Advances in Textile Biotechnology (Woodhead Publishing Series in Textiles)

Advances in textile biotechnology © Woodhead Publishing Limited, 2010 The Textile Institute and Woodhead Publishing ...

Author: Vincent Nierstrasz | Artur Cavaco-Paulo

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Advances in textile biotechnology

© Woodhead Publishing Limited, 2010

The Textile Institute and Woodhead Publishing The Textile Institute is a unique organisation in textiles, clothing and footwear. Incorporated in England by a Royal Charter granted in 1925, the Institute has individual and corporate members in over 90 countries. The aim of the Institute is to facilitate learning, recognise achievement, reward excellence and disseminate information within the global textiles, clothing and footwear industries. Historically, The Textile Institute has published books of interest to its members and the textile industry. To maintain this policy, the Institute has entered into partnership with Woodhead Publishing Limited to ensure that Institute members and the textile industry continue to have access to high calibre titles on textile science and technology. Most Woodhead titles on textiles are now published in collaboration with The Textile Institute. Through this arrangement, the Institute provides an Editorial Board which advises Woodhead on appropriate titles for future publication and suggests possible editors and authors for these books. Each book published under this arrangement carries the Institute’s logo. Woodhead books published in collaboration with The Textile Institute are offered to Textile Institute members at a substantial discount. These books, together with those published by The Textile Institute that are still in print, are offered on the Woodhead web site at: www.woodheadpublishing.com. Textile Institute books still in print are also available directly from the Institute’s website at: www. textileinstitutebooks.com. A list of Woodhead books on textile science and technology, most of which have been published in collaboration with The Textile Institute, can be found towards the end of the contents pages.


Woodhead Publishing Series in Textiles: Number 107

Advances in textile biotechnology Edited by V. A. Nierstrasz and A. Cavaco-Paulo

Oxford

Cambridge

Philadelphia

New Delhi


Published by Woodhead Publishing Limited in association with The Textile Institute Woodhead Publishing Limited, Abington Hall, Granta Park, Great Abington Cambridge CB21 6AH, UK www.woodheadpublishing.com Woodhead Publishing, 525 South 4th Street #241, Philadelphia, PA 19147, USA Woodhead Publishing India Private Limited, G-2, Vardaan House, 7/28 Ansari Road, Daryaganj, New Delhi – 110002, India www.woodheadpublishingindia.com First published 2010, Woodhead Publishing Limited © Woodhead Publishing Limited, 2010 The authors have asserted their moral rights. This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. Reasonable efforts have been made to publish reliable data and information, but the authors and the publisher cannot assume responsibility for the validity of all materials. Neither the authors nor the publisher, nor anyone else associated with this publication, shall be liable for any loss, damage or liability directly or indirectly caused or alleged to be caused by this book. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming and recording, or by any information storage or retrieval system, without permission in writing from Woodhead Publishing Limited. The consent of Woodhead Publishing Limited does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from Woodhead Publishing Limited for such copying. Trademark notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library. ISBN 978-1-84569-625-2 (print) ISBN 978-0-85709-023-2 (online) ISSN 2042-0803 Woodhead Publishing Series in Textiles (print) ISSN 2042-0811 Woodhead Publishing Series in Textiles (online) The publisher’s policy is to use permanent paper from mills that operate a sustainable forestry policy, and which has been manufactured from pulp which is processed using acid-free and elemental chlorine-free practices. Furthermore, the publisher ensures that the text paper and cover board used have met acceptable environmental accreditation standards. Typeset by Toppan Best-set Premedia Limited, Hong Kong Printed by TJI Digital, Padstow, Cornwall, UK


Contents

Contributor contact details Woodhead Publishing Series in Textiles Preface

Part I

Technologies involved in textile biotechnology

1

Design and engineering of novel enzymes for textile applications R. Araújo, M. Casal and A. Cavaco-Paulo, University of Minho, Portugal

1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8 2

2.1 2.2 2.3 2.4 2.5

Basic principles of recombinant deoxyribonucleic acid (DNA) molecular cloning Production of enzymes: searching for efficient production systems Enzyme engineering Design and engineering of novel enzymes for textile applications Advantages and limitations Future trends Sources of further information and advice References Developments in processes and equipment for enzymatic textile treatments M. M. C. G. Warmoeskerken and G. H. Bouwhuis, University of Twente, The Netherlands Introduction Current processes for the pre-treatment of cotton Mass transfer limitations in textiles The squeezing model A mass transfer model

xi xv xxi

1

3

3 7 8 10 19 21 22 22

32

32 33 36 39 43 v


vi

Contents

2.6 2.7 2.8 2.9 2.10

Adsorption limitation in textile pores The application of ultrasound Future trends Acknowledgements References

3

Smart textiles and biomaterials containing enzymes or enzyme substrates E. Wehrschütz-Sigl, A. Hasmann and G. M. Guebitz, Graz University of Technology, Austria

3.1 3.2 3.3 3.4 3.5 3.6

Introduction Smart materials containing enzymes Smart materials responding to enzymes as triggers Conclusion and future trends Acknowledgements References

Part II Modification of particular fibres through the use of biotechnology 4

4.1 4.2

4.3 4.4 4.5 4.6 4.7 5

5.1 5.2

Enzymatic hydrolysis and modification of core polymer fibres for textile and other applications C. Silva and A. Cavaco-Paulo, University of Minho, Portugal; and V. A. Nierstrasz, Ghent University, Belgium Introduction Enzymatic hydrolysis and modification of poly(ethylene terephthalate) and polyamide: basic principles, methods and technologies Applications Advantages and limitations Conclusion and future trends Acknowledgements References Enzymatic modification of polyacrylonitrile and cellulose acetate fibres for textile and other applications T. Matamá and A. Cavaco-Paulo, University of Minho, Portugal Introduction Basic concepts and principles


47 52 53 54 54

56

56 58 63 67 67 68

75

77

77

80 90 90 91 91 92

98

98 100

Contents 5.3 5.4 5.5 5.6 5.7 5.8 6

From theory to practice: current methodologies for the assessment of fibre biomodification Polyacrylonitrile biomodification Cellulose acetate biomodification Future trends Sources of further information and advice References

vii

106 109 120 124 126 126

Enzymatic treatment versus conventional chemical processing of cotton I. R. Hardin, The University of Georgia, USA

132

6.1 6.2 6.3 6.4 6.5 6.6 6.7

Introduction Chemistry and structure of the cotton fiber Cotton seed coat fragments Conventional chemical treatments and conditions Enzymatic treatments of cotton Future trends References

132 133 135 138 140 144 145

7

Engineering of plants for improved fibre qualities M. Wróbel-Kwiatkowska, Medical University in Wrocław, Poland; J. Szopa, University of Wrocław, Poland; and S. Hawkins, Université de Lille, France

150

7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.8

Introduction Defining plant fibres Plant engineering methods Some examples of engineering in fibre species Future trends Conclusions Acknowledgements References

150 151 155 156 161 163 163 163

8

Enzymatic treatment of wool and silk fibres J. Shen, De Montfort University, UK

171

8.1 8.2 8.3 8.4 8.5 8.6 8.7

Introduction Enzymes used for processing of protein fibres Application of enzymatic treatments to wool fibres Application of enzymatic treatments to silk fibres Future trends Acknowledgements References

171 172 174 182 186 187 187


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9

Functionalisation of wool and silk fibres using nonproteolytic enzymes G. Freddi, Stazione Sperimentale per la Seta, Italy

193

9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8

Introduction Transglutaminases Functionalisation of protein fibres using transglutaminases Tyrosinases Functionalisation of protein fibres using tyrosinases Other enzymes for protein fibre functionalisation Conclusions and future trends References

193 195 201 205 213 218 223 226

10

Developments in recombinant silk and other elastic protein fibers for textile and other applications J. C. Rodríguez-Cabello, C. García-Arévalo, L. Martín, M. Santos and V. Reboto, University of Valladolid, Spain

10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 11

11.1 11.2 11.3 11.4

Introduction Principles of recombinant DNA methods applied in the production of recombinant proteins Biomimetic design of recombinant proteins Expression systems Purification of recombinant proteins Experimental characterization of recombinant proteins Examples and applications of recombinant protein fibers (silk, elastin, collagen, resilin) Advances and limitations in the biosynthetic production of recombinant proteins Future trends Acknowledgments References Enzymatic functionalization of cellulosic fibres for textile and other applications: xyloglucan as a molecular anchor H. Brumer, Royal Institute of Technology (KTH), Sweden Introduction Xyloglucans: a family of functional plant polysaccharides Technical uses of native xyloglucans in cellulose modification Chemo-enzymatic modification of xyloglucans to functionalize cellulose surfaces


