported, as shown in Fig. 12.2 [69]. When chitin was used in the higher amount of 7% (w/w) with AMIMBr and heated up to 100 C, upon cooling at room temperature a gel-like material with higher viscosity was obtained, as shown in Fig. 12.2B. However, when a 5% (w/w) concentration of chitin was used with AMIMBr, dissolved chitin was obtained, as can be seen in Fig. 12.2A. 12.2.1.4 Enhanced solubility of chitin and chitosan on their modification The solubility of chitin and chitosan can be improved/enhanced by appropriate chemical modification [39,75
79]. A common practice involves the introduction of water-soluble functional groups, hydrophilic moieties and hydrophobic bulky groups to enhance the solubility [44,80
84]. An increased solubility of chitosan is reported by Sashiwa and coworkers after its simple acylation [85,86]. Water-soluble chitosan is prepared by simple N-acetylation on treatment with acetic anhydride [87]. The introduction of fatty acids with a hydrophobic nature with polar end groups are considered to enhance the solubility of polymeric

Handbook of Chitin and Chitosan

382

12. Miscibility, properties, and biodegradability of chitin and chitosan

FIGURE 12.2 Images of 5% (w/w) chitin (A) 7% (w/w) chitin (B) with 1-allyl-3methylimidazolium bromide (AMIMBr). Source: Reproduced with permission from Elsevier.

materials. Substitution (N-acylation) of chitosan with fatty acid chlorides with different chain length imparts a hydrophobic character making important changes in its structural features [88]. Treatment of chitosan with n-fatty acid anhydrides using homogeneous aqueous solution of 2 vol.% acetic acid
methanol has been reported to obtain water-soluble polymers [89,90]. Although the introduction of bulky groups increases the solubility, the solubility remains poorer on the substitution of shorter chains of fatty acids. The production of carboxymethyl derivatives of chitin and chitosan, which are highly water soluble, has great potential for various applications. The introduction of hydrophobic methyl groups by this modification disrupts the hydrogen bonding network and provides greater solubility [84,91
94]. Keeping cellulose structural features in mind, the carboxymethylation of chitin was carried out by treating it with monochloroacetic acid using a high concentration of sodium hydroxide [92,94]. Enzymatic hydrolysis is another way to produce low-molecularweight chitosan with high water solubility [95
97]. This method is environment-friendly and mild as it does not need harsh conditions (strong acid, bases, and oxidants) to reduce the crystallinity and molecular weight of chitin. Chitosan-hydrolyzing enzymes (chitinases or chitosanases) have been studied and reported by different authors for partial enzymatic hydrolysis of chitosan as an alternative method to chemical depolymerization [98,99]. The production of chitosan oligomers by the enzymatic deacetylation of chitin has also been studied and reported by Hamer et al. [100].

Handbook of Chitin and Chitosan

12.2 Physicochemical properties of chitin and chitosan

383

12.2.2 Dissolution mechanism 12.2.2.1 Acid-catalyzed hydrolysis A plausible mechanism for the hydrolysis of glycosidic linkage in the presence of acid catalyst is described [101] in Fig. 12.3. In an initial step, glycosidic oxygen atoms get protonated to form a conjugate acid, which further undergo heterolysis of exocyclic O-5 to C-1 bond to make a cyclic carbonium
oxonium ion that exists in a half chair conformation, where C-2, C-1, O-5, and C-5 are in one plane. Further, on contact with a water molecule the carbonium
oxonium ion gets protonated and as a result a reducing sugar is formed. With this mechanism, nonrandom acid-catalyzed hydrolysis of polysaccharides can also be explained, as hydrolytic cleavage of the glycosidic bond at the nonreducing end occurs very rapidly as compared to the glycosidic bond in the chain. From this, it was concluded that the formation of the carbonium
oxonium ion is due to the hydrolysis of the internal glycosidic linkage. 12.2.2.2 Base-catalyzed hydrolysis Dissolution of chitin using base as a catalyst is illustrated [1] in Fig. 12.4. The procedure/mechanism involves the soaking of chitin in an aqueous solution of NaOH (8 wt.%) and urea (4 wt.%) at an ambient temperature. Sodium hydroxide assisted the penetration of water molecules into the chitin molecular chains. Further freezing and expanding of water molecules at the freezing temperature results in the separation of water and NaOH molecules, and the breakdown of inter- and intramolecular hydrogen bonding occurs. This phenomenon enhances the solubility of chitin. However, if the freezing and expansion is carried out below
30 C, it actually shortens the process of freezing and expansion, making the effect of

FIGURE 12.3 Proposed reaction mechanism for acid-catalyzed hydrolysis of glycosidic linkage. Source: Reproduced with permission from Elsevier.

Handbook of Chitin and Chitosan

384

12. Miscibility, properties, and biodegradability of chitin and chitosan

FIGURE 12.4

An illustration of the dissolution process of chitin, including: (A) chitin soaked in an aqueous solution with 8 wt.% NaOH and 4 wt.% urea at ambient temperature; (B) penetration of water molecules into chitin’s molecular chain promoted by NaOH; (C) freezing and then expanding of water molecules at the freezing temperature, breaking the inter- and intrahydrogen bondings; and (D) increasing solubility of the chitin. chitin chain,

hydrogen bond,

sodium ion,

water molecular.

Source: Reproduced with permission from Elsevier.

expansion weaker compared to the treatment carried out at freezing temperature and as a result its solubility gets poor. 12.2.2.3 Cleavage/dissolution by ionic liquids Uto et al. studied the dissolution behavior of chitin crystals in imidazolium-based ILs by adopting a molecular dynamics approach [66]. The dissolution process involves two stages (Fig. 12.5) in the presence of ionic solvents such as AMIMBr, EMIMOAc, BMIMOAc, and AMIMOAc. Initially, anions of ILs play a role to cleave hydrogen bonds among chitin chains by making bridges with their acetamido and hydroxyl functional groups. As a result, chitin chains peel off from the surface of the chitin crystal and begin to disperse into ILs, while the cations of ILs help to prevent their return back onto the crystalline phase after the peeling process occurs in the presence of AMIMBr. However, in the presence of imidazolium acetate solvents, these peeled chains sometimes return back onto the crystalline surface/phase. While with the use of poor solvents like 1-butyl-3-methylmidazoium bromide (BMIMBr), 1-ethyl-3methylimidazolium bromide (EMIMBr), 1-ethyl-3-methylimidazolium chloride (EMIMCl), BMIMCl, and 1-allyl-3-methylimidazolium chloride

Handbook of Chitin and Chitosan

12.3 Biological properties of chitin and chitosan

385

FIGURE 12.5 Dissolution process of chitin crystal models in imidazolium-based ILs. Source: Reproduced with permission from RSC publishers.

(AMIMCl), no chain peeling was observed in chitin crystals as strong hydrogen bonding is maintained by acetamido functional groups (Fig. 12.5).

12.3 Biological properties of chitin and chitosan The biological properties of chitin and chitosan mainly depend upon their physicochemical properties which are determined by the source of the chitin and the conditions used during the production process of chitin and chitosan [102]. Among the physicochemical properties, molecular weight, polydispersity, degree of N-acetylation, crystallinity, and pattern of acetylation are the main key factors that need to be considered [103]. Chitin and chitosan have no solubility in water and in most of the organic solvents. Chitin, chitosan, and their derivatives [chitooligosaccharides (COS)]—obtained by depolymerization of chitin and chitosan via chemical or enzymatic hydrolysis have shown interesting biological properties (antioxidant, anticoagulant, antidiabetic, antiallergic, antiobesity, antihypertensive, antimicrobial, antiinflammatory, neuroprotective, and anticancer) because of their biocompatible, biodegradable, nonallergenic, and nontoxic nature. Hence a number of applications have been found in various industries such as the environmental, pharmaceutical, agriculture, food, and biomedical sectors [104]. It also has been reported that not every chitin or chitosan produced has all these biological activities [103].

Handbook of Chitin and Chitosan

386

12. Miscibility, properties, and biodegradability of chitin and chitosan

Several biological activities of chitin, chitosan, and their derivatives obtained under different conditions are described here.

12.3.1 Antioxidant property Reactive oxygen species such as hydrogen peroxide or hydroxyl radicals are generally produced during normal metabolism, and oxidize biomolecules, such as carbohydrates, proteins, lipids, and DNA, creating oxidative stress [105]. In the absence of cellular defense, this oxidative stress may lead to various health disorders such as aging, rheumatoid arthritis, cancer, and inflammation [106
110]. Among antioxidants, chitosan and its derivatives also have good free radical-scavenging capability to prevent the oxidative damage by terminating free radical chain reaction during the oxidation process [104]. Moreover, COS with less than 1 kDa were found to retard or terminate the formation of intracellular radical species in B16F1, a murine melanoma cell line, indicating the prevention of oxidative stress-related disease [111]. An increased antioxidant defense capacity was observed in a sow’s diet upon addition of COS as a supplement and also facilitated the placental amino acid transport. As a result, fetus development and sow’s health was improved during gestation [112]. Chang et al. investigated seven chitosan derivatives obtained by enzymatic degradation with molecular weight ranging from 2.2 to 300 kDa exhibiting antioxidant and antimutagenic properties. It was also mentioned that these properties of chitosan derivatives were inversely related to their molecular weight [113]. Kang et al. prepared a chitosan-based interpenetrating polymer network (IPN) hydrogel by coupling gallic acid with the amino functional group on its surface via an amide coupling reaction and reported it to be antioxidative. They mentioned that hydrogels modified with gallic acid with a longer chain of chitosan backbone demonstrated higher antioxidant activity compared with those with a shorter chitosan moiety [114]. The protective effect of COS against H2O2-induced oxidative stress on human embryonic hepatocytes (L02 cells) as well as its in vitro scavenging activity against DPPH (1,1-diphenyl-2-picrylhydrazyl) radical was investigated by Xu et al. It was suggested from obtained results that COS can be helpful in the clinical setting for the treatment of oxidative stress-related liver damage [115].

12.3.2 Anticancer/antitumor property Cancer is a chronic disease in humans, arising due to changes in nutrition, lifestyle, and global warming. Free radicals are responsible for

Handbook of Chitin and Chitosan

12.3 Biological properties of chitin and chitosan

387

the proliferation of cancer in human. Chemopreventive compounds as anticancer drugs have attained considerable popularity to treat cancer diseases [116]. The preparation of chitin
glucan
aldehyde
quercetin conjugate has been recently reported by a condensation process and was evaluated for antioxidant and anticancer activity. The prepared conjugate displayed strong DPPH-scavenging activity and high cytotoxicity against J774 cell lines without affecting peripheral blood mononuclear cells [117]. In another study, production of COS from chitosan hydrogel via degradation was reported and investigated for anticancer activity. The prepared water-soluble chitosan derivatives were found to exhibit the best inhibitory effect against human lung cancer cells (H460) [118]. Chien et al. obtained chitosan by N-deacetylation of chitin from shiitake stipes and crab shells under alkaline conditions and evaluated its antiproliferative effect on IMR 32 and Hep G2 cells. At a concentration of 5 mg/mL, 68.8%
85.0% cell viability of IMR 32 and 60.4%
82.9% of Hep G2 cells was shown when incubated with chitosan [119].

12.3.3 Antimicrobial property Chitin, chitosan, and their derivatives have gained noticeable attention due to their potential as antimicrobial agents against various microorganisms such as yeast, bacteria, and fungi. Rahman et al. [120] reported the antifungal activity of chitosan with different degrees of polymerization obtained by partial N-deacetylation of chitin. This study showed that COS have shown the potential to inhibit fungal growth as well as germination. They ascribed this effect to the level of polymerization of the prepared oligomers. Chien et al. reported high antimicrobial activities of N-deacetylated chitin against eight different species of Gram-negative and Gram-positive pathogenic bacteria showing inhibition zones of 11.4
26.8 mm at a concentration of 0.5 mg/mL [119]. Zhong et al. prepared 12 different kinds of new hydroxylbenzenesulfonailides derivatives by reacting chitosan, carboxymethyl chitosan, and chitosan sulfates of different molecular weight with 2-hydroxyl-5-chloride1,3-benzene-disulfochloride or 4-hydroxyl-5-chloride-1,3-benzene-disulfochloride and evaluating their antimicrobial activity. They reported increased antimicrobial activity of the prepared derivatives against four bacteria (Escherichia coli, Staphylococcus aureus, Pseudomonas aeruginosa, and Sarcina) and five crop-threatening pathogenic fungi named as Phomopsis asparagi, Fusarium oxysporum, Aiternaria solani, Vasinfectum, and Colletotrichum gloeosporioides [121]. Yang et al. prepared N-alkylated disaccharides from chitosan and studied their antibacterial activity against Staphylococcus aureus and

Handbook of Chitin and Chitosan

388

12. Miscibility, properties, and biodegradability of chitin and chitosan

Escherichia coli. An increase in antibacterial activity of chitosan derivatives (30%
40% degree of substitution) against Escherichia coli was observed when pH was raised from 5 to 7
7.5, while no significant effect of pH was observed in the antibacterial activity of N-alkylated disaccharides against Staphylococcus aureus.

12.3.4 Antiinflammatory property Numerous studies have been reported on the biological activities of chitin, chitosan, and their derivatives, while a few investigations have been published recently on their antiinflammatory activity. Inflammation is an immune response of the body against toxic chemicals, pathogens, and physical damage. Prolonged inflammation can be dangerous and result in the development of various diseases, particularly rheumatoid arthritis, chronic asthma, inflammatory bowel disease, multiple sclerosis, psoriasis, and cancer [104]. Preparation of chitosan
alginate nanoparticles loaded with zaltoprofen (a nonsteroidal antiinflammatory drugs) using an ionotropic gelation technique were reported by Hirva et al. [122]. The purpose of the study was to release the drug in a sustained manner to minimize side effects and enhance bioavailability. Their results indicated that the chitosan
alginate nanoparticles loaded with zaltoprofen displayed prolonged (10 h) antiinflammatory activity compared with the free zaltoprofen. Chung et al. reported the preparation of low-molecular-weight chitosan derivatives ( , 1 kDa) by enzymatic digestion process and evaluated their in vitro and in vivo study against allergic asthma and allergic reaction. In their study, they suggested low-molecular-weight chitosan derivatives as a potential candidate in the development of therapeutic drugs to treat allergic asthma [123]. The therapeutic potential of COS with more than 90% degree of deacetylation with molecular weight in the range of 5
10 kDa for the treatment of inflammatory bowel disease has been investigated by Yousef et al. The COS were found to be effective in hindering intestinal inflammation and mortality in mice of 5% dextran sulfate sodium (DSS)induced acute colitis [124].

12.3.5 Neuroprotective property Neuronal cells are highly vulnerable and have a confined capacity of self-repair when they come under the influence of physical damage or injury. Procedures and strategies utilized in order to provide protection

Handbook of Chitin and Chitosan

12.4 Biodegradability of chitin and chitosan

389

against neural injury, dysfunction, apoptosis, and degeneration in the central nervous system are known as neuroprotection [125]. Chitosan and its derivatives (COS) are found to exhibit enzyme inhibitory activities and are considered to be novel therapeutic agents for the treatment of Alzheimer’s and Parkinson’s diseases in the pharmaceutical industry [104]. Acetylcholinesterase inhibitory property was evaluated for chitosan derivatives obtained by grafting different ratios of gallic acid, as studied by Cho et al. Gallate chitosan derivatives displayed potent acetylcholinesterase inhibitory activity with IC50 values in the range of 138.5 6 2.5 to 397.6 6 5.2 μg/mL [126]. Huang et al. investigated the neuroprotective property of COS (with a degree of polymerization , 10) in a neuronal cell line of humans and reported them to be potential therapeutic agents against Alzheimer’s disease [127]. Jiang et al. prepared five different deacetylated COS by degradation of chitosan and evaluated their neuroprotective activity. Among them, chitotriose was found to induce the highest increase in Schwann cell survival [128]. Dai et al. reported that COS derivatives with 2
7 degree of polymerization, and more than 90% degree of deacetylation remarkably inhibited the cell death induced by β-amyloidpeptide (Aβ) exposure as it was measured by lactate dehydrogenase release assay and cell viability assay [129]. In a recent study, Moriano et al. reported the synthesis of COS by enzymatic hydrolysis of chitosan and evaluated their in vitro neuroprotective (toward human SH-S5Y5 neurons) and antiinflammatory activities. They concluded that a mixture of COS demonstrated moderate biological activity and no toxicity to neurons and RAW macrophage cells [130]. Je et al. reported the synthesis of heterochitooligosaccharides and sulfonated COS (SCOS) varying in their molecular sizes and degree of depolymerization (50%, 75%, and 90%) by using an ultrafiltration membrane reactor; they investigated their prolyl endopeptidase (PEP) inhibitory activity (to treat mental disorders such as Alzheimer’s and Parkinson’s diseases). Higher PEP inhibitory activity was observed in the case of SCOS with 50% deacetylation compared with those SCOS with 75% and 90% deacetylation. From this study, they concluded that SCOS with a molecular weight of 1
5 kDa and 50% deacetylation that showed PEP inhibitory activity have the potential to be considered for the development of new PEP inhibitors from carbohydrate material [131].

12.4 Biodegradability of chitin and chitosan Biodegradation is a process which involves the decomposition of organic material/polymer in aerobic and anaerobic conditions by the action of microorganisms. This process completely spoils the physical

Handbook of Chitin and Chitosan

390

12. Miscibility, properties, and biodegradability of chitin and chitosan

and chemical properties of the polymer as a result CO2, H2O, CH4, and other low-molecular-weight products are formed [132]. Biodegradation involves physical (sorption, swelling, dissolution, crystallization, mineralization, impact facture, and wear) or chemical (hydrolysis, enzymolysis, oxidation, and oxidation chain cleavage) mechanisms, but is not limited to only one kind of process [133]. Chitin, chitosan, and their derivatives have been reported as biocompatible and biodegradable materials. Degradation of chitosan can be achieved by enzymes which hydrolyze components (glucosamine
glucosamine, glucosamine
N-acetyl-glucosamine, and N-acetyl-glucosamine
N-acetylglucosamine) linkages. Generally in microorganisms, chitinase enzymes, known as endo-chitinases (EC 3.2.1.14), hydrolyze N-acetyl-β-1,4-glucosaminide linkages randomly [134]. An efficient degradation was observed by lysozyme for chitosan, where a 66% loss in viscosity was obtained in the case of acetylated chitosan (50% acetylation) when incubated in vitro for 4 h at 37 C and pH of 5.5 (0.1 M phosphate buffer, 0.2 M NaCl) [135]. In addition to the extent of DA, other modifications, including covalent cross-linking and thiolation, have also displayed an effect on the degradation phenomenon [136,137]. A number of protease enzymes have shown different rates of degradation on chitosan films and leucine aminopeptidase was found to be most effective, displaying 38% degradation of film in 30 days [138]. Degradation of chitosan to low-molecular-weight chitosan was also observed by pectinase isozyme (from Aspergilus niger) at low pH [139,140]. Ibrahim et al. reported the degradation of films prepared by using 10% (by weight) of chitin and chitosan individually with polyethylene. Films were incubated at 25 C in a basal medium with no source of nitrogen and carbon under two different types of environment (using pure microbial culture and soil environment). In this study, they concluded that in the soil environment, both polyethylene
chitin and polyethylene
chitosan films showed a higher rate of degradation compared with commercial starch-based film, suggesting the use of chitin-based films as promising material for the development of biodegradable packaging [141]. Another author studied and reported the biodegradation behavior of films named as chitosan-graft-polymethylmethacrylate (C-g-PMMA), prepared by graft copolymerization of methylmethacrylate on chitosan. They reported 40%
45% degradation of graft films by fungus (Aspergillus flavus) over 5
25 days of aerobic cultivation. SEM images of the prepared films taken during the biodegradation process are shown in Fig. 12.6. It can be seen clearly from the images that the chitosan in the treated samples started degrading by Aspergillus flavus within the first 24 h and reached the maximum in around 20 days, whereas no morphological changes were observed in the case of uninoculated film [142] (Fig. 12.6).

Handbook of Chitin and Chitosan

12.4 Biodegradability of chitin and chitosan

391

SEM ( 3 650) biodegradation images of C-g-PMMA films incubated with Aspergillus flavus. Source: Reproduced with permission from Elsevier.

FIGURE 12.6

12.4.1 factors affecting the biodegradability Several factors have been reported and discussed above which can affect the degradation rate of chitin, chitosan, and their derivatives. Those factors include but are not limited to: 1. 2. 3. 4. 5. 6.

degree of depolymerization; molecular weight or extent of crystallinity; DA or N-substitution; temperature/environmental conditions; chemical modification and cross-linking agents; and type of enzymes/microorganism.

The degradation procedure of chitin/chitosan could be considered easier to target compared with other polymers such as humic acid, cellulose, and structurally heterogeneous lignin. This is only because of chitin/chitosan’s simple structure and the existence of a primer system targeting chitin-modifying enzymes. Therefore the degradation of chitin

Handbook of Chitin and Chitosan

392

12. Miscibility, properties, and biodegradability of chitin and chitosan

can be considered as a general model to be explored for understanding microbial mechanisms of biopolymers degradation in the biosphere [134,143].

12.5 Concluding remarks Chitin and chitosan are natural polysaccharides, which can be modified or altered either chemically, physically, or enzymatically to overcome their limitations, such as poor solubility, high crystallinity, higher molecular weight, and DA, in order to enhance their potential application in various industries, especially in the biomedical or pharmaceutical sectors. Various factors have been discussed in this chapter which affect the solubility of chitin including DA, molecular weight or size of the polymer, chemical modification, solvent nature, and temperature. Polymers with a lower DA, molecular weight, and crystallinity have been reported to have higher solubility in water and most of the solvents. Water-soluble derivatives of chitin and chitosan are biodegradable, renewable, and biocompatible, thus displaying unique biological properties. Various characteristics of polysaccharides (chitin and chitosan), such as their physical, chemical, and biological properties, as well as their biodegradability, have been described in detail to provide readers a better insight into the importance of chitin and chitosan.

References [1] X. Hu, Y. Du, Y. Tang, Q. Wang, T. Feng, J. Yang, et al., Solubility and property of chitin in NaOH/urea aqueous solution, Carbohydr. Polym. 70 (2007) 451
458. [2] P.R. Rege, L.H. Block, Chitosan processing: influence of process parameters during acidic and alkaline hydrolysis and effect of the processing sequence on the resultant chitosan’s properties, Carbohydr. Res. 321 (1999) 235
245. [3] A. Tolaimate, J. Desbrie`res, M. Rhazi, A. Alagui, M. Vincendon, P. Vottero, On the influence of deacetylation process on the physicochemical characteristics of chitosan from squid chitin, Polymer 41 (2000) 2463
2469. [4] L. Chen, Y. Du, H. Wu, L. Xiao, Relationship between molecular structure and moisture-retention ability of carboxymethyl chitin and chitosan, J. Appl. Polym. Sci. 83 (2002) 1233
1241. [5] S.M. Hudson, C. Smith, Polysaccharides: chitin and chitosan: chemistry and technology of their use as structural materials, in: D.L. Kaplan (Ed.), Biopolymers From Renewable Resources, Springer Berlin Heidelberg, Berlin, Heidelberg, 1998, pp. 96
118. [6] T.R.A. Sobahi, M.Y. Abdelaal, M.S.I. Makki, Chemical modification of chitosan for metal ion removal, Arab. J. Chem. 7 (2014) 741
746. [7] Z. Karim, A.P. Mathew, M. Grahn, J. Mouzon, K. Oksman, Nanoporous membranes with cellulose nanocrystals as functional entity in chitosan: removal of dyes from water, Carbohydr. Polym. 112 (2014) 668
676.

Handbook of Chitin and Chitosan

References

393

[8] T. Uragami, T. Saito, T. Miyata, Pervaporative dehydration characteristics of an ethanol/water azeotrope through various chitosan membranes, Carbohydr. Polym. 120 (2015) 1
6. [9] P. Chantarasataporn, P. Tepkasikul, Y. Kingcha, R. Yoksan, R. Pichyangkura, W. Visessanguan, et al., Water-based oligochitosan and nanowhisker chitosan as potential food preservatives for shelf-life extension of minced pork, Food Chem. 159 (2014) 463
470. [10] D. Chen, B. Hu, C. Huang, Chitosan modified ordered mesoporous silica as microcolumn packing materials for on-line flow injection-inductively coupled plasma optical emission spectrometry determination of trace heavy metals in environmental water samples, Talanta 78 (2009) 491
497. [11] W. Zhang, J. Zhang, Q. Jiang, W. Xia, The hypolipidemic activity of chitosan nanopowder prepared by ultrafine milling, Carbohydr. Polym. 95 (2013) 487
491. [12] E.P. Minet, C. O’Carroll, D. Rooney, C. Breslin, C.P. McCarthy, L. Gallagher, et al., Slow delivery of a nitrification inhibitor (dicyandiamide) to soil using a biodegradable hydrogel of chitosan, Chemosphere 93 (2013) 2854
2858. ´ .M.H. Caballero, C. Schmidt, C.P. Covas, N, [13] J.P. Quin˜ones, K.V. Gothelf, J. Kjems, A O6-partially acetylated chitosan nanoparticles hydrophobically-modified for controlled release of steroids and vitamin E, Carbohydr. Polym. 91 (2013) 143
151. [14] Y.A. Gomaa, L.K. El-Khordagui, N.A. Boraei, I.A. Darwish, Chitosan microparticles incorporating a hydrophilic sunscreen agent, Carbohydr. Polym. 81 (2010) 234
242. [15] N. Mati-Baouche, P.-H. Elchinger, H. de Baynast, G. Pierre, C. Delattre, P. Michaud, Chitosan as an adhesive, Eur. Polym. J. 60 (2014) 198
212. [16] J.-P. Wang, Y.-Z. Chen, S.-J. Yuan, G.-P. Sheng, H.-Q. Yu, Synthesis and characterization of a novel cationic chitosan-based flocculant with a high water-solubility for pulp mill wastewater treatment, Water Res. 43 (2009) 5267
5275. [17] K. Madhumathi, P.T. Sudheesh Kumar, S. Abhilash, V. Sreeja, H. Tamura, K. Manzoor, et al., Development of novel chitin/nanosilver composite scaffolds for wound dressing applications, J. Mater. Sci.: Mater. Med. 21 (2010) 807
813. [18] R. Jayakumar, M. Prabaharan, R.L. Reis, J.F. Mano, Graft copolymerized chitosan— present status and applications, Carbohydr. Polym. 62 (2005) 142
158. [19] K. Madhumathi, N.S. Binulal, H. Nagahama, H. Tamura, K.T. Shalumon, N. Selvamurugan, et al., Preparation and characterization of novel β-chitin
hydroxyapatite composite membranes for tissue engineering applications, Int. J. Biol. Macromol. 44 (2009) 1
5. [20] G. Borchard, Chitosans for gene delivery, Adv. Drug. Deliv. Rev. 52 (2001) 145
150. [21] R. Jayakumar, M. Prabaharan, S.V. Nair, H. Tamura, Novel chitin and chitosan nanofibers in biomedical applications, Biotechnol. Adv. 28 (2010) 142
150. [22] H. Nagahama, N. Nwe, R. Jayakumar, S. Koiwa, T. Furuike, H. Tamura, Novel biodegradable chitin membranes for tissue engineering applications, Carbohydr. Polym. 73 (2008) 295
302. [23] Y. Wang, H. Chen, J. Wang, L. Xing, Preparation of active corn peptides from zein through double enzymes immobilized with calcium alginate
chitosan beads, Process. Biochem. 49 (2014) 1682
1690. [24] K.T. Shalumon, N.S. Binulal, N. Selvamurugan, S.V. Nair, D. Menon, T. Furuike, et al., Electrospinning of carboxymethyl chitin/poly(vinyl alcohol) nanofibrous scaffolds for tissue engineering applications, Carbohydr. Polym. 77 (2009) 863
869. [25] R. Jayakumar, K.P. Chennazhi, R.A.A. Muzzarelli, H. Tamura, S.V. Nair, N. Selvamurugan, Chitosan conjugated DNA nanoparticles in gene therapy, Carbohydr. Polym. 79 (2010) 1
8.

Handbook of Chitin and Chitosan

394

12. Miscibility, properties, and biodegradability of chitin and chitosan

[26] K. Madhumathi, P.T. Sudheesh Kumar, K.C. Kavya, T. Furuike, H. Tamura, S.V. Nair, et al., Novel chitin/nanosilica composite scaffolds for bone tissue engineering applications, Int. J. Biol. Macromol. 45 (2009) 289
292. [27] M. Peter, N.S. Binulal, S.V. Nair, N. Selvamurugan, H. Tamura, R. Jayakumar, Novel biodegradable chitosan
gelatin/nano-bioactive glass ceramic composite scaffolds for alveolar bone tissue engineering, Chem. Eng. J. 158 (2010) 353
361. [28] M. Peter, P.T. Sudheesh Kumar, N.S. Binulal, S.V. Nair, H. Tamura, R. Jayakumar, Development of novel α-chitin/nanobioactive glass ceramic composite scaffolds for tissue engineering applications, Carbohydr. Polym. 78 (2009) 926
931. [29] K. Muramatsu, S. Masuda, Y. Yoshihara, A. Fujisawa, In vitro degradation behavior of freeze-dried carboxymethyl-chitin sponges processed by vacuum-heating and gamma irradiation, Polym. Degrad. Stab. 81 (2003) 327
332. [30] M. Prabaharan, J.F. Mano, Chitosan-based particles as controlled drug delivery systems, Drug. Deliv. 12 (2004) 41
57. [31] A. Anitha, V.V. Divya Rani, R. Krishna, V. Sreeja, N. Selvamurugan, S.V. Nair, et al., Synthesis, characterization, cytotoxicity and antibacterial studies of chitosan, O-carboxymethyl and N,O-carboxymethyl chitosan nanoparticles, Carbohydr. Polym. 78 (2009) 672
677. [32] J. Cai, J. Yang, Y. Du, L. Fan, Y. Qiu, J. Li, et al., Purification and characterization of chitin deacetylase from Scopulariopsis brevicaulis, Carbohydr. Polym. 65 (2006) 211
217. [33] A. Ghosh, M. Azam Ali, R. Walls, Modification of microstructural morphology and physical performance of chitosan films, Int. J. Biol. Macromol. 46 (2010) 179
186. [34] A. Inmaculada, M. Marian, H. Ruth, P. Ines, M. Beatriz, A. Niuris, et al., Functional characterization of chitin and chitosan, Curr. Chem. Biol. 3 (2009) 203
230. [35] K. Kurita, Chitin and chitosan: functional biopolymers from marine crustaceans, Mar. Biotechnol. 8 (2006) 203
226. [36] K. Kurita, S. Mori, Y. Nishiyama, M. Harata, N-alkylation of chitin and some characteristics of the novel derivatives, Polym. Bull. 48 (2002) 159
166. [37] S. Santhosh, T.K. Sini, P.T. Mathew, Variation in properties of chitosan prepared at different alkali concentrations from squid pen and shrimp shell, Int. J. Polym. Mater. 59 (2010) 286
291. [38] D.N.S. Hon, Chitin and chitosan: medical applications, Polysaccharides medicinal appl. part II (1996) 631
651. [39] H. Yi, L.Q. Wu, W.E. Bentley, R. Ghodssi, G.W. Rubloff, J.N. Culver, et al., Biofabrication with chitosan, Biomacromolecules 6 (2005) 2881
2894. [40] K.M. Kim, J.H. Son, S.K. Kim, C.L. Weller, M.A. Hanna, Properties of chitosan films as a function of pH and solvent type, J. Food Sci. 71 (2006) E119
E124. [41] M. Rinaudo, G. Pavlov, J. Desbrie`res, Influence of acetic acid concentration on the solubilization of chitosan, Polymer 40 (1999) 7029
7032. [42] C. Kienzle-Sterzer, D. Rodriguez-Sanchez, C. Rha, Dilute solution behavior of a cationic polyelectrolyte, J. Appl. Polym. Sci. 27 (1982) 4467
4470. [43] M. Rinaudo, G. Pavlov, J. Desbrie`res, Solubilization of chitosan in strong acid medium, Int. J. Polym. Anal. Charact. 5 (1999) 267
276. [44] C.K.S. Pillai, W. Paul, C.P. Sharma, Chitin and chitosan polymers: chemistry, solubility and fiber formation, Prog. Polym. Sci. 34 (2009) 641
678. [45] A. Domard, M. Rinaudo, Preparation and characterization of fully deacetylated chitosan, Int. J. Biol. Macromol. 5 (1983) 49
52. [46] N. Kubota, Y. Eguchi, Facile preparation of water-soluble N-acetylated chitosan and molecular weight dependence of its water-solubility, Polym. J. 29 (1997) 123
127.

Handbook of Chitin and Chitosan

References

395

[47] K. Kurita, M. Kamiya, S.I. Nishimura, Solubilization of a rigid polysaccharide: controlled partial N-acetylation of chitosan to develop solubility, Carbohydr. Polym. 16 (1991) 83
92. [48] C. Peniche, W. Argu¨elles-Monal, Overview on structural characterization of chitosan molecules in relation with their behavior in solution, Macromol. Symp. 168 (2001) 1
20. [49] G. Ru, S. Wu, X. Yan, B. Liu, P. Gong, L. Wang, et al., Inverse solubility of chitin/ chitosan in aqueous alkali solvents at low temperature, Carbohydr. Polym. 206 (2019) 487
492. [50] A. Chenite, M. Buschmann, D. Wang, C. Chaput, N. Kandani, Rheological characterisation of thermogelling chitosan/glycerol-phosphate solutions, Carbohydr. Polym. 46 (2001) 39
47. [51] A. Chenite, C. Chaput, D. Wang, C. Combes, M.D. Buschmann, C.D. Hoemann, et al., Novel injectable neutral solutions of chitosan form biodegradable gels in situ, Biomaterials 21 (2000) 2155
2161. [52] J. Cho, M.C. Heuzey, A. Be´gin, P.J. Carreau, Physical gelation of chitosan in the presence of β-glycerophosphate: the effect of temperature, Biomacromolecules 6 (2005) 3267
3275. [53] J. Du, Y.L. Hsieh, PEGylation of chitosan for improved solubility and fiber formation via electrospinning, Cellulose 14 (2007) 543
552. [54] R.A.A. Muzzarelli, Chitin and its derivatives: new trends of applied research, Carbohydr. Polym. 3 (1983) 53
75. [55] R. Seoudi, A.M.A. Nada, Molecular structure and dielectric properties studies of chitin and its treated by acid, base and hypochlorite, Carbohydr. Polym. 68 (2007) 728
733. [56] P.P.V. Weimarn, Conversion of fibroin, 1chitin, casein, and similar substances into the ropy-plastic state and colloidal solution2, Ind. Eng. Chem. 19 (1927) 109
110. [57] G. Kunike, Chitin and chitin silk, J. Soc. Dyers Colour. 42 (1926) 318
342. [58] I.V. Serov, A.M. Bochek, N.P. Novoselov, N.M. Zabivalova, V.K. Lavrent0 ev, E.N. Vlasova, et al., Chitin in aqueous alkaline solutions with urea and thiourea additives and the structures of films obtained from them, Fibre Chem. 47 (2015) 247
250. [59] C.J. Brine, P.R. Austin, Renatured chitin fibrils, films, and filaments, ACS Symp. Ser. 18 (1975) 505
518. [60] S. Tokura, N. Nishi, J. Noguchi, Studies on chitin. III. Preparation of chitin fibers, Polym. J. 11 (1979) 781
786. [61] M. Isik, H. Sardon, D. Mecerreyes, Ionic liquids and cellulose: dissolution, chemical modification and preparation of new cellulosic materials, Int. J. Mol. Sci. 15 (2014) 11922. [62] M.M. Jaworska, T. Kozlecki, A. Gorak, Review of the application of ionic liquids as solvents for chitin, J. Polym. Eng. 32 (2012) 67
69. [63] S.S. Silva, J.F. Mano, R.L. Reis, Ionic liquids in the processing and chemical modification of chitin and chitosan for biomedical applications, Green. Chem. 19 (2017) 1208
1220. [64] W.-T. Wang, J. Zhu, X.-L. Wang, Y. Huang, Y.-Z. Wang, Dissolution behavior of chitin in ionic liquids, J. Macromol. Sci., Part B 49 (2010) 528
541. [65] M.E. Zakrzewska, E. Bogel-Łukasik, R. Bogel-Łukasik, Solubility of carbohydrates in ionic liquids, Energ. Fuel. 24 (2010) 737
745. [66] T. Uto, S. Idenoue, K. Yamamoto, J.-i. Kadokawa, Understanding dissolution process of chitin crystal in ionic liquids: theoretical study, Phys. Chem. Chem. Phys. 20 (2018) 20669
20677. [67] J.-I. Kadokawa, Ionic liquid as useful media for dissolution, derivatization, and nanomaterial processing of chitin, Green Sustain. Chem. 03 (02) (2013) 7. [68] J.-i. Kadokawa, Fabrication of nanostructured and microstructured chitin materials through gelation with suitable dispersion media, RSC Adv. 5 (2015) 12736
12746.