235

235 236 237 239 244 245 247 258 259 260 260

266 266 268 270 272

Contents

ix

11.5 11.6 11.7 11.8

Conclusion and future trends Sources of further information and advice Acknowledgements References

281 281 282 282

12

Developments in the processing of chitin, chitosan and bacterial cellulose for textile and other applications A. Francesko, M. Díaz González, G. R. Lozano and T. Tzanov, Universitat Politécnica de Catalunya, Spain

288

12.1 12.2 12.3 12.4 12.5 12.6 12.7 12.8 13

13.1 13.2 13.3 13.4 13.5 13.6 13.7 13.8 13.9

Introduction Types of new fibres: chitin, chitosan and bacterial cellulose Basic principles, methods and technologies Applications Advantages and limitations Future trends Sources of further information and advice References Hydrolysis of regenerated cellulose fibres for textile and other applications T. Bechtold and C. B. Schimper, Leopold-Franzens University Innsbruck, Austria

288 289 293 300 303 304 305 306

312

Introduction Regenerated cellulose fibres Cellulases Cellulase hydrolysis of regenerated cellulose fibres Restructuring by heat and steam treatment Treatment in alkaline swelling solutions Technical processing Conclusion References

312 314 315 316 319 321 324 324 325

Index

328


Contributor contact details

(* = main contact)

Editors V. A. Nierstrasz* Department of Textiles Ghent University Technologiepark 907 9052 Gent/Zwijnaarde Belgium E-mail: [email protected] A. Cavaco-Paulo Department of Textile Engineering University of Minho 4800-058 Guimarães Portugal E-mail: [email protected]

M. Casal CBMA-Centre of Molecular and Environmental Biology Department of Biology University of Minho 4710-057 Braga Portugal

Chapter 2 M. M. C. G. Warmoeskerken* and G. H. Bouwhuis Faculty of Engineering Technology (CTW) Postbus 217 7500 AE Enschede University of Twente The Netherlands E-mail: m.m.c.g.warmoeskerken@ utwente.nl

Chapter 1 R. Araújo and A. Cavaco-Paulo* Department of Textile Engineering University of Minho 4800-058 Guimarães Portugal E-mail: [email protected]; [email protected]

Chapter 3 E. Wehrschütz-Sigl, A. Hasmann and G. M. Guebitz* Department of Environmental Biotechnology Graz University of Technology Petersgasse 12 8010 Graz Austria E-mail: [email protected] xi


xii


Chapter 4

Chapter 7

C. Silva and A. Cavaco-Paulo Department of Textile Engineering University of Minho 4800-058 Guimarães Portugal

M. Wróbel-Kwiatkowska* Department of Pharmaceutical Biology and Botany Medical University in Wrocław Al. J. Kochanowskiego 10 51-601 Wrocław Poland

E-mail: [email protected]; [email protected] V. A. Nierstrasz* Department of Textiles Ghent University Technologiepark 907 9052 Gent/Zwijnaarde Belgium E-mail: [email protected]

E-mail: magdalena.wrobel@ibmb. uni.wroc.pl J. Szopa Faculty of Biotechnology University of Wrocław Przybyszewskiego 63/77 51-148 Wrocław Poland

Chapter 5

E-mail: [email protected]

T. Matamá* and A. Cavaco-Paulo Department of Textile Engineering University of Minho 4800-058 Guimarães Portugal

S. Hawkins UMR USTL INRA 1281 Stress abiotiques et différenciation des végétaux cultivés Université de Lille 1 Bâtiment SN2 F-59650 Villeneuve 8217 Ascq cedex France

E-mail: [email protected]; [email protected]

Chapter 6 Ian R. Hardin Georgia Power Professor of Textile Science The University of Georgia Athens GA 30602-3622 USA E-mail: [email protected]


Chapter 8 Jinsong Shen Textile Engineering and Materials Research Group De Montfort University The Gateway Leicester LE1 9BH UK E-mail: [email protected]



xiii

Chapter 9

Chapter 12

G. Freddi Stazione Sperimentale per la Seta Via Giuseppe Colombo 83 20133 Milano Italy

A. Francesko, M. Díaz González, G. R. Lozano and T. Tzanov* Department of Chemical Engineering Universitat Politécnica de Catalunya Escola Universitária d’Enginyeria Tècnica Industrial Colom 1 08222 Terrassa Spain


Chapter 10 J. C. Rodríguez-Cabello*, C. García-Arévalo, L. Martín, M. Santos and V. Reboto GIR BIOFORGE (Group for Advanced Materials and Nanobiotechnology) University of Valladolid (I+D) Paseo de Belén 11 47011 Valladolid Spain E-mail: [email protected]

Chapter 11 H. Brumer Division of Glycoscience School of Biotechnology Royal Institute of Technology (KTH) AlbaNova University Centre 106 91 Stockholm Sweden


Chapter 13 T. Bechtold* and C. B. Schimper Research Institute for Textile Chemistry and Textile Physics Leopold-Franzens University Innsbruck Hoechsterstrasse 73 6850 Dornbirn Austria E-mail: [email protected]



Woodhead Publishing Series in Textiles

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

Watson’s textile design and colour Seventh edition Edited by Z. Grosicki Watson’s advanced textile design Edited by Z. Grosicki Weaving Second edition P. R. Lord and M. H. Mohamed Handbook of textile fibres Vol 1: Natural fibres J. Gordon Cook Handbook of textile fibres Vol 2: Man-made fibres J. Gordon Cook Recycling textile and plastic waste Edited by A. R. Horrocks New fibers Second edition T. Hongu and G. O. Phillips Atlas of fibre fracture and damage to textiles Second edition J. W. S. Hearle, B. Lomas and W. D. Cooke Ecotextile ’98 Edited by A. R. Horrocks Physical testing of textiles B. P. Saville Geometric symmetry in patterns and tilings C. E. Horne Handbook of technical textiles Edited by A. R. Horrocks and S. C. Anand Textiles in automotive engineering W. Fung and J. M. Hardcastle Handbook of textile design J. Wilson High-performance fibres Edited by J. W. S. Hearle Knitting technology Third edition D. J. Spencer

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xvi 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38

Woodhead Publishing Series in Textiles Medical textiles Edited by S. C. Anand Regenerated cellulose fibres Edited by C. Woodings Silk, mohair, cashmere and other luxury fibres Edited by R. R. Franck Smart fibres, fabrics and clothing Edited by X. M. Tao Yarn texturing technology J. W. S. Hearle, L. Hollick and D. K. Wilson Encyclopedia of textile finishing H-K. Rouette Coated and laminated textiles W. Fung Fancy yarns R. H. Gong and R. M. Wright Wool: Science and technology Edited by W. S. Simpson and G. Crawshaw Dictionary of textile finishing H-K. Rouette Environmental impact of textiles K. Slater Handbook of yarn production P. R. Lord Textile processing with enzymes Edited by A. Cavaco-Paulo and G. Gübitz The China and Hong Kong denim industry Y. Li, L. Yao and K. W. Yeung The World Trade Organization and international denim trading Y. Li, Y. Shen, L. Yao and E. Newton Chemical finishing of textiles W. D. Schindler and P. J. Hauser Clothing appearance and fit J. Fan, W. Yu and L. Hunter Handbook of fibre rope technology H. A. McKenna, J. W. S. Hearle and N. O’Hear Structure and mechanics of woven fabrics J. Hu Synthetic fibres: nylon, polyester, acrylic, polyolefin Edited by J. E. McIntyre Woollen and worsted woven fabric design E. G. Gilligan Analytical electrochemistry in textiles P. Westbroek, G. Priniotakis and P. Kiekens


Woodhead Publishing Series in Textiles 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Bast and other plant fibres R. R. Franck Chemical testing of textiles Edited by Q. Fan Design and manufacture of textile composites Edited by A. C. Long Effect of mechanical and physical properties on fabric hand Edited by Hassan M. Behery New millennium fibers T. Hongu, M. Takigami and G. O. Phillips Textiles for protection Edited by R. A. Scott Textiles in sport Edited by R. Shishoo Wearable electronics and photonics Edited by X. M. Tao Biodegradable and sustainable fibres Edited by R. S. Blackburn Medical textiles and biomaterials for healthcare Edited by S. C. Anand, M. Miraftab, S. Rajendran and J. F. Kennedy Total colour management in textiles Edited by J. Xin Recycling in textiles Edited by Y. Wang Clothing biosensory engineering Y. Li and A. S. W. Wong Biomechanical engineering of textiles and clothing Edited by Y. Li and D. X-Q. Dai Digital printing of textiles Edited by H. Ujiie Intelligent textiles and clothing Edited by H. R. Mattila Innovation and technology of women’s intimate apparel W. Yu, J. Fan, S. C. Harlock and S. P. Ng Thermal and moisture transport in fibrous materials Edited by N. Pan and P. Gibson Geosynthetics in civil engineering Edited by R. W. Sarsby Handbook of nonwovens Edited by S. Russell Cotton: Science and technology Edited by S. Gordon and Y-L. Hsieh Ecotextiles Edited by M. Miraftab and A. R. Horrocks


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xviii 61 62 63 64 65 66 67 68 69 70 71 72 73 74

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Woodhead Publishing Series in Textiles Composite forming technologies Edited by A. C. Long Plasma technology for textiles Edited by R. Shishoo Smart textiles for medicine and healthcare Edited by L. Van Langenhove Sizing in clothing Edited by S. Ashdown Shape memory polymers and textiles J. Hu Environmental aspects of textile dyeing Edited by R. Christie Nanofibers and nanotechnology in textiles Edited by P. Brown and K. Stevens Physical properties of textile fibres Fourth edition W. E. Morton and J. W. S. Hearle Advances in apparel production Edited by C. Fairhurst Advances in fire retardant materials Edited by A. R. Horrocks and D. Price Polyesters and polyamides Edited by B. L. Deopura, R. Alagirusamy, M. Joshi and B. S. Gupta Advances in wool technology Edited by N. A. G. Johnson and I. Russell Military textiles Edited by E. Wilusz 3D fibrous assemblies: Properties, applications and modelling of three-dimensional textile structures J. Hu Medical and healthcare textiles Edited by S. C. Anand, J. F. Kennedy, M. Miraftab and S. Rajendran Fabric testing Edited by J. Hu Biologically inspired textiles Edited by A. Abbott and M. Ellison Friction in textile materials Edited by B .S. Gupta Textile advances in the automotive industry Edited by R. Shishoo Structure and mechanics of textile fibre assemblies Edited by P. Schwartz Engineering textiles: Integrating the design and manufacture of textile products Edited by Y.E. El-Mogahzy


Woodhead Publishing Series in Textiles 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103

Polyolefin fibres: industrial and medical applications Edited by S. C. O. Ugbolue Smart clothes and wearable technology Edited by J. McCann and D. Bryson Identification of textile fibres Edited by M. Houck Advanced textiles for wound care Edited by S. Rajendran Fatigue failure of textile fibres Edited by M. Miraftab Advances in carpet technology Edited by K. Goswami Handbook of textile fibre structure Volume 1 and Volume 2 Edited by S. J. Eichhorn, J. W. S. Hearle, M. Jaffe and T. Kikutani Advances in knitting technology Edited by K-F. Au Smart textile coatings and laminates Edited by W. C. Smith Handbook of tensile properties of textile and technical fibres Edited by A. R. Bunsell Interior textiles: Design and developments Edited by T. Rowe Textiles for cold weather apparel Edited by J. T. Williams Modelling and predicting textile behaviour Edited by X. Chen Textiles, polymers and composites for buildings Edited by G. Pohl Engineering apparel fabrics and garments J. Fan and L. Hunter Surface modification of textiles Edited by Q. Wei Sustainable textiles Edited by R. S. Blackburn Advances in yarn spinning technology Edited by C. A. Lawrence Handbook of medical textiles Edited by V. T. Bartels Technical textile yarns Edited by R. Alagirusamy and A. Das Applications of nonwovens in technical textiles Edited by R. A. Chapman Colour measurement: Principles, advances and industrial applications Edited by M. L. Gulrajani


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Woodhead Publishing Series in Textiles Textiles for civil engineering Edited by R. Fangueiro New product development in textiles Edited by B. Mills Improving comfort in clothing Edited by G. Song Advances in textile biotechnology Edited by V. A. Nierstrasz and A. Cavaco-Paulo Textiles for hygiene and infection control Edited by B. McCarthy Nanofunctional textiles Edited by Y. Li Joining textiles: principles and applications Edited by I. Jones and G. Stylios Soft computing in textile engineering Edited by A. Majumdar Textile design Edited by A. Briggs-Goode and K. Townsend. Biotextiles as medical implants Edited by M. King and B. Gupta Textile thermal bioengineering Edited by Y. Li Woven textile structure B. K. Behera and P. K. Hari Handbook of textile and industrial dyeing. Volume 1: principles processes and types of dyes Edited by M. Clark Handbook of textile and industrial dyeing. Volume 2: Applications of dyes Edited by M. Clark Handbook of natural fibres. Volume 1: Types, properties and factors affecting breeding and cultivation Edited by R. Kozlowski Handbook of natural fibres. Volume 2: Processing and applications Edited by R. Kozlowski Functional textiles for improved performance, protection and health Edited by N. Pan and G. Sun Computer technology for textiles and apparel Edited by Jinlian Hu Advances in military textiles and personal equipment Edited by E. Sparks Specialist yarn, woven and fabric structure: Developments and applications Edited by R. H. Gong


Preface

This book is a follow-up to the 2003 publication Textile processing with enzymes. Whereas the first publication was based on enzyme processes in textiles and aimed to be a comprehensive guide for students, newcomers and professionals interested in the area, this book aims to give an updated vision of biotechnology in textiles. The impact of biotechnology in the production of textiles is increasingly significant. Enzyme technology has already proven to be advantageous in industrial textiles processing of natural fibres. The Novozymes Report 2009 discloses that the major share of the industrial enzyme market is divided between Novozymes (47%), Danisco Genencor (21%), and DSM (6%), with the remaining 26% shared between other minor players. Sales of industrial enzymes have increased by 32% since 2005 and the worldwide value of the industrial enzyme markets in 2009 was estimated to be about 12000 million. Detergents are the largest application of industrial enzymes representing more than 32% of the market size. Food represents 21% of the market, feed 8%, and technical (including textiles, pulp and paper) and others 31%. The exact share of textiles is not given in the report, but is estimated to be about 10% of industrial enzymes. Protease is the main enzyme class employed in detergents along with amylase added for improved removal of proteinaceous and starchy stains. Cellulases are included in some detergents for improved brightness as well as for soil removal. The most recent development is the addition of mannanase in detergent formulations to remove stains based on guar gum. A future development could be the addition of redox enzymes to reduce interfabric staining. Most of the enzymes used in detergents have been tailored to meet requirements on operational and storage stability. Enhanced resistance to high ionic forces and/or bleaching agents is a common trend in detergent enzymes; in addition, new enzymes exist that act at low temperatures for washing with cold water. The major enzymes in textile applications include amylases (desizing), catalases (bleach removal/termination) and cellulases (pilling removal, xxi © Woodhead Publishing Limited, 2010

xxii

Preface

softening and ageing effects). Other less significant enzymes used in textile applications include laccases (bleaching and over-dyeing effects), pectate lyases (cotton scouring), proteases (degumming of silk, sand washing of silk, wool finishing) and enzymes for retting of flax (pectinases, xylanases, hemicellulases). Advances in textile biotechnology provides an overview of the state of the art and recent developments in biotechnology for modification, functionalisation and processing of textile fibres and polymers (natural, synthetic, or man-made). Advances in textile biotechnology consists of two parts: Part I addresses Technologies involved in textile biotechnology, and Part II focuses on Modification of particular fibres through the use of biotechnology. In Part I, detailed views are provided on methods, processes and equipment where enzymes can be used in textiles. A range of examples are given using rational and random mutation techniques as well as fusion protein approaches for designing engineered enzymes with better properties for textile end uses. Smart wound dressings used as sensors are described for identification of high enzyme activities normally associated with infected open wounds. In Part II, two comprehensive reviews and updates on the state of the art in enzyme modification and functionalisation of synthetic fibres, namely polyester, polyamide, acrylics and cellulose acetate materials, are presented. Advantages and limitations are addressed. The state of the art is reviewed for enzyme modification of wool, silk and cotton fibrous materials as well as biomimetic approaches to functionalise cellulosic materials. Genetically modified protein and materials based on plant polymers complement the range of biotechnologies. The production of polyhydroxyalkanoate (PHA) biopolymers including polyhydroxybutyrate (PHB) and protein polymers completes the review in this area. It is believed that this book will promote biotechnology for the modification, functionalisation and processing of textile fibres in various applications, and increase understanding and acceptance of biotechnology in industry and society. V. A. Nierstrasz A. Cavaco-Paulo


1 Design and engineering of novel enzymes for textile applications R. A R AÚJO, M. C A SA L and A. C AVAC O - PAU L O, University of Minho, Portugal

Abstract: The principles of recombinant DNA molecular cloning and transformation of host cells are outlined, and applications for protein engineering in the textile industry are described. High production yields of enzymes can be achieved at competitive costs and enzymes can be redesigned with novel properties adapted to suit industrial conditions. Protein engineering techniques, such as site-directed mutagenesis and directed evolution, are described in detail. Despite their complexity and disadvantages, these techniques for enzyme design have been successfully applied at the industrial level and examples of applications are discussed in this chapter. Key words: enzyme engineering, molecular genetics, site-directed mutagenesis, directed evolution.