Handbook of Chitin and Chitosan

396

12. Miscibility, properties, and biodegradability of chitin and chitosan

[69] K. Prasad, M.-a. Murakami, Y. Kaneko, A. Takada, Y. Nakamura, J.-i. Kadokawa, Weak gel of chitin with ionic liquid, 1-allyl-3-methylimidazolium bromide, Int. J. Biol. Macromol. 45 (2009) 221
225. [70] J.-i. Kadokawa, Dissolution, derivatization, and functionalization of chitin in ionic liquid, Int. J. Biol. Macromol. 123 (2019) 732
737. [71] Y. Qin, X. Lu, N. Sun, R.D. Rogers, Dissolution or extraction of crustacean shells using ionic liquids to obtain high molecular weight purified chitin and direct production of chitin films and fibers, Green. Chem. 12 (2010) 968
971. [72] Y. Wu, T. Sasaki, S. Irie, K. Sakurai, A novel biomass-ionic liquid platform for the utilization of native chitin, Polymer 49 (2008) 2321
2327. [73] P. Walther, A. Ota, A. Mu¨ller, F. Hermanutz, F. Ga¨hr, M.R. Buchmeiser, Chitin foils and coatings prepared from ionic liquids, Macromol. Mater. Eng. 301 (2016) 1337
1344. [74] H. Xie, S. Zhang, S. Li, Chitin and chitosan dissolved in ionic liquids as reversible sorbents of CO2, Green. Chem. 8 (2006) 630
633. [75] R. Jayakumar, R.L. Reis, J.F. Mano, Chemistry and applications of phosphorylated chitin and chitosan, E-Polymers (2006) 1
16. [76] M.N.V. Ravi Kumar, A review of chitin and chitosan applications, React. Funct. Polym. 46 (2000) 1
27. [77] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. (Oxf.) 31 (2006) 603
632. [78] H. Sashiwa, S.I. Aiba, Chemically modified chitin and chitosan as biomaterials, Prog. Polym. Sci. (Oxf.) 29 (2004) 887
908. [79] V.A. Vasnev, A.I. Tarasov, G.D. Markova, S.V. Vinogradova, O.G. Garkusha, Synthesis and properties of acylated chitin and chitosan derivatives, Carbohydr. Polym. 64 (2006) 184
189. [80] S. Hirano, Chitin and chitosan as novel biotechnological materials, Polym. Int. 48 (1999) 732
734. [81] N. Ma, Q. Wang, S. Sun, A. Wang, Progress in chemical modification of chitin and chitosan, Prog. Chem. 16 (2004) 643
653. [82] M. Morimoto, H. Saimoto, Y. Shigemasa, Control of functions of chitin and chitosan by chemical modification, Trends Glycosci. Glyc. 14 (2002) 205
222. [83] K.V.H. Prashanth, R.N. Tharanathan, Preparation and properties of reduced chitooligomers, Trends Food Sci. Tech. 18 (2007) 117. [84] M. Zhang, H.X. Ren, Structural modification and application of chitosan, J. Clin. Rehab. Tissue Eng. Res. 11 (2007) 9817
9820. [85] H. Sashiwa, N. Kawasaki, A. Nakayama, E. Muraki, N. Yamamoto, S.I. Aiba, Chemical modification of chitosan. 14:1 synthesis of water-soluble chitosan derivatives by simple acetylation, Biomacromolecules 3 (2002) 1126
1128. [86] H. Sashiwa, Y. Shigemasa, Chemical modification of chitin and chitosan 2: preparation and water soluble property of N-acylated or N-alkylated partially deacetylated chitins, Carbohydr. Polym. 39 (1999) 127
138. [87] G. Qun, W. Ajun, Z. Yong, Effect of reacetylation and degradation on the chemical and crystal structures of chitosan, J. Appl. Polym. Sci. 104 (2007) 2720
2728. [88] C. Le Tien, M. Lacroix, P. Ispas-Szabo, M.A. Mateescu, N-acylated chitosan: hydrophobic matrices for controlled drug release, J. Control. Rel. 93 (2003) 1
13. [89] S. Hirano, Y. Yamaguchi, M. Kamiya, Novel N-saturated-fatty-acyl derivatives of chitosan soluble in water and in aqueous acid and alkaline solutions, Carbohydr. Polym. 48 (2002) 203
207. [90] S. Hirano, Y. Yamaguchi, M. Kamiya, et al., N-(n-Fatty acyl)chitosans, Macromol. Biosci. 3 (2003) 629
631.

Handbook of Chitin and Chitosan

References

397

[91] Z. Li, X. Zhuang, X. Liu, Y. Guan, K. Yao, Rheological study on O-carboxymethylated chitosan/ cellulose polyblends from LiCl/N,N-dimethylacetamide solution, J. Appl. Polym. Sci. 88 (2003) 1719
1725. [92] S. Santhosh, P.T. Mathew, Preparation and properties of glucosamine and carboxymethylchitin from shrimp shell, J. Appl. Polym. Sci. 107 (2008) 280
285. [93] T.K. Sini, S. Santhosh, P.T. Mathew, Study of the influence of processing parameters on the production of carboxymethylchitin, Polymer 46 (2005) 3128
3131. [94] J.M. Wasikiewicz, H. Mitomo, N. Nagasawa, T. Yagi, M. Tamada, F. Yoshii, Radiation crosslinking of biodegradable carboxymethylchitin and carboxymethylchitosan, J. Appl. Polym. Sci. 102 (2006) 758
767. [95] C. Qin, Y. Du, L. Xiao, Z. Li, X. Gao, Enzymic preparation of water-soluble chitosan and their antitumor activity, Int. J. Biol. Macromol. 31 (2002) 111
117. [96] G. Villa-Lerma, H. Gonza´lez-Ma´rquez, M. Gimeno, S. Trombotto, L. David, S. Ifuku, et al., Enzymatic hydrolysis of chitin pretreated by rapid depressurization from supercritical 1,1,1,2-tetrafluoroethane toward highly acetylated oligosaccharides, Bioresour. Technol. 209 (2016) 180
186. [97] A. Zhang, G. Wei, X. Mo, N. Zhou, K. Chen, P. Ouyang, Enzymatic hydrolysis of chitin pretreated by bacterial fermentation to obtain pure N-acetyl-D-glucosamine, Green. Chem. 20 (2018) 2320
2327. [98] A. Sørbotten, S.J. Horn, V.G.H. Eijsink, K.M. Va˚rum, Degradation of chitosans with chitinase B from Serratia marcescens, FEBS J. 272 (2005) 538
549. [99] K.M. Va˚rum, H. Kristiansen Holme, M. Izume, B. Torger Stokke, O. Smidsrød, Determination of enzymatic hydrolysis specificity of partially N-acetylated chitosans, Biochim. Biophys. Acta 1291 (1996) 5
15. [100] S.N. Hamer, S. Cord-Landwehr, X. Biarne´s, A. Planas, H. Waegeman, B.M. Moerschbacher, et al., Enzymatic production of defined chitosan oligomers with a specific pattern of acetylation using a combination of chitin oligosaccharide deacetylases, Sci. Rep. 5 (2015) 8716. [101] A. Einbu, H. Grasdalen, K.M. Va˚rum, Kinetics of hydrolysis of chitin/chitosan oligomers in concentrated hydrochloric acid, Carbohydr. Res. 342 (2007) 1055
1062. [102] J. Kumirska, M.X. Weinhold, J. Tho¨ming, P. Stepnowski, Biomedical activity of chitin/chitosan based materials—influence of physicochemical properties apart from molecular weight and degree of N-acetylation, Polymers 3 (2011) 1875. [103] M.H. Struszczyk, Chitin and chitosan: part I. Properties and production, Polimery/ Polymers 47 (2002) 316
325. [104] D.-H. Ngo, T.-S. Vo, D.-N. Ngo, K.-H. Kang, J.-Y. Je, H.N.-D. Pham, et al., Biological effects of chitosan and its derivatives, Food Hydrocoll. 51 (2015) 200
216. [105] I. Rahman, S.K. Biswas, A. Kode, Oxidant and antioxidant balance in the airways and airway diseases, Eur. J. Pharmacol. 533 (2006) 222
239. [106] M.V. Blagosklonny, Aging: ROS or TOR, Cell Cycle 7 (2008) 3344
3354. [107] P.S. Leung, Y.C. Chan, Role of oxidative stress in pancreatic inflammation, Antioxid. Redox Sign. 11 (2009) 135
166. [108] S. Maynard, S.H. Schurman, C. Harboe, N.C. de Souza-Pinto, V.A. Bohr, Base excision repair of oxidative DNA damage and association with cancer and aging, Carcinogenesis 30 (2009) 2
10. [109] A. Mirshafiey, M. Mohsenzadegan, The role of reactive oxygen species in immunopathogenesis of rheumatoid arthritis, Iran. J. Allergy, Asthm. 7 (2008) 195
202. [110] S. Pillai, C. Oresajo, J. Hayward, Ultraviolet radiation and skin aging: roles of reactive oxygen species, inflammation and protease activation, and strategies for prevention of inflammation-induced matrix degradation—a review, Int. J. Cosmet. Sci. 27 (2005) 17
34.

Handbook of Chitin and Chitosan

398

12. Miscibility, properties, and biodegradability of chitin and chitosan

[111] E. Mendis, M.-M. Kim, N. Rajapakse, S.-K. Kim, An in vitro cellular analysis of the radical scavenging efficacy of chitooligosaccharides, Life Sci. 80 (2007) 2118
2127. [112] Y. Xie, S.H. Ho, C.N.N. Chen, C.Y. Chen, K. Jing, I.S. Ng, et al., Disruption of thermo-tolerant Desmodesmus sp. F51 in high pressure homogenization as a prelude to carotenoids extraction, Biochem. Eng. J. 109 (2016) 243
251. [113] S.-H. Chang, C.-H. Wu, G.-J. Tsai, Effects of chitosan molecular weight on its antioxidant and antimutagenic properties, Carbohydr. Polym. 181 (2018) 1026
1032. [114] B. Kang, T.P. Vales, B.-K. Cho, J.-K. Kim, H.-J. Kim, Development of gallic acidmodified hydrogels using interpenetrating chitosan network and evaluation of their antioxidant activity, Molecules 22 (2017) 1976. [115] Q. Xu, P. Ma, W. Yu, C. Tan, H. Liu, C. Xiong, et al., Chitooligosaccharides protect human embryonic hepatocytes against oxidative stress induced by hydrogen peroxide, Mar. Biotechnol. 12 (2010) 292
298. [116] A. Gurib-Fakim, Medicinal plants: traditions of yesterday and drugs of tomorrow, Mol. Asp. Med. 27 (2006) 1
93. [117] A. Singh, P.K. Dutta, H. Kumar, A.K. Kureel, A.K. Rai, Synthesis of chitin-glucanaldehyde-quercetin conjugate and evaluation of anticancer and antioxidant activities, Carbohydr. Polym. 193 (2018) 99
107. [118] C. Chayanaphat, R. Ratana, T. Sewan, S. Nagahiro, Degradation of chitosan hydrogel dispersed in dilute carboxylic acids by solution plasma and evaluation of anticancer activity of degraded products, Jpn. J. Appl. Phys. 57 (2018) 0102B0105. [119] R.-C. Chien, M.-T. Yen, J.-L. Mau, Antimicrobial and antitumor activities of chitosan from shiitake stipes, compared to commercial chitosan from crab shells, Carbohydr. Polym. 138 (2016) 259
264. [120] M.H. Rahman, L.G. Hjeljord, B.B. Aam, M. Sørlie, A. Tronsmo, Antifungal effect of chito-oligosaccharides with different degrees of polymerization, Eur. J. Plant. Pathol. 141 (2015) 147
158. [121] Z. Zhong, P. Li, R. Xing, S. Liu, Antimicrobial activity of hydroxylbenzenesulfonailides derivatives of chitosan, chitosan sulfates and carboxymethyl chitosan, Int. J. Biol. Macromol. 45 (2009) 163
168. [122] H.A. Shah, R.P. Patel, Statistical modeling of zaltoprofen loaded biopolymeric nanoparticles: characterization and anti-inflammatory activity of nanoparticles loaded gel, Int. J. Pharm. Investig. 5 (2015) 20
27. [123] M.J. Chung, J.K. Park, Y.I. Park, Anti-inflammatory effects of low-molecular weight chitosan oligosaccharides in IgE-antigen complex-stimulated RBL-2H3 cells and asthma model mice, Int. Immunopharmacol. 12 (2012) 453
459. [124] M. Yousef, R. Pichyangkura, S. Soodvilai, V. Chatsudthipong, C. Muanprasat, Chitosan oligosaccharide as potential therapy of inflammatory bowel disease: therapeutic efficacy and possible mechanisms of action, Pharmacol. Res. 66 (2012) 66
79. [125] W.H. Organization, Mental Health Action Plan 2013
2020, WHO, Geneva, 2013, in 2015. [126] Y.-S. Cho, S.-K. Kim, C.-B. Ahn, J.-Y. Je, Inhibition of acetylcholinesterase by gallic acid-grafted-chitosans, Carbohydr. Polym. 84 (2011) 690
693. [127] H.C. Huang, L. Hong, P. Chang, J. Zhang, S.Y. Lu, B.W. Zheng, et al., Chitooligosaccharides attenuate Cu2 1 -induced cellular oxidative damage and cell apoptosis involving Nrf2 activation, Neurotox. Res. 27 (2015) 411
420. [128] M. Jiang, Z. Guo, C. Wang, Y. Yang, X. Liang, F. Ding, Neural activity analysis of pure chito-oligomer components separated from a mixture of chitooligosaccharides, Neurosci. Lett. 581 (2014) 32
36.

Handbook of Chitin and Chitosan

References

399

[129] X. Dai, P. Chang, Q. Zhu, W. Liu, Y. Sun, S. Zhu, et al., Chitosan oligosaccharides protect rat primary hippocampal neurons from oligomeric β-amyloid 1-42-induced neurotoxicity, Neurosci. Lett. 554 (2013) 64
69. [130] P. Santos-Moriano, L. Fernandez-Arrojo, M. Mengibar, E. Belmonte-Reche, P. Pen˜alver, F.N. Acosta, et al., Enzymatic production of fully deacetylated chitooligosaccharides and their neuroprotective and anti-inflammatory properties, Biocatal. Biotransfor. 36 (2018) 57
67. [131] J.-Y. Je, E.-K. Kim, C.-B. Ahn, S.-H. Moon, B.-T. Jeon, B. Kim, et al., Sulfated chitooligosaccharides as prolyl endopeptidase inhibitor, Int. J. Biol. Macromol. 41 (2007) 529
533. [132] M. Bhattacharya, R.L. Reis, V. Correlo, L. Boesel, Material properties of biodegradable polymers, in: R. Smith (Ed.) Biodegradable Polymers for Industrial Applications, Woodhead Publishing, 2005, pp. 336-356. [133] J.M. Anderson, Biodegradation of polymers, in: K.H.J. Buschow, R.W. Cahn, M.C. Flemings, B. Ilschner, E.J. Kramer, S. Mahajan, et al. (Eds.), Encyclopedia of Materials: Science and Technology, Elsevier, Oxford, 2001, pp. 560
563. [134] T. Kean, M. Thanou, Biodegradation, biodistribution and toxicity of chitosan, Adv. Drug. Deliv. Rev. 62 (2010) 3
11. [135] H. Onishi, K. Inaoka, K. Matsubara, K. Suzuki, Numerical analysis of flow and conjugate heat transfer of a two-row plate-finned tube heat exchanger, in: Proceedings of the Second International Conference on Compact Heat Exchangers and Enhancement Technology for the Process Industries, 1999, pp. 175
182. [136] K. Kafedjiiski, F. Fo¨ger, H. Hoyer, A. Bernkop-Schnu¨rch, M. Werle, Evaluation of in vitro enzymatic degradation of various thiomers and cross-linked thiomers, Drug. Dev. Ind. Pharm. 33 (2007) 199
208. [137] G. Lu, B. Sheng, G. Wang, Y. Wei, Y. Gong, X. Zhang, et al., Controlling the degradation of covalently cross-linked carboxymethyl chitosan utilizing bimodal molecular weight distribution, J. Biomater. Appl. 23 (2009) 435
451. [138] S.B. Rao, C.P. Sharma, Use of chitosan as a biomaterial: studies on its safety and hemostatic potential, J. Biomed. Mater. Res. 34 (1997) 21
28. [139] F.S. Kittur, A.B. Vishu Kumar, R.N. Tharanathan, Low molecular weight chitosans Preparation by depolymerization with Aspergillus niger pectinase, and characterization, Carbohydr. Res. 338 (2003) 1283
1290. [140] F.S. Kittur, A.B. Vishu Kumar, M.C. Varadaraj, R.N. Tharanathan, Chitooligosaccharides—preparation with the aid of pectinase isozyme from Aspergillus niger and their antibacterial activity, Carbohydr. Res. 340 (2005) 1239
1245. [141] I. Makarios-Laham, T.-C. Lee, Biodegradability of chitin- and chitosan-containing films in soil environment, J. Environ. Polym. Degrad. 3 (1995) 31
36. [142] K.V. Harish Prashanth, K. Lakshman, T.R. Shamala, R.N. Tharanathan, Biodegradation of chitosan-graft-polymethylmethacrylate films, Int. Biodeterior. Biodegrad. 56 (2005) 115
120. [143] S. Beier, S. Bertilsson, Bacterial chitin degradation-mechanisms and ecophysiological strategies, Front. Microbiol. 4 (2013). 149.

Handbook of Chitin and Chitosan

C H A P T E R

13 Chitin and chitosan: current status and future opportunities Ruchi Mutreja1, Abhijeet Thakur2 and Arun Goyal2 1

Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India, 2Carbohydrate Enzyme Biotechnology Laboratory, Department of Biosciences and Bioengineering, Indian Institute of Technology Guwahati, Guwahati, India O U T L I N E

13.1 Introduction

402

13.2 Properties of chitin and chitosan 13.2.1 Structural properties 13.2.2 Biological properties

403 403 404

13.3 Chitin, chitosan, and their derivatives 13.3.1 Biosynthesis of chitin 13.3.2 Chitin and chitosan production

405 405 405

13.4 Applications of chitin and chitosan 13.4.1 Industrial applications 13.4.2 Biomedical applications of chitosan

407 407 410

13.5 Conclusion and future perspectives

412

References

413

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00013-4

401

© 2020 Elsevier Inc. All rights reserved.

402

13. Chitin and chitosan: current status and future opportunities

13.1 Introduction Chitin is the most abundant biopolymer after cellulose and starch. Chitin and its deacetylated form, chitosan, are widely used in textile processing, tissue engineering, drug delivery, wastewater treatment, the food industry, etc. Chitin, a long-chain polymer of N-acetylglucosamine, is mostly found in the form of crystalline macrofibrils and is the major structural component of the exoskeletons of arthropods and the cell walls of fungi. Chitin is nitrogenous polysaccharide, white in color, hard, inelastic, insoluble in water, and it is commercially produced generally from arthropod’s exoskeleton shells. The exoskeleton shells are treated first with acid and then alkali to dissolve the calcium carbonate and proteins, respectively, followed by decolorization, for pigment removal. For exploring the applicability of chitin, it is deacetylated to form chitosan, a copolymer of N-acetylglucosamine and glucosamine (Fig. 13.1). The history of chitosan dates back to the 19th century, when Rouget discussed the deacetylated form of chitosan in 1859 [1]. The wide application of chitin in diverse fields is attributed to its unique biochemical and biophysical properties, such as biocompatibility, biodegradability, nontoxicity, ability to form films, and ready availability. The degree of N-acetylation of chitin affects its solubility in water, swelling properties, molecular weight, and presence of inter- and intramolecular hydrogen bonds [2]. The presence of amino and hydroxyl groups in chitosan opens the door for many industrial and biomedical applications. Researchers across the globe are working on the production of chitin and its derivatives from different sources and exploring them for various applications and functions. The global chitosan market is projected to grow from US$553.6 million in 2017 to US$1088.0 million by 2022, at a compound annual growth rate (CAGR) of 14.5% between 2017 and 2022. This chapter aims to present the state-of-the-art knowledge for the bioextraction or isolation of chitin and chitosan from different sources, their properties, and their applications, along with the challenges encountered and future prospective.

FIGURE 13.1 Chitin and its deacetylation to chitosan.

Handbook of Chitin and Chitosan

13.2 Properties of chitin and chitosan

403

13.2 Properties of chitin and chitosan 13.2.1 Structural properties Chitin and chitosan are basic polysaccharides. Chitin is hydrophobic in nature and is found to be soluble in hexafluoroisopropanol, hexafluoroacetone, and chloroalcohols in combination with mineral acids [3] and dimethylacetamide [4]. The solubility of chitosan in N-methyl morpholine-N-oxide (NMMO)/H2O was reported [5]. Depending upon the bonding between the chains, chitin occurs in different polymorphic forms, that is, α, β, and γ, which can be easily differentiated by infrared spectroscopy, solid-state nuclear magnetic resonance, and X-ray diffraction. The α-chitin form is most abundant, followed by the β- and γ-forms [6]. α-Chitin is found in arthropods shell, whereas β-chitin is found in squid pens and worms tubes [7]. α-Chitin is also found in fungi and yeast [8] and also can be synthesized by chitin recrystallization [9], in vitro biosynthesis [10], or polymerization [11]. α-Chitin consists of two antiparallel molecules per unit cell with intersheet hydrogen bonds, whereas only one molecule is present in the parallel arrangement in β-chitin, and γ-chitin consists of mixed parallel and antiparallel orientations (Fig. 13.2). Chitosan is a linear polysaccharide composed of randomly distributed β-(1 4)-linked D-glucosamine and N-acetyl-D-glucosamine with different molecular weights, protein content, crystallinity, moisture, solubility, flexibility, polymer conformation, viscosity, surface area, porosity, tensile strength, conductivity, and photoluminescence [12]. It can undergo N-acylation and Schiff reaction owing to the presence of the amino groups. Due to its negative charge, it can also form complexes with polymers or macromolecules with a positive charge. It is also known to chelate metal ions [13].

FIGURE 13.2 γ-chitin.

Different structural forms of chitin: (A) α-chitin, (B) β-chitin, and (C)

Handbook of Chitin and Chitosan

404

13. Chitin and chitosan: current status and future opportunities

FIGURE 13.3 Biological properties of chitin.

13.2.2 Biological properties Chitin and its derivatives are biocompatible, natural nontoxic polymers and are known to bind to mammalian and microbial cells. They are known to have hemostatic, fungistatic, spermicidal, antitumor, and anticholesteremic activities, to accelerate bone formation, and be a central nervous system depressant and immunoadjuvant (Fig. 13.3). The biological properties of chitin and its derivatives depend on their solubility in water or solvents along with its degree of acetylation (DA) [14].

Handbook of Chitin and Chitosan

13.3 Chitin, chitosan, and their derivatives

405

Therefore extensive work is being done on its solubility for its maximum exploitation [15]. The effects of molecular weight, polydispersity, and DA on the physicochemical and biological properties of chitin and chitosan have been reported [16].

13.3 Chitin, chitosan, and their derivatives 13.3.1 Biosynthesis of chitin Chitin biosynthesis is a highly complex process involving a series of events. Chitosomes, cytoplasmic microvesicles, are known to display chitin synthase (CS) activity. Chitosomes are known for translocating CS at predetermined locations after activation through proteolytic reactions [17]. Different polymeric forms of chitin are synthesized by CS by using N-acetylglucosamine (GlcNAc) as the starting material [18].

13.3.2 Chitin and chitosan production Chemical or biological processes known for chitin and chitosan production are discussed in the following section. 13.3.2.1 Chemical process The economical and eco-friendly production of chitin and chitosan by a chemical process using shelfish waste, fungi, and insect cuticles has been reported (Fig. 13.4) [19]. For this chemical process, deproteinization, demineralization, and decolorization are the major steps. Deproteinization is done by using sodium hydroxide, while the demineralization involves treatment with an acid at high temperature. The arthropods’ exoskeleton or cell wall of fungi are covantly associated with other constituents of the shells. Therefore treating them with a harsh alkali or acid under high temperature may result in depolymerization and partial deacetylation, releasing undesired by-products with undesirable properties [20]. In order to avoid the abovementioned undesirable by-products, the use of ambient temperature and the use of fermenters have been reported [21]. Chitin was chemically extracted by dissolving crustacean shells in 1-ethyl-3methyl-imidazolium acetate [22]. The use of organic acids (lactic and acetic) produced by cheese whey fermentation for the demineralization of microbially deproteinized shrimp shells has also been reported [23]. Insect biomass is also used for the industrial production of chitin and its derivatives due to the richness of the biopolymer and the higher availability of dry matter [24]. For chitin extraction from insect cuticle, demineralization and deproteinization is performed almost in similar way as for arthropods shells, though much stronger treatment is

Handbook of Chitin and Chitosan

406

13. Chitin and chitosan: current status and future opportunities

FIGURE 13.4 Chemical synthesis of chitin and its derivatives.

required here. Chitin from seven different Orthoptera species has been isolated previously and their structures were compared; it was found that the chitin content of the different species varied between 5.3% and 8.9% with low molecular weights (5.2 6.8 kDa). The study concluded that the insects after insecticide treatment should be collected and can be exploited for the extraction of chitin and its derivatives [25]. The extraction of chitin up to 15% from Holotrichia parallela Motschulsky was performed using 1 M HCl and 1 M NaOH treatment followed by decolorization using 1% potassium permanganate [26]. In another study, chitosan was isolated from the American cockroach by treatment with 1%

Handbook of Chitin and Chitosan

13.4 Applications of chitin and chitosan

407

HCl followed by 4% (w/v) NaOH and then characterization by FTIR analysis [27]. The structure of isolated chitosan was found to be similar to that of commercially available chitosan. The extraction of chitin from different aquatic invertebrates [28] and grasshopper species [29], and chitosan from different local insects [30] exhibited alternative chitin sources for different industrial and biomedical applications. Other than arthropods and insects, the fungal cell wall is also known as a source of chitin. Chitin and chitosan from Aspergillus terreus were extracted, characterized, and their applications were shown [30]. The biomass from mushrooms can also be a promising source of chitin [31]. 13.3.2.2 Biological process The biological process for the extraction of chitin and its derivatives is gaining popularity because of the limitations associated with the chemical process, such as the high energy requirement and the production of extremely hazardous and undesirable by-products [19]. Biotechnological processes have been explored for the commercial production of chitin and its derivatives along with their high-value by-products, such as lactic acid, antioxidants, protein hydrolysates, and carotenoids (astaxanthin) [32]. The production of chitin from crab shell waste by fermentation using Lactobacillus paracasei and Serratia marcescens FS-3 was achieved [33]. In another study, chitin and chitosan production was accomplished by fermentation of shrimp shell using Bacillus subtilis, where the proteases of Bacillus helped in deproteinization and around 84% protein and 72% minerals were removed using Bacillus, thus producing good quality chitin, which was further deacetylated for the production of chitosan [34]. The production of chitin by the fermentation of shrimp biowaste and its deacetylation to chitosan was also reported [35].

13.4 Applications of chitin and chitosan Chitin and its derivatives are used for a diversity of applications owing to their unique physical and biochemical properties. They can be easily molded into films, fibers, sponges, beads, powder, gel, and solutions. Different industrial and biomedical applications of chitin and chitosan are discussed in the subsequent sections.

13.4.1 Industrial applications Depending upon the DA and molecular weight, chitin and chitosan can be used for a variety of applications, such as pharmaceuticals,

Handbook of Chitin and Chitosan

408

13. Chitin and chitosan: current status and future opportunities

cosmetic products, wastewater treatment, paper industry, textile industry, and food industry. 13.4.1.1 Cosmetics A preparation that is applied outside the human body, that is, external parts or teeth or oral cavity mucous membranes, with an intention to clean or change or improve the appearance is known as a cosmetic product. Chitin and its derivatives are polycation hydrocolloids that become viscous when neutralized with acid. Thus they can act as abrasive filmformers and can interact with integuments (skin covers) and hair. Chitosan is compatible with substances that can absorb UV radiation, so different dyes can be covalently linked to amino groups of chitosan. These can be used as antioxidants, antiallergic, and antiinflammatory substances. Chitin and its derivatives are used in cosmetics for hair, skin, and oral care [36]. The use of chitin and its derivatives in haircare products is gaining attention due to its unique properties, such as high thermostability, good solubility in acidic media and cosmetic bases, stability in pH range, neutral or pleasant odor with low volatility. It interacts with keratin forming transparent films on hair fibers and can also improve the rheological behavior and adhesion. In solution form, chitin forms a clear, elastic film on hair, thereby increasing its elasticity, appearance, mechanical properties, softness, and strength [37], and thus can be used in shampoos, rinses, colorants, hair lotions, spray, and tonics [38]. The use of chitin and its derivatives have been reported in skincare products such as sunscreens, moisturizer foundation, eyeshadow, lipstick, cleansing materials, and bath agent. The encapsulation of sunscreen particles in chitosan gel was reported [39]. A study showed the use of chitosan oligosaccharides for reducing skin photoaging in a mouse model after exposure to UV radiation for 10 weeks [40]. The effects of chitin nanofibrils on the functioning of skin using skin models was observed and it was found that the nanofibrils and nanocrystals help in skin structure preservation and also reduce the production of transforming growth factor-β [41]. Chitosan argininamide has been identified for preparing skin cleansing products [42]. The use of chitin and its derivatives as humectants, antioxidants, emulsifiers, stabilizers, antimicrobials, viscosifiers, surfactants etc. has been reported [36]. Chitin and its derivatives have been used in toothpaste, mouthwashes, and chewing gum as a dental filler, to prevent plaque formation [43], dental abrasion reduction [44], and for cleaning purposes by acting either as a carrier or vehicle. 13.4.1.2 Water engineering Chitin and its derivatives have been used as a heavy metal trapping, flocculating, and chelating agent [45]. They have been used in

Handbook of Chitin and Chitosan

13.4 Applications of chitin and chitosan

409

wastewater treatment for removing heavy metal ions [46]. Due to the interaction of chitosan with anionic wastes, agglomeration along with the formation of precipitates and floe can be seen, and thus can be used for the recycling of food processing waste. The use of chitosan for removing color from effluents has been also reported [47]. Chitosan can compete effectively with synthetic resins for the capture of heavy metals from processing wastewater. The use of chitin for the decontamination of wastewater containing plutonium and mercury ion has been documented [48]. Studies also showed the use of chitin and its derivatives for removing arsenic from drinking water and petroleum or petroleum products from wastewater [49]. 13.4.1.3 Paper and textile industry Chitin and chitosan are known to resemble cellulose, and thus can be used for strengthening recycled paper and packaging material. Therefore less chemical additives are required with better results, giving a smoother surface and resistance to moisture. Hydroxymethyl chitin is used for the production of toilet paper and wrapping paper [3]. The use of chitin and its derivatives has been reported in the preparation of antistatic, soil repellent, printing, and finishing material, as well as for dye removal and for preparing textile sutures, threads, and fibers [50]. 13.4.1.4 Food industry Chitosan is used in the food industry as a nonabsorbable carrier, thickener, and gelling agent. Chitosan is used as a food wrap owing to its ability to form semipermeable tough, long-lasting, flexible films, thus extending the shelf life of food [51]. It has been shown that when the chitosan film is applied on the surface of tomatoes, quince, pears, and anquito squash, it resulted in enhanced quality, reduced maturation, and longer decay times, with prolonged shelf life [52]. A study also showed the use of chitosan films in packaging for inhibiting microbial growth [53]. The use of microcrystalline chitin as a flavoring, antifungal and coloring agent was demonstrated [54]. Chitin and its derivatives have been applied for enzyme immobilization or entrapping enzymes, microbial, animal or plant cells. This has been possible due to the availability of reactive functional groups, their ease of modification, nontoxicity, low cost, and insolublility in water and most ordinary solvents. Chitosan in the form of membranes, beads, gels, powders, and fibers can be used for enzyme immobilization [55]. The immobilization of rennet from Kluyveromyces lactis using pectin chitosan porous beads for milk coagulation was reported [56]. Chitin and its derivatives can be used as additives in food items to impart color and flavor, and as a blending agent [57]. Dyes diffuse in chitosan at the same rate as in cellulose [58]. They have also been used

Handbook of Chitin and Chitosan

410

13. Chitin and chitosan: current status and future opportunities

as emulsifying agents [59]. The emulsification of sunflower oil was carried out using chitosan solutions and it was found that the viscosity, stability, and aging of the emulsion depends upon the concentration of chitosan [60]. The use of chitosan as an antioxidant agent and gelling agent has also been documented [61]. 13.4.1.5 Other industrial applications Chitin and chitosan have been used in agriculture as antifungal agents and also to accelerate the growth of plant and decelerate root knot worm infestations [62]. Use of chitosan as a fixing agent and to improve diffusion in photography has also been reported [63]. Due to the presence of amino and hydroxyl groups, chitosan is also used as chromatographic support material. The use of chitin and chitosan as a sorbent material in solid-phase extraction of phenol and chlorophenols by using high-performance liquid chromatography (HPLC) was reported [64]. Chitosan hydrohobicity is well explored for its use in the construction of solid-state proton conducting polymer batteries due to the generation of protons transported through microvoids in chitin polymer [65].

13.4.2 Biomedical applications of chitosan 13.4.2.1 Tissue engineering Tissue engineering deals with the repair, replacement, maintenance, or enhancement of the function of a particular tissue or organ. For this, the polymer scaffolds are designed to have properties such as high porosity, appropriate pore size distribution, high surface area, nontoxicity, biocompatibility, biodegradability, and structural integrity with appropriate mechanical properties. They should also interact with the cells to promote cell adhesion, proliferation, migration, and differentiated cell function. Use of chitosan as a biomaterial for tissue engineering applications has intensified during the last three decades [66]. The use of chitin and chitosan composite-based scaffolds for tissue engineering has been reported. Their extraordinary properties, such as their cationic nature and ability to form interconnected porous structures, have led to the formation of scaffolds [67]. The use of chitin and chitosan has been reported for bone repair and reconstruction upon its bonding with other biomaterials, such as hydroxyapatite (HAp), bioactive glass ceramic (BGC), etc., to form a carbonated apatite layer to enhance the mechanical properties [68]. BGC in the nanoform for bonding with chitin can be produced by the sol gel method [69]. The cells are attached to the pore walls of the scaffolds without any toxicity and thus, can be used for tissue engineering applications. Chitosan

Handbook of Chitin and Chitosan

13.4 Applications of chitin and chitosan

411

(CS)-gelatin (CG) scaffolds with nBGC for tissue engineering applications have been reported [70]. Sugar-bound chitosans have been reported for their interaction with cells [71]. The synthesis of galactosylated chitosan (GC) lactobionic acid using EDC/NHS chemistry was carried out for its use as extracellular matrix for hepatocyte attachment [72]. Mannosylated chitosan (MC) was synthesized for specific recognition of B-cell, dendritic cell, and macrophage [73]. A carboxymethyl chitin (CMC)/polyvinyl alcohol (PVA) blend containing 7% CMC and 8% PVA was reported in the form of nanofibers as a scaffold for tissue engineering applications [74]. The bioactive and biocompatible nanofibrous scaffold thus formed was cross-linked with glutaraldehyde followed by heat treatment, and was tested for cytotoxicity and cell attachment studies. The ability of CMC/PVA nanofibrous scaffolds for cell adhesion and proliferation was demonstrated and thus proposed for tissue engineering applications [75]. The use of chitin tubes for tissue engineering applications has been reported [76]. For this, mechanically strong chitin tubes were prepared using acylation chemistry and mold-casting techniques, whereas alkaline hydrolysis was employed for chitosan tubes synthesis. The study showed nerve cell adhesion and neurite outgrowth using biodegradable and biocompatible chitin and chitosan tubes, thus showing them to be promising candidates for neural tissue engineering [77]. Composite membranes of chitin hydroxyapatite were prepared and evaluated for tissue engineering applications [78]. The presence of hydroxyapatite helps in the attachment and spreading of cells [78]. Another study showed the application of composite scaffolds of chitin and nanosilica using MG 63 cell line for bone tissue engineering [79]. Adding silica to chitin helps in increasing its bioactivity and biocompatibility [79]. Cross-linked chitosan with dimethyl-3,3-dithio-bis-propionimidate (DTBP) was synthesized to avoid chitosan degradation and also to enhance its mechanical strength [80]. The composite of chitosan with gelatin was synthesized to improve the stiffness of the scaffold [81]. A composite of chitosan with poly(ε-caprolactone) [82], γ-poly(glutamic acid) [83], and alginate [84] has also been reported for tissue engineering applications. The immobilization of specific sequences such as ArgGly-Asp (RGD) by chemical cross-linking to promote cell adhesion has been reported [85]. Polyglycolic acid (PGA) chitosan hybrid composite was synthesized and tested on cultured fibroblast cells showing better cell attachment and spreading as compared with chitosan films, thus it has been proposed for tissue engineering [86]. 13.4.2.2 Wound healing/wound dressing Chitin and chitosan in the form of fibers, mats, sponges, etc. have been reported for wound healing/wound dressing [87]. A study showed the

Handbook of Chitin and Chitosan

412

13. Chitin and chitosan: current status and future opportunities

application of water-soluble chitin for wound healing in rats and demonstrated complete healing and reepithelialization of tissues in 7 days after the initial wounding [88]. The effect of chitin and its derivatives was investigated on wound healing in rats [89]. A composite of chitin with fucoidan and alginate was used for wound healing [90]. Upon histological examination, advanced granulation with tissue and capillary formation was observed in the wounds treated with composite compared with those treated with calcium alginate fiber. The effects of chitin and its derivatives on the synthesis of collagen for wound healing were studied [91] and it was found that chitin at higher concentration induces stable collagen synthesis in the early wound healing process. Another study showed that the photocrosslinkable chitosan hydrogel enhances wound healing upon the addition of fibroblast growth factor-2 [92]. Chitin and its derivatives have also been used for burn treatment and wound dressings owing to their unique properties, such as toughness, biocompatibility, and oxygen permeability. 13.4.2.3 Drug delivery Controlled release of drugs regulates the release rate and thus enhances efficiency, safety, and fidelity of drugs. Several natural and synthetic degradable polymers are used to achieve this purpose [93]. Chitosan, being an adsorbable and nontoxic polymer, can be used for drug encapsulatation or can be adsorbed covalently to chitosan, which can be released slowly as the chitosan is degraded. Chitosan is also favored in drug delivery because of antiulcer and antacid properties, which help in preventing drug irritation [94]. The DA and the molecular weight of chitosan influence the hydrophobicity and solubility and thus affect the drug encapsulation [95]. Chitosan can be modified differently to make it a more efficient drug delivery vehicle, such as 5-fluorouracil loaded chitosan [96], chitosan cross-linked with pectin [97], and chitosan with β-cyclodextrin [98].

13.5 Conclusion and future perspectives Chitin and chitosan are natural nontoxic polymers with unique structural and functional characteristics. Chitin and chitosan are currently used in various industrial and biomedical processes. The reactivity of the amino and hydroxyl groups makes it more interesting for diverse applications in tissue engineering, wound healing, drug delivery, cosmetics, water treatment, and agriculture. An initiative to synthesize new chitin and chitosan-based eco-friendly polymers for food packaging, membrane separation, hydrogels, and biomaterials should be undertaken. The reduction in pollution and the promotion of green chemistry

Handbook of Chitin and Chitosan

References

413

can be achieved by the use of chitin and chitosan-based polymers with molecular imprinting techniques.

References [1] C. Rouget, Des substances amylace´es dans les tissus des animaux, spe´cialement des Articule´s (chitine), Comp. Rend. 48 (1859) 792 795. [2] M. Hayes, Chitin, chitosan and their derivatives from marine rest raw materials: potential food and pharmaceutical applications, Marine Bioactive Compounds, Springer, 2012. [3] P.K. Dutta, M.N.V. Ravikumar, J. Dutta, Chitin and chitosan for versatile applications, J. Macromol. Sci. Part C: Polym. Rev. 42 (2002) 307 354. [4] P.K. Dutta, M.K. Khatua, J. Dutta, R. Prasad, Use of chitosan-DMAc/LiCl gel as drug carriers, Int. J. Chem. Sci. 1 (2003) 93. [5] M.N.V.R. Kumar, P.K. Dutta, S. Nakamura, Chitosan-amine oxide: a new gelling system, characterization and in vitro evaluations, Indian. J. Pharm. Sci. 62 (2000) 55. [6] K. Rudall, Chitin and its association with other molecules, J. Polym. Sci. Part C: Polym. Symp 28 (1969) 83 102. [7] F. Gaill, J. Persson, J. Sugiyama, R. Vuong, H. Chanzy, The chitin system in the tubes of deep sea hydrothermal vent worms, J. Struct. Biol. 109 (1992) 116 128. [8] S.M. Bowman, S.J. Free, The structure and synthesis of the fungal cell wall, Bioessays 28 (2006) 799 808. [9] W. Helbert, J. Sugiyama, High-resolution electron microscopy on cellulose II and α-chitin single crystals, Cellulose 5 (1998) 113 122. [10] S. Bartnickigarcia, J. Persson, H. Chanzy, An electron microscope and electron diffraction study of the effect of calcofluor and congo red on the biosynthesis of chitin in vitro, Arch. Biochem. Biophys. 310 (1994) 6 15. [11] J. Sakamoto, J. Sugiyama, S. Kimura, T. Imai, T. Itoh, T. Watanabe, et al., Artificial chitin spherulites composed of single crystalline ribbons of α-chitin via enzymatic polymerization, Macromolecules 33 (2000) 4155 4160. [12] M. Bajaj, J. Winter, C. Gallert, Effect of deproteination and deacetylation conditions on viscosity of chitin and chitosan extracted from Crangon crangon shrimp waste, Biochem. Eng. J. 56 (2011) 51 62. [13] B. Singh, S. Maharjan, Y.-J. Choi, T. Akaike, C.-S. Cho, Marine materials: gene delivery, Springer Handbook of Marine Biotechnology, Springer, 2015. [14] C.K.S. Pillai, W. Paul, C.P. Sharma, Chitin and chitosan polymers: chemistry, solubility and fiber formation, Prog. Polym. Sci. 34 (2009) 641 678. [15] I. Aranaz, R. Harris, A. Heras, Chitosan amphiphilic derivatives. Chemistry and applications, Curr. Org. Chem. 14 (2010) 308 330. [16] I. Aranaz, M. Mengı´bar, R. Harris, I. Pan˜os, B. Miralles, N. Acosta, et al., Functional characterization of chitin and chitosan, Curr. Chem. Biol. 3 (2009) 203 230. [17] K. Suresh, C. Subramanyam, A putative role for calmodulin in the activation of Neurospora crassa chitin synthase, FEMS Microbiol. Lett. 150 (1997) 95 100. [18] E. Cohen, In vitro chitin synthesis in an insect: formation and structure of microfibrils, Eur. J. Cell Biol. 26 (1982) 289 294. [19] S. Kaur, G.S. Dhillon, Recent trends in biological extraction of chitin from marine shell wastes: a review, Crit. Rev. Biotechnol. 35 (2015) 44 61. [20] M.C. Gortari, R.A. Hours, Biotechnological processes for chitin recovery out of crustacean waste: a mini-review, Electron. J. Biotechnol. 16 (2013) 1 18. [21] C.T. Pires, J.A.P. Vilela, C. Airoldi, The effect of chitin alkaline deacetylation at different condition on particle properties, Procedia Chem. 9 (2014) 220 225.

Handbook of Chitin and Chitosan

414

13. Chitin and chitosan: current status and future opportunities

[22] Y. Qin, X. Lu, N. Sun, R.D. Rogers, Dissolution or extraction of crustacean shells using ionic liquids to obtain high molecular weight purified chitin and direct production of chitin films and fibers, Green. Chem. 12 (2010) 968 971. [23] N.S. Mahmoud, A.E. Ghaly, F. Arab, Unconventional approach for demineralization of deproteinized crustacean shells for chitin production, Am. J. Biochem. Biotechnol. 3 (2007) 1 9. [24] A.T. Dossey, Insects and their chemical weaponry: new potential for drug discovery, Nat. Prod. Rep. 27 (2010) 1737 1757. [25] M. Kaya, S. Erdogan, A. Mol, T. Baran, Comparison of chitin structures isolated from seven Orthoptera species, Int. J. Biol. Macromol. 72 (2015) 797 805. [26] S. Liu, J. Sun, L. Yu, C. Zhang, J. Bi, F. Zhu, et al., Extraction and characterization of chitin from the beetle holotrichia Parallela motschulsky, Molecules 17 (2012). [27] D. Wanule, J.V. Balkhande, P.U. Ratnakar, A.N. Kulkarni, C.S. Bhowate, Extraction and FTIR analysis of chitosan from American cockroach, Periplaneta americana, Extraction 3 (2014). [28] M. Kaya, T. Baran, A. Mentes, M. Asaroglu, G. Sezen, K.O. Tozak, Extraction and characterization of α-chitin and chitosan from six different aquatic invertebrates, Food Biophys. 9 (2014) 145 157. ˙ I. Sargin, G. Arslan, A. Mol, et al., [29] M. Kaya, E. Leleˇsius, R. Nagrockaite, Differentiations of chitin content and surface morphologies of chitins extracted from male and female grasshopper species, PLoS One 10 (2015) e0115531. [30] N.H. Marei, E.A. El-Samie, T. Salah, G.R. Saad, A.H.M. Elwahy, Isolation and characterization of chitosan from different local insects in Egypt, Int. J. Biol. Macromol. 82 (2016) 871 877. [31] S.P. Ospina a´lvarez, D.A. Ramı´rez cadavid, D.M. Escobar sierra, C.P. Ossa orozco, D.F. Rojas vahos, P. Zapata ocampo, et al., Comparison of extraction methods of chitin from Ganoderma lucidum mushroom obtained in submerged culture, BioMed. Res. Int. 2014 (2014). [32] M.C. Gortari, R.A. Hours, Biotechnological processes for chitin recovery out of crustacean waste: a mini-review, Electron. J. Biotechnol. 16 (2013). 14. [33] W.J. Jung, G.H. Jo, J.H. Kuk, Y.J. Kim, K.T. Oh, R.D. Park, Production of chitin from red crab shell waste by successive fermentation with Lactobacillus paracasei KCTC3074 and Serratia marcescens FS-3, Carbohydr. Polym. 68 (2007) 746 750. [34] T.K. Sini, S. Santhosh, P.T. Mathew, Study on the production of chitin and chitosan from shrimp shell by using Bacillus subtilis fermentation, Carbohydr. Res. 342 (2007) 2423 2429. [35] M.S. Rao, W.F. Stevens, Chitin production by Lactobacillus fermentation of shrimp biowaste in a drum reactor and its chemical conversion to chitosan, J. Chem. Technol. Biotechnol. 80 (2005) 1080 1087. [36] I. Aranaz, N. Acosta, C. Civera, B. Elorza, J. Mingo, C. Castro, et al., Cosmetics and cosmeceutical applications of chitin, chitosan and their derivatives, Polymers 10 (2018) 213. [37] A. Sionkowska, B. Kaczmarek, M. Michalska, K. Lewandowska, S. Grabska, Preparation and characterization of collagen/chitosan/hyaluronic acid thin films for application in hair care cosmetics, Pure Appl. Chem. 89 (2017) 1829 1839. [38] P.K. Dutta, J. Dutta, V.S. Tripathi, Chitin and chitosan: chemistry, properties and applications, J. Sci. Ind. Res. India 63 (2004) 20 31. [39] Y.A. Gomaa, L.K. El-Khordagui, N.A. Boraei, I.A. Darwish, Chitosan microparticles incorporating a hydrophilic sunscreen agent, Carbohydr. Polym. 81 (2010) 234 242. [40] S.-Z. Kong, D.-D. Li, H. Luo, W.-J. Li, Y.-M. Huang, J.-C. Li, et al., Anti-photoaging effects of chitosan oligosaccharide in ultraviolet-irradiated hairless mouse skin, Exp. Gerontol. 103 (2018) 27 34.

Handbook of Chitin and Chitosan

References

415

[41] I. Ito, T. Osaki, S. Ifuku, H. Saimoto, Y. Takamori, S. Kurozumi, et al., Evaluation of the effects of chitin nanofibrils on skin function using skin models, Carbohydr. Polym. 101 (2014) 464 470. [42] I. Aranaz, N. Acosta, C. Civera, B. Elorza, J. Mingo, C. Castro, et al., Cosmetics and cosmeceutical applications of chitin, chitosan and their derivatives, Polymers 10 (2018). [43] K. Shibasaki, H. Sano, T. Matsukubo, Y. Takaesu, Effects of low molecular chitosan on pH changes in human dental plaque, Bull. Tokyo Dent. Coll. 35 (1994) 33 39. [44] S. Ozalp, O. Tulunoglu, SEM EDX analysis of brushing abrasion of chitosan and propolis based toothpastes on sound and artificial carious primary enamel surfaces, Int. J. Paediatr. Dent. 24 (2014) 349 357. [45] S. Pokhrel, P.N. Yadav, R. Adhikari, Applications of chitin and chitosan in industry and medical science: a review, Nepal. J. Sci. Technol. 16 (2015) 99 104. [46] T.R. Sridhari, P.K. Dutta, Synthesis and characterization of maleilated chitosan for dye house effluent, Indian J. Chem. Technol. 7 (2000) 198 201. [47] K. Durga bhavani, P.K. Dutta, Physico-chemical adsorption properties on chitosan for dyehouse effluent, Am. Dyest. Report. 88 (1999) 53 58. [48] C. Jeon, W.H. Ho¨ll, Chemical modification of chitosan and equilibrium study for mercury ion removal, Water Res. 37 (2003) 4770 4780. [49] G. Crini, Recent developments in polysaccharide-based materials used as adsorbents in wastewater treatment, Prog. Polym. Sci. 30 (2005) 38 70. ¨ Zogul, J.M. Regenstein, Industrial applications of crustacean by[50] I. Hamed, F. O products (chitin, chitosan, and chitooligosaccharides): a review, Trends Food Sci. & Technol. 48 (2016) 40 50. [51] R.A.A. Muzzarelli, S. Aiba, Y. Fujiwara, T. Hideshima, C. Hwang, M. Kakizaki, et al., Filmogenic properties of chitin/chitosan, Chitin in Nature and Technology, Springer, 1986. [52] M.S. Shin, S.I. Kim, I.Y. Kim, N.G. Kim, C.G. Song, S.J. Kim, Characterization of hydrogels based on chitosan and copolymer of poly (dimethylsiloxane) and poly (vinyl alcohol), J. Appl. Polym. Sci. 84 (2002) 2591 2596. [53] R.-K. Bai, M.-Y. Huang, Y.-Y. Jiang, Selective permeabilities of chitosan-acetic acid complex membrane and chitosan-polymer complex membranes for oxygen and carbon dioxide, Polym. Bull. 20 (1988) 83 88. [54] D. Jianglian, Z. Shaoying, Application of chitosan based coating in fruit and vegetable preservation: a review, J. Food Process. Technol. 4 (2013). [55] S. Dumitriu, E. Chornet, Immobilization of xylanase in chitosan—xanthan hydrogels, Biotechnol. Prog. 13 (1997) 539 545. [56] E. Agullo´, M.S. Rodrı´guez, V. Ramos, L. Albertengo, Present and future role of chitin and chitosan in food, Macromol. Biosci. 3 (2003) 521 530. [57] D. Knorr, Recovery and utilization of chitin and chitosan in food processing waste management, Food Technol, 1991. [58] M.N.V.R. Kumar, A review of chitin and chitosan applications, React. Funct. Polym. 46 (2000) 1 27. [59] M.V. Tzoumaki, T. Moschakis, V. Kiosseoglou, C.G. Biliaderis, Oil-in-water emulsions stabilized by chitin nanocrystal particles, Food Hydrocoll. 25 (2011) 1521 1529. [60] M.S. Rodriguez, L.A. Albertengo, E. Agullo, Emulsification capacity of chitosan, Carbohydr. Polym. 48 (2002) 271 276. [61] K. Li, Y. Hwang, T. Tsai, S. Chi, Chelation of iron ion and antioxidative effect on cooked salted ground pork by N-carboxymethylchitosan (NCMC), Food Sci. Taiwan. 23 (1996) 608 616. [62] R.G. Sharp, A review of the applications of chitin and its derivatives in agriculture to modify plant-microbial interactions and improve crop yields, Agronomy 3 (2013) 757 793.

Handbook of Chitin and Chitosan

416

13. Chitin and chitosan: current status and future opportunities

[63] N.K. Mathur, C.K. Narang, Chitin and chitosan, versatile polysaccharides from marine animals, J. Chem. Educ. 67 (1990) 938. [64] J.-S. Rhee, M.-W. Jung, K.-J. Paeng, Evaluation of chitin and chitosan as a sorbent for the preconcentration of phenol and chlorophenols in water, Anal. Sci. 14 (1998) 1089 1092. [65] D. Permana, L.O. Ahmad, Enhanced conductivity and ion exchange capacity of chitosan membranes through modification with lithium for lithium polymer battery application, WSEAS Transac. Power Syst. 11 (2017) 183 189. [66] I.-Y. Kim, S.-J. Seo, H.-S. Moon, M.-K. Yoo, I.-Y. Park, B.-C. Kim, et al., Chitosan and its derivatives for tissue engineering applications, Biotechnol. Adv. 26 (2008) 1 21. [67] K.S. Chow, E. Khor, Novel fabrication of open-pore chitin matrixes, Biomacromolecules 1 (2000) 61 67. [68] T. Kokubo, Bioactive glass ceramics: properties and applications, Biomaterials 12 (1991) 155 163. [69] W. Xia, J. Chang, Preparation and characterization of nano-bioactive-glasses (NBG) by a quick alkali-mediated sol gel method, Mater. Lett. 61 (2007) 3251 3253. [70] M. Peter, P.T.S. Kumar, N.S. Binulal, S.V. Nair, H. Tamura, R. Jayakumar, Development of novel α-chitin/nanobioactive glass ceramic composite scaffolds for tissue engineering applications, Carbohydr. Polym. 78 (2009) 926 931. [71] X. Li, Y. Tushima, M. Morimoto, H. Saimoto, Y. Okamoto, S. Minami, et al., Biological activity of chitosan sugar hybrids: specific interaction with lectin, Polym. Adv. Technol. 11 (2000) 176 179. [72] I.-K. Park, J. Yang, H.-J. Jeong, H.-S. Bom, I. Harada, T. Akaike, et al., Galactosylated chitosan as a synthetic extracellular matrix for hepatocytes attachment, Biomaterials 24 (2003) 2331 2337. [73] T.H. Kim, J.W. Nah, M.-H. Cho, T.G. Park, C.S. Cho, Receptor-mediated gene delivery into antigen presenting cells using mannosylated chitosan/DNA nanoparticles, J. Nanosci. Nanotechnol. 6 (2006) 2796 2803. [74] K. Shalumon, N. Binulal, N. Selvamurugan, S. Nair, D. Menon, T. Furuike, et al., Electrospinning of carboxymethyl chitin/poly (vinyl alcohol) nanofibrous scaffolds for tissue engineering applications, Carbohydr. Polym. 77 (2009) 863 869. [75] K.T. Shalumon, N.S. Binulal, N. Selvamurugan, S.V. Nair, D. Menon, T. Furuike, et al., Electrospinning of carboxymethyl chitin/poly(vinyl alcohol) nanofibrous scaffolds for tissue engineering applications, Carbohydr. Polym. 77 (2009) 863 869. [76] T. Freier, R. Montenegro, H.S. Koh, M.S. Shoichet, Chitin-based tubes for tissue engineering in the nervous system, Biomaterials 26 (2005) 4624 4632. [77] T. Freier, R. Montenegro, H. Shan koh, M.S. Shoichet, Chitin-based tubes for tissue engineering in the nervous system, Biomaterials 26 (2005) 4624 4632. [78] K. Madhumathi, N.S. Binulal, H. Nagahama, H. Tamura, K.T. Shalumon, N. Selvamurugan, et al., Preparation and characterization of novel β-chitin hydroxyapatite composite membranes for tissue engineering applications, Int. J. Biol. Macromol. 44 (2009) 1 5. [79] K. Madhumathi, P.T. Sudheesh kumar, K.C. Kavya, T. Furuike, H. Tamura, S.V. Nair, et al., Novel chitin/nanosilica composite scaffolds for bone tissue engineering applications, Int. J. Biol. Macromol. 45 (2009) 289 292. [80] I. Adekogbe, A. Ghanem, Fabrication and characterization of DTBP-crosslinked chitosan scaffolds for skin tissue engineering, Biomaterials 26 (2005) 7241 7250. [81] Y. Huang, S. Onyeri, M. Siewe, A. Moshfeghian, S.V. Madihally, In vitro characterization of chitosan gelatin scaffolds for tissue engineering, Biomaterials 26 (2005) 7616 7627. [82] A. Sarasam, S.V. Madihally, Characterization of chitosan polycaprolactone blends for tissue engineering applications, Biomaterials 26 (2005) 5500 5508.

Handbook of Chitin and Chitosan

References

417

[83] C.-Y. Hsieh, S.-P. Tsai, D.-M. Wang, Y.-N. Chang, H.-J. Hsieh, Preparation of γ-PGA/ chitosan composite tissue engineering matrices, Biomaterials 26 (2005) 5617 5623. [84] T.W. Chung, J. Yang, T. Akaike, K.Y. Cho, J.W. Nah, S.I. Kim, et al., Preparation of alginate/galactosylated chitosan scaffold for hepatocyte attachment, Biomaterials 23 (2002) 2827 2834. [85] C.H. Schugens, C.H. Grandfils, R. Je´roˆme, P.H. Teyssie, P. Delree, D. Martin, et al., Preparation of a macroporous biodegradable polylactide implant for neuronal transplantation, J. Biomed. Mater. Res. 29 (1995) 1349 1362. [86] Y.-C. Wang, M.-C. Lin, D.-M. Wang, H.-J. Hsieh, Fabrication of a novel porous PGAchitosan hybrid matrix for tissue engineering, Biomaterials 24 (2003) 1047 1057. [87] K. Azuma, R. Izumi, T. Osaki, S. Ifuku, M. Morimoto, H. Saimoto, et al., Chitin, chitosan, and its derivatives for wound healing: old and new materials, J. Funct. Biomater. 6 (2015) 104 142. [88] Y.-W. Cho, Y.-N. Cho, S.-H. Chung, G. Yoo, S.-W. Ko, Water-soluble chitin as a wound healing accelerator, Biomaterials 20 (1999) 2139 2145. [89] T. Minagawa, Y. Okamura, Y. Shigemasa, S. Minami, Y. Okamoto, Effects of molecular weight and deacetylation degree of chitin/chitosan on wound healing, Carbohydr. Polym. 67 (2007) 640 644. [90] K. Murakami, H. Aoki, S. Nakamura, S.-I. Nakamura, M. Takikawa, M. Hanzawa, et al., Hydrogel blends of chitin/chitosan, fucoidan and alginate as healing-impaired wound dressings, Biomaterials 31 (2010) 83 90. [91] K. Kojima, Y. Okamoto, K. Kojima, K. Miyatake, H. Fujise, Y. Shigemasa, et al., Effects of chitin and chitosan on collagen synthesis in wound healing, J. Vet. Med. Sci. 66 (2004) 1595 1598. [92] K. Obara, M. Ishihara, T. Ishizuka, M. Fujita, Y. Ozeki, T. Maehara, et al., Photocrosslinkable chitosan hydrogel containing fibroblast growth factor-2 stimulates wound healing in healing-impaired db/db mice, Biomaterials 24 (2003) 3437 3444. [93] M.N.V. Ravi kumar, N. Kumar, Polymeric controlled drug-delivery systems: perspective issues and opportunities, Drug. Dev. Ind. Pharm. 27 (2001) 1 30. [94] S. Miyazaki, K. Ishii, T. Nadai, The use of chitin and chitosan as drug carriers, Chem. Pharm. Bull. 29 (1981) 3067 3069. [95] T. Sannan, K. Kurita, Y. Iwakura, Studies on chitin, 2. Effect of deacetylation on solubility, Die Makromol.Chem. Macromol. Chem. Phys. 177 (1976) 3589 3600. [96] E.B. Denkba¸s, M. Seyyal, E. PI¸SKin, Implantable 5-fluorouracil loaded chitosan scaffolds prepared by wet spinning, J. Membr. Sci. 172 (2000) 33 38. [97] H.-Y. Lin, C.-T. Yeh, Controlled release of pentoxifylline from porous chitosan-pectin scaffolds, Drug. Deliv. 17 (2010) 313 321. [98] M. Prabaharan, R. Jayakumar, Chitosan-graft-β-cyclodextrin scaffolds with controlled drug release capability for tissue engineering applications, Int. J. Biol. Macromol. 44 (2009) 320 325.

Handbook of Chitin and Chitosan

C H A P T E R

14 Fungal chitosan: prospects and challenges Joseph Sebastian1, Tarek Rouissi1 and Satinder Kaur Brar1,2 1

INRS-ETE, Universite´ du Que´bec, Que´bec, QC, Canada, 2Department of Civil Engineering, Lassonde School of Engineering, York University, Toronto, ON, Canada O U T L I N E

14.1 Introduction 14.1.1 Chitin and chitosan

420 421

14.2 Current commercial production and its disadvantages

426

14.3 Green synthesis of chitosan

427

14.4 Fungal chitosan 14.4.1 Significance of fungal sources of chitosan 14.4.2 Production of fungal chitosan

429 429 431

14.5 Future prospects

446

14.6 Conclusion

447

14.7 Acknowledgments

448

References

448

Online resource

452

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00014-6

419

© 2020 Elsevier Inc. All rights reserved.

420

14. Fungal chitosan: prospects and challenges

14.1 Introduction Natural biopolymers are biocompatible, biodegradable, extremely bioactive, and replenishable, due to being obtained from renewable sources, and these qualities have rendered biopolymers highly desirable for a wide variety of industrial applications. The suitability and flexibility of natural biopolymers to chemical and enzymatic modifications have led to derivatives with diverse applications and the general awareness about their safety and biodegradability has fueled their demand. Cellulose, starch, pectin, chitin, and chitosan are some examples of biopolymers that have been identified to have applications in diverse industries. Most of the natural polysaccharides are either neutral or acidic in nature, whereas chitin and chitosan are examples of highly basic polysaccharides [1]. Chitin and chitosan are natural polymers, widely distributed in nature, that play a key protective role, similar to cellulose in plants, for many crustaceans, insects, and lower eukaryotes such as diatoms and fungi. Even though the polymers are widely distributed in nature, currently crustacean waste and fish scales are used for the commercial production of chitin and chitosan, which require large amount of chemicals for extraction. The lower eukaryotic organisms have been considered as an alternate source for the isolation of chitin and chitosan but due to reasons similar to production from shellfish waste, wherein the amount of toxic waste produced and quantity of chemicals required for extraction and purification of the polymer, no significant progress has been made in using them. Moreover, the risk of overexploitation of these organisms for commercial production could lead to detrimental environmental impacts [2,3]. The current advances in fermentation technology provide an alternate means of eco-friendly production of the polymer. Chitin and chitosan are present as an integral part of the cell wall of fungi and significant quantities of chitosan can be obtained by culturing fungi on suitable media, most likely residues from the food processing industry as well as organic residues. This method of chitin and chitosan production is more sustainable and environment-friendly compared with the current method. Moreover, polymers of consistent quality and quantity can be produced year-round, as they are not subject to seasonal variabilities. In addition, the fungal cultures can be utilized for biotransformation of chitin, present in shellfish waste as well as waste mycelium obtained after the industrial production of compounds like citric acid and antibiotics, to chitosan [4,5]. This review provides a brief summary about chitin and chitosan biopolymers, their properties, applications, and production from fungal sources.

Handbook of Chitin and Chitosan

14.1 Introduction

421

14.1.1 Chitin and chitosan Chitin is a natural polysaccharide of major importance made up of β-(1
4)-N-acetyl-D-glucosamine (GlcNAc) residues. It is synthesized by a wide variety of living organisms and considered to be the most abundant polymer after cellulose. The polymer occurs in nature as ordered crystalline microfibrils that form the structural component in the exoskeleton of arthropods as well as the cell walls of fungi and yeast. Chitosan on the other hand is obtained after the deacetylation of chitin and is made up of glucosamine (GlcN) residues. Both chitin and chitosan, being renewable natural polymers, are currently being explored intensively for their applications in pharmaceutical, cosmetics, biomedical, biotechnological, agricultural, food, and nonfood industries (water treatment, paper, and textile). These unique polymers have been recommended for various applications in diverse fields due to their versatile biological activity, excellent biocompatibility, complete biodegradability, and low toxicity. To exploit their unique properties and to realize full potential of these versatile polysaccharides, attempts are being made to derivatize them [1,6]. 14.1.1.1 Chitin Chitin is a natural biodegradable polymer that has low toxicity and is inert in the gastrointestinal tract of mammals. The biodegradability of chitin owes to the presence of chitinases enzyme, widely distributed in nature and found in bacteria, fungi, and plants, and in the digestive systems of many animals. Moreover, chitinases are involved in host defense against microbial invasion. For example, lysozymes from egg white, fig, and papaya plants, degrade chitin and bacterial cell walls [7,8]. Chitin is an important natural polysaccharide made up of β-(1
4)-Nacetyl-D-glucosamine (GlcNAc) monomer units. It is synthesized by a wide variety of living organisms, represented in Table 14.1, and is present as a structural component in the exoskeleton of arthropods, as well as the cell walls of fungi and yeast. In fungi, chitin is present in the inner layer of the cell wall close to the plasma membrane and responsible for the maintenance of its shape, strength, and integrity. The biopolymer is also produced by higher organisms, mostly invertebrates, such as arthropods, protozoa, nematodes, mollusks, coelenterates, and rotifers, as well as produced by a number of living organisms in the plant and animal kingdoms, where reinforcement and strength are required. Due to the abundant presence of the polymer, it is considered as the second most abundant polymer after cellulose [2,8]. Chitin is synthesized at the plasma membrane, by chitin synthase, and extruded out but the precursor uridine diphosphate-N-acetylglucosamine is synthesized in the cytoplasm from glucose-6-phosphate. The glucose-6phosphate, obtained from glucose by the action of hexokinase, is

Handbook of Chitin and Chitosan

422 TABLE 14.1

14. Fungal chitosan: prospects and challenges

Examples of organisms that have shown presence of chitin.

Organism

Location in organism

Examples

Fungi

Cell wall

Most yeasts and filamentous fungi- Rhizopus oryzae, Mucor rouxii, Gongronella butleri

Nematodes

Eggshells

Ascaris suum

Pharynx

Ascaris lumbricoides, Caenorhabditis elegans

Microfillarial sheath

Brugia Malawi, Onchocerca volvulus

Arthropods

Cuticle

Insects, Arachnids and Crustaceans

Protozoa

Cyst wall

Entamoeba, Giardia

Cell surface

Phytomonas, Trichomonas, Blastocystis

Algae

Cell wall

Diatoms and green/brown algae

Molluscs

Squid pens, shells, radula

Gastropods, Cephalopods, Solenogastres and Polyplacophora

Adapted from M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (2006) 603
632. doi:10.1016/j.progpolymsci.2006.06.001; J.P. Martı´nez, M.P. Falomir, D. Gozalbo, Chitin: A Structural Biopolysaccharide With Multiple Applications, ELS, John Wiley & Sons, Ltd, Chichester, UK, 2014. , https://doi.org/10.1002/9780470015902.a0000694.pub3 . .

transformed to fructose-6-phosphate by the enzyme phosphoglucoisomerase. This is then converted to glucosamine-6-phosphate in the presence of glutamine. The glucosamine-6-phosphate so formed is converted, in the presence of the coenzyme acetyl-CoA, to N-acetylglucosamine-6-phosphate, which is then transformed to N-acetylglucosamine-1-phosphate by the action of mutase enzyme. This glucosamine-phosphate, in the presence of uridine triphosphate (UTP), is converted to uridine diphosphateN-acetylglucosamine. The chitin synthase enzyme acts on this to produce chitin, which is then deacetylated by the action of chitin deacetylase to from chitosan. This metabolic pathway of chitin and chitosan synthesis is represented in Fig. 14.1 and their structures represented in Fig. 14.2 [2]. Currently, chitin is commercially sourced from crab and shrimp shells, even though it is considered as the second most abundant polymer. It is extracted from the shells of crustaceans initially by acid treatment to dissolve calcium carbonate, followed by alkaline extraction to solubilize proteins. This is followed by a decolorization step which is often added to remove leftover pigments and to obtain a colorless product. These treatments have to be adapted to each chitin source, owing to differences in the composition of the initial materials. The harshness of the isolation process and the variability in the raw materials cause inconsistencies in the physical and chemical properties of chitin, such as degree of polymerization, and molecular weight, thereby limiting its

Handbook of Chitin and Chitosan

14.1 Introduction

423

FIGURE 14.1 Pathway of chitin and chitosan biosynthesis. Source: Adapted from J.P. Martı´nez, M.P. Falomir, D. Gozalbo, Chitin: A Structural Biopolysaccharide with Multiple Applications, ELS, John Wiley & Sons, Ltd, Chichester, UK, 2014. ,https://doi.org/10.1002/ 9780470015902.a0000694.pub3..

FIGURE 14.2 Structure of chitin (A) and chitosan (B).

Handbook of Chitin and Chitosan

424

14. Fungal chitosan: prospects and challenges

applications. Moreover, the presence of even a minute quantity of residues, other than chitin, has the potential to induce allergic reactions in people sensitive to shellfish and risk their well-being [1,6]. 14.1.1.2 Chitosan Chitin becomes soluble in aqueous acidic media when the degree of deacetylation reaches about 50% (depending on the origin of the polymer) and this deacetylated form is called chitosan. It is the only pseudonatural cationic polymer and hence it finds many applications that utilize this unique feature such as protein recovery and depollution. Being soluble in aqueous solutions, chitosan can be used in diverse applications as solutions, gels, or films and fibers. In the solid state, chitosan is semicrystalline in nature and has a polymorphous morphology [1,8]. The first step in the characterization of chitosan is purification of the sample. The sample is dissolved in excess acid and filtered on porous membranes of different pore diameters down to 0.45 μm. The pH of the solution is then adjusted to 7.5 by adding NaOH or NH4OH. This causes flocculation due to deprotonation and insolubility of the polymer at neutral pH. The polymer is then washed with water and dried [8]. Chitosan (structure represented in Fig. 14.3) is an important derivative of chitin and is obtained after the deacetylation of chitin, represented in Fig. 14.4, and is made up of glucosamine (GlcN) residues. The deacetylation renders chitin soluble in aqueous acidic media and the solubilization occurs by protonation of the amine (
NH2) group. This converts the polysaccharide to a polyelectrolyte and makes it a more applicable and valuable product than the chitin. The biopolymers have been recommended for various applications in diverse fields due to their versatile biological activity, excellent biocompatibility, complete biodegradability, and low toxic nature. Due to its unique property, it finds many applications in numerous industries, such as food industries (preservative, antimicrobial, coating, antioxidant), cosmetology (hair additives, lotions, facial and body creams), biotechnology (emulsifier, chelator, flocculent), agriculture (fungicide, films, soil modifier, elicitor) in addition to its pharmacological and medical uses (fabrics, fibers, artificial organs, drugs and DNA delivery, membranes) [1,5,6,9].

FIGURE 14.3 Structure of chitosan.

Handbook of Chitin and Chitosan

14.1 Introduction

425

FIGURE 14.4 Conversion of chitin to chitosan. TABLE 14.2 Various applications of chitosan in different area. Area

Application

References

Agriculture

Defensive mechanism in plants, stimulation of plant growth, seed coating, frost protection, time release of nutrients and fertilizers

[1,2,8,10]

Water and waste treatment

Flocculant, metal ion removal, odor removal, dye removal

[11
14]

Food and beverages

Bind lipids, preservative, thickener, protective, fungistatic, antibacterial coating, nutritional supplement

Cosmetics and toiletries

Maintain skin moisture, acne treatment, improve suppleness of hair, tone skin

Pharmaceutical

Immunologic, antitumoral, hemostatic, fungistatic, bacteriostatic, DNA and drug delivery, wound dressing

[2,10,15
17]

[1,2,8] [1,2,15,17
22]

14.1.1.3 Applications of chitosan The positive charge of chitosan provides the polymer with numerous and unique physiological and biological characteristics with application in a wide range of industries. These diverse applications, represented in Table 14.2, in multiple fields make chitosan a valuable compound of commercial significance [1,2,5,23]. The versatile physicochemical properties have made chitosan a unique and important polysaccharide, with wide variety of applications

Handbook of Chitin and Chitosan

426

14. Fungal chitosan: prospects and challenges

especially in the pharmaceutical, biomedical and food industry. Chitosan can be used for the delivery of drug molecules, as well as protein, nucleic acid, and virus particles, due to the presence of multiple attachment sites present on the polysaccharide chain. Moreover, the nonallergic, nontoxic, and biocompatible properties of chitosan and its derivatives have led to its wide use in the medical field. Chitosan and its derivatives have been shown to possess antimicrobial, antiviral, antitumor, antioxidant, hypocholesterolemic, antiasthmatic, and antiinflammatory properties as well. These properties have furthered the importance of chitosan for the pharmaceutical industry [1,2,15,17
22]. Chitosan is a biocidal agent against a wide range of target organisms. The activity considerably varies with the type of chitosan, the target organism, and the environment in which it is applied. In general, yeasts and molds are the most sensitive group, followed by Gram-positive bacteria and finally Gram-negative bacteria. The antimicrobial activity of chitosan is demonstrated to correlate with its cationic nature (positively charged), whereas many of the microbial cell components (cell wall, DNA, RNA) are negatively charged. The interaction between chitosan and microbial cells could be on the cell surface, which leads to increased permeability of the cell wall and leakage of intracellular components, or inside the cell, which inhibits DNA and RNA synthesis and directs cells into death. The exact mechanism of the antimicrobial effect of chitosan, chitin, and any other derivatives is not known yet but several mechanisms have been proposed. Most of these mechanisms involve some kind of damage or interaction with the cell membrane and DNA, thereby preventing proliferation [22]. In addition to the current and potential applications of chitosan as a pharmaceutical compound, it has shown versatile applications in the food and beverage industry as well as in agriculture. Another important sector in which chitosan can have a significant impact is the agricultural sector, as chitosan exhibits antivirus and antiphage activities [2,8,10,15
17]. Additionally, chitosan is known as the best heavy metal adsorbent among all the polymers and has been used as a chelating agent. This chelating ability is mainly due to the presence of the high number of amino and hydroxyl groups present on the polymer chain. Hence, it can be used for the removal of hazardous heavy metal ions, radioactive isotopes, and dyes from contaminated water and sewage efficiently [11
14].