1.1

Basic principles of recombinant deoxyribonucleic acid (DNA) molecular cloning

The manipulation of genetic information involves the identification and purification of a particular coding sequence of interest, i.e. DNA or ribonucleic acid (RNA), which can be isolated from a biological specimen or obtained by chemical synthesis. After the target nucleic acid molecule is obtained in purified form, the further steps involve the construction of a recombinant DNA molecule by covalent ligation to a well characterized replicative system and further cloning and amplification into a desirable cell host. The construction of a recombinant DNA molecule makes use of restriction endonucleases to create a linear double strand of DNA fragments of both the target gene and the vector, in such way that they can recombine in vitro becoming covalently linked. The enzyme ligase promotes the phosphodiester ligation of the ends of any two DNA fragments that display blunt ends or complementary sticky ends. The resulting recombinant DNA molecule is then transferred into a host cell in a process called transformation. 3 © Woodhead Publishing Limited, 2010

4


1.1.1 Obtaining, cutting and joining the DNA Because a DNA cloning strategy initially requires identification of the genetic sequence coding for a specific protein, the first step is to consider the source of such a DNA molecule, which represents a specific gene or portion of a gene. Typical fragments of interest include genomic DNA, cDNA, polymerase chain reaction (PCR) products and chemically synthesized oligonucleotides. Two major types of enzymes are crucial for the isolation and construction of recombinant DNA molecules: restriction endonucleases and DNA ligases. Restriction endonucleases can be grouped into three classes according to their molecular structure, need for specific co-factors, the types of sequences recognized, and the nature of the cut made in the DNA. We will focus on type II endonucleases, the most commonly used in recombinant DNA technology. These enzymes recognize a specific site on doublestranded DNA, called a restriction site, which comprises a short sequence of nucleotides, and cleave within it without requiring ATP hydrolysis for nucleolytic activity (Marx, 1973; Sambrook and Maniatis, 1989). Type II restriction endonucleases exhibit a high degree of sequence specificity and a single base pair change in the recognition site of a restriction endonuclease essentially hinders and blocks enzymatic activity. In contrast, type I and III restriction endonucleases cause random cleavage patterns in the DNA which are useless for gene cloning (Gorbalenya and Koonin, 1991). So far, around 3000 type II restriction enzymes have been identified and only a small fraction of those has been biochemically characterized. Orthodox type II endonucleases are composed of two identical polypeptide subunits that join together to form a homodimer. These homodimers recognize palindromic DNA sequences of 4–8 bp, although 6 bp DNA sequences are the most recognized by restriction endonucleases used in molecular genetics. Mg2+ ions are required for catalysis. First, the restriction endonuclease contacts with DNA sugar–phosphate backbone in a non-specific way for efficient recognition of the target site location. Once the restriction site is located, the recognition process triggers conformational changes in the enzyme and the DNA, in turn, that leads to the activation of the catalytic center. In contrast to the non-specific contact, specific binding involves direct interaction of the enzyme with DNA nitrogenous bases. Type II restriction enzymes cut the DNA double strand disrupting the covalent, phosphodiester bond between the phosphate of one nucleotide and the sugar of an adjacent nucleotide, producing 3′-hydroxyl and 5′phosphate ends (detailed reviews of target site location, recognition and catalysis mechanism of type II endonucleases have been given by Pingoud and Jeltsch (2001) and Pingoud et al. (2005)).


Design and engineering of novel enzymes for textile applications

5

Some of these enzymes cleave the DNA creating overhangs, i.e. stretches of unpaired nucleotides at the ends of DNA molecules. These unpaired nucleotides can be in either strand, creating either 3′ or 5′ overhangs. The simplest case of an overhang is a single nucleotide. Longer overhangs are called cohesive ends or sticky ends (Fig. 1.1a). Other type II enzymes cut both strands of the DNA at the same position and generate blunt ends i.e. with no unpaired nucleotides when they cleave the DNA (Roberts, 1980; Roberts and Macelis, 1996) (Fig. 1.1b). The DNA molecules of the cloning vector and of the coding sequence of interest must be cut with one or more of these enzymes, creating linear molecules having ends suitable to be correctly joined by the enzyme ligase. The DNA ligase joins two molecules of DNA by catalyzing the formation of covalent phosphodiester bonds between the 5′-phosphate end of one fragment of DNA and the 3′-hydroxyl end of another with the expenses of an ATP molecule (Cozzarelli et al., 1967).

(a)

EcoRI 5′P

PstI 3′OH

GAATTC CTTAAG

3′OH

5′P

5′P

3′OH

5′ protruding ends 5′P 3′OH

3′OH

G CTTAA

5′P 5′P

5′P 3′OH

5′P

5′P

3′ protruding ends 3′OH

AATTC G

5′P

5′P 3′OH

3′OH

CTGCA G 5′P

3′OH 3′OH

5′P

G ACGTC

3′OH 5′P

SmaI

(b)

3′OH

3′OH

CTGCA G G ACGTC

CCC GGG

CCC GGG GGG CCC

3′OH 5′P

5′P 3′OH

3′OH 5′P

GGG CCC

3′OH 5′P

1.1 Cleavage patterns of some common restriction endonucleases. The cleavage sites of EcoRI, PstI and SmaI are shown. (a) EcoRI cuts asymmetrically within the recognition site such that a short singlestranded segment extends from the 5′ ends. PstI also cuts asymmetrically within the recognition site, but the result is a singlestranded overhang from the two 3′ ends. The 5′ or 3′ overhangs generated by enzymes that cut asymmetrically are called sticky ends or cohesive ends. (b) SmaI cuts at precisely opposite sites in the two strands of DNA generating blunt ends.


6


There are several DNA ligases isolated from different sources but the most commonly used in the laboratory is derived from the bacteriophage T4 (Cozzarelli et al., 1967; Weiss et al., 1968; Weiss, 1971). T4 DNA ligase can join, through a covalent ligation, two DNA fragments with both sticky or blunt ends, although in the latter case the reaction is less efficient and requires higher enzyme concentrations and optimized reaction conditions (Murray et al., 1979). Ligation between the target DNA and the vector is a very inefficient reaction because the two cleaved ends of a cloning vector (if compatible) can self-ligate. Self-ligation can be avoided if the vector is treated with a phosphatase, an enzyme that removes both terminal 5′-phosphate groups from the vector. Without it, the vector cannot be recircularized by ligase because the phosphodiester bond does not form. When the vector is joined with an insert, the 5′-phosphate is provided by the insert. Alternatively, if the vector is digested by two different restriction endonucleases noncomplementary sticky ends, which hinder the self-ligation reaction and promote controlled ligation of the DNA insert in the desired orientation within the vector, are produced.

1.1.2 Transformation of host cells The ligation reaction mixture can be introduced into host cells in a process called transformation. Chemical transformation and electroporation are two methods used to transform Escherichia coli cells. Several chemical methods have been established that induce bacterial cell transformation. Treatment of bacterial cells with CaCl2 is still the most widely used cloning method in small laboratories (Cohen et al., 1972; Mandel and Higa, 1970; Oishi and Cosloy, 1972) and over the years its efficiency has been greatly improved (Dagert and Ehrlich, 1974; Chen et al., 2001; Fregel et al., 2008; Huff et al., 1990; Nakata et al., 1997). The permeable ‘competent’ cells are then mixed with DNA to allow the uptake of the exogenous DNA into the bacterial cell. Electroporation can also be used to introduce DNA into cells by subjecting them to a strong electric current, thus producing transient pores in their membranes. Multiplication of the vector DNA, either recombinant or nonrecombinant, occurs within each transformed bacterium. A single bacterial cell cultivated in an appropriate medium, containing the necessary nutrients, can multiply to form a visible colony made of millions of identical cells (clones). As the host cell divides, the vectors, commonly plasmids, are passed on to progeny, where they continue to replicate. Numerous cell divisions of a single transformed bacteria result in a clone of cells, that, when cultivated into an agar plate, produce a visible bacterial colony. The cloned DNA can then be isolated from the bacterial cells.



1.2

7

Production of enzymes: searching for efficient production systems

Various expression hosts (Escherichia coli, Bacillus sp., Saccharomyces cerevisiae, Pichia pastoris, filamentous fungi, insect and mammalian cell lines) have been developed to express heterologous proteins (Chelikani et al., 2006; Huynh and Zieler, 1999; Li et al., 2007; Makrides, 1996; Ogay et al., 2006; Silbersack et al., 2006). Among the many systems available for heterologous protein production, the enteric Gram-negative bacterium E. coli remains one of the most attractive. Compared with other established and emerging expression systems, E. coli offers several advantages including its capacity to grow rapidly and at high density on economical carbon sources, simple scale-up process, its well-characterized genetics and the availability of an increasingly large number of commercial cloning vectors and optimized host strains (Baneyx, 1999). However, depending on protein to be expressed, the use of E. coli is not always suitable so there is no guarantee that a recombinant gene product will accumulate in E. coli at high levels in a full-length and biologically active form (Makrides, 1996). To circumvent such constraints and as an alternative, the genes have to be cloned back into species similar to those from which they were derived, although with an expression cassette driven towards an increasing of production. In these instances, bacteria from the unrelated genera Bacillus (Biedendieck et al., 2007; Silbersack et al., 2006), Clostridium (Girbal et al., 2005) and Staphylococcus, and the lactic acid bacteria Streptococcus (Arnau et al., 2006), Lactococcus (Miyoshi et al., 2002) and Lactobacillus (Miyoshi et al., 2004), can be an alternative. If heterologous produced proteins require complex post-translational modifications and are not expressed in the soluble form using prokaryotic expression systems, yeasts can be an efficient choice because they present several advantages over bacteria for the production of eukaryotic proteins. Among yeast species, the methylotrophic yeast Pichia pastoris is a particularly well suited host for this purpose, offering a number of important benefits: (a) (b) (c) (d)

(e)

high levels of recombinant protein expression are reached under the alcohol oxidase 1 gene (aox 1) promoter; high-density cell cultures can be obtained; scaled-up fermentation methods without loss of yield have been developed; efficient secretion of the recombinant product together with a very low level of endogenous protein secretion represents a very simple and convenient pre-purification step; accurate post-translational modifications are possible (such as proteolytic processing and glycosylation).


8


Furthermore, the existence of efficient methods to integrate several copies of the expression cassette carrying the recombinant DNA into the genome, eliminating the problems associated with expression from plasmids, make this yeast the micro-organism of choice for an increasing number of biotechnologists (Cereghino and Cregg, 2000; Hollenberg and Gellissen, 1997). Once fermentation is completed, the micro-organisms are destroyed, and the enzymes are isolated and further processed for commercial use.

1.3

Enzyme engineering

In broad terms, enzyme engineering refers to the creation of mutant proteins with predetermined enzymatic characteristics. The three main concerns of enzyme optimization as industrial biocatalysts are the volumetric productivity or activity of the enzyme, the stability of the enzyme under process conditions, and the availability or cost of producing the enzyme (Marrs et al., 1999). The activity and stability of enzymes can be modified by chemical modification, immobilization, or by the use of solvents (DeSantis and Jones, 1999; Bull et al., 1999). Recently, enzyme engineering by molecular modeling, site-directed mutagenesis and directed evolution has become remarkably efficient and may generally be applied in combination to develop desired mutants.

1.3.1 Site-directed mutagenesis Site-directed mutagenesis represents a powerful technique that allows selective engineering of gene sequences and has led to rapid advances in the understanding of gene expression and function. In site-directed mutagenesis, nucleotide alterations are introduced into a target sequence by incorporating DNA base changes within an oligonucleotide utilized in the DNA synthesis step (primer). For example, deletions, insertions and point mutations can be easily introduced using suitable primers containing the desired mutations. Several protocols of site-directed mutagenesis based on polymerase chain reaction (PCR) methods have been developed and optimized in order to achieve efficient mutagenesis of a target DNA sequence (Hemsley et al., 1989; Ho et al., 1989; Ito et al., 1991) Among the PCR-based protocols, the ‘megaprimer’ method, first introduced by Kammann and collaborators (1989), is simple and cost-effective. This method involves two rounds of PCR with two ‘flanking’ primers and one internal mutagenic primer containing the desired mutation. The first PCR is performed using the mutagenic internal primer and the first flanking primer. After purification, this first PCR product, the ‘megaprimer’, is used, together with the second flanking primer, for a second PCR round from which the final PCR product,



9

containing the desired mutation, is obtained (Kammann et al., 1989). This method was later modified to increase yield and optimize time consumption (Sarkar and Sommer, 1990; Barik and Gahnski, 1991; Barik, 1995; Ke and Madison, 1997). Later, a new method was described combining both PCR technology and homologous recombination for introducing site-specific alterations in any DNA sequence cloned into a plasmidic expression vector. This approach is based on PCR amplification of the entire plasmid DNA by mutagenic primers divergently oriented but overlapping at their 5′ ends (Martin et al., 1995). This approach was optimized to overcome length limitation, owing to the plasmid size, to increase mutagenesis efficiency and to minimize the rate of undesired mutations (Ansaldi et al., 1996; Rabhi et al., 2004). Several commercial site-directed mutagenesis kits have now been developed. Muta-Gene M13 in vitro Mutagenesis Kit from Bio-Rad and the QuikChange kit from Stratagene guarantees mutation frequencies higher than 50% (Siemers et al., 1996). Enzyme stability and activity can be optimized by site-directed mutagenesis but this technique is dependent on the availability of structural and biochemical information. First, the 3D-structure of the protein of interest must be solved to allow the identification of important amino acids, then, the protein variants are constructed based on predictions derived from the analysis of the 3D-structure and, finally, these variants are biochemically characterized.

1.3.2 Directed evolution Directed evolution involves the application of repeated rounds of random mutagenesis, in vitro recombination, and selection to develop enzymes with improved properties. This strategy mimics natural evolution, as an initial parent gene is chosen and a diverse library of offspring genes is created through mutagenesis or recombination. A screening is applied to the library and the mutants that exhibit the greatest improvement in the desired properties are chosen to become the parents to the next generation (Bloom et al., 2005). Directed evolution is a more general strategy for the isolation of catalysts as it does not necessarily require structural information of the enzyme or its catalytic mechanism (Cherry and Fidantsef, 2003; Farinas et al., 2001; Jaeger et al., 2001; Powell et al., 2001; Tao and Cornish, 2002) and it can be applied to most chemical reactions in aqueous solutions (Kettling et al., 1999). However, as it results in a large number of variants, the success of this approach is dependent on an efficient screening procedure, so that variants with improved and desired properties can be identified. Directed evolution has been used to alter a diverse range of enzyme performance properties including modification of stability and


10


availability of proteases, amylases, laccases, phytases, and cellulases (Cherry and Fidantsef, 2003).

1.4


Enzymes can be obtained from any live organism, but naturally occurring enzymes are often not readily available in sufficient quantities for industrial use. The advances of the last decade in genomics, proteomics and bioinformatics created new opportunities to exploit an extremely large amount of biological data that was not formerly available, and has allowed the exploitation of such resources to create and develop innovative biotechnologybased products. More than 50% of the industrially important enzymes are engineered or produced from genetically engineered micro-organisms (Hodgson, 1994).

1.4.1 Amylases Amylases were the first and are still the most successful enzymes used in textile industry for desizing to facilitate the removal of the starch-containing size that has served as a protective coating on yarns during weaving. In order to reduce processing costs the desizing processing is sometimes combined with the scouring and bleaching steps. In such instances, nonenzymatic auxiliaries such as alkali or oxidation agents are typically used. The desizing process is still the main application of amylases in textile industry but their applications as additives in laundering detergent formulations have increased recently. The requirements for an optimal performance of amylases in both the applications referred to above, mainly concern pH, oxidative stability, chelator resistance and temperature behaviors. So as to develop modified amylases with improved performance, various strategies are used, amongst which protein engineering methods, such as random mutagenesis, homology considerations and site-directed mutagenesis are widely used. Declerck et al. (1990) and Joyet et al. (1992) increased the thermostability of Bacillus licheniformis α-amylase (BLA) by two independent aminoacid substitutions His133Tyr (or His133Ile) and Ala209Val (or Ala209Ile) using random mutagenesis. This increase in enzyme thermostability is presumably the result of entropy gain and better packing of the protein structure. Two regions that are important for the higher thermostability of BLA in comparison to Bacillus amyloliquefaciens α-amylase (BAA) were first identifed by Suzuki and collaborators (1989). Based on amino acid differences observed in these two regions, three stabilizing mutations in the BAA