14.2 Current commercial production and its disadvantages Chitin and chitosan are obtained commercially from crab and shrimp shells. It is extracted from crustaceans by acid treatment to dissolve calcium carbonate followed by alkaline extraction to solubilize proteins, followed by a decolorization step which is often added to remove

Handbook of Chitin and Chitosan

14.3 Green synthesis of chitosan

427

leftover pigments and obtain a colorless product. These treatments must be adapted to each chitin source, owing to differences in the ultrastructure of the initial materials. The most important derivative of chitin, chitosan, is obtained by partial deacetylation under alkaline conditions. These stages in production, from crab and shrimp shells, produce large quantities of highly alkaline and acidic effluents rich in proteinaceous waste materials during the extraction process, in addition to the long duration involved in the extraction process which can take up to 2 days. The current chemical approach for production of chitosan from chitin sourced from the shells of crustaceans limits its use, as the chitosan obtained shows inconsistencies in both physical and chemical properties. Moreover, the dependence on crustaceans causes environmental pollution associated with the technique used for commercial production [8,24
26]. Chitosan is currently produced from the shell of crabs and shrimps discarded by processing industry based in the United States and Japan. The production of chitosan from the shell waste is economical and sustainable to an extent but not environment-friendly as it produces large quantities of harmful effluents. About 6.3 kg of HCl and 1.8 kg of NaOH is needed for the production of every kilogram of 70% deacetylated chitosan, in addition to nitrogen and water for processing and cooling. These requirements can significantly increase the price of chitosan, in addition to producing a significant amount of toxic effluents rich in organic content, which currently stands at US$1000
1500 per kg, as quoted on Sigma-Aldrich. According to the work done by researchers of the Central Institute of Fisheries Technology, India, the average chitinous solid waste fraction of shellfish caught in India ranges from 60 to 80,000 tons. It was observed that dry prawn waste and dry squilla contained 23% and 15% chitin, respectively [1].

14.3 Green synthesis of chitosan The quantity of chemicals required, toxic effluents produced, and time required for the extraction of chitosan from crustacean waste, in addition to the issues of variabilities and purity of chitosan extracted from shellfish waste, has necessitated the use of alternate means of chitosan production either by the switch to fungal chitosan or by modern means of extraction. The switch to fungal sources produces chitosan of consistent quality as well as providing flexibility to produce chitosan of required properties by using different fungal strains or by varying the culture conditions. Moreover, the extraction of chitosan from fungal sources requires profoundly a smaller amount of toxic ingredients and hence produces a smaller amount of toxic waste [23]. The production of

Handbook of Chitin and Chitosan

428

14. Fungal chitosan: prospects and challenges

chitosan from fungal sources requires a shorter duration and concomitant lower energy requirement, and their significance is discussed in detail in the following section. The switch to a biological means of extraction by using enzymes or by different microorganisms is more effective and more environmentfriendly than chemical methods of extraction. Microorganisms, such as Lactobacillus sp., Pseudomonas aeruginosa, Bacillus licheniformis, Bacillus subtilis, and Bacillus cereus, mediate deproteinization and demineralization, and can curtail the amount of chemicals required for the extraction of chitosan, thereby reducing the amount of toxic effluent produced. The biological means of extraction can also be performed using pure enzymes, such as trypsin and alcalase, for deproteinization, followed by lactic acidmediated demineralization. The degree of deproteinization and demineralization can range between, 60%
95% and 70%
95%, respectively, based on the biological means used for the stages. In addition to the biological means of deproteinization and demineralization, deacetylation can also be achieved by use of either deacetylase enzyme obtained from fungal strains such as R. oryzae or by biotransformation of chitin to chitosan, by culturing fungus in the presence of the chitin obtained from crustacean waste [23,27,28]. These means of chitosan production cause little detrimental impact on the environment and greatly improve the green credentials of chitosan production but the coculture method needs further optimization before being adopted at a commercial scale. The advent of microwave-assisted extraction has revolutionized chemical extraction and the use of microwave heating in place of conventional heating has the potential to reduce the extraction time of chitosan to minutes from hours or days. The use of microwave radiation is a more efficient, environment-friendly, and energy-saving way to extract chemicals, in this case chitosan. During conventional heating, the reactants are slowly activated, not uniformly, whereas microwave heating happens at molecular level leading to a uniform rapid rise in temperature. The conventional method of chitosan extraction takes time and consumes a lot of energy. Hence microwave technology is an attractive alternative, as chitosan produced from microwave heating reduces to a few minutes the time taken to reach the same degree of deacetylation as traditional methods. In addition, the use of microwave heating, in place of conventional heating, during the stages of deproteinization and demineralization has been shown to significantly reduce the time further from a day to a few hours. Furthermore, it has been observed that the use of microwave-assisted extraction produced chitosan of higher molecular weight and crystallinity [29
32]. The shift from conventional heating to the more efficient extraction technique can lead to a reduction in the quantity of chemicals used, in addition to a lower energy requirement, thereby leading to a greener synthesis.

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

429

14.4 Fungal chitosan Chitin is an important component of fungal and yeast cell wall, as well as being present in other organisms. The presence of chitin in fungus, in the cell wall, was initially reported, in Mucor rouxii, by Hopkins in the year 1929. It was later, only in the year 1954, that the presence of natural chitosan was first identified by Kreger in the hyphae and sporangiophores of Phycomyces blakesleeanus. The interest in microbial chitosan production was propelled by the identification by BartnickiGarcia and Nickerson that chitosan was the most abundant component in the cell wall of the fungus M. rouxii. They observed that even though the fungus exhibits dimorphism, that is, filamentous and yeast-like forms, chitosan is the major component of the cell walls with concentrations of 32.7% and 27.9%, of the total cell wall composition, respectively. This observation led other investigators to look into the possibility of isolating chitosan from other fungal species, for commercial production, as well as using waste mycelia from industrial fungal cultures for the isolation of chitosan [33
35]. The unique physical, chemical, and biological properties and the versatility of chitin and chitosan, has led to the identification of a plethora of applications for the polymer and hence its importance.

14.4.1 Significance of fungal sources of chitosan Chitosan is generally produced from chitin that is a waste product of the seafood processing industry [1]. However, it has heterogeneous and inconsistent physiochemical properties since supplies of the seafood residues are subject to seasonal variabilities. Hence several yeasts and filamentous fungi that contain chitin and chitosan in their cell wall, for example, Schizosaccharomyces pombe, Candida albicans, Saccharomyces cerevisiae, M. rouxii, P. blakesleeanus, Coprinus cinereus, Neurospora crassa, Trichoderma reesei, Rhizopus spp., Absidia spp., Mucor spp., Mortierella isabelina, and Lentinus edodes, are being investigated for the production of chitosan. These microorganisms can be readily cultured in simple nutrients and used as an alternative source of chitosan. Also, the chitosan obtained from them show better uniformity in molecular weight and deacetylation and are free from potential life-threatening anaphylactic shock-inducing allergens [36]. The production of chitosan from fungus, which does not require stages of demineralization, deproteinization, and decolorization, provides an attractive alternate route for obtaining chitosan of consistent quality. The current advances in fermentation technology provide an alternate means of eco-friendly production of the polymer. Several yeast

Handbook of Chitin and Chitosan

430

14. Fungal chitosan: prospects and challenges

and fungal species, such as C. albicans, S. cerevisiae, M. rouxii, Cunninghamella elegans, Gongronella butleri, P. blakesleeanus, Rhizopus spp., and Absidia spp., have been investigated for chitosan production and discussed in the following sections [4,5,24,25,36
39]. The biotransformation of chitin, present in the cell wall, to chitosan can be achieved by the action of the enzyme chitin deacetylase present in fungi. The enzyme hydrolyzes the N-acetamido groups in the N-acetylglucosamine units of chitin and chitosan, thus forming glucosamine units and acetic acid. Chitin deacetylase (EC 3.5.1.4) was first identified and characterized in M. rouxii, and since then identified in other fungal, bacterial, and arthropod species. It is highly stable even at high temperatures and is not inhibited by acetate, a product of deacetylation, especially the one produced by C. lindemuthianum and A. nidulans. The enzyme is considered to have basically two biological roles, namely cell wall formation and interaction of pathogens with host organism, such as plants during infection. Based on the location of the enzyme during fungal growth, chitin deacetylase is classified into two groups, intracellular, located in the periplasm, and extracellular, secreted into the culture medium [40
43]. Fungal species such as, M. rouxii and Absidia coerulea, synthesize chitin deacetylases that are secreted into the periplasm, while species like Colletotrichum lindemuthianum and Aspergillus nidulans produce chitin deacetylases that are secreted into the culture medium [40]. The chitosan obtained by the action of the enzyme has a more regular and controllable deacetylation pattern than those obtained after treatment with hot NaOH [42]. The deacetylase enzyme based on the mechanism of action can be classified as exo-type and endo-type. Exo-type enzyme hydrolyze the acetyl groups according to a multiple attack mechanism, where the binding of the enzyme on a chitosan polymer or chitin oligomer substrate leads to a number of sequential deacetylations and then the enzyme binds to another new chain. The exo-type enzymes, from M. rouxii, are able to act only on oligomers with a degree of polymerization greater than 2 and are not able to act on the nonreducing end, as observed by Martinou and coworkers [44]. The endo-type enzyme, like the one isolated from C. lindemuthianum, catalyzes the removal of the acetyl group according to a multiple chain mechanism [45]. The enzyme is not able to deacetylate the reducing end residue, as observed by Tsigos et al. Moreover, it has been observed that the enzyme is effective against soluble chitosan and ineffective against insoluble chitin crystals. Pretreatment of crystalline chitin seems to be important for the enzyme to be effective [40]. The enzyme chitin deacetylase is synthesized by the microorganisms only during the period corresponding to their special biological roles [43]. For example, the fungus, C. lindemuthianum secreted the enzyme

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

431

while infecting the plant to prevent the plant’s immune system from recognizing the chitin present on the cell wall of the fungus. On the other hand, the chitin deacetylase was produced by M. rouxii during cell wall formation, whereas in the case of S. cerevisiae, it was produced only during sporulation. The chitin deacetylase synthesized by the marine bacteria, Vibrio cholera, help it to utilize chitin as a carbon, nitrogen, and energy source [40]. The enzyme can be used for the transformation of chitosan from chitin, sourced from either the waste of shellfish or waste mycelium obtained after industrial production of products like citric acid and antibiotics, as reported by Cai and team [46]. Use of the enzyme chitin deacetylase can help by pass the requirement of hot NaOH, used during chemical conversion, thereby significantly reducing the amount of harmful effluent produced during chitosan production. In addition to the use for the production of chitosan, the dependence of organisms, such as V. cholera, plant and human fungal pathogens and pest insects, on chitosan deacetylase can be exploited to develop biological control measures. Inhibitors of the enzyme can be used to prevent pathogens from infecting host organisms, like plants and animals, as well as control insect pests by modifying their gut [45].

14.4.2 Production of fungal chitosan Fungi are a promising alternative source of chitosan, in addition to their application as a biotransformer of chitin, sourced from shellfish waste, to chitosan. Garcia and Nickerson observed that the dimorphic fungus, M. rouxii, contains significant quantities of chitosan in the cell wall. They concluded that the fungus can be cultured on a simple medium and that the chitosan can be extracted from the cell wall [33]. These observations fueled the interest in investigating the possibility of obtaining chitosan from other fungal cultures, optimization of culture conditions to maximize production, and the use of alternate low-cost media for culturing the fungus. Chitosan prepared from fungi has received much attention, especially belonging to Zygomycetes species, as they are known to contain chitosan as natural components of their cell wall [13,47]. Fungi from Ascomycetes, Basidiomycetes, and Deuteromycetes also contain chitin as the principal structural component of their cell walls, but the presence of native chitosan has not been reported for these fungi. However, treatment with concentrated sodium hydroxide can simultaneously deacetylate the chitinous fraction, dissolve the proteins, remove the soluble glucan, and hydrolyze the lipids. Hence, by this treatment chitosan can be extracted from the chitinous, but native-chitosan-free strains of fungi [13]. Moreover, they can be treated to give chitosan of more consistent and desired

Handbook of Chitin and Chitosan

432

14. Fungal chitosan: prospects and challenges

physicochemical properties compared with chitosan obtained from crustacean sources. Also, the advances in fermentation technology provided us with an alternative route for eco-friendly production of the biopolymer [22]. A flowchart of the procedure for extraction of chitosan from mycelia of M. rouxii, used during the initial early days of chitosan production from fungus is represented in Fig. 14.5. The mycelia obtained after the culture of the fungus were harvested and washed with distilled water. These mycelia are homogenized and then autoclaved in the presence of 1 N NaOH (Biomass: NaOH::1:20 v/v). The alkali-treated fungal mycelia were then filtered and the supernatant discarded. The alkali-insoluble material (AIM) was collected and washed with ethanol and distilled water. The AIM is then homogenized in the

FIGURE 14.5 Flowchart for the extraction of chitosan from fungal mycelia. Source: Adapted from S. Arcidiacono, D.L. Kaplan, Molecular weight distribution of chitosan isolated from Mucor rouxii under different culture and processing conditions, Biotechnol. Bioeng. 39 (1992) 281
286. doi:10.1002/bit.260390305.

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

433

presence of 2% acetic acid, where the AIM:acetic acid ratio is 1:100. It is then centrifuged and the supernatant collected. The chitosan present in the supernatant is then precipitated by adjusting the pH to 8.5 and the precipitate is washed, centrifuged, and then lyophilized [48]. This extraction was slightly modified by increasing the amount of sodium hydroxide used to obtain alkaline-insoluble material. The modified method uses a biomass to NaOH ratio of 1:40 to obtain AIM [13]. Several yeast and fungal species have been investigated for chitosan production and one such large-scale investigation was performed by Hu and coworkers. They screened 33 fungal strains for their ability to produce chitosan. It is generally considered that chitin is the only structural polymer of Ascomycetes, Basidiomycetes, and Deuteromycetes and has no native chitosan, whereas strains belonging to Zygomycetes class are chitosan-containing fungi [13]. They observed that chitosan could be extracted from all the 33 fungal strains irrespective of the class they belonged to but native glucan-free chitosan was present in the fungus that belonged to the order Mucorales. The Mucorales strain, Absidia glauca was found to be a promising chitosan producer that can be used for commercial production of chitosan. The other strains that were identified for commercial chitosan production were A. nidulans, M. rouxii, Mucor hiemalis, Penicillium digitatum, and G. butleri. It was also concluded that chitosan is extractable from some common industrial fungal mycelia, such as A. niger, A. gossypii, P. chrysogenum, and Aspergillus spp. and hence waste mycelia of these fungi can be used for chitosan production [13]. The other strains that have been investigated for their ability to produce chitosan are Rhizopus oryzae, A. coerulea, Cunninghamella echinulata, and C. elegans [24,36,47,49,50]. The concentration of chitosan obtained by researchers from culturing fungal strains on various media in their studies is represented in Table 14.3. M. rouxii, R. oryzae, and G. butleri are the three fungal species, belonging to the Mucoraceae family, that have been studied extensively for their ability to produce chitosan and suitability for commercial production. Some of the previous studies performed using these strains are discussed below. 14.4.2.1 Mucor rouxii M. rouxii was used as the model organism for most of the initial studies on the production of fungal chitosan. One of the initial investigations that studied the feasibility of fungal chitosan production was performed by White, Farina, and Fulton. They developed a method for laboratory-scale production and isolation of chitosan (polyglucosamine) from the mycelium. The biomass yield obtained was in the range of 16%
22% of which 35%
40% was glucosamine and a chitosan yield of 4%
8% of the weight of dry biomass material was obtained. Due to higher power requirements

Handbook of Chitin and Chitosan

434

14. Fungal chitosan: prospects and challenges

TABLE 14.3 Chitosan concentration obtained from various fungal strains (SMF, submerged fermentation; SSF, solid-state fermentation). Fungal strain/fermentation type

Chitosan yield

Reference

Rhizopus oryzae/SSF

4.3 g/kg of soyabean residue

[36]

Mucor rouxii/SMF

0.61 g/L for molasses salt medium

[39]

A. glauca/SMF

7:4% of dry cell weight 3:9% of dry cell weight 3:8% of dry cell weight

A. nidulans/SMF M. rouxii/SMF G. butleri USDB 0201/SMF

g

[13] PGY salt broth

93:4 mg=200 mL medium 79:3 mg=200 mL medium 76:6 mg=200 mL medium

C. echinulate/SMF G. butleri USDB 0489/SMF

g

[49] Nutrient broth

M. rouxii/SMF

7.3% of dry mycelial weight—synthetic medium

[51]

G. butleri USDB 0201/SSF

4
6 g/100 g mycelia—sweet potato

[52]

Mucor racemosus/SMF

35:1 mg=g dry mycelia YPD 20:5 mg=g dry mycelia

[24]

Mucor rouxii/SMF

4%
8% dry mycelial weight—YPD

[4]

Mucor rouxii/SMF

12:7 6 5:0%dry mycelia YM ðDIFCOÞ 10:1 6 2:9% dry mycelia

[5]

Gongronella butleri CCT4274/ SMF

1.19 g/L of chitosan per liter of apple pomace extract culture medium or 21% of the dry cell weight

[39]

R. oryzae/SMF

1.13 6 0.10 g/L of whey medium (in presence of gibberellic acid)

[38]

R. oryzae TISTR 3189/SMF

138 mg/g or 14% of dry mycelial weight— PDB medium

[50]

G. butleri IF08081/SMF

730 mg/L of Shochu distillery effluent

[37]

M. rouxii/SMF

5%
10% of total biomass dry weight— defined and complex synthetic media (YPG, BG and TVB)

[48]

A. coerulea/SMF

4.11 6 0.08 g/L of synthetic medium containing soybean pomace as nitrogen source

[53]

g g

Cunninghamella elegans/SMF

P. blakesleeanus/SMF

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

435

during industrial-scale production, due to mycelial growth they suggested screening for alternate thermophilic, anaerobic pellet-forming cultures [4]. Knor and Klein studied the suitability of using P. blakesleeanus in addition to M. rouxii for the production of fungal chitosan. They obtained yields of 7.4%
23.3% of dry biomass from M. rouxii and 9.0%
12% of dry cell material from P. blakesleeanus. They also reported that the two strains were able to convert commercial chitin obtained from shellfish and chitin from waste mycelia of A. niger into chitosan. This observation can be exploited to obtain chitosan of the desired degree of acetylation from commercially sourced chitosan via fermentation using these organisms [5]. The work by White et al. and Knor and Klein focused primarily on the feasibility of production of chitosan by M. rouxii and detailed investigations on the physicochemical properties were not carried out. The molecular weight and degree of deacetylation plays a crucial role in determining the application of chitosan and hence understanding of these parameters is vital. Arcidiacono and Kaplan investigated the yield, molecular weight distribution, and degree of acetylation of chitosan isolated from the M. rouxii mycelial cell wall under different growth and extraction conditions. The weight average molecular weight of chitosan obtained was determined by gel permeation chromatography and found to range from 2.0 3 l05 to 1.4 3 l06 Da. The chitosan yield ranged from 5%
10% of total biomass dry weight while the degree of deacetylation ranged between 87%
92%. It was also observed that the duration of incubation and medium composition are the two parameters that affected biomass production and molecular weight of chitosan. In addition, it was concluded that chitosan obtained directly from the cell wall of the fungus had a higher degree of deacetylation than commercial chitosan from the chemical conversion process and that the dissolved oxygen level in the medium is a crucial factor that influences biomass yield in addition to mixing [48]. Arcidiacono and Kaplan observed that the molecular weight of chitosan increased during the first 3 days of fermentation and then declined with further incubation. They concluded that this decrease in molecular weight might be due to chitosan being degraded by the action of chitosanases during autolysis or modified during arthrospore formation. This observation was not in accordance with the observation made by Synowiecki and Al-Khateeb (1997). They concluded in their work, which investigated the production of chitosan by culturing of M. rouxii on YPG media, as well as the effect of prolonged incubation on chitosan, that no significant influence was observed in the amount of available chitosan, even after prolonged cultivation. They were able to obtain, after 2 days of incubation, 8.9% and 7.3% (dried biomass weight) of chitin and chitosan, respectively [51]. These amounts of chitosan obtained were in agreement with the observations made during earlier works.

Handbook of Chitin and Chitosan

436

14. Fungal chitosan: prospects and challenges

Chatterjee and coworkers studied chitosan production by fungus M. rouxii cultured in three different media, molasses salt medium, potato dextrose broth, and yeast extract glucose, and concluded that molasses salt medium was the most suitable media for the commercial production of chitosan as it was the cheapest of the three media investigated. The three media yielded almost the same amount of chitosan, 0.6 g/L for MSM, 0.56 g/L for YPG and 0.51 g/L for PDB but upon investigation of the physicochemical properties it was found that the chitosan obtained from MSM was less polydispersed and more crystalline compared with those from YPG and PDB. The degree of deacetylation of the chitosan obtained was observed to be similar irrespective of the media used. It was also observed that the addition of excess sucrose in molasses salt medium did not improve growth of the fungus but the highest amount of biomass yield was obtained when YPG media was used, 18% higher biomass yield was obtained when compared to that obtained using MSM media but it took longer to achieve the biomass yield. This indicates that amount of nitrogen in the media might be the limiting factor that dictates biomass yield and ability of the media to support fungal growth for longer periods [25]. The studies on production of fungal chitosan using M. rouxii and their findings are represented in Table 14.4.

TABLE 14.4

Studies performed using Mucor rouxii and chitosan yield.

Culture media/ fermentation type

Chitosan yield/molecular weight

Degree of acetylation/ deacetylation*

References

Synthetic YPD medium/SMF

4%
8% dry mycelial weight




[4]

Synthetic medium— YM/SMF

9%
18% dry mycelial weight

0.5%
5%

[5]

Synthetic medium/SMF

5%
10% of total biomass dry weight/200
1400KDa

8%
13%

[48]

Synthetic medium/SMF

7.3% of dry mycelial weight

27.3%

[51]

Molasses salt mediuma/ PDBb/YPGc/SMF

0.61g/L
2.48 3 104 Daa/0.51 g/L
4.58 3 104 Dab/0.56 g/ L
5.59 3 104 Dac

87.2a/89.8b/ 82.8c*

[25]

Synthetic medium— YPD/SMF

12.49% mycelial dry weight

19.5%

[54]

*

Corresponds to when degree of deacetylation is presented. In the study three different media were used and each media gave different chitosan yield, molecular weight and degree of deacetylation. SMF, Submerged fermentation; SSF, solid-state fermentation.

a,b,c

Handbook of Chitin and Chitosan

437

14.4 Fungal chitosan

14.4.2.2 Rhizopus oryzae R. oryzae is a filamentous fungus that belongs to the division of Zygomycota. The strains belonging to this species of fungus are generally considered as safe (GRAS). It is one of the fungal strains that is generally used for the production of industrially important platform chemicals, such as L-lactic acid, fumaric acid, and ethanol, at high concentrations, in addition to a wide variety of commercial enzymes, such as amylase, xylanase, pectinase, and cellulase. The fungus is also used in the production of traditional fermented food products, such as tempeh, famous in Indonesia and Malaysia, as well as alcoholic beverages. The fungal strain requires only simple media for its growth and is able to grow in temperatures ranging from 25 C to 45 C and pH ranges of 4
9, but requires higher humidity [55,56]. These observations and wide use in the industry make the fungal strain ideal for the production of fungal chitosan both from the waste mycelium and by culturing the fungus in simple media. Table 14.5 provides a summary of the research on the potential use of the fungal strain for chitosan production. The initial work that studied the feasibility of using R. oryzae mycelia as a possible source of chitosan was performed by Hang (1990). Chitosan yield of 700 mg/L was obtained by culturing the fungus on simple rice under submerged fermentation (SMF) conditions. This work showed that R. oryzae can be used as a source of fungal chitosan in addition to M. rouxii. It was observed by Tan et al. that the highest concentration of chitosan was obtained during the late exponential growth TABLE 14.5 yield.

Studies on using Rhizopus oryzae in chitosan production and chitosan Degree of acetylation/ deacetylation*

Reference

138 mg/g dry weight/ 6.9 3 104 Da

87.9 6 2.1*

[50]

Whey medium with plant growth hormones/SMF

0.749
1.131 g/L of medium/120
270 kDa

87.2
87.5 6 0.5*

[38]

Synthetic medium/SMF

55.70 6 8.53 and 43.98 6 3.05 mg/200 mL substrate




[49]

Soybean and mungbean residues/SSF

4.3 g/kg of soyabean residue




[36]

Rice medium/SMF

700 mg/L




[57]

Rice straw/SSF

5.63 g/kg of medium

90%*

[18]

Culture media/ fermentation type

Chitosan yield/molecular weight

Potato dextrose broth/ SMF

SMF, submerged fermentation; SSF, solid-state fermentation.

Handbook of Chitin and Chitosan

438

14. Fungal chitosan: prospects and challenges

phase, by culturing the fungus on a synthetic medium under submerged conditions, and a similar observation was made for other fungal strains [48,49,58]. The observations made point to the significance of identifying the period during the growth phase that provide maximum chitosan yield, which can vary based on the fermentation conditions and the media used for production. Pochanavanich and Suntornsuk were able to obtain highest yield of chitosan, 138 mg/g cell dry weight, from R. oryzae cultured with potato dextrose broth. The molecular weight and degree of acetylation of the chitosan obtained were 6.9 3 104 Da and 87.9%, respectively, in the study, which investigated four different filamentous fungi representing four different species, A. niger TISTR 3245, R. oryzae TISTR 3189, L. edodes, and Pleurotus sajo-caju, and two yeast strains, Zygosaccharomyces rouxii TISTR5058 and C. albicans TISTR 5239 [50]. The ability of R. oryzae to grow on a simple medium, especially on industrial and agricultural residues, has been proven by multiple studies and chitosan of high quality and yield were obtained during these studies. Chatterjee et al., was able to produce chitosan of varying properties, such as molecular weight and degree of acetylation, by culturing the fungus on a whey medium supplemented with different plant hormones. The molecular weight of the chitosan obtained ranged from 120 to 270 kDa and the degree of deacetylation of 87%. Chitosan yield of 5.63 g/kg of medium and degree of deacetylation of 90% was obtained by culturing the fungus on rice straw [18]. The ability of the fungus to grow on residues and produce chitosan was further shown by Suntornsuk and team, by utilizing soybean and mungbean residues as the media for the production of chitosan. They were able to obtain yields of 4.3 g/kg of soyabean residue and 1.6 g/kg of mungbean residue [36]. These findings and the possibility of utilizing the mycelial biomass left after the production of commercial enzymes and platform chemicals make R. oryzae an ideal source of fungal chitosan. 14.4.2.3 Gongronella butleri G. butleri is another fungal species, belonging to the Mucoraceae family, that has been studied for its chitosan production ability. In most of the studies performed, where this species of fungus was used, the culture medium was made up mainly of agricultural and industrial residues. G. butleri has been studied for its chitosan production ability in both SMF and solid-state fermentation modes. In general, solid-state fermentation yielded a higher amount of chitosan but took longer. It has been observed that different strains of the fungus show significant differences in the quantity of chitosan produced. Two strains, G. butleri 0489 and G. butleri 0201, of the fungus grown on synthetic media produced different quantities, 76.63 6 9.00 and 93.38 6 6.69 mg/200 mL substrate,

Handbook of Chitin and Chitosan

439

14.4 Fungal chitosan

respectively, of chitosan [49]. This observation shows the importance of choosing the right strain for commercial chitosan production, even within species. The chitosan obtained during the studies showed a high degree of deacetylation (82%
96%) but varied in molecular weight, which ranged from 25 to 220 kDa, probably due to the medium composition, fermentation conditions, and the fungal strain used for the study [37,39,49,58,59]. Table 14.6 provides a summary of the chitosan yield, its properties, obtained during fermentation studies performed using the fungal species of G. butleri. 14.4.2.4 Other fungal strains The summary of the research on fungal chitosan production using other fungal species is provided in Table 14.7 with their corresponding yield and properties like molecular weight and degree of acetylation. The diversity of chitosan produced, with varying degrees of acetylation and molecular weight, by different fungal strains can be observed in the tables and discussions above. This flexibility, rendered by the use of

TABLE 14.6 yield.

Studies on using Gongronella butleri in chitosan production and chitosan Degree of acetylation/ deacetylation*

Reference

1.19 g/L of chitosan per liter medium




[39]

Shochu distillery effluent/SMF

730 mg/L

82%*

[37]

Sweet potato/SMF and SSF with N-supplementation

12.7/9.2% biomass SSF/ SMF with Urea 25
38 kDa SSF/ 70
120 kDa SMF

92
96%* (SSF)/94%
96%* (SMF)

[59]

Sweet potato pieces/SSF

4
6 g/100 g mycelia 160
220 kDa based on harvest time

13%

[59]

Sweet potato pieces with urea supplementation and varied pH/SSF

4.31 6 0.65 g/kg substrate (14.3 g urea)/mol. wt. increased with increase in pH and urea concentration

11%

[58]

Synthetic media/SMF G. butleri 0489 G. butleri 0201

76.63 6 9.00 and 93.38 6 6.69 mg/200 mL substrate




[49]

Culture media/fermentation type

Chitosan yield/Molecular weight

Apple pomace extract/SMF

SMF, submerged fermentation; SSF, solid-state fermentation.

Handbook of Chitin and Chitosan

440

14. Fungal chitosan: prospects and challenges

TABLE 14.7 Studies on using other fungal strains for chitosan production and their chitosan yield. Fungal strain/culture media/fermentation type

Chitosan yield/ molecular weight

Degree of acetylation/ deacetylation*

M. racemosus and C. elegans/synthetic media/SMF

35.1 and 20.5 mg/g dry mycelia

49% and 20% respectively

[24]

Absidia coerulea/potato pieces/SSF

6.12 g/kg substrate/ 6.4 kDa

85%*

[47]

Cunninghamella echinulate/synthetic media/SMF

79.73 6 6.04 mg/ 200 mL substrate




[49]

Absidia glauca/synthetic media/SMF

65.24 6 10.81 mg/ 200 mL substrate




[49]

A. coerulea/synthetic media/SMF

51 mg/100 mL of medium 4.5 3 105 Da

6.9% 6 1.5%

[12]

A. coerulea/synthetic-YM medium/SMF

9.4% dry mycelia/ 140 kDa

35%

[11]

C. blakesleeana/syntheticYM medium/SMF

10% dry mycelia/ 450 kDa

5%

[11]

Lentinus edodes/ SMF—synthetic media/ SSF—wheat straw

SMF—5.5%
12.5% varies with incubation time SSF—10%
12.5%

[60]

120 mg/L medium 6.18 g/kg substrate

A. butleri/synthetic medium/SMF

1 g/L of media

79.89%*

[61]

Aspergillus niger/ SMF—synthetic medium/ SSF—soybean residue

0.8455 g/L medium 17.05 6 0.95 g/kg dry substrate




[62]

References

SMF, Submerged fermentation; SSF, solid-state fermentation.

different fungal strains as well as media, can be exploited at industrial scale for the production of chitosan of desired properties. The observations made above show that the use of simple media, in addition to industrial residues, is crucial for commercial fungal chitosan production. Some of the observations regarding the use of industrial residues as potential media for chitosan production is discussed below. One of the main factors that decides the feasibility of the commercial production of a chemical

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

441

compound is the raw material cost and the use of industrial residues can significantly reduce the cost associated with the culture media. 14.4.2.5 Use of industrial residues as culture medium for fungal chitosan production The main factor that drives cost of production at commercial scale is the cost of the raw material. In the case of the commercial production of fungal chitosan, the main factor that determines the cost of production is the raw materials used for medium preparation. Hence, it becomes imperative that the media used be economical and composition kept simple to improve feasibility as well as to prevent issues during further processing of fungal biomass to obtain chitosan. Fungal culture media and fermentation condition can be modified to obtain chitosan of more consistent physicochemical properties as well as chitosan of various physical properties, such as molecular weight, molecular size, polydispered nature, crystallinity, and glucosamine content, by varying media composition and fermentation conditions [1,5,6,47,58
60]. Industrial by-products, such as molasses from the sugar industry, can be used as an inexpensive carbon source to grow fungi for the production of fungal chitosan, as shown by Chatterjee and coworkers. They were able to obtain chitosan yield of 0.6 g/L when molasses salt medium was used and the chitosan so obtained was found to be less polydispersed and more crystalline [25]. Several by-products from the food processing industry have been investigated for use as culture media for the production of chitosan. The suitability of soybean and mungbean residues as the media for production of chitosan was studied [36] and it was observed that the highest amount of chitosan, 4.3 g/kg of substrate, was obtained from R. oryzae grown on soybean residues. Another food processing industry byproduct that was investigated for its suitability to be used for the production of fungal chitosan is apple pomace left after juice extraction. Streit and team used the aqueous extract of apple pomace supplemented with sodium nitrate as a nitrogen source as the culture medium for fungus G. butleri CCT4274. It was found that the nitrogen-supplemented aqueous extract of apple pomace is a good medium for the development of microorganisms, due to its high content of reducing sugars (60 g/L). The chitosan yield obtained was 1.19 gm per liter of culture medium after 72.5 hours of cultivation, which represent around 21% of the biomass content [39]. The use of these food industry by-products has significance for commercial production, as it enables us to keep the cost of raw materials required for production low, thereby profoundly impacting the feasibility of the production of the biopolymer. Another possible medium that can be considered for fungal chitosan production is industrial organic effluents. Yokoi et al. studied chitosan production from shochu distillery wastewater and proved the feasibility

Handbook of Chitin and Chitosan

442

14. Fungal chitosan: prospects and challenges

of producing chitosan using sewage as the media for commercial production. The shochu distillery wastewater contained high concentrations of organic matters, where the chemical oxygen demand (COD) of the effluents were in the range of 19,500
21,200 mg/L, and can be used as culture medium. Fungal strains Absidia atrospora IF09471, G. butleri IF08080, and G. butleri IF08081 were investigated for their ability to produce chitosan by utilizing the organic matter present in the sewage. Two different shochu distillery wastewater media were investigated, namely barley
buckwheat
shochu distillery wastewater (BBS medium) and sweet potato
shochu wastewater (SPS medium) [37]. It was observed that the supernatant of both these media supported rich growth of the fungus but in the SPS medium the three strains grew in a wide pH range (4
8.5) and produced the most amounts of chitosan (730 mg/L). Hence, it was concluded that G. butleri IF08081 grown in SPS medium is most suitable for chitosan production. The added benefit of using organic sewage waste as the medium for production is that the fungi were able to remove 49% of COD, 51% of total sugar, 43% of reducing sugar, 45% of protein, 61% of total nitrogen, and 88% of total phosphorus [37]. This observation can be exploited and used as a stage in the treatment of sewage waste but the suitability of chitosan produced for commercial use needs to be investigated and as an alternative the chitosan obtained can be used in the sewage plant for removing heavy metal ions from the treated water. 14.4.2.6 Avenues and challenges of commercial production of fungal chitosan In addition to media composition, the key factor affecting both quantity and quality of chitosan produced is the fermentation conditions used for culturing the fungus. Fermentation conditions, such as pH, nitrogen content of the media, and harvest time of fungal biomass have an impact on fungal chitosan production. Chitosan is a nitrogen-containing biopolymer, which is the deacetylated form of chitin, and hence the fungus requires an inorganic or organic nitrogen source as a nutrient to synthesize the biopolymer for their cell wall, and this is one of the important factors for the production of chitosan by fungi [58,59]. Nitrogen can be supplied directly in the form of ammonium ions or other organic forms of nitrogen, such as urea. Nwe et al. studied the effect of various nitrogen sources, both organic and inorganic, and identified urea to be the most ideal nitrogen source for chitosan production using G. butleri. This observation is not in accordance with the observation made by Jiang et al. They studied the effect of various nitrogen sources on A. coerulea and observed that soybean pomace was the ideal nitrogen source compared to nitrogen supplied in the forms of sodium nitrate, ammonium

Handbook of Chitin and Chitosan

14.4 Fungal chitosan

443

sulfate, ammonium carbonate, and urea. They obtained 4.11 g/L chitosan by using soy pomace, whereas only 1.97 g/L was obtained when urea was used. Both these works do suggest that different molecular weight chitosan can be produced by using different nitrogen sources and that the molecular weight of chitosan increased with the increased concentration of nitrogen in the medium [53]. These observations suggest that the choice of nitrogen source has to depend on the fungal strain used for the production and molecular weight of the chitosan to be obtained. Nwe and Stevens investigated the effectiveness of using urea as a nitrogen source for production of fungal chitosan under solid-substrate fermentation conditions. They also suggested that the molecular weight of chitosan could be increased by using more N-source (peptone) in the SMF medium. Nwe and Stevens observed that pH 5.5
6.5 is best for the production of fungal chitosan from mycelia grown on solid-substrate fermentation medium supplemented with urea [58]. They observed that the weight average molecular weight and polydispersity of the chitosan were lower at lower pH and higher at pH about 5.5. The weight average molecular weight increased from 235 to 317 kDa with increasing pH (3.7
5.6). A similar observation was made in the case of polydispersity and it varied from 4.2 to 5.2 under the same pH range as mentioned earlier. These observations are in accordance with the observation made by Arcidiacono and Kaplan that the biomass of M. rouxii and molecular weight of chitosan reduced at pH 3 and molecular weight slightly increased at pH 6 in SMF. Hence factors, such as pH and nitrogen content, have a crucial role in chitosan synthesis and the type of chitosan produced. Therefore, needs to be monitored and controlled closely [48]. Another important factor that has a significant impact on fungal biomass and in turn on chitosan production is the fermentation technique used for the culture. Fungal biomass can be produced either by solidsubstrate fermentation (SSF) or SMF and a comparison of their advantages and disadvantages is presented in Table 14.8. The comparison of chitosan production in SSF and SMF processes has shown that the yield of chitosan using SSF (w/w) is higher than that in SMF (w/v) due to the low quantity of mycelia produced in SMF. Fifty times more chitosan (6 g/kg substrate), when compared to liquid fermentation, was obtained by SSF from L. edodes by Crestini, Kovac, and Sermanni [60]. Suntornsuk et al. were able to obtain 4.3 and 1.6 g of chitosan (per kg of substrate) by culturing R. oryzae on soybean residue and mungbean residue, respectively, which is significantly higher than that obtained by other researchers culturing the fungus on various liquid media, such as corn, rice, and complex media [36]. These observations are not in accordance with those of Nwe et al., who observed that the optimal production of biomass and chitosan by SMF was around 1.5
2.5 times higher than solid-state fermentation [59].

Handbook of Chitin and Chitosan

444

14. Fungal chitosan: prospects and challenges

TABLE 14.8 Comparison of advantages and disadvantages of solid-state and submerged fermentation. Solid-state fermentation

Submerged fermentation

Advantages:

Advantages:

• Better suited for fungal fermentation • Cost effective • Cheap raw materials like lignocellulosic waste can be used as culture media • Substrate used slowly and steadily, support long fermentation durations • Support mycelial growth

• • • • • •

Disadvantages:

Disadvantages:

• Difficult to control process parameters • Temperature build-up due to poor heat transfer • Uniform mixing not possible • Media composition not defined

• Media can be expensive • Large quantity of effluents • Substrate utilized fast, so supplementation required • Not ideal for fungal fermentation due to mycelium formation which leads to difficulty in mixing • Low mycelia formation

Better control over process parameters Defined media composition Better heat transfer Uniform mixing can be achieved Faster fermentation Downstream processing easier

Adapted from V. Maghsoodi, S. Yaghmaei, Comparison of solid substrate and submerged fermentation for chitosan production by Aspergillus niger, Sci. Iran. (2010) 153
157; R. Subramaniyam, R. Vimala, Solid state and submerged fermentation for the production of bioactive substances: a comparative study, Int. J. Sci. Nat. 3 (2012) 480
486 [63].

Different fungal strains have their own requirements to achieve maximum biomass yield and morphology development. Therefore a universal fermentation condition applicable to all fungal strains would not be possible and the fermentation parameters should be optimized for the fungus used for the production of chitosan. A brief summary of the effect of some of the important fermentation parameters observed by researchers is presented in Table 14.9. It was concluded by researchers that temperature in the range of 25 C
30 C and pH of 5.5
6.5 were ideal for fungal growth and hence possibly the optimum production of chitosan. They also observed that the molecular weight of chitosan can be modified by varying the pH of fermentation. The other media components that can have an impact on fungal chitosan production are nitrogen content of the media. Chitosan is made up of glucosamine residues and a suitable nitrogen source is essential for its synthesis as well as for biomass production. The amount of dissolved oxygen in the media has been shown to affect mycelial growth of the fungus M. rouxii and might have a similar effect on other fungal species as well. To this effect, mixing might have a crucial role in

Handbook of Chitin and Chitosan

445

14.4 Fungal chitosan

TABLE 14.9 Fermentation parameter

Impact of various fermentation parameters and conditions. Impact on chitosan yield

References

Temperature

25 C
30 C ideal for fungal culture for optimum chitosan production Fungal strains: M. rouxii, M. racemosus, C. elegans, R. oryzae

[24,25,48,50]

pH

pH 5.5
6.5 is best for the production of fungal chitosan using fungal strains, M. racemosus, C. elegans, and G. butleri, in SSF and molecular weight slightly increased at pH 6 in SMF

[48,58]

Incubation time

Late exponential phase harvest of mycelium to obtain maximum chitosan yield from fungal strains M. racemosus, C. elegans, G. butleri, and C. echinulata

[48,49,58]

Nitrogen

Nitrogen essential for chitosan production and chitosan from fungal strains, G. butlerii, A. coerulea, M. rouxii, and R. oryzae, of varied mol. wt. can be obtained by using more different N-source (like peptone)

[39,47,58,59]

Mixing

Low speed led to lower growth and chitosan yield by A. coerulea

[64]

Dissolved oxygen

Mucor rouxii requires oxygen for mycelial growth

[48]

Mode of fermentation: • SSF and SMF • Batch and continuous

[36,58,59,64,65] SSF led to higher chitosan yield from fungus, G. butlerii and R. oryzae, and ideal for low-molecular-weight chitosan Continuous fermentation led to mycelial accretion and lower chitosan yield from A. coerulea and A. glauca

SMF, submerged fermentation; SSF, solid-state fermentation.

chitosan production and its possible effect and significance are discussed in the following section [48]. The use of high inoculum size (5.8 3 107 spores) can lead to an increased initial growth rate but this growth leveled off earlier due to the agglomeration of fungal mycelium, as observed by Davoust and Hanson. Mixing or agitation can play a crucial role in overcoming this issue. Filamentous fungi are adapted to grow on solid substrates and when cultured in liquid medium cause an increase in viscosity. This leads to poor maintenance of dissolved oxygen levels as well as inadequate mass

Handbook of Chitin and Chitosan

446

14. Fungal chitosan: prospects and challenges

transfer, leading to poor utilization of media components. The use of traditional disc turbines, which are designed for low viscosity liquids, does not provide adequate mixing when used for fungal cultures. The presence of mycelium increases viscosity and hampers mixing, especially fungus with nonseptate hyphae like Zygomycetes, whereas fungus with septate hyphae fragment easily and lead to a reduction in viscosity. It was observed by Davoust and Hanson that low stirrer speed led to the formation of a pulp with decline in growth rate whereas increased stirrer speed caused the formation of small pellets. This can be overcome by using an open paddle impeller-type agitator with a large diameter, run at a high stirrer speed, which allows for satisfactory growth of most of the fungal species investigated and no viscous pulp formation [64]. Harvesting time is another important factor, in addition to fungal strain, fermentation type, medium composition, and physical parameters, that decides the quality and quantity of chitosan extracted from the fungal mycelia. Most often the fungal mycelia are harvested after a fixed incubation time, 3
4 days for SMF and 10
12 days for solid-state fermentation, because it becomes difficult to extract chitosan from the mycelia after its active growth phase. The amount of extractable chitosan decreases and becomes more difficult to obtain due to physiological changes in the fungal cell wall toward the end of fermentation. Tan et al. observed that after 74 h of fermentation the amount of chitosan from R. oryzae decreased gradually. A similar observation was made for Absidia butleri by Shimahara et al. (1988), as cited by Tan et al., and they observed that even though growth continued until day 7, the maximum amount chitosan was extracted on day 4 [49]. During active growth phase or exponential phase, the amount of free chitosan is high due to active growth. Once the cells enter stationary phase the chitosan is bound to chitin and other polysaccharides and becomes more difficult to extract. The best time to harvest mycelium, to obtain the highest chitosan yield, will be during late exponential phase, which for Mucor sp. was 74 h and for C. echinulata, G. butleri USDB 0201, and Zygorhynchus moelleri was 96 h, as suggested by Tan et al. They also concluded that the harvest time and late exponential phase has to be identified for individual fungal strains because different fungi have different growth rates and harvesting at a fixed incubation time may not yield the maximum amount of chitosan [48,49,58].

14.5 Future prospects Chitin and chitosan sourced from crustacean waste is not of a consistent quality owing to inherent variabilities in the raw materials used.

Handbook of Chitin and Chitosan

14.6 Conclusion

447

The review here provides a brief outline of the research activities performed on chitosan production from fungus so far. These literatures support and suggest that fungi can be used as an alternate source of chitosan but have not led to the commercial use of fungi to obtain chitosan. It was found that currently mushroom (Agaricus bisporus) and to an extent the fungus Aspergilus niger, obtained after citric acid production, are used as alternate sources of chitosan and marketed as vegan chitosan (https://chitosanlab.com/vegan/). Hence there is a huge potential for the use of fungi as a commercial source of chitosan. Being a versatile compound with applications in diverse fields, such as agriculture, food and beverage, pharmaceuticals, cosmetics, and wastewater treatment, the demand for chitosan is bound to increase in the near future. In addition, the identification of novel applications, such as self-healing paint, proton-conducting transistors, coatings for keeping fruits fresh for longer periods, and biodegradable plastic, will further put stress on the existing methods of production of chitosan, making it costly as well as leading to overexploitation of shrimp and crab stocks and the generation of a large amount of toxic waste. Hence avenues for alternate sources of chitosan, such as fungal culture, will have to be utilized to meet the demand. To this effect, the understandings provided by laboratory-scale studies need to be translated into the commercial production of chitosan. Moreover, to be economically feasible the medium used for culturing will have to be as inexpensive. Residues containing a high quantity of organic compounds, such as sewage, or by-products from industries that are generally discarded or used as animal feed, are suitable alternate feedstocks. The use of these materials as culture media will keep the cost of production low as well as prevent environmental pollution caused by the introduction of these organic residues into the environment. This necessitates further investigation to identify the suitable media for the production, as well as ideal fungal strains and fermentation parameters to obtain maximum chitosan yield.

14.6 Conclusion The chemical and biological properties of chitin and chitosan make them valuable compounds of diverse applications for both plants and animals. The increase in diverse application of these polymers has the potential to lead to an ever-increasing demand for the polymers but the limited supply and complex extraction process will lead to an increased price for the polymers. Also, the limited and dwindling supply of the current raw material, shellfish, can further increase the price of the polymers. This necessitates the need for alternate sources for chitin and

Handbook of Chitin and Chitosan

448

14. Fungal chitosan: prospects and challenges

chitosan. The switch to fungal chitin and chitosan has a pivotal role in meeting the demand for these polymers. Moreover, the switch to fungal sources that have reduced environmental impact, due to reduced use of toxic chemicals being required or associated waste generated during the extraction from crustaceans, can significantly improve the green credentials of the production process. The use of modern techniques, such as microwave heating, enzymatic extraction, and deacetylation, has the potential to further the environmental friendliness of chitin and chitosan biopolymer. To achieve this objective, the exploitation of advances in fermentation technology is required. Moreover, the identification and optimization of cheap culture medium, suitable strains, and fermentation conditions is essential for its feasibility. The use of fungus for production has the advantages of ease of handling, harvesting, and production of highquality chitosan with low variance. It will enable the production of polymers with diverse physicochemical properties, like molecular weight and degree of deacetylation, by using different fungal strains and fermentation parameters, like nutritional composition of the media and physical parameters. The research works so far have proven the feasibility of the production of chitin and chitosan biopolymer from fungal sources at the laboratory scale but have not led to commercial production. The stumbling blocks are the low yield and the cost associated with the culture media. To this effect, the use of molecular techniques for strain improvement and the use of industrial residues as culture media can improve the productivity and reduce the cost associated with production at commercial scale.

14.7 Acknowledgments The authors are sincerely thankful to the Natural Sciences and Engineering Research Council of Canada (Discovery Grant 355254 and Strategic Grants), and Ministe`re des Relations Internationales du Que´bec (project 07.302) (Coope´ration Que´bec-Catalanya 2012
14) for financial support. The views or opinions expressed in this chapter are those of the authors.

References [1] M.N.V. Ravi Kumar, A review of chitin and chitosan applications, React. Funct. Polym. 46 (2000) 1
27. Available from: https://doi.org/10.1016/S1381-5148(00)00038-9. [2] J.P. Martı´nez, M.P. Falomir, D. Gozalbo, Chitin: A Structural Biopolysaccharide With Multiple Applications, ELS, John Wiley & Sons, Ltd, Chichester, 2014. Available from: http://doi.org/10.1002/9780470015902.a0000694.pub3. [3] M. Kannan, M. Nesakumari, K. Rajarathinam, Production and characterization of mushroom chitosan under solid-state fermentation conditions, Adv. Biol. Res. 4 (2010) 10
13.

Handbook of Chitin and Chitosan

References

449

[4] S.A. White, P.R. Farina, I. Fulton, Production and isolation of chitosan from Mucor rouxii, Appl. Environ. Microbiol. 38 (1979) 323
328. [5] D. Knorr, J. Klein, Production and conversion of chitosan with cultures of Mucor rouxii or Phycomyces blakesleeanus, Biotechnol. Lett. 8 (1986) 691
694. Available from: https://doi.org/10.1007/BF01032563. [6] V.K. Mourya, N.N. Inamdar, Chitosan-modifications and applications: opportunities galore, React. Funct. Polym. 68 (2008) 1013
1051. Available from: https://doi.org/ 10.1016/j.reactfunctpolym.2008.03.002. [7] K.M. Rudall, W. Kenchington, The chitin system, Biol. Rev. 48 (1973) 597
633. Available from: https://doi.org/10.1111/j.1469-185X.1973.tb01570.x. [8] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (2006) 603
632. Available from: https://doi.org/10.1016/j.progpolymsci.2006.06.001. [9] S. Rajeshkumar, C. Venkatesan, M. Sarathi, V. Sarathbabu, J. Thomas, K. Anver Basha, et al., Oral delivery of DNA construct using chitosan nanoparticles to protect the shrimp from white spot syndrome virus (WSSV), Fish. Shellfish. Immunol. 26 (2009) 429
437. Available from: https://doi.org/10.1016/j.fsi.2009.01.003. [10] R.A.A. Muzzarelli, J. Boudrant, D. Meyer, N. Manno, M. DeMarchis, M.G. Paoletti, Current views on fungal chitin/chitosan, human chitinases, food preservation, glucans, pectins and inulin: a tribute to Henri Braconnot, precursor of the carbohydrate polymers science, on the chitin bicentennial, Carbohydr. Polym. 87 (2012) 995
1012. Available from: https://doi.org/10.1016/j.carbpol.2011.09.063. [11] H. Miyoshi, K. Shimura, K. Watanabe, K. Onodera, Characterization of some fungal chitosans, Biosci. Biotechnol. Biochem. 56 (1992) 1901
1905. Available from: https:// doi.org/10.1271/bbb.56.1901. [12] K.D. Rane, D.G. Hoover, An evaluation of alkali and acid treatments for chitosan extraction from fungi, Process. Biochem. 28 (1993) 115
118. Available from: https:// doi.org/10.1016/0032-9592(93)80016-A. [13] K.-J. Hu, J.-L. Hu, K.-P. Ho, K.-W. Yeung, Screening of fungi for chitosan producers, and copper adsorption capacity of fungal chitosan and chitosanaceous materials, Carbohydr. Polym. 58 (2004) 45
52. Available from: https://doi.org/10.1016/j.carbpol.2004.06.015. [14] A. Bhatnagar, M. Sillanpa¨a¨, Applications of chitin- and chitosan-derivatives for the detoxification of water and wastewater—a short review, Adv. Colloid Interface Sci. 152 (2009) 26
38. Available from: https://doi.org/10.1016/j.cis.2009.09.003. [15] H.S. Kas, Chitosan: Properties, preparations and application to microparticulate systems, J. Microencapsul. 14 (1997) 689
711. Available from: https://doi.org/10.3109/ 02652049709006820. [16] F. Shahidi, J.K.V. Arachchi, Y.-J. Jeon, Food applications of chitin and chitosans, Trends Food Sci. Technol. 10 (1999) 37
51. Available from: https://doi.org/10.1016/ S0924-2244(99)00017-5. [17] O. Cota-Arriola, M.O. Cortez-Rocha, E.C. Rosas-Burgos, A. Burgos-Herna´ndez, Y.L. Lo´pez-Franco, M. Plascencia-Jatomea, Antifungal effect of chitosan on the growth of Aspergillus parasiticus and production of aflatoxin B1, Polym. Int. 60 (2011) 937
944. Available from: https://doi.org/10.1002/pi.3054. [18] S.A. Khalaf, Production and characterization of fungal chitosan under solid-state fermentation conditions, Int. J. Agri. Biol. 6 (2004) 1033
1036. [19] J.C. Fernandes, M.J. Tiera, F.M. Winnik, DNA
chitosan nanoparticles for gene therapy: current knowledge and future trends, in: C.S.S.R. Kumar (Ed.), Nanotechnologies for the Life Sciences, Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim, 2007. Available from: http://doi.org/10.1002/9783527610419.ntls0015. [20] W.E. Rudzinski, T.M. Aminabhavi, Chitosan as a carrier for targeted delivery of small interfering RNA, Int. J. Pharm. 399 (2010) 1
11. Available from: https://doi. org/10.1016/j.ijpharm.2010.08.022.

Handbook of Chitin and Chitosan

450

14. Fungal chitosan: prospects and challenges

[21] L. Plapied, G. Vandermeulen, B. Vroman, V. Pre´at, A. Des Rieux, Bioadhesive nanoparticles of fungal chitosan for oral DNA delivery, Int. J. Pharm. 398 (2010) 210
218. Available from: https://doi.org/10.1016/j.ijpharm.2010.07.041. [22] A.A. Tayel, S. Moussa, K. Opwis, D. Knittel, E. Schollmeyer, A. Nickisch-Hartfiel, Inhibition of microbial pathogens by fungal chitosan, Int. J. Biol. Macromol. 47 (2010) 10
14. Available from: https://doi.org/10.1016/j.ijbiomac.2010.04.005. [23] G.S. Dhillon, S. Kaur, S.K. Brar, M. Verma, Green synthesis approach: extraction of chitosan from fungus mycelia, Crit. Rev. Biotechnol. 33 (2013) 379
403. Available from: https://doi.org/10.3109/07388551.2012.717217. [24] R.V. da, S. Amorim, W. de Souza, K. Fukushima, G.M. de Campos-Takaki, Faster chitosan production by mucoralean strains in submerged culture, Braz. J. Microbiol. 32 (2001) 20
23. Available from: https://doi.org/10.1590/S1517-83822001000100005. [25] S. Chatterjee, M. Adhya, A.K. Guha, B.P. Chatterjee, Chitosan from Mucor rouxii: production and physico-chemical characterization, Process. Biochem. 40 (2005) 395
400. Available from: https://doi.org/10.1016/j.procbio.2004.01.025. [26] S. Kumari, P.K. Rath, Extraction and characterization of chitin and chitosan from (Labeo rohit) fish scales, Procedia Mater. Sci. 6 (2014) 482
489. Available from: https://doi.org/10.1016/j.mspro.2014.07.062. [27] J. Synowiecki, N.A. Al-Khateeb, Production, properties, and some new applications of chitin and its derivatives, Crit. Rev. Food Sci. Nutr. 43 (2003) 145
171. Available from: https://doi.org/10.1080/10408690390826473. [28] S. Kaur, G.S. Dhillon, Recent trends in biological extraction of chitin from marine shell wastes: a review, Crit. Rev. Biotechnol. 35 (2015) 44
61. Available from: https://doi.org/10.3109/07388551.2013.798256. [29] F.A.A. Sagheer, M.A. Al-Sughayer, S. Muslim, M.Z. Elsabee, Extraction and characterization of chitin and chitosan from marine sources in Arabian Gulf, Carbohydr. Polym. 77 (2009) 410
419. Available from: https://doi.org/10.1016/j.carbpol.2009. 01.032. [30] A. Alishahi, A. Mirvaghefi, M.R. Tehrani, H. Farahmand, S.A. Shojaosadati, F.A. Dorkoosh, et al., Enhancement and characterization of chitosan extraction from the wastes of shrimp packaging plants, J. Polym. Environ. 19 (2011) 776
783. Available from: https://doi.org/10.1007/s10924-011-0321-5. [31] M. Mahdy Samar, M.H. El-Kalyoubi, M.M. Khalaf, M.M. Abd El-Razik, Physicochemical, functional, antioxidant and antibacterial properties of chitosan extracted from shrimp wastes by microwave technique, Ann. Agric. Sci. 58 (2013) 33
41. Available from: https://doi.org/10.1016/j.aoas.2013.01.006. [32] H. El Knidri, R. El Khalfaouy, A. Laajeb, A. Addaou, A. Lahsini, Eco-friendly extraction and characterization of chitin and chitosan from the shrimp shell waste via microwave irradiation, Process. Saf. Environ. Prot. 104 (2016) 395
405. Available from: https://doi.org/10.1016/j.psep.2016.09.020. [33] S. Bartnicki-Garcia, W.J. Nickerson, Nutrition, growth, and morphogenesis of mucor rouxii, J. Bacteriol. 84 (1962) 841
858. [34] R.A.A. Muzzarelli, F. Tanfani, G. Scarpini, Chelating, film-forming, and coagulating ability of the chitosan-glucan complex from Aspergillus niger industrial wastes, Biotechnol. Bioeng. 22 (1980) 885
896. Available from: https://doi.org/10.1002/ bit.260220412. [35] L.L. Davis, S. Bartnicki-Garcia, The co-ordination of chitosan and chitin synthesis in Mucor rouxii, Microbiology 130 (1984) 2095
2102. Available from: https://doi.org/ 10.1099/00221287-130-8-2095. [36] W. Suntornsuk, P. Pochanavanich, L. Suntornsuk, Fungal chitosan production on food processing by-products, Process. Biochem. 37 (2002) 727
729. Available from: https://doi.org/10.1016/S0032-9592(01)00265-5.

Handbook of Chitin and Chitosan

References

451

[37] H. Yokoi, T. Aratake, S. Nishio, J. Hirose, S. Hayashi, Y. Takasaki, Chitosan production from shochu distillery wastewater by funguses, J. Ferment. Bioeng. 85 (1998) 246
249. Available from: https://doi.org/10.1016/S0922-338X(97)86777-3. [38] S. Chatterjee, S. Chatterjee, B.P. Chatterjee, A.K. Guha, Enhancement of growth and chitosan production by Rhizopus oryzae in whey medium by plant growth hormones, Int. J. Biol. Macromol. 42 (2008) 120
126. Available from: https://doi.org/ 10.1016/j.ijbiomac.2007.10.006. [39] F. Streit, F. Koch, M.C.M. Laranjeira, J.L. Ninow, Production of fungal chitosan in liquid cultivation using apple pomace as substrate, Braz. J. Microbiol. 40 (2009) 20
25. Available from: https://doi.org/10.1590/S1517-83822009000100003. [40] I. Tsigos, N. Zydowicz, A. Martinou, A. Domard, V. Bouriotis, Mode of action of chitin deacetylase from Mucor rouxii on N-acetylchitooligosaccharides: mode of action of chitin deacetylase, Eur. J. Biochem. 261 (2001) 698
705. Available from: https://doi. org/10.1046/j.1432-1327.1999.00311.x. [41] S.M. Bowman, S.J. Free, The structure and synthesis of the fungal cell wall, BioEssays 28 (2006) 799
808. Available from: https://doi.org/10.1002/bies.20441. [42] H. Zhang, S. Yang, J. Fang, Y. Deng, D. Wang, Y. Zhao, Optimization of the fermentation conditions of Rhizopus japonicus M193 for the production of chitin deacetylase and chitosan, Carbohydr. Polym. 101 (2014) 57
67. Available from: https://doi.org/ 10.1016/j.carbpol.2013.09.015. [43] H. Me´lida, D. Sain, J.E. Stajich, V. Bulone, Deciphering the uniqueness of Mucoromycotina cell walls by combining biochemical and phylogenomic approaches: the unique features of Mucoromycotina cell walls, Environ. Microbiol. 17 (2015) 1649
1662. Available from: https://doi.org/10.1111/1462-2920.12601. [44] A. Martinou, D. Kafetzopoulos, V. Bouriotis, Chitin deacetylation by enzymatic means: monitoring of deacetylation processes, Carbohydr. Res. 273 (1995) 235
242. Available from: https://doi.org/10.1016/0008-6215(95)00111-6. [45] Y. Zhao, R.-D. Park, R.A.A. Muzzarelli, Chitin deacetylases: properties and applications, Mar. Drugs 8 (2010) 24
46. Available from: https://doi.org/10.3390/md8010024. [46] J. Cai, J. Yang, Y. Du, L. Fan, Y. Qiu, J. Li, et al., Purification and characterization of chitin deacetylase from Scopulariopsis brevicaulis, Carbohydr. Polym. 65 (2006) 211
217. Available from: https://doi.org/10.1016/j.carbpol.2006.01.003. [47] W. Wang, Y. Du, Y. Qiu, X. Wang, Y. Hu, J. Yang, et al., A new green technology for direct production of low molecular weight chitosan, Carbohydr. Polym. 74 (2008) 127
132. Available from: https://doi.org/10.1016/j.carbpol.2008.01.025. [48] S. Arcidiacono, D.L. Kaplan, Molecular weight distribution of chitosan isolated from Mucor rouxii under different culture and processing conditions, Biotechnol. Bioeng. 39 (1992) 281
286. Available from: https://doi.org/10.1002/bit.260390305. [49] S.C. Tan, T.K. Tan, S.M. Wong, E. Khor, The chitosan yield of zygomycetes at their optimum harvesting time, Carbohydr. Polym. 30 (1996) 239
242. Available from: https://doi.org/10.1016/S0144-8617(96)00052-5. [50] P. Pochanavanich, W. Suntornsuk, Fungal chitosan production and its characterization, Lett. Appl. Microbiol. 35 (2002) 17
21. Available from: https://doi.org/ 10.1046/j.1472-765X.2002.01118.x. [51] J. Synowiecki, N.A.A.Q. Al-Khateeb, Mycelia of Mucor rouxii as a source of chitin and chitosan, Food Chem. 60 (1997) 605
610. Available from: https://doi.org/10.1016/ S0308-8146(97)00039-3. [52] N. Nwe, W.F. Stevens, Production of fungal chitosan by solid substrate fermentation followed by enzymatic extraction, Biotechnol. Lett. 24 (2002) 131
134. [53] L. Jiang, S. Pan, J.M. Kim, Influence of nitrogen source on chitosan production carried out by Absidia coerulea CTCC AF 93105, Carbohydr. Polym. 86 (2011) 359
361. Available from: https://doi.org/10.1016/j.carbpol.2011.04.045.

Handbook of Chitin and Chitosan

452

14. Fungal chitosan: prospects and challenges

[54] T. Wu, S. Zivanovic, F.A. Draughon, W.S. Conway, C.E. Sams, Physicochemical properties and bioactivity of fungal chitin and chitosan, J. Agric. Food Chem. 53 (2005) 3888
3894. Available from: https://doi.org/10.1021/jf048202s. [55] B.J. Meussen, L.H. de Graaff, J.P.M. Sanders, R.A. Weusthuis, Metabolic engineering of Rhizopus oryzae for the production of platform chemicals, Appl. Microbiol. Biotechnol. 94 (2012) 875
886. Available from: https://doi.org/10.1007/s00253-0124033-0. [56] I. Cantabrana, R. Perise, I. Herna´ndez, Uses of Rhizopus oryzae in the kitchen, Int. J. Gastron. Food Sci. 2 (2015) 103
111. Available from: https://doi.org/10.1016/j. ijgfs.2015.01.001. [57] Y.D. Hang, Chitosan production from Rhizopus oryzae mycelia, Biotechnol. Lett. 12 (1990) 911
912. Available from: https://doi.org/10.1007/BF01022589. [58] N. Nwe, W.F. Stevens, Effect of urea on fungal chitosan production in solid substrate fermentation, Process. Biochem. 39 (2004) 1639
1642. Available from: https://doi. org/10.1016/S0032-9592(03)00301-7. [59] N. Nwe, S. Chandrkrachang, W.F. Stevens, T. Maw, T.K. Tan, E. Khor, et al., Production of fungal chitosan by solid state and submerged fermentation, Carbohydr. Polym. 49 (2002) 235
237. Available from: https://doi.org/10.1016/ S0144-8617(01)00355-1. [60] C. Crestini, B. Kovac, G. Giovannozzi-Sermanni, Production and isolation of chitosan by submerged and solid-state fermentation from Lentinus edodes, Biotechnol. Bioeng. 50 (1996) 207
210. Available from: https://doi.org/10.1002/bit.260500202. [61] P.N. Vaingankar, A.R. Juvekar, Fermentative production of mycelial chitosan from zygomycetes: media optimization and physico-chemical characterization, Adv. Biosci. Biotechnol. 05 (2014) 940
956. Available from: https://doi.org/10.4236/ abb.2014.512108. [62] V. Maghsoodi, S. Yaghmaei, Comparison of solid substrate and submerged fermentation for chitosan production by Aspergillus niger, Sci. Iran. (2010) 153
157. [63] R. Subramaniyam, R. Vimala, Solid state and submerged fermentation for the production of bioactive substances: a comparative study, Int. J. Sci. Nat. 3 (2012) 480
486. [64] N. Davoust, G. Hansson, Identifying the conditions for development of beneficial mycelium morphology for chitosan-producing Absidia spp. in submersed cultures, Appl. Microbiol. Biotechnol. 36 (1992). Available from: https://doi.org/10.1007/ BF00183238. [65] T. Kleekayai, W. Suntornsuk, Production and characterization of chitosan obtained from Rhizopus oryzae grown on potato chip processing waste, World J. Microbiol. Biotechnol. 27 (2011) 1145
1154. Available from: https://doi.org/10.1007/s11274010-0561-x.

Online resource https://chitosanlab.com/vegan/ (accessed 09.06.18).

Handbook of Chitin and Chitosan

C H A P T E R

15 Preparation, properties, and application of low-molecularweight chitosan Nguyen Cong Minh1, Nguyen Van Hoa2 and Trang Si Trung2 1

Institute of Biotechnology and Environment, Nha Trang University, Nha Trang, Vietnam, 2Faculty of Food Technology, Nha Trang University, Nha Trang, Vietnam O U T L I N E 15.1 Introduction

454

15.2 Preparation of low-molecular-weight chitosan 15.2.1 Chemical methods 15.2.2 Physical methods 15.2.3 Biological methods

454 455 457 458

15.3 Properties of low-molecular-weight chitosan 15.3.1 Physicochemical properties 15.3.2 Biological activities

460 460 460

15.4 Applications of low-molecular-weight chitosan

461

15.5 Agriculture 15.5.1 Aquaculture 15.5.2 Food technology

463 463 463

15.6 Conclusions

464

References

464

Handbook of Chitin and Chitosan DOI: https://doi.org/10.1016/B978-0-12-817970-3.00015-8

453

© 2020 Elsevier Inc. All rights reserved.

454

15. Preparation, properties, and application of low-molecular-weight chitosan

15.1 Introduction Chitosan is a polysaccharide copolymer of β-(1-4)-D-glucosamine and acetyl-β-(1-4)-D-glucosamine and can be prepared by alkaline deacetylation of chitin. Chitosan has been widely applied in various fields such as wastewater treatment, agriculture, aquaculture, pulp and paper industry, food industry, cosmetics, tissue engineering, and wound healing [1 4]. Moreover, chitosan can be easily processed into gels, membranes, films, nanofibers, microbeads, microparticles, nanoparticles, scaffolds, and sponges [3]. However, the physical, chemical, and biological properties of chitosan mainly depend on the degree of deacetylation (DD) and molecular weight (Mw) [2]. Chitosans are commercially available from crustacean shells with a DD of about 70% 90% and a Mw of between 50 and 2000 kDa [5,6]. Based on the range of its Mw, chitosan can be classified in three different types, namely, high-molecular-weight chitosan (HMWC, .700 kDa), medium-molecular-weight chitosan (MMWC, 150 700 kDa), and low-molecular-weight chitosan (LMWC, less than 150 kDa) [7]. To date, HMWC and MMWC have been widely used for bioplastics flocculation, thin films, and biofiber applications [1,8 10]. It is reported that the use of HMWC is necessary to form big aggregates in the solid/ liquid separation processes [11,12]. However, a poor solubility at neutral pH values and high viscosity of HMWC solutions make it difficult to use and therefore limit its industrial applications. In contrast, the low viscosity LMWC solutions were found to be much easier to use in largescale applications [13]. Moreover, LMWC was reported to penetrate more quickly and easily than HMWC into bacterial membranes, suggesting its higher antibacterial and antioxidative activity. Taking this unique property, huge studies have been carried out by using LMWC as a biological stimulant, antibacterial material, antifungal agent, and as an antioxidant [14 19]. The aim of this chapter is to provide the reasons why we need to prepare and use LMWC. The different degradation technologies for fabrication of LMWC will be also discussed, including physical, chemical, and biological methods. In particular, the solid-state degradation of chitosan is presented as well. Moreover, the properties and potential applications of LMWC for the food industry, agriculture, cosmetics, pharmacy, etc. will be discussed.

15.2 Preparation of low-molecular-weight chitosan LMWC can be prepared either by depolymerizing natural HMWC or synthesizing it from its building unit of D-glucosamine. However, the

Handbook of Chitin and Chitosan

15.2 Preparation of low-molecular-weight chitosan

FIGURE 15.1

455

Three general strategies for the preparation of LMWC.

latter process is costly and thus rarely used. The natural chitosan is a linear polysaccharide consisting of randomly β-(1-4)-linked D-glucosamine (GlcNAc; A-unit) and N-acetyl-D-glucosamine units (GlcN; Dunit). The glycosidic bonds are rather unstable and can be cleaved by hydrolyzing agents to produce chitosan with variable low Mw [20,21]. Along with the DD, the Mw of chitosan decides the chemical, physical, and biological properties of chitosan. Practically, LMWC showed higher antibacterial, antioxidative, and soluble properties compared with HMWC. Therefore the production of LMWC from HMWC has been widely studied for its potential applications as antibacterial materials, antioxidants, and drug delivery agents. Generally, the methods for preparing LMWC can be classified into physical, chemical, and enzymatic strategies, as described in Fig. 15.1. In addition, Table 15.1 shows a rough comparison of the main treatment methods of the preparation of LMWC. Each strategy has its own advantages and disadvantages according to the used reagents and treatment conditions. For example, chemical depolymerization seems to more preferable in the industrial production of LMWC but it may cause pollution issues. By contrast, physical degradation such as X-rays or gamma irradiation does not practically change the DD of chitosan but special equipment is required. This information should be carefully considered for further production and applications.

15.2.1 Chemical methods The chemical degradation of chitosan to produce LMWC is normally conducted using strong acids (HCl, HNO2, HF, H3PO4) [22 26] or weak

Handbook of Chitin and Chitosan

456

15. Preparation, properties, and application of low-molecular-weight chitosan

TABLE 15.1 LMWC.

Comparison of various degradation methods for the preparation of

Methods

Treatment conditions

Advantages

Disadvantages

Physical

Using gamma irradiation (ex. Co-60), microwave, electrochemical degradation, etc.

Scale-up easily

Low quantity of products.

Using strong acids (HCl, HNO2, HF, H3PO4, etc.) or oxidants (H2O2, K2S2O4, etc.)