11

sequence were designed: the substitution Lys269Ala and the deletion of Arg176 and Gly177. Both mutations have drastically increased thermostability of BAA. The Arg176-Gly177 deletion was further applied to LAMY α-amylase derived from alkaliphilic Bacillus species with similar effects on the thermostability (Igarashi et al., 1998). The same group also reported for the first time the improvement of LAMY stability by proline substitution using site-directed mutagenesis (Igarashi, 1999). One of the most important criteria for the use of amylases in detergents is to maintain optimal activity under the very oxidizing washing environment (Ito et al., 1998). Oxidative stability of α-amylases was achieved by site-directed mutagenesis, replacing oxidation-sensitive aminoacids such as cysteines and methionines (Brosnan et al., 1992) with non-oxidizable residues as shown in the case of BLA (Brzozowski et al., 2000) and AmyK, α-amylase from an alkaliphilic Bacillus strain KSM-1378 (Igarashi et al., 2003). The commercial tradenames of the two currently available oxidatively stable α-amylases are Purastar OxAm® from Genencor Intl and Duramyl® from Novozymes. Another drawback related to the use of amylase in liquid detergents is the presence of calcium-containing chelating agents, known as builders. The primary function of these additives is to soften the wash water by binding calcium ions. Amylases in direct contact with these builders lose the calcium ions that support the enzyme molecule thus affecting long-term storage stability. Several groups have attempted to produce calcium-independent and acid-stable α-amylases by protein engineering. The x-ray structure of the α-amylase from Aspergillus oryzae was used as a template to study the importance of the region vicinal to the structural calcium-binding site. This region was modified by amino acid substitution so that the negative potential around the site was increased and made more attractive for cations such as calcium. The result was an amylase that bound calcium more tightly and was more thermostable than the wild-type parent enzyme from B. licheniformis (Van der Laan and May, 1995). Termamyl LCTM, which was produced by site-directed mutagenesis, also showed high calcium independence (Hashida and Bisgaard-Frantzen, 2000). The amino acids that lead to the tightening of calcium binding between the A domain and the B domain of the enzyme were substituted reducing, in this way, the destabilizing electrostatic interactions at low pH (Fig. 1.2). Ben Ali et al. (2001) reported the cloning and characterization of an atypically thermostable α-amylase from Geobacillus stearothermophilus US100 strain (AmyUS100), and also used site-directed mutagenesis to create an AmyUS100ΔIG mutant, which exhibited improved thermostability and very low calcium requirement, by the deletion of residues Ile214 and Gly215. (Ben Ali et al., 2006). This improved enzyme was also engineered with the aim of enhancing its resistance towards chemical oxidation


12


Ser264 Val209 Tyr156 Thr181 Phe190

1.2 A model structure of calcium independent α-amylase–Termamyl LCTM representing the five single substitutions His156Tyr, Ala181Thr, Asn190Phe, Ala209Val and Gln264Ser.

(Khemakhem et al., 2009). The same strategy of methionine replacement by non-oxidizable amino acids was followed and the mutant AmyUS100ΔIG/ Met197Ala was created by substituting methionine 197 to alanine. The mutant α-amylase was found to be very stable in the presence of solid detergents and, in some instances, the activity was increased by 10–50% after incubation of the enzyme at 40 °C with Ariel, Skip, Dixan, Det, NewDet, Nadhif and OMO. These studies indicate that this enzyme is an important candidate for potential application in detergent industry (Khemakhem et al., 2009).



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1.4.2 Pectinases Pectinolytic enzymes or pectinases are a complex group of enzymes involved in the degradation of pectic substances. They are primarily produced in nature by saprophytes and plant pathogens (bacteria and fungi) for degradation of plant cell walls (Lang and Dörenberg, 2000). There are three major classes of pectin-degrading enzymes: pectin esterases (PEs), polygalacturonases (PGs) and polygalacturonate lyases (PGLs), and their potential for bioscouring have been exploited (Choe et al., 2004; Ibrahim et al., 2004; Karapinar and Sariisik, 2004; Li and Hardin, 1997; Tzanov et al., 2001). Despite all the research done to develop an efficient bioscouring process, there is still no broad commercial application on the industrial scale. There is thus still a demand for a pectinase with higher activity and stability at high temperatures and alkaline conditions. Solbak and collaborators developed a novel pectate lyase using directed evolution that has improved thermostability. The new enzyme contains eight point mutations (Ala118His, Tyr190Leu, Ala197Gly, Ser208Lys, Ser263Lys, Asn275Tyr, Tyr309Trp, and Ser312Val). Compared to the wild-type enzyme, it presented a 16 °C higher melting temperature and exhibited improved bioscouring performance at low enzyme dosage in a high-temperature bioscouring process (Solbak et al., 2005).

1.4.3 Cellulases Cellulases are hydrolytic enzymes that catalyse the breakdown of cellulose to smaller oligosaccharides and finally glucose. Cellulase activity refers to a multicomponent enzyme system combining at least three types of cellulases, endoglucanases or endocellulases, cellobiohydrolases or exocellulases and β-4-glucosidase working synergistically together (Teeri, 1997). These enzymes are commonly produced by soil-dwelling fungi and bacteria, the most important being Trichoderma, Penicillium and Fusarium (Jorgensen et al., 2005; Kuhad et al., 1999; Verma et al., 2007). Cellulases from bacteria and fungi were first introduced in textile industry for denim finishing and are now also successfully used in laundry detergents. The yield of bacterial cellulases is normally low (Knowles et al., 1987) making fungal cellulases more promising and suitable for industrial applications. However, fungal cellulases have optimum activities at acidic pH. Their activities are greatly reduced at pH > 6, thus limiting the application of fungal cellulases under neutral or alkaline conditions. Therefore, protein engineering of cellulases has predominantly been used as a tool to increase the catalytic efficiency of these enzymes at higher pH. Becker et al., (2001) reported a variant of T. reesei Cel7A containing five point mutations (Glu223Ser/ Ala224His/Leu225Val/Thr226Ala/Asp262Gly) and having a more alkaline


14


optimum pH. They designed the mutations based on the sequence comparisons of family 7 cellulases with different pH behaviors. The comparison studies revealed that a histidine residue adjacent to the acid/base catalyst could account for the higher pH optimum of the Humicola insolens Cel7B endoglucanase and therefore a mutation Ala224His was designed. Modeling studies further suggested that four additional amino acid changes (Glu223Ser/Leu225Val/ Thr226Ala/Asp262Gly) would be required in order to fit the bulkier histidine side-chain. As the optimum pH of the Cel7A mutant enzyme became more alkaline, however, the overall activity on both soluble and insoluble substrates decreased (Becker et al., 2001). Wang et al. (2005) demonstrated that a single substitution of Asn342 by Thr in T. reesei endoglucanase Cel5A causes a pH optimum shift from 4.8 to 5.4. More recently, the same enzyme was successfully engineered not only to a more alkaline pH optimum but also to a higher catalytic efficiency (Qin et al., 2008). Residue 342 was subjected to saturation mutagenesis, and the enzyme was further changed by random mutagenesis and two rounds of DNA shuffling. A mutant form Asn342Val, with an optimal activity at pH 5.8, was obtained. This corresponds to a basic shift of one pH unit compared with the wild-type enzyme. The same mutant exhibited also improved catalytic efficiency for the main substrates at pH 6.2. Other variants obtained, namely Leu218His, Gln139Arg/Asn342Thr and Gln139Arg/Leu218His/Trp276Arg/ Asn342Thr, presented more than a 4.5-fold higher activity in reactions compared with the wild-type at pH 7.0. The increase in activities and pH optima of the variants was attributed to the existence of more stable helixes and to the changed electrostatic interactions between the catalytic residues and substrates (Qin et al., 2008). Voutilainen et al. (2007) recently reported the successful expression of the single-module cellobiohydrolase Cel7B from the thermophilic fungus Melanocarpus albomyces (Ma Cel7B) in Saccharomyces cerevisiae (Sc Cel7B) and used random mutagenesis to improve thermostability of the enzyme. Three interesting mutants (Ala30Thr, Gly184Asp and Ser290Thr) with increases in thermostability from 1–3 °C were obtained. Two different strategies were used to improve the hydrolytic activity of this enzyme towards crystalline cellulose at elevated temperatures: structure-guided protein engineering was used to introduce an additional tenth disulfide bridge to the Ma Cel7B catalytic module, and a fusion protein was also constructed by linking a cellulose-binding module (CBM) and a linker from the T. reesei Cel7A to the C terminus of Ma Cel7B. Both approaches proved to be successful (Voutilainen et al., 2009). Moreover, the disulfide bridge mutation Gly4Cys/Met70Cys was added to the most thermostable mutant (Ser209Thr) created previously and the double mutant Gly4Cys/Met70Cys/ Ser209Thr appears to be a promising candidate for textile applications requiring high temperatures (Voutilainen et al., 2009).



15

Other workers have followed the same strategy of construction of fusion proteins using CBMs from different sources. The application of CBMs in several areas of biotechnology has increased recently because CBMs are independently folded domains and a priori they can function perfectly when fused to other proteins. Moreover, their ligands are abundant and renewable materials with good properties, and they are normally low-priced (Shoseyov and Warren, 1997). One of the most important areas is bioprocessing, because large-scale purification and recovery of biological molecules continues to be a challenge. CBMs can be expressed as an affinity tag for protein immobilization, processing and purification using cellulose as a matrix in many formats, from affinity chromatography to two-phase liquid separations (McCormick and Berg, 1997). Targeting of compounds to polysaccharides that are present in many daily products, in particular cellulose, is also an important application of CBMs. This area of application includes, among others, denim stonewashing and the targeting of enzymes that do not possess a natural affinity to cotton in laundry detergents, (Berry et al., 2001; Cavaco-Paulo, 1995; Kalum and Andersen, 2000; von der Osten et al., 2000a,b). Degani and collaborators (2004) developed a new method to measure the scouring efficiency of cotton based on the fusion of a cellulose-binding domain with the β-glucuronidase protein (CBD-GUS). The chimeric enzyme was used as an effective tool for measuring scouring and wettability of cotton fabrics. CBD-GUS binding increased with hydrophilicity of the fabric owing to the increasing harshness of the scouring treatment. This quantitative method combines both simplicity and sensitivity in contrast to qualitatively inaccurate techniques or those requiring expensive instrumentation. Matamá et al. (2010) presented a novel and promising approach to increase cutinase adsorption on cellulose acetate fibers by fusing cutinase with two distinct (bacterial or fungal) CBM. They found an increase in reactivity and hydrophilicity of fibers treated with chimeric enzyme compared with the native cutinase. The application of CBMs is far from being exhausted and will be further expanded as the understanding of these binding domains increases.

1.4.4 Proteases: subtilisins Subtilisins are a family of alkaline serine proteases secreted by a variety of Bacillus species (Siezen and Leunissen, 1997). These enzymes catalyze the hydrolysis of peptide and ester bonds through formation of an acyl-enzyme intermediate. The objective of cloning bacterial protease genes was mainly the overproduction of enzymes for the detergent and leather industries and, more


16


recently, for the treatment of wool. Most subtilisin protein engineering continues to involve enhancement of catalytic activity and thermostability, as well as, substrate specificity and oxidation resistance. The first improvement of a detergent protease was performed in B. amyloliquefaciens subtilisin (subtilisin BPN′), to achieve oxidative stability. Met in position 222, adjacent to the Ser221 in the enzyme active site was susbtituted by non-oxidizable amino acids preserving the three-dimensional configuration of the protein (Estell et al., 1985). The mutant form of the enzyme became a commercial detergent protease under the tradename Maxapem®. A similar variant with the substitution Met222Ala was later introduced in the market under the name Durazyme®. The industrial use of proteases in detergents or for leather processing also requires an enzyme that is stable at higher temperatures. Several approaches have been taken to increase the thermostability of detergent proteases in order to achieve improved storage stability. Bryan et al. (1986) reported the thermal stabilization of subtilisin BPN′ as a result of the single substitution of Ser for Asn at position 218. The results of x-ray analysis indicated that slight improvements in the hydrogen-bond parameters enhanced the enzyme’s thermal stability. Potential salt bridges have been introduced into subtilisin BPN′ by protein engineering to improve the thermal stability (Erwin et al., 1990). Takagi et al. (1990) succeeded in stabilizing subtilisin E by introduction of a disulfide bond between residues Cys61 and Cys98. The two cysteines were introduced by protein engineering based on structural comparison with a thermophilic serine protease. The structure of subtilisin BPN′ was compared with that of subtilisin Carlsberg and the presence of a leucine at position 217 in subtilisin Carlsberg was identified as responsible for the high specific activity of this enzyme for synthetic substrates (Wells et al., 1987). By replacing the tyrosine that was at position 217 in subtilisin BPN′ by leucine, the resulting mutant enzyme (BPN′ Tyr217Leu) exhibited a ten-fold increase in efficiency, compared with the wild-type subtilisin BPN′, for the hydrolysis of synthetic substrates. When tested for performance in neutral pH liquid detergents, the Tyr217Leu variant was found to be twice as efficient as subtilisin BPN′ and also more stable (Wolff et al., 1996). Thus, a single amino acid change in subtilisin BPN′ yielded an enzyme that is significantly better for laundry applications than the parent enzyme. In a similar study, the structure of the subtilisin produced by Bacillus lentus (subtilisin BL) was compared with the structure of subtilisin BPN′ and it was found that both enzymes differ in 103 out of 275 positions. One of the sequence differences between the two enzymes is that subtilisin BL has valine at position 104 whereas subtilisin BPN′ has tyrosine at this position. The amino acid at position 104 in the sequence of subtilisin BPN′ was previously identified and reported as important for performance of the enzyme and that a tyrosine was preferred



17

over valine at this position (Estell and Wells, 1988). In separate mutagenesis experiments on the subtilisin BL enzyme, it was found that mutations at positions 123 and 274 were also important for performance of this enzyme (Caldwell et al., 1991). A variant of subtilisin BL was therefore constructed in which tyrosine replaced valine at position 104, serine replaced asparagine at position 123, and alanine replaced threonine at position 274. The resulting enzyme was more than twice as efficient as the parent subtilisin BL in the cleaning of enzyme sensitive stains in a granular alkaline detergent matrix (Caldwell et al., 1991). SDM was also used to introduce a Asn218Ser mutation that increased the thermostability of the enzyme (Wang et al., 1993). SDM was also used by Yang et al. (2000a) to generate a Ser236Cys mutant subtilisin E with a half-life, at 60 °C, four-fold longer than that of native subtilisin E. Using this mutant, thermostability could also be increased, by forming a disulfide bridge between two molecules of Ser236Cys subtilisin E. Yang et al. (2000b) also used random mutagenesis PCR technique to develop a thermally stable and oxidation-resistant mutant. The new Met222Ala/Asn118Ser subtilisin E was five-fold more thermally stable than native enzyme. In another report, the thermal stability of subtilisin E was increased using directed evolution to convert B. subtilis subtilisin E into an enzyme functionally equivalent to its thermophilic homolog thermitase from Thermoactinomyces vulgaris (Zhao and Arnold, 1999). Mainly for ecological reasons, proteases of the subtilisin type are also being studied as an alternative for chemical pre-treatment of wool. However, the increase in subtilisin molecular weight is crucial for its successful application in wool finishing because, owing to its small size, the enzyme is able to penetrate into the fibre cortex causing the destruction of the inner parts of wool structure (Shen et al., 1999). Araújo et al., (2008a) attempted to create recombinant subtilisin E with high molecular weights using two novel approaches: the construction of two polysubtilisins, (fusing two and four subtilisin E genes in frame), and the formation of a subtilisin trimer by fusion of native prosubtilisin E gene with the gene coding for SP-D neckdomain. Both approaches resulted in the expression of three modified subtilisins although no activity was recovered for these enzymes. They also reported a different approach to increase the subtilisin E molecular weight based on the fusion of the DNA sequences coding for B. subtilis proSubtilisin E and for an elastin-like polymer (ELP) (Araújo et al., 2009). The ELP gene used is based on 220 repetitions of the monomeric sequence Val Pro Ala Val Gly (VPAVG), and it was constructed in the same laboratory. The recombinant protein, exhibiting a molecular weight above 116 kDa (an increase of more than four-fold in the weight of the native enzyme), was biologically produced in E. coli, purified and used for wool-finishing assays. Both yarns and fabrics treated with genetically engineered enzyme


18


presented higher tensile strength and lower felting and pilling, along with lower weight loss, which indicates that the new developed enzyme only hydrolyzed the cuticle layer of wool (Araújo et al., 2009). This new high molecular weight subtilisinE-VPAVG220 represents a breakthrough in the wool-finishing process, promising to be an alternative to the traditional highly polluting chlorine/Hercosett treatment. In addition, this enzyme can be included in new detergent formulations that can be used to wash all types of garments, including silk and wool. Finally, the cost of enzyme production is a major obstacle to the successful application of proteases in textile industry. Protease yields have been improved by screening for hyperproducing strains and/or by optimization of the fermentation medium. Strain improvements either by conventional mutagenesis or recombinant-DNA technology have been useful in improving the production of proteases. Most, if not all, Bacillus detergent proteases currently are recombinant, genetically engineered products, secreted by overproducing strains.