Low cost of production

Chemical

Special equipment required Difficult control process High cost of associate steps

Enzymatic

Using enzymes (cellulases, lipases, proteases, chitosanases, etc.) at ambient temperature

High quantity of products

High cost of production

Safety and ease of control of production

acids (acetic acid, ascorbic acid), but a mixture of organic acids (hydroxyacetic acid, lactic acid, and hydrated citric acid) has been used as well [27 29]. In these acid-based processes, chitosan is first dissolved in a diluted acidic solution before reacting with agents in the hydrolysis step at room or ambient temperature. Afterward, the products are collected by being precipitated with ethanol or alkaline solutions and then washed with ethanol or water to recover the LMWC. In the acid hydrolysis, the degraded rates of the glycosidic bonds are of the order A A B A D .. D A B D D, suggesting that chitosan with a lower degree of acetylation is more stable in an acidic medium [30]. The chemical method is widely used for the preparation of LMWC at the industrial scale due to its rapid process, ease, and low cost of large production. However, the chemical structure of the prepared LMWC may be damaged under acidic treatment. Moreover, it has a low yield and a high cost of associate steps because the production process normally discharges a large amount of acidic solution that needs to be treated before being disposed of in the environment. The oxidation-based methods have been applied for the preparation of LMWC by using strong oxidative agents such H2O2 [31] or a combination of using H2O2 with microwave [32] and gamma rays [33]. Some other oxidants have been used, such as nitrous acid and its salt [34], NaClO2/H2O2/HCl, NaClO/H2O2, sodium perborate, sodium

Handbook of Chitin and Chitosan

15.2 Preparation of low-molecular-weight chitosan

457

percarbonate, and ozone [35,36]. Even through this oxidation-based degradation showed quite successfully, they still require a three-step process including dissolution, degradation, and precipitation as acid-based processes, which is complex and limits industrial scale production. In addition, these methods also produce a large amount of chemicals and wastewater, which may cause environmental problems. To overcome these problems, the solid-state method has been developed [37,38]. This method is facile, efficient, and easily scaled up in industry. It neglects the dissolution of chitosan, thus avoiding complicated precipitation and purification steps. Moreover, the product can be easily collected [15]. Belamie et al. prepared LMWC in the solid state by using gaseous HCl as a reagent [37]. It allows the production of LMWC directly with relatively low polydispersity after washing of the hydrolyzed products. However, the final Mw of the product is uncontrolled. Recently, Minh et al. [38] soaked chitosan in 0.2 wt.% NaOH solution for 8 h to allow it to swell completely before the degradation process under gaseous HCl at room temperature. By this treatment, the Mw of chitosan can be degraded partially but it can be done in a controlled way. Fig. 15.2 shows photos of the increased volume of chitosan soaked in a water and alkaline solution, which apparently facilitated the H2O2 degradation of the chitosan. Fig. 15.2B and C showing microscopy measurements confirm the increase of the chitosan particle size before and after soaking. This process provides LMWC with a high purity and can be scaled up for the industrial production in solid state.

15.2.2 Physical methods Physical methods have been considered as environment-friendly processes due to their reactions based on the supply of certain energy to break bonds between monomers in the chitosan structure. High-energy sources can be obtained from gamma radiation, X-ray, ultraviolet radiation, microwave radiation, electrochemical process, and pulsed electric fields [39 49]. The physical degradation can be also carried out at high temperature and pressure [40]. Moreover, the degradation process can be accelerated by combining the physical agents with the control of temperature or adding chemical reagents such as H2O2, HCl, acetic acid, formic acid, and FeSO4 [43,44,48]. However, the physical preparation method has some disadvantages, which limit it being applied for industrial production, such as (1) the reaction rates are too fast; (2) the large distribution of prepared product is obtained; (3) this method normally requires special equipment. Gamma irradiation using a Co-60 source has been widely studied for degradation of chitosan in solid state or in solution without/with adding certain reagents such as acetic acid, HCl,

Handbook of Chitin and Chitosan

458

15. Preparation, properties, and application of low-molecular-weight chitosan

H2O2, and K2S2O8 [49 52]. The irradiation process could be carried out at room temperature and be more effective by adding an oxidant. However, this degradation may cause the loss of the main backbone structure and the physical properties of prepared products [52].

15.2.3 Biological methods Enzymatic depolymerization methods, based on either nonspecific enzymes (cellulase, lipases, pectinase, papain, protease, etc.) or specific enzymes (chitinase, chitosanases, and lysozyme), have received great interest since the 1980s for the production of LMWC. Enzymes are extracted from different sources such as bacteria, virus, microfuges, etc. [53]. Specific enzymes are expensive and/or unavailable for commercial application. The degradation of chitosan using these enzymes prefers to form chitooligomers monomers. By contrast, nonspecific enzymes are often cheap and commercially available and result in the formation of LMWC [54,55]. The biological methods have been attracting increasing attention for the industrial production of LMWC because they are safety and offer easy control of reactions [53,56 58]. Table 15.2 presents the preparation of LMWC using enzymatic methods. The three most effective and used enzymes for degradation of chitosan are cellulase, chitinase, and chitosanase [56]. Chitinases have a unique ability to hydrolyze A-A bonds and do not hydrolyze D-D bonds, while chitosanases can cleave A-D and D-D linkages [71]. Cellulase is found to cut randomly the A-A bonds in chitosan [72]. At the same hydrolysis

FIGURE 15.2 (A)Photos of chitosan in powder (100 mesh size) and swollen in water and 0.2 wt.% NaOH solution after 8 h; (B, C) the microscopy images of the chitosan before and after soaking in alkaline solution after 8 h, respectively (Olympus BX41 microscope, 910, magnification bar 100 μm). Source: Reproduced from N.C. Minh, H.N. Cuong, P.T. Phuong, S. Schwarz, W.F. Stevens, V.N. Hoa, et al. Swelling-assisted reduction of chitosan molecular weight in the solid state using hydrogen peroxide. Polym. Bull. 74 (8) (2017) 3077 3087. with permission from Springer.

Handbook of Chitin and Chitosan

459

15.2 Preparation of low-molecular-weight chitosan

TABLE 15.2 methods.

Some published studies on the preparation of LMWC using enzymatic

pH

Temp. ( C)

Time (h)

Yield (%)

Ref.

7

30

48

40.3

[59]

Chitinase

4.5

45

25

92.6

[60]

Chitin

Chitinase from Paenibacillus barengoltzii

5.5

55

12

89.5

[61]

Chitosan

Cellulase

5.3

45

6

79.8

[62]

Chitosan

Chitosanase from Ficus awkeotsang makino

4.5

50

NA

94.0

[63]

Chitosan

α-Amylase

5.5

55

4

96.2

[64]

Chitosan

Pectinase from Aspergillus niger

3,0

38

6

86.0

[65]

Chitin

Chitinase C and Nacetylhexosaminidase from Streptomyces coelicolor A3 (2)

5.0

55

8

90.0

[66]

Chitin and chitosan

Chitinase from Pyrococcus furiosus and Trichoderma viride

5.0

40

24

67.0

[67]

Chitosan

Chitosanase from Streptomyces sp.

6.8

40

2

40.0

[68]

Chitosan

Cellulase

5.2

55

18

90.0

[69]

Chitosan

Neutral protease

5.4

50

4

93.5

[70]

Sources

Enzymes

Chitin

Vihrio anyurllarum strain E-383a

Chitin

conditions, cellulose displayed a stronger activity than chitinase and lysozyme [73]. Moreover, LMWCs produced by using these different enzymes, even though they had a similar MW, may have significantly different Mw distribution. In addition, the properties of prepared LMWC depend on the DD of initial chitosan. The higher the DD of chitosan used, the higher the Mw and antibacterial activity of the prepared LMWC were observed to be [73]. It was noted that the presence of trace transition metal ions is a critical factor for the cleaving of β-1,4-glycosidic linkages and further applications of chitosan due to its safety and purity [74,75]. In order to improve the efficiency of the degradation of chitosan and to reduce the production cost, a mixture of enzymes has been used [53,73,76,77]. A series of LMWC products was successfully prepared by using a mixture of chitinase, lysozyme, and cellulase [73]. Similarly, by catalysis with a mixture of cellulase, lipase, and bromelain, the hydrolysis rate is higher than that with unspecific enzymes [77].

Handbook of Chitin and Chitosan

460

15. Preparation, properties, and application of low-molecular-weight chitosan

15.3 Properties of low-molecular-weight chitosan Generally, after the degradation process by any technique, the LMWC product is a complicated mixture of different DD and Mw and often has a quite large Mw distribution (PDI). Practically, the properties of LMWC are very dependent on these parameters, especially in biological activities. Moreover, by using heterogeneous chitosan mixtures, it is difficult to determine clearly which chitosan molecules are responsible for the observed properties. Therefore three parameters (DD, Mw, and PDI) should be first determined before other characterizations of LMWC.

15.3.1 Physicochemical properties The physicochemical properties of LMWC include mostly properties of the initial chitosan such as linear aminopolysaccharide with a large nitrogen content; weak base with deprotonated amino groups as nucleophiles (pKa 6.3); able to form hydrogen bonds intermolecularly; great reactive groups for cross-linking and chemical activation; formation of salts with organic and inorganic acids; chelating and complexing properties; ionic conductivity as polyelectrolytes (at acidic pH) [3]. In addition, the low Mw results in high solubility, low viscosity aqueous solutions and high permeability, which make give it potential and allow it to be easily used in the fields of food, health, and agriculture [72]. Most animal intestines, especially the human gastrointestinal tract, cannot directly cleave the β-glucosidic linkage in chitosan chains [70]. The crystalline structure of LMWC was observed to be lower than that of HMWC after the partial hydrolysis of chitosan [38,78]. The low Mw was reported to influence the physical properties of chitosan membranes, including thermal, mechanical, and permeability [79]. LMWC have lower tensile strength and melting point, and higher permeability than those of HMWC. Similarly, the reduction in Mw of chitosan resulted in a decrease of thermostability [78]. Recently, Minh et al. reported that hydrochloric chitosan salts with a narrow DP could be prepared from LMWC [80]. Moreover, the degradation of chitosan was minimized by using low Mw of chitosan reagent, and even exposure to strong acid. The original chitosan weight (Mw of 127 kDa) was slightly reduced to form LMWC (Mw of 94 kDa) after degradation in a gaseous HCl medium. In contrast, in the same medium, the Mw of HMWC (228 kDa) was largely decreased to form oligochitosan (38 kDa) [37].

15.3.2 Biological activities Like other chitosan derivatives with hydroxyl, amine, and acetylated amine groups, LMWC can interact readily with a range of cell receptors in

Handbook of Chitin and Chitosan

15.4 Applications of low-molecular-weight chitosan

461

living organisms, resulting in diverse biological properties, such as nontoxic, biocompatible, biodegradable, mucoadhesive, antimicrobial (fungi, bacteria, viruses), antioxidant, antitumor, blood anticoagulants, hypolipidemic, and hypocholesterolemic activities [3,77,79,81,82]. These bioactivities are very dependent on the DD, Mw, and PDI of LMWC. Among them, the chain length of chitosan and its distribution are considered to be principal factors influencing its biological activities. Practically, the short chains of LMWC with a narrow distribution are more easily absorbed on the surface of the substrate compared to the long chitosan chains [55]. For example, LMWC of 20 kDa prevented the progression of diabetes mellitus and showed a higher affinity for lipopolysaccharide than chitosan of 140 kDa [83]. Evidence is beginning to accumulate, suggesting that LMWC shows higher biological activities than those of chitooligomers and HMWC [79]. Unfortunately, so far most of the biological activity mechanisms are controversial. It may be due to the use of the LMWC mixture with various Mw and PDI which was obtained directly from the degradation process [82]. Therefore in order to understand in-depth knowledge on the mode of the bioactivity, a highly purify and defined size of LMWC should be used. The heterogeneous LMWC molecules could be separated by various chromatographic techniques [82]. The biological activities are also known to be influenced by DD of LMWC. Generally, the higher DD provide an enhanced antibacterial activity. For example, LMWC of high DD (92%) possessed higher antibacterial activity against Escherichia coli and Staphylococcus aureus than LMWC with a low DD (80%) [73]. The antioxidant activity of LMWC, which is active against 1,1-diphenyl-2-picrylhydrazyl, hydroxyl, superoxide, and carboncentered radicals, is dependent on its DD [84,85]. It can be explained based on the scavenging mechanism of chitosan [86]. The free radicals can react with H1 ion from the (NH31) ions, which has been formed by NH2 groups absorbing H1 ions from the solution, to form a stable molecule. The cytotoxicity of chitosan has been widely studied, especially in the relationship between Mw and cytotoxicity. However, these findings are still controversial. Some works reported that chitosan was toxic and the toxicity was dependent on their Mw and DD [87 89]. By contrast, other studies indicated that the toxicity of chitosan was negligible [90 93]. Chitosan was nontoxic when its concentration was as high as 1 mg/mL [34]. The LMWC of 20 kDa showed a significantly higher transfection efficiency and less cytotoxicity than poly-L-lysine [94].

15.4 Applications of low-molecular-weight chitosan By having many useful properties as described in detail above, LMWC has been widely studied for applications in a variety of fields

Handbook of Chitin and Chitosan

462

15. Preparation, properties, and application of low-molecular-weight chitosan

such as agriculture, aquaculture, food technology, wastewater treatment, biomedicine, and cosmetology [95,96]. Table 15.3 presents some applications of LMWC. In this section, we would like to provide more details on agriculture, aquaculture, and food technology applications. TABLE 15.3

Some applications of LMWC.

Fields

Specifications and functions

Agriculture

• Seed-coating and germination agents • Plant growth simulators • Defense elicitors • Chemical pesticides replacement • Controlled release of fertilizers and nutrients • Feed additives for animals

[1,17,50,52,86,93,97 105]

Aquaculture

• Supplements and binders in feed of fishes and shrimps • Vaccine and vitamin delivery agents • Water quality improvement • Immunomodulatory effects

[88,90,106 112]

Food technology

• Improvement of quality and shelf life of foods • Food and drinks preservation • Fish preservation • Protective, fungistatic, antibacterial coating for fruits

[6,16,113 115]

Wastewater treatment

• Flocculants for wastewater treatment • Removal of heavy metal ions • Ecological polymer • Reduce odors

[11,12,116,117]

Biomedical and pharmaceutical materials

• • • • •

Cosmetics

• Retinol encapsulation • Weathering films • Maintain skin moisture, treat acne • Improve suppleness of hair

Antidiabetic action Obesity inhibition Antitumoral agents Hemostatic and anticoagulant Healing, bacteriostatic materials

References

[71,74,78,83,87,91,94,118,119]

Handbook of Chitin and Chitosan

[120 122]

15.5 Agriculture

463

15.5 Agriculture In the field of agriculture, LMWC is applied as plant defense elicitors, plant growth promoters, seed-coating agents, soil amendments, and animal feed additives [13,97 99]. The plant protective properties of LMWC are mainly based on their abilities against fungi, viruses, bacterial diseases, and nematodes [99]. As a potent elicitor, LMWC can actively induce a set of defense reactions such as depolarization of membrane potential, ion fluxes, production of reactive oxygen species, and phytoalexin synthesis [100 102]. Due to its nontoxic, biodegradable, and antimicrobial properties, LMWC is recommended to be used against crop pests and microbials, instead of using chemical pesticides [105]. In addition, to improve the utilization of fertilizer and water resources, LMWC-coated fertilizer was developed [123]. This product showed a good controlled release and water retention capacity and was degradable in soil and environment-friendly, suggesting its potential use in agricultural and horticultural applications.

15.5.1 Aquaculture In aquaculture applications, LMWC can be used as diet supplement, controlled diseases, water quality improvement, and wastewater treatment [88,90,106 110]. LMWC-coated feed helped to control the health condition of the olive flounder via the enhancement of immunostimulation [109]. This product also effectively reduced water pollution by preventing the collapse of the feed. Dietary supplementation of LMWC enhanced the growth of common carp (Cyprinus carpio) under field conditions [111]. LMWC nanoparticles-encapsulated vitamin C could maintain the immune-inducing property and regulate the release of vitamin C up to 48 h in the gastrointestinal tract of rainbow trout [108]. Moreover, the innate immunity indices, lysozyme, and complement were considerably increased in the fish serum. LWMC was also used as a flocculant for waste and algae and as chelator of heavy metal ions [112].

15.5.2 Food technology In food technology, LMWC is used as food additives, specialty ingredients, and antimicrobial packaging materials [41]. As a health food ingredient, the effect of LMWC on the management of hypercholesterolemia was conducted in human clinical trials [115]. LMWC is efficacious and safe in lowering LDL cholesterol concentrations in treatment naive patients with low-to-moderate hypercholesterolemia

Handbook of Chitin and Chitosan

464

15. Preparation, properties, and application of low-molecular-weight chitosan

[115]. In comparison to its Mw of 21 and 130 kDa, LMWC of 46 kDa exhibited the highest activity at inhibiting pancreatic lipase activity and plasma triacylglycerol elevation and thus was considered as a safe functional food ingredient for obesity [124]. Based on its nontoxic and antibacterial properties, LMWC has been used as a packaging material to improve food safety and shelf life [16,113,114]. Normally, LMWC has been widely used in antimicrobial films to provide edible protective coating, and in dipping and spraying for the quality preservation of a variety of food products.

15.6 Conclusions As a product of the hydrolysis of chitosan, LMWC shows a higher solubility than its initial polymer. Taking into account the quality of product and environment concerns, the enzymatic degradation of chitosan is preferred for the preparation of LMWC. The biological activities of LMWC are very dependent on its molecular size. As have many unique properties, LMWC has been successfully tested for potential applications in agricultural, aquaculture, food technology, wastewater treatment, biomedicine, and so on. However, the active mechanisms of LMWC are still unclear and thus more in-depth studies should be done. Moreover, the commercial products of LMWC are comparatively low at present and should be developed in the near future, especially for dietary supplements or health food ingredients.

References [1] M. Rinaudo, Chitin and chitosan: properties and applications, Prog. Polym. Sci. 31 (7) (2006) 603 632. [2] A. Domard, A perspective on 30 years research on chitin and chitosan, Carbohydr. Polym. 84 (2) (2011) 696 703. [3] V. Zargar, M. Asghari, A. Dashti, A review on chitin and chitosan polymers: structure, chemistry, solubility, derivatives, applications, ChemBioEng Rev. 2 (3) (2015) 204 226. [4] H.K. No, S.P. Meyers, Preparation and characterization of chitin and chitosan—a review, J. Aquat. Food Product. Technol. 4 (2) (1995) 27 52. [5] Applications of chitin and chitosan, in: M.F.A. Goosen (Ed.), CRC Press, 1996. [6] F. Seyfarth, S. Schliemann, P. Elsner, U.C. Hipler, Antifungal effect of high- and lowmolecular-weight chitosan hydrochloride, carboxymethyl chitosan, chitosan oligosaccharide and N-acetyl-d-glucosamine against Candida albicans, Candida krusei and Candida glabrata, Int. J. Pharm 353 (1 2) (2008) 139 148. [7] M. Mishra (Ed.), Handbook of Encapsulation and Controlled Release, CRC Press, 2015. [8] D.W. Lee, H. Lim, H.N. Chong, W.S. Shim, Advances in chitosan material and its hybrid derivatives: a review, Open Biomater. J. 1 (1) (2009) 10 20. [9] R. Hudson, S. Glaisher, A. Bishop, J.L. Katz, From lobster shells to plastic objects: a bioplastics activity, J. Chem. Educ. 92 (11) (2015) 1882 1885.

Handbook of Chitin and Chitosan

References

465

[10] J.G. Fernandez, D.E. Ingber, Manufacturing of large-scale functional objects using biodegradable chitosan bioplastic, Macromol. Mater. Eng. 299 (8) (2014) 932 938. [11] R. Rojas-Reyna, S. Schwarz, G. Heinrich, G. Petzold, S. Schu¨tze, J. Bohrisch, Flocculation efficiency of modified water soluble chitosan versus commonly used commercial polyelectrolytes, Carbohydr. Polym. 81 (2) (2010) 317 322. [12] S. Schwarz, S. Bratskaya, W. Jaeger, B.R. Paulke, Effect of charge density, molecular weight, and hydrophobicity on polycations adsorption and flocculation of polystyrene latices and silica, J. Appl. Polym. Sci. 101 (5) (2006) 3422 3429. ¨ zogul, J.M. Regenstein, Industrial applications of crustacean by-pro[13] I. Hamed, F. O ducts (chitin, chitosan, and chitooligosaccharides): a review, Trends Food Sci. Technol. 48 (2016) 40 50. [14] P.J. Park, J.Y. Je, H.G. Byun, S.H. Moon, S.K. Kim, Antimicrobial activity of heterochitosans and their oligosaccharides with different molecular weights, J. Microbiol. Biotechnol. 14 (2) (2004) 317 323. [15] Y.C. Chung, Y.P. Su, C.C. Chen, G. Jia, H.L. Wang, J.G. Wu, et al., Relationship between antibacterial activity of chitosan and surface characteristics of cell wall, Acta Pharmacol. Sin. 25 (7) (2004) 932 936. [16] G.U. Tsai, W.H. Su, H.C. Chen, C.L. Pan, Antimicrobial activity of shrimp chitin and chitosan from different treatments, Fish. Sci. 68 (1) (2002) 170 177. [17] R. Sharp, A review of the applications of chitin and its derivatives in agriculture to modify plant-microbial interactions and improve crop yields, Agronomy. 3 (4) (2013) 757 793. [18] H.K. No, N.Y. Park, S.H. Lee, S.P. Meyers, Antibacterial activity of chitosans and chitosan oligomers with different molecular weights, Int. J. Food Microbiol. 74 (1 2) (2002) 65 72. [19] H. Liu, Y. Du, X. Wang, L. Sun, Chitosan kills bacteria through cell membrane damage, Int. J. Food Microbiol. 95 (2) (2004) 147 155. [20] S.K. Kim, N. Rajapakse, Enzymatic production and biological activities of chitosan oligosaccharides (COS): a review, Carbohydr. Polym. 62 (4) (2005) 357 368. [21] F. Liaqat, R. Eltem, Chitooligosaccharides and their biological activities: a comprehensive review, Carbohydr. Polym. 184 (2018) 243 259. [22] K. Tømmeraas, K.M. Va˚rum, B.E. Christensen, O. Smidsrød, Preparation and characterisation of oligosaccharides produced by nitrous acid depolymerisation of chitosans, Carbohydr. Res. 333 (2) (2001) 137 144. [23] S.T. Horowitz, S. Roseman, H.J. Blumenthal, The preparation of glucosamine oligosaccharides. I. Separation 1, 2, J. Am. Chem. Soc. 79 (18) (1957) 5046 5049. [24] S.A. Barker, A.B. Foster, M. Stacey, J.M. Webber, Amino-sugars and related compounds. Part IV. Isolation and properties of oligosaccharides obtained by controlled fragmentation of chitin, J. Chem. Soc. (1958) 2218 2227. [25] A. Einbu, H. Grasdalen, K.M. Va˚rum, Kinetics of hydrolysis of chitin/chitosan oligomers in concentrated hydrochloric acid, Carbohydr. Res. 342 (8) (2007) 1055 1062. [26] S. Trombotto, C. Ladavie`re, F. Delolme, A. Domard, Chemical preparation and structural characterization of a homogeneous series of chitin/chitosan oligomers, Biomacromolecules. 9 (7) (2008) 1731 1738. [27] Z. Jia, D. Shen, Effect of reaction temperature and reaction time on the preparation of low-molecular-weight chitosan using phosphoric acid, Carbohydr. Polym. 49 (4) (2002) 393 396. [28] M. Hamdine, M.C. Heuzey, A. Be´gin, Effect of organic and inorganic acids on concentrated chitosan solutions and gels, Int. J. Biol. Macromol. 37 (3) (2005) 134 142. [29] I.M. Lipatova, N.A. Kornilova, Effect of hydroacoustic treatment on the rate of hydrolytic degradation of chitosan in acetic acid solutions, Russ. J. Appl. Chem. 81 (5) (2008) 815 819.

Handbook of Chitin and Chitosan

466

15. Preparation, properties, and application of low-molecular-weight chitosan

[30] K.M. Va˚rum, M.H. Ottøy, O. Smidsrød, Acid hydrolysis of chitosans, Carbohydr. Polym. 46 (1) (2001) 89 98. [31] F. Tian, Y. Liu, K. Hu, B. Zhao, Study of the depolymerization behavior of chitosan by hydrogen peroxide, Carbohydr. Polym. 57 (1) (2004) 31 37. [32] T. Sun, D. Zhou, J. Xie, F. Mao, Preparation of chitosan oligomers and their antioxidant activity, Eur. Food Res. Technol. 225 (3 4) (2007) 451 456. [33] B. Kang, Y.D. Dai, H.Q. Zhang, D. Chen, Synergetic degradation of chitosan with gamma radiation and hydrogen peroxide, Polym. Degrad. Stab. 92 (3) (2007) 359 362. [34] S. Mao, X. Shuai, F. Unger, M. Simon, D. Bi, T. Kissel, The depolymerization of chitosan: effects on physicochemical and biological properties, Int. J. Pharm. 281 (1 2) (2004) 45 54. [35] W. Yue, P. Yao, Y. Wei, Influence of ultraviolet-irradiated oxygen on depolymerization of chitosan, Polym. Degrad. Stab. 94 (5) (2009) 851 858. [36] S. Seo, J.M. King, W. Prinyawiwatkul, Simultaneous depolymerization and decolorization of chitosan by ozone treatment, J. Food Sci. 72 (9) (2007) C522 C526. [37] E. Belamie, A. Domard, M.M. Giraud-Guille, Study of the solid-state hydrolysis of chitosan in presence of HCl, J. Polym. Sci. Part A Polym. Chem. 35 (15) (1997) 3181 3191. [38] N.C. Minh, H.N. Cuong, P.T. Phuong, S. Schwarz, W.F. Stevens, V.N. Hoa, et al., Swelling-assisted reduction of chitosan molecular weight in the solid state using hydrogen peroxide, Polym. Bull. 74 (8) (2017) 3077 3087. [39] S.K. Kim (Ed.), Chitin, Chitosan, Oligosaccharides and their Derivatives: Biological Activities and Applications, CRC Press, 2010. [40] K. Sato, T. Kishimoto, M. Morimoto, H. Saimoto, Y. Shigemasa, Hydrolysis of acetals in water under hydrothermal conditions, Tetrahedron Lett. 44 (47) (2003) 8623 8625. [41] H. Yin, Y. Du, J. Zhang, Low molecular weight and oligomeric chitosans and their bioactivities, Curr. Top. Med. Chem. 9 (16) (2009) 1546 1559. [42] A. Sahu, P. Goswami, U. Bora, Microwave mediated rapid synthesis of chitosan, J. Mater. Sci. Mater. Med. 20 (1) (2009) 171 175. [43] R. Xing, S. Liu, H. Yu, Z. Guo, P. Wang, C. Li, et al., Salt-assisted acid hydrolysis of chitosan to oligomers under microwave irradiation, Carbohydr. Res. 340 (13) (2005) 2150 2153. [44] R. Xing, S. Liu, H. Yu, Q. Zhang, Z. Li, P. Li, Preparation of low-molecular-weight and high-sulfate-content chitosans under microwave radiation and their potential antioxidant activity in vitro, Carbohydr. Res. 339 (15) (2004) 2515 2519. [45] E.S. Tang, M. Huang, L.Y. Lim, Ultrasonication of chitosan and chitosan nanoparticles, Int. J. Pharm. 265 (1 2) (2003) 103 114. [46] G. Cravotto, S. Tagliapietra, B. Robaldo, M. Trotta, Chemical modification of chitosan under high-intensity ultrasound, Ultrason. Sonochem. 12 (1 2) (2005) 95 98. [47] T. Wu, S. Zivanovic, D.G. Hayes, J. Weiss, Efficient reduction of chitosan molecular weight by high-intensity ultrasound: underlying mechanism and effect of process parameters, J. Agric. Food Chem. 56 (13) (2008) 5112 5119. [48] N.Q. Hien, D. Van Phu, N.N. Duy, N.T. Lan, Degradation of chitosan in solution by gamma irradiation in the presence of hydrogen peroxide, Carbohydr. Polym. 87 (1) (2012) 935 938. [49] J.M. Wasikiewicz, F. Yoshii, N. Nagasawa, R.A. Wach, H. Mitomo, Degradation of chitosan and sodium alginate by gamma radiation, sonochemical and ultraviolet methods, Radiat. Phys. Chem. 73 (5) (2005) 287 295. [50] C. Siri-Upathum, Radiation degradation of chitosan and its application for young orchid plant growth promotion. Advance in chitin, Science. 5 (2002) 475 478.

Handbook of Chitin and Chitosan

References

467

[51] R. Yoksan, M. Akashi, M. Miyata, S. Chirachanchai, Optimal γ-ray dose and irradiation conditions for producing low-molecular-weight chitosan that retains its chemical structure, Radiat. Res. 161 (4) (2004) 471 480. [52] L.Q. Luan, V.T. Ha, N. Nagasawa, T. Kume, F. Yoshii, T.M. Nakanishi, Biological effect of irradiated chitosan on plants in vitro, Biotechnol. Appl. Biochem. 41 (1) (2005) 49 57. [53] H. Zhang, W. Zhang, Induction and optimization of chitosanase production by Aspergillus fumigatus YT-1 using response surface methodology, Chem. Biochem. Eng. Q. 27 (3) (2013) 335 345. [54] F.S. Kittur, A.V. Kumar, L.R. Gowda, R.N. Tharanathan, Chitosanolysis by a pectinase isozyme of Aspergillus niger—a non-specific activity, Carbohydr. Polym. 53 (2) (2003) 191 196. [55] A.B. Kumar, L.R. Gowda, R.N. Tharanathan, Non-specific depolymerization of chitosan by pronase and characterization of the resultant products, Eur. J. Biochem. 271 (4) (2004) 713 723. [56] M. Naveed, L. Phil, M. Sohail, M. Hasnat, M.M.F.A. Baig, A.U. Ihsan, et al., Chitosan oligosaccharide (COS): an overview, Int. J. Biol. Macromol. 129 (2019) 827 843. [57] W. Xia, P. Liu, J. Zhang, J. Chen, Biological activities of chitosan and chitooligosaccharides, Food Hydrocoll. 25 (2) (2011) 170 179. [58] H. Zhang, S.H. Neau, In vitro degradation of chitosan by a commercial enzyme preparation: effect of molecular weight and degree of deacetylation, Biomaterials. 22 (12) (2001) 1653 1658. [59] Y. Takiguchi, K.N. Shimahara, N-Diacetylchitobiose production from chitin by Vibrio anguillarum strain E-383a, Lett. Appl. Microbiol. 6 (6) (1988) 129 131. [60] X. Fu, Q. Yan, S. Yang, X. Yang, Y. Guo, Z. Jiang, An acidic, thermostable exochitinase with β-N-acetylglucosaminidase activity from Paenibacillus barengoltzii converting chitin to N-acetyl glucosamine, Biotechnol. Biofuels 7 (1) (2014) 174 185. [61] S. Yang, X. Fu, Q. Yan, Y. Guo, Z. Liu, Z. Jiang, Cloning, expression, purification and application of a novel chitinase from a thermophilic marine bacterium Paenibacillus barengoltzii, Food Chem. 192 (2016) 1041 1048. [62] H. Dong, Y. Wang, L. Zhao, J. Zhou, Q. Xia, Y. Qiu, Key technologies of enzymatic preparation for DP 6 8 chitooligosaccharides, J. Food Process. Eng. 38 (4) (2015) 336 344. [63] C.T. Chang, Y.L. Lin, S.W. Lu, C.W. Huang, Y.T. Wang, Y.C. Chung, Characterization of a Chitosanase from jelly fig (Ficus awkeotsang Makino) latex and its application in the production of water-soluble low molecular weight Chitosans, PLoS One. 11 (3) (2016) e0150490. [64] S. Wu, Preparation of chitooligosaccharides from Clanis bilineata larvae skin and their antibacterial activity, Int. J. Biol. Macromol. 51 (5) (2012) 1147 1150. [65] F.S. Kittur, A.B. Kumar, M.C. Varadaraj, R.N. Tharanathan, Chitooligosaccharides— preparation with the aid of pectinase isozyme from Aspergillus niger and their antibacterial activity, Carbohydr. Res. 340 (6) (2005) 1239 1245. [66] T.N. Nguyen, N. Doucet, Combining chitinase C and N-acetylhexosaminidase from Streptomyces coelicolor A3 (2) provides an efficient way to synthesize N-acetylglucosamine from crystalline chitin, J. Biotechnol. 220 (2016) 25 32. [67] M. Mahata, S. Shinya, E. Masaki, T. Yamamoto, T. Ohnuma, R. Brzezinski, et al., Production of chitooligosaccharides from Rhizopus oligosporus NRRL2710 cells by chitosanase digestion, Carbohydr. Res. 383 (2014) 27 33. [68] S. Sinha, S. Chand, P. Tripathi, Microbial degradation of chitin waste for production of chitosanase and food related bioactive compounds, Appl. Biochem. Microbiol. 50 (2) (2014) 125 133.

Handbook of Chitin and Chitosan

468

15. Preparation, properties, and application of low-molecular-weight chitosan

[69] G.J. Tsai, S.L. Zhang, P.L. Shieh, Antimicrobial activity of a low-molecular-weight chitosan obtained from cellulase digestion of chitosan, J. Food Prot. 67 (2) (2004) 396 398. [70] J. Li, Y. Du, J. Yang, T. Feng, A. Li, P. Chen, Preparation and characterisation of low molecular weight chitosan and chito-oligomers by a commercial enzyme, Polym. Degrad. Stab. 87 (3) (2005) 441 448. [71] B.B. Aam, E.B. Heggset, A.L. Norberg, M. Sørlie, K.M. Va˚rum, V.G. Eijsink, Production of chitooligosaccharides and their potential applications in medicine, Mar. Drugs. 8 (5) (2010) 1482 1517. [72] W. Xia, P. Liu, J. Liu, Advance in chitosan hydrolysis by non-specific cellulases, Bioresour. Technol. 99 (15) (2008) 6751 6762. [73] S.B. Lin, Y.C. Lin, H.H. Chen, Low molecular weight chitosan prepared with the aid of cellulase, lysozyme and chitinase: characterisation and antibacterial activity, Food Chem. 116 (1) (2009) 47 53. [74] S.B. Qasim, S. Husain, Y. Huang, M. Pogorielov, V. Deineka, M. Lyndin, et al., Invitro and in-vivo degradation studies of freeze gelated porous chitosan composite scaffolds for tissue engineering applications, Polym. Degrad. Stab. 136 (2017) 31 38. [75] A.D. Nugraheni, D. Purnawati, Chotimah, A. Kusumaatmaja, K. Triyana, Study of thermal degradation of PVA/chitosan/gelatin electrospun nanofibers, in: AIP Conference Proceedings 2016. AIP Publishing, 1755 (1), 150017. [76] H. Zhang, Y. Du, X. Yu, M. Mitsutomi, S.I. Aiba, Preparation of chitooligosaccharides from chitosan by a complex enzyme, Carbohydr. Res. 320 (3 4) (1999) 257 260. ˇ unek, ˚ [77] G. Tishchenko, J. Sim J. Brus, M. Netopilı´k, M. Peka´rek, Z. Walterova´, et al., Low-molecular-weight chitosans: preparation and characterization, Carbohydr. Polym. 86 (2) (2011) 1077 1081. [78] C. Qin, B. Zhou, L. Zeng, Z. Zhang, Y. Liu, Y. Du, et al., The physicochemical properties and antitumor activity of cellulase-treated chitosan, Food Chem. 84 (1) (2004) 107 115. [79] F.S. Kittur, A.B. Kumar, R.N. Tharanathan, Low molecular weight chitosans— preparation by depolymerization with Aspergillus niger pectinase, and characterization, Carbohydr. Res. 338 (12) (2003) 1283 1290. [80] N.C. Minh, N.V. Hoa, S. Schwarz, W.F. Stevens, T.S. Trung, Preparation of water soluble hydrochloric chitosan from low molecular weight chitosan in the solid state, Int. J. Biol. Macromol 121 (2019) 718 726. [81] V.K. Thakur, M.K. Thakur, Recent advances in graft copolymerization and applications of chitosan: a review, ACS Sustain. Chem. Eng. 2 (12) (2014) 2637 2652. [82] K. Li, R. Xing, S. Liu, P. Li, Advances in preparation, analysis and biological activities of single chitooligosaccharides, Carbohydr. Polym. 139 (2016) 178 190. [83] Y. Kondo, A. Nakatani, K. Hayashi, M. Ito, Low molecular weight chitosan prevents the progression of low dose streptozotocin-induced slowly progressive diabetes mellitus in mice, Biol. Pharm. Bull. 23 (12) (2000) 1458 1464. [84] K.W. Kim, R.L. Thomas, Antioxidative activity of chitosans with varying molecular weights, Food Chem. 101 (1) (2007) 308 313. [85] J.Y. Je, P.J. Park, S.K. Kim, Free radical scavenging properties of heterochitooligosaccharides using an ESR spectroscopy, Food Chem. Toxicol. 42 (3) (2004) 381 387. [86] S.N. Kulikov, S.N. Chirkov, A.V. Il’ina, S.A. Lopatin, V.P. Varlamov, Effect of the molecular weight of chitosan on its antiviral activity in plants, Appl. Biochem. Microbiol. 42 (2) (2006) 200 203. [87] D. Sgouras, R. Duncan, Methods for the evaluation of biocompatibility of soluble synthetic polymers which have potential for biomedical use: 1—use of the tetrazolium-based

Handbook of Chitin and Chitosan

References

469

colorimetric assay (MTT) as a preliminary screen for evaluation of in vitro cytotoxicity, J. Mater. Sci: Mater. Med. 1 (2) (1990) 61 68. [88] J. Heller, L.S. Liu, R. Duncan, S. Richardson, Alginate/chitosan microporous microspheres for the controlled release of proteins and antigens, Proc. Int. Symp. Control. Rel. Bioact. Mater. 23 (1996) 269 270. [89] B. Carreno-Gomez, R. Duncan, Evaluation of the biological properties of soluble chitosan and chitosan microspheres, Int. J. Pharm 148 (2) (1997) 231 240. [90] T.J. Aspden, J. Adler, S.S. Davis, L. Illum, Chitosan as a nasal delivery system: evaluation of the effect of chitosan on mucociliary clearance rate in the frog palate model, Int. J. Pharm 122 (1 2) (1995) 69 78. [91] T.J. Aspden, L. Illum, Ø. Skaugrud, Chitosan as a nasal delivery system: evaluation of insulin absorption enhancement and effect on nasal membrane integrity using rat models, Eur. J. Pharm. Sci. 4 (1) (1996) 23 31. [92] T.J. Aspden, J.D. Mason, N.S. Jones, J. Lowe, Ø. Skaugrud, L. Illum, Chitosan as a nasal delivery system: the effect of chitosan solutions on in vitro and in vivo mucociliary transport rates in human turbinates and volunteers, J. Pharm. Sci. 86 (4) (1997) 509 513. [93] T. Aspden, L. Illum, Ø. Skaugrud, The effect of chronic nasal application of chitosan solutions on cilia beat frequency in guinea pigs, Int. J. Pharm. 153 (2) (1997) 137 146. [94] M. Lee, J.W. Nah, Y. Kwon, J.J. Koh, K.S. Ko, S.W. Kim, Water-soluble and low molecular weight chitosan-based plasmid DNA delivery, Pharm. Res. 18 (4) (2001) 427 431. [95] A. Muxika, A. Etxabide, J. Uranga, P. Guerrero, K. De La Caba, Chitosan as a bioactive polymer: processing, properties and applications, Int. J. Biol. Macromol. 105 (2017) 1358 1368. [96] L. Phil, M. Naveed, I.S. Mohammad, L. Bo, D. Bin, Chitooligosaccharide: an evaluation of physicochemical and biological properties with the proposition for determination of thermal degradation products, Biomed. Pharmacother. 102 (2018) 438 451. [97] L.A. Hadwiger, Multiple effects of chitosan on plant systems: solid science or hype, Plant Sci. 208 (2013) 42 49. [98] N.A. Dzung, V.T. Khanh, T.T. Dzung, Research on impact of chitosan oligomers on biophysical characteristics, growth, development and drought resistance of coffee, Carbohydr. Polym. 84 (2) (2011) 751 755. [99] M.A. Ramı´rez, A.T. Rodrı´guez, L. Alfonso, C. Peniche, Chitin and its derivatives as biopolymers with potential agricultural applications, Biotechnol. Apl. 27 (4) (2010) 270 276. [100] K. Kuchitsu, H. Kosaka, T. Shiga, N. Shibuya, EPR evidence for generation of hydroxyl radical triggered by N-acetylchitooligosaccharide elicitor and a protein phosphatase inhibitor in suspension-cultured rice cells, Protoplasma. 188 (1 2) (1995) 138 142. [101] K. Kuchitsu, Y. Yazaki, K. Sakano, N. Shibuya, Transient cytoplasmic pH change and ion fluxes through the plasma membrane in suspension-cultured rice cells triggered by N-acetylchitooligosaccharide elicitor, Plant. Cell Physiol. 38 (9) (1997) 1012 1018. [102] R. Takai, K. Hasegawa, H. Kaku, N. Shibuya, E. Minami, Isolation and analysis of expression mechanisms of a rice gene, EL5, which shows structural similarity to ATL family from Arabidopsis, in response to N-acetylchitooligosaccharide elicitor, Plant. Sci. 160 (4) (2001) 577 583. [103] X.J. Li, X.S. Piao, S.W. Kim, P. Liu, L. Wang, Y.B. Shen, et al., Effects of chitooligosaccharide supplementation on performance, nutrient digestibility, and serum composition in broiler chickens, Poult. Sci. 86 (6) (2007) 1107 1114.

Handbook of Chitin and Chitosan

470

15. Preparation, properties, and application of low-molecular-weight chitosan

[104] P.L. Kashyap, X. Xiang, P. Heiden, Chitosan nanoparticle based delivery systems for sustainable agriculture, Int. J. Biol. Macromol. 77 (2015) 36 51. [105] M.E. Badawy, E.I. Rabea, A biopolymer chitosan and its derivatives as promising antimicrobial agents against plant pathogens and their applications in crop protection, Int. J. Carbohydr. Chem. (2011) 1 29. [106] R.L. Huang, Z.Y. Deng, C.B. Yang, Y.L. Yin, M.Y. Xie, G.Y. Wu, et al., Dietary oligochitosan supplementation enhances immune status of broilers, J. Sci. Food Agri. 87 (1) (2007) 153 159. [107] G.J. Wu, G.J. Tsai, Cellulase degradation of shrimp chitosan for the preparation of a water-soluble hydrolysate with immunoactivity, Fish. Sci. 70 (6) (2004) 1113 1120. [108] A. Alishahi, A. Mirvaghefi, M.R. Tehrani, H. Farahmand, S. Koshio, F.A. Dorkoosh, et al., Chitosan nanoparticle to carry vitamin C through the gastrointestinal tract and induce the non-specific immunity system of rainbow trout (Oncorhynchus mykiss), Carbohydr. Polym. 86 (1) (2011) 142 146. [109] S.H. Cha, J.S. Lee, C.B. Song, K.J. Lee, Y.J. Jeon, Effects of chitosan-coated diet on improving water quality and innate immunity in the olive flounder, Paralichthys olivaceus, Aquaculture 278 (1 4) (2008) 110 118. [110] S. Lin, S. Mao, Y. Guan, L. Luo, L. Luo, Y. Pan, Effects of dietary chitosan oligosaccharides and Bacillus coagulans on the growth, innate immunity and resistance of koi (Cyprinus carpio koi), Aquaculture 342 (2012) 36 41. [111] A. Gopalakannan, V. Arul, Immunomodulatory effects of dietary intake of chitin, chitosan and levamisole on the immune system of Cyprinus carpio and control of Aeromonas hydrophila infection in ponds, Aquaculture 255 (1 4) (2006) 179 187. [112] Y.C. Chung, Y.H. Li, C.C. Chen, Pollutant removal from aquaculture wastewater using the biopolymer chitosan at different molecular weights, J. Environ. Sci. Health 40 (9) (2005) 1775 1790. [113] C.R. Huei, H.D. Hwa, Effect of molecular weight of chitosan with the same degree of deacetylation on the thermal, mechanical, and permeability properties of the prepared membrane, Carbohydr. Polym. 29 (4) (1996) 353 358. [114] S.B. Schreiber, J.J. Bozell, D.G. Hayes, S. Zivanovic, Introduction of primary antioxidant activity to chitosan for application as a multifunctional food packaging material, Food Hydrocoll. 33 (2) (2013) 207 214. [115] S. Jaffer, J.S. Sampalis, Efficacy and safety of chitosan HEP-40 [TM] in the management of hypercholesterolemia: a randomized, multicenter, placebo-controlled trial, Altern. Med. Rev. 12 (3) (2007) 265 274. [116] Z.F. Jiang, J.M. Geng, H.Y. Yang, Low molecular weight chitosans as chelators of Cu (II) in vitro and in vivo decreased neurotoxicity of amyloid beta-peptide bound with Cu (II), Ann. Neurol. 62 (2007) S26. [117] J.P. Wang, Y.Z. Chen, S.J. Yuan, G.P. Sheng, H.Q. Yu, Synthesis and characterization of a novel cationic chitosan-based flocculant with a high water-solubility for pulp mill wastewater treatment, Water Res. 43 (20) (2009) 5267 5275. [118] K. Hayashi, M. Ito, Antidiabetic action of low molecular weight chitosan in genetically obese diabetic KK-Ay mice, Biol. Pharm. Bull. 25 (2) (2002) 188 192. [119] H.T. Yao, S.Y. Huang, M.T. Chiang, A comparative study on hypoglycemic and hypocholesterolemic effects of high and low molecular weight chitosan in streptozotocin-induced diabetic rats, Food Chem. Toxicol. 46 (5) (2008) 1525 1534. [120] D.G. Kim, Y.I. Jeong, C. Choi, S.H. Roh, S.K. Kang, M.K. Jang, et al., Retinolencapsulated low molecular water-soluble chitosan nanoparticles, Int. J. Pharm. 319 (1 2) (2006) 130 138. [121] A. Sionkowska, A. Planecka, J. Kozlowska, J. Skopinska-Wisniewska, P. Los, Weathering of chitosan films in the presence of low-and high-molecular weight additives, Carbohydr. Polym. 84 (3) (2011) 900 906.

Handbook of Chitin and Chitosan

References

471

[122] S. Hirano, K. Hirochi, K.I. Hayashi, T. Mikami, H. Tachibana, Cosmetic and pharmaceutical uses of chitin and chitosan, Cosmetic and Pharmaceutical Applications of Polymers, Springer, Boston, MA, 1991, pp. 95 104. [123] L. Wu, M. Liu, Preparation and properties of chitosan-coated NPK compound fertilizer with controlled-release and water-retention, Carbohydr. Polym. 72 (2) (2008) 240 247. [124] M. Sumiyoshi, Y. Kimura, Low molecular weight chitosan inhibits obesity induced by feeding a high-fat diet long-term in mice, J. Pharm. Pharmacol 58 (2) (2006) 201 207.

Handbook of Chitin and Chitosan

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

A Abiotic reactions, 248
249 Abscisic acid, 236 Absidia butleri, 446 Absidia coerulea, 229
230, 430, 442
443 Absidia glauca, 433 ACC method. See Aqueous counter collision (ACC) method N-Acetamido bonds, 384 in chitin, 138
139 groups, 290, 430 2-Acetamido-2-deoxy-β-D-glucose, 3
4, 168, 290 Acetic acid solutions, 12, 48, 201, 253
254 Acetone/organic solvent mixtures, 39 Acetylation, 61
62, 171, 250 of chitin, 104, 186 of chitosan, 134
145 N-Acetyl-d-glucosamine (GlcNAc), 40, 139
142, 227
228, 336, 348, 378, 403, 454
455 N-Acetyl glucosamine, 132, 138
141, 144f, 227
228, 290, 402, 405 Acetyl groups, molecular weight and distribution, 379
380 Acetyl xylan esterase, 139 Acid-based processes, 455
457 Acid catalyst method, 120
121 Acid-catalyzed hydrolysis, 383, 383f Acid chloride, 122
124 Acid hydrolysis, 141, 455
456 Acid titration DA, 141 Acryloyl chloride, in dichloromethane, 69 Activation of chitin, 121 N-Acylation method, 40, 381
382 Adenosine monophosphate-activated protein kinase (AMPK) pathway, 112
113 Adsorption process, 24
25, 357
358 Aerogels, chitin and chitosan, 286
287, 298f Au(III)
chitosan
silica, 291
292

characterization of, 296
324 morphological/microscopic analysis, 296
304 porosity, 304
309 scanning electron microscope, 296
302 thermal properties, 309
312 transmission electron microscopy, 302
304 classification, 288
290, 289f CS-PDA, 309 definition of, 287 Fourier transform infrared spectroscopy, 317
319, 318f, 319f future aspects of, 324
327 morphology and physical properties, 296f nuclear magnetic resonance spectroscopy, 322
323, 322f preparation process, 292
295, 293f production, drying technologies for, 294t Raman spectroscopy, 312
313 sources and properties, 290
292 structure formation for, 295f tensile property of mechanical property, 314
317, 314f thermal properties, 309
312 differential scanning calorimetry, 310
312 thermogravimetric analysis, 309
310 water contact angle, 323
324 X-ray diffraction, 319, 320f X-ray photoelectron spectroscopy, 319
322 Agaricus bisporus, 229, 446
447 Aggregation, 85
87 AgNP. See Silver nanoparticles (AgNP) Agr-CS-G. See Genipin crosslinked blend (Agr-CS-G) Agriculture, 256 industrial applications, of chitosan, 358
359 sector, 426

473

474

Index

AIM. See Alkali-insoluble material (AIM) α-(1-4)-linked-2-amino-2-deoxy-β-Dglucopyranose, 61, 290
291 N-aliphatic-O-dicinnamoyl-chitosans, 149
151 Alkali chitin, 135
136 Alkali deacetylation process, 344 Alkali-insoluble material (AIM), 432
433 Alkaline hydrolysis methods, 344
345, 411 Alkaline N-deacetylation, 132 Alkyl-acylated chitin, 124 N-alkylated disaccharides, 387
388 Alkylating agents, 67 Allomorphs, 62
63 All-trans retinoic acid (ATRA), 87
88 1-Allyl-3-methylimidazolium acetate (AMIMOAc), 380
381 1-Allyl-3-methylimidazolium bromide (AMIMBr), 104
105, 107, 117, 124, 184, 380
381, 382f, 384
385 α-chitin, 4
5, 14
16, 37, 62
63, 102, 108, 168, 226
227, 336
337, 349, 349f, 403, 403f Amide I in, 109 FTIR spectrum of, 109f powder and nanofiber, 103 solid 13C NMR, 110, 111f TGA curves of, 20f XRD spectrum of, 110f Ambient parameters, electrospinning process, 44
45 Ambient pressure drying, 293
295 Amide groups, 268
269 Amide I band, 14 AMIMBr. See 1-Allyl-3-methylimidazolium bromide (AMIMBr) AMIMOAc. See 1-Allyl-3methylimidazolium acetate (AMIMOAc) Amino and hydroxyl groups, 426 2-Amino-2-deoxy-β-D-glucopyranose, 337 2-Amino-2-deoxy-(1-4)-β-D-glucopyranose (ß-(1,4)-2-amino-2- deoxy-D-glucopyranose), 249 2-Amino-2-dexoy-D-glucopyranose, 18
19 Amino groups, 67
68 in chitosan, 22, 351 substitution, 66
67 Ammonium peroxy disulphate (APS), 117 Ammonium salts of chitosan, 152
153 Amphiphilic sodium salt, 322 AMPK pathway. See Adenosine monophosphate-activated protein kinase (AMPK) pathway

Amylopectin, 258 Amylose, 258 Anhydrous acetic acid, 186 Anionic functional groups, 187 Antiaging cosmetics, 231
232 Anticancer/antitumor property, 386
387 Antiinflammatory property, 388 Antimicrobial agents, 346 Antimicrobial chitosan
polyethylene oxide mats, 214
215 Antimicrobial property, 387
388 Antioxidant property, 386 Antiparallel hydrogen bonding, formation of, 227
228 Antipsoriatic drugs, 177 APS. See Ammonium peroxy disulphate (APS) Aquaculture, LMWC in, 463 Aqueous acid solutions, 40 Aqueous counter collision (ACC) method, 180 Aqueous organic acid PEG-aldehyde in, 92
93 solutions, 253
254 Aqueous solution chitin, 102 2-chloroethylamine hydrochloride, 117
118 KOH, 184 NaOH/urea, 102
103 sodium hydroxide, 121 Arthropods, 226, 228 exoskeleton of, 421 Artificial skin graft, 355 Aspergillus flavus, 234, 390, 391f Aspergillus nidulans, 430 Aspergillus niger, 229 Aspergillus terreus, 405
407 Aspergilus niger, 446
447 Atomic force microscopy, 91 Atom transfer radical polymerization (ATRP), 117, 119 ATRA. See All-trans retinoic acid (ATRA) ATRP. See Atom transfer radical polymerization (ATRP) AuNPs, 304

B Bacillus coagulans, 236 Bacillus licheniformis, 235 Bacillus subtilis, 9, 115, 139, 407 Barrett
Joyner
Halenda (BJH) approach, 304
305

Index

Base-catalyzed hydrolysis, 383
384 Bead or granule type, 47 Benzoylated chitin, 123 β-chitin, 4
5, 14
16, 37, 62
63, 102, 108, 114, 168, 226
227, 250, 336
337, 349, 349f, 403, 403f crystalline structure of, 103 TGA curves of, 20f β-(1-4)-linked 2-acetamido-2-deoxy-β-Dglucose, 61 β-(1,4)-linked 2-acetamino-2-deoxy-β-Dglucopyranose, 337 β-(1
4)-linked 2-amino-2-deoxy-β-Dglucopyranose, 132 β-(1
4)-linked D-glucosamine, 403 β-(1
4)-N-acetyl-D-glucosamine (GlcNAc), 3
4, 168, 336, 421
426 (1,4)-β-N-acetyl glycosaminoglycan, 166 β-ChNF [acid] aerogels, 309, 317 β-CD-CS@HMTA-Cr, 300
301 β-(1-4)-D-glucosamine and acetyl-β-(1-4)-Dglucosamine, 454 β-1,4-glycosidic bonding, 107
108 BET theory. See Brunauer
Emmett
Teller (BET) theory Beverage industry, 232 Bilirubin, 325
326 Bio/active material-loaded chitosan-based nanofiber coating, 214
215 Bioaerogels possess, 304
305 Biocompatibility, 258, 271 Biodegradation, 248
249, 389
392 factors affecting, 391
392 Bioextraction, of chitosan, 402 Biological extraction, of chitin, 402 enzymatic demineralization, 7 enzymatic deproteinization process, 7 fermentation, 7 Biological methods, 38 chitin synthesis by, 342
344 LMWC production, 458
459 Biological process, chitin production, 407 Biological properties anticancer/antitumor property, 386
387 antioxidant property, 386 of chitin, 404
405, 404f of chitosan, 23f, 254
255, 351
354 of LMWC, 460
461 Biomedical applications, 46
47, 354
357, 410
412 burn treatment/artificial skin graft, 355 drug delivery, 356, 412

475

ophthalmology, 357 tissue engineering, 355
356, 410
411 wound dressing/wound healing, 354
355, 411
412 Biomimetics, 174 Bionanocomposite of chitin, 175
176 Bioplastics, 27
28 Biopolymers, 28, 37, 46, 261, 289
290, 424 intrinsic advantages of, 289
290 large-scale production of, 362 Biosynthesis, of chitin, 405 Biotechnological processes, 407 Biotransformation of chitin, 430 BJH approach. See Barrett
Joyner
Halenda (BJH) approach Black soldier fly, 230
231 Blending, 256 of chitosan, 257
258 of two polymers, 258 Blood contact applications, 257
258 BMIMCl. See 1-Butyl-3-methylimidazolium chloride (BMIMCl) Bone marrow mesenchymal stem cell, 260
261 Bromide ions, 107 Brunauer
Emmett
Teller (BET) theory, 304
305, 308f Burn treatment/artificial skin graft, 355 1-Butyl-3-methylimidazolium acetate ([Bmim][OAc]), 105 1-Butyl-3-methylimidazolium chloride (BMIMCl), 104
106, 380
381 1-Butyl-3-methylmidazoium bromide (BMIMBr), 384
385

C CAC. See Critical aggregation concentration (CAC) Calcium bromide dihydrate, 103 Calcium chloride dihydrate, 103 Capillary viscometry, 9
10 Carbonium
oxonium ion, 383 Carbon nanotubes, and chitin, 175 Carboxyethyl chitosan, 172 Carboxylic (hydroxyl) group, 68 Carboxylic groups, 145
146 Carboxymethylation of chitin, 381
382 Carboxymethyl chitin (CMC), 112
113, 179
180, 410
411 nanoparticle, 113 structure of, 113f

476

Index

Carboxymethyl groups, 167
168, 346 Casting and evaporating technique, 185 Cationic
lipid systems, 90 Cell membrane bound enzyme, 228 Cellulase, 458
459 Cellulose, 166, 361 chitin hybrid gel, 105 structures of, 167f whisker preparation process, 172
173 Centrifugal spinning technology, 46 Ceric ammonium nitrate, 116
117 Ceric ion, 116
117 C-g-PMMA. See Chitosan-graftpolymethylmethacrylate (C-g-PMMA) Chain characterization, of chitin, 107
112 configuration of, 108
111 Fourier-transform infrared spectroscopy, 109 solid 13C NMR, 110
111 X-ray powder diffraction spectroscopy, 110 length of, 111
112 ChCl. See Choline chloride (ChCl) Chemical deacetylation, of chitin, 135
138, 137f Chemical extraction, of chitin decolorization, 6 demineralization process, 5
6 deproteinization process, 6 Chemical methods, 38 chitin synthesis by, 340
342 decolorization, 342 demineralization, 341
342 deproteinization, 342 LMWC production, 455
457 Chemical modifications chitin nanofibers, 171 chitosan, 40, 147 Chemical process for chitin and chitosan production, 405
407 Chemical properties, chitosan, 254 Chemopreventive compounds, 386
387 Chitin (Ch), 2
5, 36, 60
63, 132, 166, 249
250, 336, 402, 421
426 N-acetamido bonds in, 138
139 acetylation of, 104 acid hydrolyzes of, 169 acylation reaction of, 123 antiinflammatory property, 388 antimicrobial property, 387
388 applications, 407
412

biomedical application, 22
24 bioplastics, 27
28 fuel cell, 25 industrial applications, 407
410. See also Industrial applications nanocomposite, 28 packaging of food, 25
26 textile industries, 26
27 wastewater treatment, 24
25 binary blend nanofibers, 179
180 biodegradability of, 389
392, 421 biological deacetylation of, 8 biological extraction enzymatic demineralization, 7 enzymatic deproteinization process, 7 fermentation, 7 biological properties, 385
389 anticancer/antitumor property, 386
387 antioxidant property, 386 biosynthesis, 405, 423f carbon nanotubes and, 175 chain characterization of. See Chain characterization, of chitin characterization of, 13
22 13 C Nuclear magnetic resonance analysis, 17
19 Fourier-transform infrared spectroscopy (FTIR) analysis, 14
16 scanning electron microscope, 20
22 thermogravimetric analysis, 19
20 x-ray diffraction analysis, 16 chemical deacetylation of, 8, 39f chemical extraction decolorization, 6 demineralization process, 5
6 deproteinization process, 6 chemical isolation of, 340f chemical modification of, 186
187 chemical structure of, 37f, 61f, 251f chemical synthesis of, 406f chemistry, 36
38 chitosan and. See Chitosan (CS) and chitosan-based aerogels. See Aerogels, chitin and chitosan to chitosan, conversion, 425f commercial production and disadvantages, 426
427 contents of, 251t crystalline structure, 133
134 crystal models, 385f

Index

deacetylation of, 3f, 252. See also Deacetylated chitin N-deacetylation of, 37 and deacetylation to chitosan, 402f degree of acetylation of, 110
111 derivatives of, 112
125 different structural forms of, 403f dissolution mechanism, 104. See also Dissolution process economic potential of, 361
364 emerging applications of, 363 enzymatic deacetylation of, 382 etherification, 112
115 carboxymethyl chitin, 112
113 quaternized chitin, 113
115 extraction process, 38
39, 38f, 229, 251
252 features, 290 fiber formation. See Fiber formation grafting of, 115
119 chitin nanofiber film, 118
119 hydrophilization of, 188 hydrophobization of, 187
188 hydroxyl groups of, 171 insolvability of, 62
63 major units in, 134f marine-based sources of, 338f morphology of, 20
22 nanofibrous hydrogel, 307f nanomaterials. See Nanomaterials, of chitin neuroprotective property, 388
389 O-acylation reaction. See O-acylated chitin origin and sources of, 337
338 oxidative modification, 186
187 physicochemical properties of, 8
9, 378
385. See also Physicochemical properties polymorphic forms, 37 poor solubility of, 378 possible reaction sites in, 136f potential applications of, 354
361 preparation process, 292
295 presence of, 422t production, 226
231, 343
344, 405
407 biological process, 407 biomass resource for, 240 chemical process, 405
407 quality control of, 237
240 raw materials, 229
231 synthesis of, 227
228

477

properties, 168
169, 290
292, 348
354, 403
405 biological properties of, 404
405, 404f physiochemical, 349
351 structural properties, 403 reactive functional groups in, 50 repeating unit of, 253f semicrystalline structure of, 39 solubility of, 12, 39
41. See also Solubility sources of, 3f, 4
5, 251t, 290
292, 405
407 structure of, 107f, 133f, 167f, 168
169, 172
173, 336f, 423f surface modification of, 185
189. See also Surface modification of chitin synthesis of, 168
169, 339
348 biological methods, 342
344 chemical method. See Chemical methods derivatives, 346
348 water-binding capacity and fat-binding capacity, 13 Chitin deacetylase, 430
431 Chitin dissolution systems, 236
237 Chitin-g-acrylic acid mixture, 117 Chitin
glucan
aldehyde
quercetin conjugate, 387 Chitin-g-poly(aminoethyl), 117
118 Chitin-g-polypyrrole, 117 Chitin-g-polystyrene, 117 Chitin nanocomposite, 174
175 Chitin nanofiber film (CNF), 118
119 Chitin nanofiber-g-poly(c-benzyl Lglutamate) (CNF-g-PBLG) film, 118
119 Chitin nanofibers, 170
172, 184 aggregation of, 181
182 blended nanofiber, 171 from chitin derivatives, 172 fabrication of, 183 modified, 171 pure, 170
171 using micro-contact printing method, 182f using self-assembly method, 181f Chitin nanogel, 176
177, 177f Chitin nanowhiskers, 172
174 extraction of, 184 surface modification of, 186 Chitin synthase, 228 Chitooligosaccharides (COS), 229, 385
389 by enzymatic hydrolysis, 389 neuroprotective property of, 389

478

Index

ChitoPEG copolymer, 73, 83f Chitosan (CS), 2
5, 60
63, 132, 166, 172, 202, 227, 337, 378, 421
426, 454 acetylation of, 134
145 aerogels. See Aerogels, chitin and chitosan antiinflammatory property, 388 antimicrobial activity of, 426 antimicrobial property, 387
388 applications, 3f, 16, 255
256, 407
412, 425
426, 425t agriculture, 256 biomedical applications of, 22
24, 410
412 bioplastics, 27
28 drug delivery. See Drug delivery systems food industry, 255
256 food packaging systems, 25
26 fuel cells, 25 industrial applications, 407
410. See also Industrial applications nanocomposite materials, 28 textile industry, 26
27 wastewater treatment, 24
25, 255 N-azidation of, 69 -based hydrogels, 274 biochemical properties of, 64
65 biodegradability of, 389
392 biological properties, 5
7, 385
389 anticancer/antitumor property, 386
387 antioxidant property, 386 biomaterial, 249
256 biomedical application of. See Biomedical applications biosynthesis, 423f blends, 257
258 with synthetic polymers, 264
274 characterization of, 13
22, 378, 424 13 C Nuclear magnetic resonance analysis, 17
19 Fourier-transform infrared spectroscopy (FTIR) analysis, 14
16 scanning electron microscope, 20
22 thermogravimetric analysis, 19
20 x-ray diffraction analysis, 16 chemical modifications of, 147 chemical structure of, 37f, 62f, 135f, 251f chemistry, 36
38 and structure of, 132
134 chitin deacetylases for, 236

clay hybrid materials, 307 commercial production and disadvantages, 426
427 composition in membranes, 268f conversion of chitin to, 425f cross-linking of, 66
67, 70
71, 147
148, 149f degree of acetylation. See Degree of acetylation (DA) with degree of deacetylation, 63 degree of solubility of, 145 dissolution mechanism. See Dissolution process economic potential of, 361
364 electrospinnability of, 200
201 emerging applications of, 363 enzymatic grafting of, 150f enzymatic hydrolysis of, 382 extraction methods, 38
39, 38f, 252f, 432f fiber formation. See Fiber formation fungal. See Fungal Chitosan in fungal cell wall, 236 by glycerol, 137f grafting copolymerization of, 151
152 green synthesis of, 427
428 hydrophilic property of, 257
258 hydroxyl groups of, 64
65, 70 industrial applications of. See Industrial applications, of chitosan low-molecular weight, 10 major units in, 134f metal binding by, 351 modification, 256
258 polymer blending technique, 256
257 molecular weight, 9
10, 253
254 morphology of, 20
22 nanofiber mats, 48 nanofibers, 200
201 nanoparticles, 215
216 natural polymers blends with. See Natural polymers neuroprotective property, 388
389 origin and sources of, 337
338 oxidization of, 151
152 peaks for, 73 PEGylation of. See PEGylation physicochemical properties of, 8
13, 64
65, 378
385. See also Physicochemical properties polysaccharides, 255 possible reaction sites in, 136f potential applications of, 354
361

Index

in powder, 458f preparation, 8, 137f, 249
253 production, 231
240, 405
407 alternative methods of, 233
237 biological process, 407 biotechnological method for, 236
237 chemical process, 405
407 combined biological methods, 236 enzymic methods, 235
236 fermentations, 233
235 properties, 253
255, 348
354, 403
405 biological properties, 254
255, 290
292, 351
354, 404
405, 404f chemical properties, 254 physical properties, 253
254 physiochemical, 349
351 structural properties, 403 ratio, effects of, 259f, 260f reactive functional groups in, 50 solubility, 12, 39
41, 63, 253
254. See also Solubility in dilute acids, 253
254 sources of, 250f, 290
292 structural modification, 3
4 structure, 133f, 167f, 252
253, 336f, 423f, 424f synthesis of, 339
348 derivatives, 346
348 viscosity, 10
11 water-binding capacity and fat-binding capacity, 13 water-soluble, 145
151, 150f Chitosan-based face mask, 48
49 Chitosan-based films, 26 Chitosan-based nanofibers coating, 212
214 bio/active material-loaded, 214
215 liquid smoke-loaded, 210
211 thymol-loaded, 214
215 Chitosan-based nanomaterial controlled release property of, 210 electrospinning. See Electrospinning process encapsulation efficiency, 209
210 morphological characteristics of, 207
209 thermal decomposition of, 210
211 zeta potential and size of, 211
212 Chitosan
collagen nanofiber, 196
198 Chitosan electrospun fibers, 199
200 Chitosan-folate nanoparticles, 215
216 Chitosan-graft-polymethylmethacrylate (Cg-PMMA), 390, 391f

479

Chitosan-L-glutamic acid (CL-GA) aerogel derivative, 302, 302f, 310
312 Chitosan nanoscaffold, 174 Chitosan
silica composite aerogel, 297 hybrid aerogel, 293
295 Chitosan
thioglycolic acid (chitosan-TGA), 356 Chitosan
transition metal ion complex fibers, 49 Chitosomes, 405 2-Chloroethylamine hydrochloride aqueous solution, 117
118 (3-Chloro-2-hydroxypropyl) trimethylammonium chloride (CHPTAC), 115 3-Chloro-2hydroxypropyltrimethylammonium chloride (CTA), 114 Chlorophenols, 410 Choline chloride (ChCl), 124
125 Choline chloride
malonic acid, 236
237 CHPTAC. See (3-Chloro-2-hydroxypropyl) trimethylammonium chloride (CHPTAC) Cicadas, 230
231 Citric acid, 135
136 CL-GA aerogel derivative. See Chitosan-Lglutamic acid (CL-GA) aerogel derivative Click chemistry, 69, 69f CMC. See Carboxymethyl chitin (CMC) CNF. See Chitin nanofiber film (CNF) CNF-g-poly (LA-co-CL) film, 118 Coagulation
flocculation process, 357
358 COF. See Covalent organic framework (COF) Collagen, 260
261 Colletotrichum gloeosporioides, 236 Colletotrichum lindemuthianum, 430
431 Colloidal titration method, 77 Colorimetric method, 77
78 Combined biological methods, 236 Composite of chitin, 411
412 of chitosan, 411 Compound annual growth rate (CAGR), 363 Compression stress
strain curves, 316
317, 316f Contact lenses, 357 Conventional method of chitosan, 428 Conventional spinning methods, 42

480 Cosmetic applications, chitin and chitosan fibers for, 48
49 Cosmetic product, 408 Cosmetics industries, 360 Cospinning agents, 201 Covalent organic framework (COF), 291
292 Crab chitin-based 2D soft materials, 177
178 Crab shells chitin extraction, 170 exoskeleton structure of, 169f waste, 407 Crangon crangon shrimp waste, 36
37 Critical aggregation concentration (CAC), 68 Cross-linking process aerogel materials, 317 chitosan, 70
71, 147
148, 149f, 411 chitosan-based aerogels, 296
297 with FeCl3 and CaCl2, 113 with polyethylenimine, 288 Crustaceans, 228, 233, 236
238, 249
250 chitin, 236 exoskeleton of, 249
252 wastes, 252f Crustacean shells, 342 Crystalline index (CI), 110 Crystalline polymorph, 16 Crystalline polymorphic forms, 102 Crystalline polysaccharides, 39 Crystallinity of alkyl-acylated chitin, 124 β-chitin, 16 during chitosan production, 233
234 semicrystalline polymer, 349
350 CS-PDA aerogels. See Polydopaminemodified chitosan (CS-PDA) aerogels CTA. See 3-Chloro-2hydroxypropyltrimethylammonium chloride (CTA) Cultivated plants, 230
231 Culture media, for fungal production, 441
442 Culturing fungus, 428 Curcumin, 176
177 Cuticle, 228 Cyprinus carpio, 463 Cytocompatibility, 83
85 Cytotoxicity, 83
85 of chitosan, 461

Index

D DCC. See 4-Dicyclohexylcarbodimide (DCC) Deacetylated chitin, 133 catalytic action of, 138f methods, 135
139 chemical deacetylation, 135
138, 137f enzymatic deacetylation, 138
139 monitoring process for, 139 N-Deacetylation, 132 Deacetylation degree (DD), 39 Deacetylation process, 12, 429 of acetyl groups, 8 chitin, 3
4, 39, 252 Decoloration process, 38, 61, 340, 342 Decolorization, 6 Degradation procedure of chitin/chitosan, 391
392 Degree of acetylation (DA), 105, 132, 134
135, 229, 236, 348, 404
405, 435 characterization methods, 139
140 of chitin, 110
111 of chitosan, 139
145 acid hydrolysis, 141 distribution of acetyl groups, 142 elemental analysis, 140 high-performance liquid chromatography, 141 infrared, 141
142 measurement of, 140
142 nuclear magnetic resonance spectroscopy, 142 pyrolysis GC-MS, 141 relationship between acetylation and properties, 142
145 titration methods, 141 Degree of deacetylation (DD/DDA), 8
9, 12, 61
63, 133
134, 137
140, 142
145, 252
254, 345, 352, 435
436, 454, 458
460 Degree of deproteinization, 428 Degree of hydrolysis (deacetylation), 252
253 Degree of N-acetylation, 170 Degree of PEG-g-chitosan (DEPG), 85
86 Degree of substitution (DS) determination of, 76
80 of N,O-PEGylated chitosan copolymers, 80t of N-PEGylated chitosan copolymers, 78t of O-PEGylated chitosan copolymers, 80t Degrees of solubility (DS), 67
68

Index

Demineralization process, 38, 339
342, 405
407 of chitin, 5
6, 38 enzymatic, 7 2-Deoxy-2-(acetylamino) glucose, 252
253 DEPG. See Degree of PEG-g-chitosan (DEPG) Deproteinization process, 7, 339
341, 343, 405 of chitin, 6, 39 enzymatic, 7 Derivatization method N-hydroxy succinimidyl ester, 66f p-nitrophenyl carbonate, 67f Dexorubicin, 177 D-glucosamine units, 40, 139
140 DHA. See Docosahexaenoic acid (DHA) Dibutyrylchitin, 179
180 Dichloroacetic acid (DCA), 85
86, 380 Dichloromethane (DCM), 64, 200
201 4-Dicyclohexylcarbodimide (DCC), 65
66 Differential scanning calorimetry (DSC), 75, 310
312 1,6-Diisocyanatohexane, 147
148 Dilute acetic acid, 255 Dilute acids solubility of chitosan in, 253
254 solutions, 200 2,2-Dimethoxy-2-phenyl acetophenone (DMPA), 89 N,N-Dimethylacetamide (DMAc), 104, 111
112, 123
124 Dimethyl-3,3-dithio-bis-propionimidate (DTBP), 411 Dimethylformamide, 379
380 Dimethyl sulfoxide, 379
380 3-(4,5-Dimethylthiazol-2-yl)-2,5diphenyltetrazoliumbromide assay, 91 1,4-Dioxane/water mixtures, 269
270 1,1-Diphenyl-2-picrylhydrazyl (DPPH) radicals, 386 2,2-Diphenyl-1-picrylhydrazyl (DPPH) radicals, 352
353 Diphosphate-N-acetylglucosamine, 421
422 Dissolution process of acetyl groups, 142 of chitin crystal models, 385f by inorganic reagents, 380 mechanism, 383
385 acid-catalyzed hydrolysis, 383, 383f base-catalyzed hydrolysis, 383
384

481

cleavage/dissolution by ionic liquids, 384
385 by polar solvents and strong acids, 380 DMAc. See N,N-Dimethylacetamide (DMAc) Docosahexaenoic acid (DHA), 215 DPPH radical. See 1,1-Diphenyl-2picrylhydrazyl (DPPH) radicals Drug delivery systems, 356, 412 nanoparticles in, 86
87 PEGylated chitosan in, 66
67 Drum collector, 206
207, 206f Drying technologies for aerogel production, 294t Dry spinning method, 41
42 Dry
wet spinning technique, 48 DSC. See Differential scanning calorimetry (DSC) DTBP. See Dimethyl-3,3-dithio-bispropionimidate (DTBP) Ductility of chitosan, 81 Duration of fermentation, 239 Dye removal and wastewaters treatment, 47
48

E Eco-friendly production, 420 EDC. See 1-Ethyl-3-(3dimethylaminopropyl) carbodiimide (EDC) EDTA. See Ethylene-diamine-tetracetic acid (EDTA) Eicosapentaenoic acid (EPA), 215 Electron microscopic techniques, 207
208 Electrospinning dope solutions preparation, 202 Electrospinning method, 113 Electrospinning process, 42, 42f, 201 ambient parameters, 44
45 of chitin, 178
180, 179f determination of, 202
207 adjustment proper distance, 203
204 applied voltage, 202
203 environmental conditions, 205
206 flow rate, 204
205 selection of collector, 206
207 drum collector used in, 206f factors affecting, 42
45 humidity on, 45 processing parameters, 43 solution parameters, 44 zeta potential, 211
212

482 Electrospinning unit, 196, 197f response surface methodology for, 196
198 Electrospun chitosan, 205
206 elemental analysis of, 208
209 nanofibers, 205
206, 213
214, 213f nanomaterials, 200, 212 oil-loaded, 215
216 Electrospun fibers, of chitin and chitosan, 46 Electrospun polymer fibers, 48 Electrospun skin mask, 48 Elemental analysis, 140 Encapsulation efficiencies (EE), 198
199, 209
210 Endo-chitinases, 390 Endo-type enzyme, 430 Energy dispersive X-ray spectroscopy (EDX), 208
209 Energy sources, 238 Enzymatic deacetylation, 138
139, 382 Enzymatic depolymerization methods, 458
459, 459t Enzymatic hydrolysis, 290, 382 chitooligosaccharides by, 389 Enzymes, 138
139, 430 chitin deacetylase, 430
431 in commercial sources, 227
228 exo-type and endo-type, 430
431 Enzymic methods, 235
236 EPA. See Eicosapentaenoic acid (EPA) 2,3-Epoxypropyl trimethyl ammonium chloride (EPTMAC), 46
47 2,3-Epoxypropyl trimethylammonium chloride (EPTMAC), 115 EPTMAC. See 2,3-Epoxypropyl trimethyl ammonium chloride (EPTMAC) Erwinia chrysanthemi, 235
236 Escherichia coli, 26
27, 46, 387
388, 461 Etherification of chitin, 112
115 carboxymethyl chitin, 112
113 quaternized chitin, 113
115, 114f 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC), 67 Ethylene-diamine-tetracetic acid (EDTA), 342 1-Ethyl-3-methyl-imidazolium acetate, 405 1-Ethyl-3-methylimidazolium alkanoates, 380
381 1-Ethyl-3-methylimidazolium chloride ([Emim][Cl]), 106
107 Evaporation method, 187

Index

Exocuticle, 228 Exoskeletons, 228 of arthropods, 290 shells, 402 Exo-type enzyme, 430 Extraction process, 12, 234 of chitin, 233 of chitin nanowhisker, 184 of chitosan, 38
39, 38f, 432f

F Fabrication of chitin nanofibers, 183 nanoscale material, characterization of, 207
212 of PEGylated chitin/chitosan derivatives, 65
71 amino group substitution, 66
67, 69 cross-linked chitosan network, 70
71 miscellaneous approaches, 69 O-substitution, 67
68 solubilization of, 69
70 Fatty acids, 381
382 Fenneropenaeus indicus, 234 Fenneropenaeus semisulcatus, 234 Fermentation conditions, 442, 445t, 448 Fermentation process, 7, 233
235, 343
344 Fermentation technology, 420, 448 FESEM. See Field emission SEM (FESEM) Fiber formation, 272 chitin and chitosan, 41
50 applications, 46
50 for biomedical applications, 46
47 characterization of, 45
46 for cosmetic applications, 48
49 in different fields, other applications for, 49
50 for dye removal and wastewaters treatment, 47
48 electrospinning process. See Electrospinning process Fibers of chitin and chitosan, 49
50 Fiber spinning techniques, 41 Field emission SEM (FESEM), 207
208 Filamentous fungi, 445
446 Fish fillets, 213 Fish meal production, 240
241 Flat/drum collector, 206
207, 206f Flocculating agent, 255
256 Flow rate, of polymeric solution, 204
205 5-Fluorouracil (5-F) sustainably, 113 Food industry, 232, 255
256, 409
410, 425

Index

Food packaging systems, 25
26 Food processing industry, 359
360, 420, 441 Food technology, LMWC in, 463
464 Formic acid, 253
254, 379
380 Fourier-transform infrared spectroscopy (FTIR), 14
16, 45, 81, 106
107, 109, 273f aerogels techniques, 317
319, 318f, 319f structural analysis, 71
72 Free amino groups, 379 Free radicals, 386
387 polymerization, 69 Freeze-drying method, 306, 309, 314
315 Freeze-drying process, 295, 301
302 Freezing/thawing method, 102 FTIR. See Fourier-transform infrared spectroscopy (FTIR) Fuchsin acid dye by adsorption, 47
48 Fuel cells, 25 Functional groups, of polysaccharide, 304
305 Fungal chitosan, 429
446, 434t commercial production of, 442
446 production, 431
446 Gongronella butleri, 438
439 industrial residues as culture medium for, 441
442 Mucor rouxii, 433
436, 436t other fungal strains, 439
441, 440t Rhizopus oryzae, 437
438, 437t significance, 429
431 synthesis of, 227
228 Fungal mycelia, 446 Fungal species, 429
430, 433 Fungi, 229
230, 250, 252
253, 337
338, 344 “Fungine”, 36

G Galactosylated chitosan (GC), 410
411 γ-chitin, 4
5, 14, 16, 19
20, 37, 62
63, 102, 108, 169, 226
227, 336
337, 349, 349f, 403, 403f TGA curves of, 20f Gamma irradiation, 457
458 Gelatin, 411 Gelation method, 184 Gel-based hydrocolloids, 292
293 Gel depot system, 88 Gelling delivery system, 88 Gel permeation chromatography (GPC), 9
10, 71, 73
75

483

Gene delivery systems, 90
92 Generally recognized as safe (GRAS), 273, 437 Genipin crosslinked blend (Agr-CS-G), 309 Gibberellic acid, 236 GlcNAc. See N-Acetyl-D-glucosamine (GlcNAc) Glucosamine, 141, 290
291, 402 Glucosamine-6-phosphate, 421
422 Glutamine-fructose-6-phosphate, 227 Glycerol, 151
152 Glycerol-2-phosphate, 63 Glycol chitin, 346 Glycol
chitosan, 151
152 Glycosidic bonds, 454
455 GMP. See Good Manufacturing Practice (GMP) GOCA. See Graphene oxide/chitosan aerogel (GOCA) Gongronella butleri, 229
230, 236, 438
439, 439t, 442
443 Good Manufacturing Practice (GMP), 239
240 GPC. See Gel permeation chromatography (GPC) Graft copolymerization, 64, 256 Grafted chitin, 115
119, 116f chitin nanofiber film, 118
119 Grafting copolymerization, 73
75 of chitosan, 151
152 Grafting-modified chitosan, 71
72 Grafting PEG, 60
61, 64
65 Gram-negative bacteria, 211
212 Gram-positive bacteria, 211
212 Graphene oxide(GO), 312
313 Graphene oxide (GO)
chitosan (CS), 302, 303f Graphene oxide/chitosan aerogel (GOCA), 309
310, 326
327 thermal stability of, 311t Graphene oxide/chitosan fibers, 47
48 Green extraction methods, 342
343 “Green” method, 181
182 Green synthesis, of chitosan, 427
428 Gryllus bimaculatus, 230
231

H Harvesting time, 446 Heavy metals, 24
25 Hemicelluloses
chitosan foam, 315
316, 316f

484

Index

Hemicelluloses citrate
chitosan foam, 315
316, 316f Heterogeneous system deacetylation, 135
136 LMWC molecules, 460
461 O-acylated chitin in, 119
121 Heteronuclear multiple quantum coherence (HMQC), 71 Hexafluoroisopropanol, 121 1,1,1,3,3,3-Hexafluoro-2-propanol (HFIP), 179, 200
201 self-assembly method using, 180
181 Hexanoyl chitosan, 274 HFIP. See 1,1,1,3,3,3-Hexafluoro-2-propanol (HFIP) Hierarchical system, 248
249 High-density lipoprotein (HDL), 352
353 High-molecular-weight chitosan (HMWC), 352
353, 454
455 High-performance liquid chromatography (HPLC), 141, 410 High-quality chitin nanofibers, 167 HMQC. See Heteronuclear multiple quantum coherence (HMQC) H-NMR spectroscopy, 87
88 HOBt. See Hydroxybenzotriazole (HOBt) Homogeneous chitosan fibers, 46 Homogeneous deacetylation, 135
136, 145
146, 344
345 Homogeneous system, O-acylated chitin in, 121
125 ionic liquid system, 124
125 lithium chloride/dimethylacetamide system, 123
124 methanesulfonic acid system, 122 trifluoroacetic anhydride system, 122
123 Humidity, on electrospinning process, 45 Hybrid nanostructures, structure morphology of, 297
300, 299f Hydration process, 290 Hydrochloric acid, 5
6, 120, 146
147 Hydrogels, 176, 304
305, 347 chitin nanofibrous, 307f chitosan-based, 274 swelling ratio of, 89 Hydrogen bonds, 12, 14, 16, 268
269 Hydrogen peroxide, 386 Hydrohobicity, 410 Hydrolysis, glycosidic linkage by, 146
147 Hydrolytic methods, 139
140 Hydrophilic drug, 177

Hydrophilic functional groups, 274 Hydrophilic outer shell, 87
88 Hydrophilic polymers, 210
211, 265
268, 266f Hydrophilic property of chitosan, 257
258 Hydrophilization of chitin surface, 188 Hydrophobic acetyl groups, 254 Hydrophobic biobased nanofibrous aerogels, 323
324 Hydrophobic chains, 274 Hydrophobic groups, 23 Hydrophobic methyl groups, 381
382 Hydrophobization of chitin surface, 187
188 N-Hydroxyalkanoyl groups, 147 Hydroxyapatite, 411 Hydroxybenzotriazole (HOBt), 70 2-Hydroxyethyl acrylate (HEA), 119 2-Hydroxyethylmethacrylate (HEMA), 188 Hydroxylbenzenesulfonailides derivatives, 387 Hydroxyl groups, 61
63, 384, 402 of chitin, 171 of chitosan, 64
65, 70 Hydroxyl radicals, 386 Hydroxyls, 351 Hydroxymethyl chitin, 409 Hypercholesterolemia, 463
464

I IBD. See Inflammatory bowel disease (IBD) Industrial applications, 314, 407
410 of chitosan, 357
361 agriculture, 358
359 cosmetics industries, 360 food processing, 359
360 gel for light-emitting device, 360
361 paper industry, 361 photography, 360 textile industry, 361 water engineering, 357
358 cosmetics, 408 food industry, 409
410 other, 410 paper and textile industry, 409 water engineering, 408
409 Industrial by-products, 441 Inflammatory bowel disease (IBD), 171 Infrared, DA of chitosan, 141
142 Inhibition antibacterial test system, 207
208 Inoculum, 238

Index

485

Kluyveromyces lactis, 409 KOH aqueous solution, 184 Kurthia gibsonii, 234

LMWSC 10K. See Low-molecular-weight water-soluble chitosan (LMWSC) 10K Low-molecular-weight chitosan (LMWC), 10, 352
353, 454 agriculture, 463
464 antioxidant activity of, 461 applications, 461
462, 462t aquaculture, 463 crystalline structure of, 460 derivatives, 388 dietary supplementation of, 463 food technology, 463
464 industrial production of, 454
455 plant protective properties of, 463 preparation, 454
459, 455f biological methods, 458
459 chemical methods, 455
457 degradation methods for, 456t oxidation-based methods, 456
457 physical methods, 457
458 using enzymatic methods, 458
459, 459t properties, 460
461 biological activities, 460
461 physicochemical properties, 460 Low-molecular-weight water-soluble chitosan (LMWSC) 10K, 72
73, 74f Lyophilization technique, 302

L

M

Lactic acid bacteria, 233
234 Lactic acid fermentation, 233, 238 Lactic acid-mediated demineralization, 428 Lactobacillus brevis, 234 Lactobacillus helveticus, 343 Lactobacillus paracasei, 407 Lactobacillus plantarum, 233, 238 Lactobacillus vannamei, 233 Lactococcuslactis, 343
344 LiCl. See Lithium chloride (LiCl) Light-emitting device, chitosan gels, 360
361 Linear aminopolysaccharide, 460 Linear polymer chain, 339 Lipopolysaccharide, 460
461 Liquid fraction, 240
241 Liquid smoke-loaded, chitosan nanofibers, 210
211 Lithium chloride (LiCl), 104, 111
112, 123
124 Lithium thiocyanate, 62
63

MALLS. See Multiangle light scattering (MALLS) Mandrel collector, 206
207 Mannosylated chitosan (MC), 410
411 Marine-based biopolymer, 361 Marine biopolymer, 354 Marine gastropods, 337
338 Mark
Houwink equations, 10 Material research, 286
287 application, 286f MBA. See Methylene bisacrylamide (MBA) MBC. See Minimum bactericidal concentration (MBC) MC. See Mannosylated chitosan (MC) MCTCS, 315, 315f MC3T3-E1 cell proliferation, 260
261 Mealworms, 230
231 Mechanical property of chitosan aerogels, 314
317, 314f of P/CA, 316
317 Mechanical treatment, 181
182 Medical applications, 86
88

Inorganic acids, 39
40, 146
147 Inorganic/organic nitrogen source, 442 Inorganic reagents, dissolution by, 380 Insects, 230
231 biomass, 405
407 wing, 337
338 In situ lactic acid production, 344 Intercalation method, 121 Intermolecular hydrogen bonds, 14
16, 85
86, 253
254 Interpenetrating polymer network (IPN), 386 Intrinsic viscosity, 9
11, 11t Ionic gelation method, 64
65 Ionic liquid (ILs) system, 104
107, 106t, 124
125 cleavage/dissolution by, 384
385 for dissolution of chitin, 381f solubility in, 380
381 Irradiation process, 457
458

J Jet solution, 203
204

K

486

Index

Medium-molecular-weight chitosan (MMWC), 10, 454 Melt spinning method, 41
42 MeO-PEG acid, 82
83, 85
86 Metapenaeusmonoceros, 235 Meth-A fibrosarcoma model, 90
91 Methanesulfonic acid system, 121
122 Methanol, 64 calcium chloride dihydrate in, 103 Methoxy-poly (ethylene glycol) (mPEG), 65
67, 69, 73f, 74f grafting of, 71
72 methyl group of, 72 specific peaks of, 72 α-Methoxy-ω-succinimidylpoly(ethylene glycol) (MSS-PEG), 91 Methylene bisacrylamide (MBA), 89 Methylmethacrylate, 390 MIC. See Minimum inhibitory concentration (MIC) Micellar systems, 87
88 Microbial fermentation, 339 Microcontact printing method, 181, 182f Microcrystalline chitin (MCC), 359
360, 409 Microorganisms, 235
236, 252
253, 428
429 Microorganisms-mediated fermentation processes, 342
343 Microwave-assisted deacetylation, 137
138 Microwave-assisted extraction, 428 Microwave technique, 345 Milk coagulation, 409 Minimum bactericidal concentration (MBC), 115 Minimum inhibitory concentration (MIC), 115 Miscibility/solubility, 379
382 MMA. See Poly(methyl methacrylate) (MMA) Modification of chitosan (CS), 256
258 procedures, 185 of synthetic polymers, 264
265 Modified chitin nanofibers, 171 Mold-casting techniques, 411 Molecular dynamics (MD), 107 Molecular self-assembly, 180
181 Molecular weight (MW), 6, 39, 103, 111
112, 386, 429 chitosan, 253
254, 435 definition, 198
199

distribution, 253 of polymer, 44, 134
135 Monochloroacetic acid, 112, 381
382 Monomethyl-modified chitosan, 40 Morphological/microscopic analysis, 296
304 mPEG. See Methoxy-poly (ethylene glycol) (mPEG) mPEG-g-PEI-grafted chitosan, 91
92 MSS-PEG. See α-Methoxyω-succinimidylpoly(ethylene glycol) (MSS-PEG) Mucorales strain, 433 Mucorrouxii, 429
431, 433
436, 436t, 444
445 Multiangle light scattering (MALLS), 73
75 Mycelia, 432
433, 432f Mycotoxinogen fungal strains, 273

N N-acetylation of chitin, 402 NaClO, 183 Nanocomposite, 28 of chitin, 174
175 Nanocrystals, 166
167 Nanoencapsulation material, 209
210, 209f Nanofiber mats, chitosan, 48 Nanofibers, 42, 167 antibacterial properties of, 212 chitin, 49
50. See also Chitin nanofibers chitosan, 49
50, 172, 200
201 electrospun chitosan, 205
206, 213f formation of, 92
93 mats, 172 Nanofibrils, 166
167 Nanogel of chitin, 176
177 Nanomaterials, 166
167 electrospun chitosan, 200 structure of, 207
208 Nanomaterials, of chitin, 166
167, 169
178 chemical modification of, 186
187 acetylation of chitin, 186 chitin nanofibers, 170
172, 178 blended nanofiber, 171 from chitin derivatives, 172 modified, 171 pure, 170
171 nanocomposite of chitin, 174
175 nanogel of chitin, 176
177, 177f nanowhiskers of chitin, 172
174 polymer/chitinbionanocomposite, 175
176

Index

preparation, 178
185 aqueous counter collision method, 180 casting and evaporating technique, 185 electrospinning of chitin, 178
180, 179f extraction of chitin nanowhiskers, 184 gelation method, 184 mechanical treatment, 181
182 microcontact printing method, 181 self-assembly method, 180
181 TEMPO-mediated oxidation, 183 ultrasonication, 182
183 two-dimensional soft nanomaterials, 177
178 Nanoparticles of chitosan, 215
216 Nanoscale chitosan, 196
198 Nanoscaled polymeric assemblies, 166
167 Nanowhiskers of chitin, 166
167, 172
174 NaOH/urea aqueous solution, 102
103 Natural biopolymers, 420 Natural carbohydrate polymer, 337 Natural polymers, 274 blending with chitosan, 258
264 collagen, 260
261 hydrophilic polymers, 265
268, 266f natural rubber latex, 263
264 nylons, 268
270 poly(lactic acid), 273
274 polyacrylamide, 271
273 proteins, 261
263 starch, 258
259 Natural polysaccharides, 420 Natural rubber latex, 263
264 NCOC, 297
300 N-deacetylation, 290
291, 387 of chitin, 37 Near-infrared (NIR), 141
142 Net enthalpy, 304
305 Neuroprotective property, 388
389 NHS. See N-hydroxysuccinimide (NHS) N-hydroxysuccinimide (NHS), 65
66, 188
189 N-hydroxysuccinimidyl ester, 67 derivatization method, 66f Ninhydrin (triketohydrinedene hydrate) method, 141 Nitrogen, 442 Nitrogen adsorption
desorption isotherms, 305f, 306
309, 306f, 307f, 308f, 310f Nitrogenous polysaccharide, 2 N-methylene phosphonic chitosan, 41 N-methyl morpholine-N-oxide (NMMO), 403

487

NMR. See Nuclear magnetic resonance (NMR) spectroscopy N,N-dimethylacetamide DMA, 39
40 Nomenclature border, 37 Novel chitosan-O-mPEG graft copolymers, 65 N-PEGylated chitosan copolymers, 78t N-phenyl-L-naphthylamine (PNA), 85
86 N-phthaloyl chitosan, 67
68, 77, 346 Nuclear magnetic resonance (NMR) spectroscopy, 72
73, 322
323, 322f, 323f Nylon 66, 268
269 Nylons, 268
270, 271f

O O-acylated chitin, 119
125 in heterogeneous system, 119
121 acid catalyst method, 120
121 activation of chitin, 121 in homogeneous system, 121
125 Ionic liquid system, 124
125 lithium chloride/dimethylacetamide system, 123
124 methanesulfonic acid system, 122 trifluoroacetic anhydride system, 122
123 structure of, 120f 20 -O-HTACCt. See 20 -O-Hydroxypropyltri methylammoniumchitin chloride (20 -OHTACCt) OHT-chitin. See O-(2-hydroxy-3trimethylammonium) propyl chitin (OHT-chitin) 20 -O-Hydroxypropyltrimethylammoniumchitin chloride (20 -OHTACCt), 114 O-(2-hydroxy-3-trimethylammonium) propyl chitin (OHT-chitin), 114 Oil-in-water emulsification, 187 Oil-loaded electrospun chitosan, 215
216 O-PEGylated chitosan copolymers, 80t Ophthalmology, 357 Optimization, 196
202, 239 chitosan molecular weight, 198
199 determination of concentrations, 199
200 of electrospinning parameters, 202
207. See also Electrospinning process electrospinning dope solutions preparation, 202

488

Index

Optimization (Continued) ofelectrospun nanoscale chitosan, 196
198 parameters, 204t solvent system, 200
202 Organic acids, 146
147 dilute solutions of, 252
253 Organic aerogels, 288 Organic pollutants, 24
25 Organic solvents, 40, 70, 85
86, 379 Orthoptera species, 405
407 Orthorhombic system, 108 O-substitution, 67
68 6-O-Triphenylmethyl-chitosan, 71
72, 82, 85
86 Oxidation-based methods, 456
457 Oxidative modification, 186
187 Oxidative stress-related disease, 386 Oxidization of chitosan, 151
152

P PAA. See Poly(acrylamide) (PAA) Packaging of food, 25
26 Paper industry, 361, 409 Parapenaeuslongirostris, 343 Partially deacetylated chitin, 61
62 Partially N-acetylated chitosan, 142, 143f Pathogenic fungi, 387 PCBs. See Polychlorinated biphenyls (PCBs) PCL. See Poly(ε 2 caprolactone) (PCL) Pectinase isozyme, 390 PEG-aldehyde, 66
67 in aqueous organic acid, 92
93 PEG-dialdehyde diethyl acetals, 66
67 PEG-g-chitosan, 76
77, 83
86 aggregation number of, 85
86 cytotoxic effect of, 83
85 DPEG of, 85
86 by hydrogen bonds in aqueous solutions, 85
86, 86f solubility of, 82
83 solution properties of, 85
86 PEGylated chitin/chitosan derivatives, 64
65. See also PEGylation all-trans retinoic acid in, 87
88 applications of, 86
93 formation of nanofibres, 92
93 gene delivery systems, 90
92 medical, 86
88 thermoresponsive hydrogel system, 88
90 characterization of, 71
86

aggregation, 85
86 cytocompatibility, 83
85 degree of substitution. See Degree of substitution (DS) ductility, 81 solubility, 82
83 structural analysis. See Structural analysis fabrication of, 65
71 amino group substitution, 66
67, 69 cross-linked chitosan network, 70
71 miscellaneous approaches, 69 O-substitution, 67
68 solubilization of, 69
70 free amino groups of, 86
87 micellization behavior of, 68 modification of, 93 synthesis of, 65
66 PEGylation, 60
61, 64
65. See also PEGylated chitin/chitosan derivatives of carboxyl derivatives, 68 of chitosan, 64
67, 69f trimethyl chitosan with, 83
85 Penicilliumnotatum, 353 Penicilliumoxalicum, 139 Perchloric acid, 120
121 Petroleum-based products, 362 PGA. See Polyglycolic acid (PGA) pH50, 82 Pharmaceutical sector, 232 Phosphoric acid mixed system, 123 Photography, 360 Phycomycesblakesleeanus, 429, 435 Physical degradation, 454
455 Physical methods, 457
458 LMWC production, 457
458 Physical properties, chitosan, 253
254 Physicochemical properties of chitin and chitosan, 349
351, 378
385 dissolution by inorganic reagents, 380 miscibility/solubility, 379
382 polar solvents and strong acids, dissolution by, 380 solubility in ionic liquids, 380
381 of LMWC, 460 Phytohormones, 236 Picric acid method, 141 Plasma treatment, 188
189 Plasmid DNA (pDNA), 90 PLR. See Poly-L-arginine (PLR) PLR-PEG-g-chitosan, 91
92

Index

PNA. See N-phenyl-L-naphthylamine (PNA) p-nitrobenzoic acid, 121 Polarizing light microscopy, 180 Polar solvents and strong acids, dissolution by, 380 Poly(3-hexylthiophene) (P3HT), 186
187 Poly(acrylamide) (PAA), 271
273 Poly(caprolactone), 175 Poly(ε 2 caprolactone) (PCL), 199
200, 208 Poly(ethylene glycol), 175 Poly(ethyleneoxide) (PEO), 199
200, 202
203, 207
208 nanofibrous membrane, 208
209 Poly(ethylenimine) (PEI), 301
302 Poly(glycolic acid) (PGA), 171 Poly(lactic acid) (PLA), 207
208, 273
274 Poly(methyl methacrylate) (MMA), 116
117 Poly(styrene-co-butyl acrylate), 175 Poly(vinyl alcohol) (PVA), 173, 199
200, 210
211 electrospinnability of, 208
209 Polyacrylamide (PAAm), 271
272 Polyamides, 268
269 Polychlorinated biphenyls (PCBs), 357
358 Polydopamine-modified chitosan (CS-PDA) aerogels, 309, 326
327 Polyethylene glycol (PEG), 60
61, 64, 70
71 acetaldehyde hydrates, 66
67 alkylating agents, 67 cross-linked product, 70
71 degree of substitution of, 83
85 derivatization method N-hydroxysuccinimidyl ester, 66f p-nitrophenyl carbonate, 67f grafting of, 71
72 modified tumor necrosis factor-α, 90
91 molecular weight of, 83
85 monomethyl ether iodide, 67
68 pure chitosan and, 76 Polyethylene oxide, 49 Polyethylenimine, cross-linked with, 288 Polyglycolic acid (PGA), 411 Poly(ethylene glycol)-grafted chitosan (PEG-g-CS), 207
208 Poly-L-arginine (PLR), 91
92 Polymer bionanocomposite, 175
176 Polymerblending technique, 256
257 Polymer membranes, 261
262 Polymer nanofibers formation, 179f

489

Polymer
polymer miscibility, 256
258 Polypeptides, 251 Polysaccharides, 8, 166, 175
176, 185, 229
230, 292, 303
304, 403, 424 aerogels, 296 chains, 228, 336
337 chitosan, 255 functional groups of, 304
305 polymer, 378 Polystyrene fibers, 43 Polyvinyl alcohol (PVA), 46, 265
266, 410
411 Porosity, 304
309 Potassium permanganate (KMnO4), 342 Powder XRD (PXRD) pattern, 319, 320f Preferred industrial method, 344
345 Preparation methods of chitosan, 8 Primary amino groups, 145
146 Prolonged inflammation, 388 Properties, relationship between acetylation and, 142
145 Protease, 139 Protein extract (PE), 262
263 Protein hydrolysates, 234, 238 Proteins, 261
263 Pseudomonas aeruginosa, 139, 235 Pure chitin nanofibers, 170
171 Purified chitin-based wound dressings, 354
355 Purified chitin powder, 102
103 PVA. See Polyvinyl alcohol (PVA) PXRD pattern. See Powder XRD (PXRD) pattern Pyrolysis GC-MS, 141 Pyrolytic graphite electrodes, 261

Q Quality control, of chitin production, 237
240 Quantitative NMR spectroscopy, 142 Quaternary ammonium groups, 151
152 Quaternization of chitin, 113
115 structure of, 114f Quaternized chitosan, 46 chloride, 88
89 Quick-freeze method, 309 Quinoa protein extract (PE), 262
263

R Raman spectroscopy, 45, 312
313, 312f, 313f Raw chitin, 188

490

Index

Raw CNF, 118 Raw materials, 238 chitin production, 229
231 fungi, 229
230 insects, 230
231 marine-based sources of, 362 Raw shrimp shells, 20
22, 21f Reaction time, 341
342 Reactive functional groups, 3
4 Reactive oxygen species (ROS), 386 Response surface methodology (RSM), 196
198 Rhizopus japonicas, 138
139 Rhizopusoligosporus, 234 Rhizopusoryzae, 437
438, 437t Rhodotorularubra, 353 Rice starch, 258 Rice starch
chitosan biodegradable blend film, 259 Riftiapachyptila, 178 RSM. See Response surface methodology (RSM)

S Saccharomyces cerevisiae, 229 Salmonella choleraesuis, 115 SBET. See Specific surface area (SBET) Scaffolds, 356 cross-linked, 262f essential properties of, 261 for tissue engineering, 410
411 Scale-up, 239 Scanning electron microscopy (SEM), 45, 207
208, 300f of chitosan, 20
22 chitosan-based aerogels, 296
302, 298f, 299f ofelectrospun chitosan-based nanofibers, 213f Seafood processing industry, 249
250, 338, 361
362, 429 Seafood products safety, electrospun nanomaterials for, 212
216 SEC. See Size exclusion chromatography (SEC) Self-assembly method, 180
181, 181f SEM. See Scanning electron microscopy (SEM) Semipermeable chitosan films, 359
360 Serratiamarcescens, 235 Serratiamarcescens FS-3, 407 Shochu distillery wastewater, 441
442

Shrimp shell waste, 21f, 342
343 Shrimp waste powder, 233, 235 Silica aerogel, 297 Silver nanoparticles (AgNP), 207
208, 304 in situ formation of, 208 Single-component aerogels, 288 Size exclusion chromatography (SEC), 73
75 Skin models, 408 SmF. See Submerged fermentation (SmF) Smoluchowski’s theory, 211
212 Sodium borohydride, 136 Sodium cyanoborohydride (NaCNBH3), 66 Sodium dodecylsulfate, 70 Sodium hydroxide, 380, 383 aqueous solution, 121 Sodium tripolyphosphate (TPP) ionic gelation technique, 88 Sol
gel method, 293
295, 301
302, 410
411 Sol
gel transition, 88 Solid 13C nuclear magnetic resonance spectroscopy, 110
111, 111f Solid-state method, 456
457 Solid substrate fermentation (SSF), 229
230, 233
234, 443 advantages and disadvantages of, 444t Solubility, 379
382 of chitin, 39
41, 102
107, 106t CaCl2/methanol solvent, 103 ionic liquid, 104
107, 107f lithium chloride, 104 modification, 381
382 NaOH/urea aqueous solution, 102
103 N,N-dimethylacetamide, 104 ofchitoPEG copolymer, 83f of chitosan, 12, 39
41, 63, 145
153, 253
254 applications and requirements, 145
146 in dilute acids, 253
254 modification method, 147
153, 381
382 in solution, 146
147 in ionic liquids, 380
381 of PEG-g-chitosan, 82
83 of PEGylated chitosans, 67
68 Solution blending, 257 Solution parameters, electrospinning process, 44 Solvent system, 104, 200
202

Index

Sonication, 188 Soy proteins, 261
262 Specific surface area (SBET), 304
305 Spectroscopic method, 72 Spherical nanogel, 177 Spinning technique, 41 SSF. See Solid substrate fermentation (SSF) Standard deacetylation method, 345 Staphylococcus aureus, 26
27, 46, 113, 214
215, 387
388, 461 Starch, 258
259 Stratum corneum, 108 Structural analysis, 71
76 differential scanning calorimetry, 75 Fourier-transform infrared spectroscopy, 71
72 gel permeation chromatography, 73
75 nuclear magnetic resonance, 72
73 size exclusion chromatography, 73
75 thermogravimetric analysis, 75 wide-angle X-ray diffraction analysis, 76 Structural modeling, 319 Structural properties, 403 Submerged fermentation (SmF), 229
230, 233
234 Succinyl chitosan, 149
151 Sugar-bound chitosans, 410
411 Sunflower oil, emulsification of, 409
410 Supercritical drying, 292
293 Superhydrophobic and superoleophilic chitosan sponge, 315 Surface modification of chitin, 167
168, 185
189 acetylation of chitin, 186 of chitin nanowhiskers, 186 hydrophilization of, 188 hydrophobization of, 187
188 oxidative modification, 186
187 physical modification of, 188 plasma treatment, 188
189 ultrasound-assisted, 188 Surface-modifying systems (SMS), 361 Sustainable environment, 248f Swelling, 274 Swelling ratio, 89 Synthesis of chitin, 339
344 by biological methods, 342
344 by chemical method, 340
342 production, 227
228 of chitosan, 339, 344
346

491

Synthetic polymers, 25, 28, 62, 258, 261
262, 271, 274 chitosan blends with, 264
274

T T-CS-OCA, 323
324 TEM. See Transmission electron microscopy (TEM) TEMPO-laccase redox system, 41 TEMPO-mediated oxidation method, 173, 183, 187 Tenebrio molitor, 230
231 Tensile property of aerogels/mechanical property, 314
317 Tensile strength, 142
145 Teredinobacterturnirae, 343
344 2,2,6,6-Tetramethylpiperidine-1-oxyl radical, 183, 187 Textile industry, 26
27, 361, 409 chitin and chitosan fibers, 49 TGA. See Thermogravimetric analysis (TGA) Thermal decomposition of chitosan, 210
211 Thermal insulators, 314
315, 326
327 Thermal properties, 309
312 differential scanning calorimetry, 310
312 thermogravimetric analysis, 309
310 Thermogravimetric analysis (TGA), 75, 210
211, 309
310 Thermoplastic elastomers, 263 Thermoresponsive PEGylated chitosan hydrogels, 88
90 Thymol-loaded chitosan-based nanofibers, 214
215 Time-consuming process, 236 Tip-to-collector distance, 203
204 Tissue engineering (TE), 46, 355
356, 410
411 applications, 261 Titration methods, 141 TMC. See Trimethyl chitosan (TMC) TNBS. See 2,4,6-Trinitrobenzenesulfonic acid solution (TNBS) Transmission electron microscopy (TEM), 91, 207
209, 296, 304, 328 aerogels, 302
304 Tributylborane (TBB), 115
116 Trichloroacetic acid (TCA), 380 Trichlorotriazine, 65 Triethylamine, 69

492

Index

Trifluoroacetic acid (TFA), 200
201 Trifluoroacetic anhydride system, 122
123 Trifluoromethane sulfonyl azide, 69 Trimethyl chitosan (TMC), 83
85 2,4,6-Trinitrobenzenesulfonic acid solution (TNBS), 77
78 6-Triphenylmethylchitosan, 68 Triton X-100, 92
93 Two-component system, 81 Two-dimensional soft nanomaterials, 177
178 Two-step SmF process, 234

U UDP-GlcNAc. See Uridine-diphosphate-Nacetylglucosamine (UDP-GlcNAc) UDP-N-acetylglucosamine, 227 Ultrafine fibers, 92
93 Ultrafine nanofibers, 207
208 Ultrasonication, 182
183 Ultrasonic shock waves, 183 U87MG cells, 87
88 Uridine-diphosphate-N-acetylglucosamine (UDP-GlcNAc), 339 US Food and Drug Administration (FDA), 64 UV radiation, 88
89

V Vancomycin, 326, 327f Versatile physicochemical properties, 425
426 Vibrio cholera, 430
431 4-Vinylpyridine (4-VP), 188 Viscometry, 9
10 Viscosity, 252
253 chitosan, 10
11 Vitamin-loaded chitosan nanoparticles, 210

W Wastewater treatment, 24
25, 47
48 chitin, 47
48 chitosan applications of, 255 demand for, 231 fibers for, 47
48

Water-binding and fat-binding capacity, 13 Water contact angle, 323
324, 324f Water engineering, 357
358, 408
409 Water-insoluble polymer, 249 Water pollution, 357 Water-soluble chitin, 186, 411
412 Water-soluble chitosan, 41, 145
153, 150f, 381
382, 387 ethylamine hydroxyethyl chitosan, 151
152 formation of, 151f hydroxyethyl chitosan, 152
153 O-carboxymethyl chitosan, 152
153 Water-soluble scaffold, 113 Wet gels, 287, 292
293, 297f Wet spinning method, 41
42, 47
48 Whole composite-making process, 176 Wide-angle X-ray diffraction analysis, 76 Wound dressing/wound healing, 354
355, 411
412 Wound infections, 24

X XPS. See X-ray photoelectron spectroscopy (XPS) X-ray diffraction (XRD), 16, 17f, 45 aerogels techniques, 319, 320f analysis, 81 wide-angle, 76 X-ray photoelectron spectroscopy (XPS), 45, 319
322, 321f, 328 X-ray powder diffraction (XRD) spectroscopy, 103, 110 ofα 2 chitin, 110f XRD. See X-ray diffraction (XRD)

Y Yeast, 252
253, 429
430, 433

Z Zeta potential (ZP), 211
212 Zygomycetes, 229
230, 250, 445
446 species, 431
432 Zygorhynchusmoelleri, 446