1.4.5 Lipases/esterases Esterases represent a diverse group of hydrolases that catalyze the cleavage and formation of ester bonds. They are widely distributed in animals, plants and micro-organisms. These enzymes show a wide substrate tolerance and high regio- and stereospecificity, which make them attractive biocatalysts for the production of optically pure compounds in fine-chemicals synthesis. They do not require cofactors, are usually rather stable and are even active in organic solvents (Bornscheuer, 2002). Two major classes of hydrolases are of most importance: lipases (triacylglycerol hydrolases) and ‘true’ esterases (carboxyl ester hydrolases). Most of the alterations introduced in esterases/lipases address detergent use and surfactant compatibility. Oxidative stability, important for proteases and amylases, is not the major interest in the case of lipases, since many lipases are already stable in oxidative reagents. In Candida antarctica B lipase the exchange of Met at position 72 by Leu resulted in an increased stability towards oxidation by peroxyoctanoic acid (Patkar et al., 1998). A few studies also report the substitution of Met in lipases from Pseudomonas sp. by other residues to prevent the inactivation by oxidation in oxidative detergents (Van der Laan et al., 1994). Regarding calcium independency, Simons et al. (1999) engineered S. hyicus lipase by site-directed mutagenesis. Based on sequence alignment to other lipase sequences, from P. glumae for example, the aspartate residues in position 354 and 357 were identified as calcium-binding ligands and replacement of Asp357 by a glutamate decreased the affinity for calcium ions by 30-fold. Introduction of a lysine, an asparagine, or an alanine at position 357 and of a lysine or an asparagine at position 354 resulted in © Woodhead Publishing Limited, 2010


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calcium-independent variants. Although the Asp357Lys, Asp357Asn, and Asp357Ala variants did not bind calcium, at elevated temperatures these calcium-independent mutants showed a reduced activity. Over the whole temperature range the activities of the Asp354Lys and Asp354Asn variants are significantly lower than the wild-type enzyme in the presence of calcium. The lipase from Humicola lanuginosa has also been the target of surface engineering in order to overcome calcium dependence. The variant Asp96Leu is less dependent of calcium presence and has an increased stability in the presence of non-ionic and anionic surfactants (Svendsen et al., 1997). The proteolytic stability of P. glumae lipase in the presence of proteases for detergent use has also been improved by protein engineering. Two different strategies, the replacement of susceptible amino acid residues at the cleavage site with residues that are not accepted as substrate and the replacement of amino acid residues with proline have been employed (Frenken et al., 1993). Proline is the only proteinogenic amino acid and is not accepted by most of the proteases at the potential cleavage site (Bromme et al., 1986). In both instances, a considerably increased stability towards proteolytic degradation was observed. Cutinase has also been exploited for the surface modification of synthetic fibres. Despite the potential of cutinase from Fusarium solani to hydrolyse and improve synthetic fibres properties, these fibres are non-natural substrates of cutinase and, consequently, turnover rates are quite low. By the use of site-directed mutagenesis recombinant cutinases, with higher specific activity to large and insoluble substrates such as polyethylene terephthalate (PET) and polyamide (PA), were developed (Araújo et al., 2007). The mutations Leu81Ala, Asn84Ala, Leu182Ala, Val184Ala and Leu189Ala were used to enlarge the active site in order to better fit a larger polymer chain (Fig. 1.3). The new cutinase, Leu181Ala mutant, was the most effective in the catalysis of amide linkages of PA and displayed a remarkable hydrolytic activity towards PET fabrics (more than five-fold compared with native enzyme) (Araújo et al., 2007). This recombinant enzyme was further used to study the influence of mechanical agitation on the hydrolytic efficiency of cutinase on PET and PA in order to design a process for successful application of enzymes to synthetic fibres (Silva et al., 2007; O’Neill et al., 2007). The use of cutinase opens up new opportunities for targeted enzymatic surface functionalisation of PET and PA, polymers formerly considered as being resistant to biodegradation.

1.5

Advantages and limitations

The tools of enzyme engineering, well developed over the last few years by many laboratories, are now being applied for the optimization of © Woodhead Publishing Limited, 2010

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Advances in textile biotechnology Ala182

Ala184 Ala189

Ala81

1.3 A model structure of Fusarium solani pisi cutinase representing the five single substitutions Leu81Ala, Asn84Ala, Leu182Ala, Val184Ala and Leu189Ala.

biocatalysts used in different processes of textile industry. The use of sitedirected mutagenesis and directed evolution has rapidly emerged and these techniques are the choice for the development and selection of mutated enzymes with altered and improved activity, specificity, stability and availability. The advantages of site-directed mutagenesis are its high reproducibility, technical simplicity and cost-effectiveness measured by time-consumption and consumable expenses. However, a lack of available structural and biochemical information limits the number of proteins that can be re-designed by this approach. On the other hand directed evolution can be used to tailor a given enzyme without the need for further structural and biochemical data, but, owing to the large number of variants obtained, this is a time-consuming and expensive method. In addition, results obtained can often not be transferred from one biocatalyst to another even if it is a structurally or functionally similar one. The high investment in time and



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costly screening in combination with an often narrow substrate spectrum of enzymes can delay the rapid development of biocatalysis. Over the past few decades, several enzymes have been identified and optimized, by either method, as catalysts for some textile industrial applications. Nevertheless, one major overall limitation for the application of enzymatic processes in the textile industry is that the public recognition of its benefits is not widespread even in developed countries. It is very important to provide information about the urgent need to reduce the use of chemicals in the textile industry to preserve environmental quality and protect biodiversity, thus also bringing economic advantages. In addition, publicly funded research bodies in developing countries need to intensify efforts to develop enzyme-based technologies of their own.

1.6

Future trends

Over the past few years, enzymes were studied and used in order to develop environmentally friendly alternative processes for almost all steps in textile fibres processing. For this reason, we have witnessed great progress in the application of enzyme engineering for the production of suitable biocatalysts for these processes. There are already some commercially successful applications, such as amylases used for desizing, cellulases and laccases for denim finishing, and proteases incorporated in detergent formulations. Although some types of enzymes already play an important role in some textile processes, their potential is much greater and their applications in future processes are likely to increase in the near future. Much work remains to be done to improve productivity and efficiency to a point at which these technologies will be as economically advantageous as the conventional approaches. Further research is still required for the implementation of commercial enzyme-based processes for the biomodification of synthetic and natural fibers. Another field of research is the search for new enzyme-producing micro-organisms and enzymes extracted from extremophilic microorganisms (Schumacher et al., 2001). Furthermore, the availability of more genomes and their better understanding will open new doors to better engineer industrial host strains and will certainly contribute to overcoming the disadvantages and limitations. New approaches are needed to better understand the metabolism and growth of host organisms. Genome-wide transcription profiling, proteomics and the elucidation of the complete metabolic networks will provide important understanding of cellular processes of organisms of industrial interest. In consequence, the classical biotechnology approaches that focus on the design and improvements of individual genes or proteins will be expanded by global approaches, all aiming at the understanding of the interaction of the complete set of


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cellular components. This concept of ‘systems biology’ and the knowledge deduced will significantly improve industrial fermentation processes and will further increase the value of known and new organisms in the future of textile biotechnology.

1.7

Sources of further information and advice

For several reviews concerning endonucleases recognition sequences see Kessler and Manta, 1990; McClelland et al., 1994; Roberts and Macelis, 1996; for applications, Williams, 2003; genetics, Wilson and Murray, 1991; biology, Bickle and Kruger, 1993; structure, Anderson, 1993; Aggarwal, 1995; protein engineering, Jeltsch et al., 1996; Lanio et al., 1998 and enzymology Pingoud and Jeltsch, 2001. For further information regarding enzymes for extremophiles and their potential applications see Fujiwara, 2002. For reviews regarding the applications of enzyme in detergency and industrial textile processes see Araújo et al., 2008b; Galante and Formantici, 2003; Guebitz and Cavaco-Paulo, 2008. For reviews concerning protein engineering of subtilisins see Bryan, 2000; of cellulases, Schulein, 2000; of amylases Nielsen and Borchert, 2000 and of lipases, Svendsen, 2000.

1.8

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2 Developments in processes and equipment for enzymatic textile treatments M. M. C. G. WA R MOE SK E R K E N and G. H. BOU W H U I S, University of Twente, The Netherlands

Abstract: The role of mechanical energy in enzymatic textile treatment is examined. The special properties and behaviour of enzyme molecules can cause poor performance on an industrial scale owing to the absence of liquid flow in the capillary pores of textile. The role of mass transfer in these systems is discussed, a squeezing factor α is introduced and a model is derived with which the mass transfer rate in textile materials can be calculated. The phenomenon of exhaustion of the capillary liquid during the adsorption of the enzymes is also discussed. A dynamic adsorption model is used to explain this phenomenon. Key words: mass transfer, textile processing, enzymatic treatment.

2.1

Introduction

The application of enzymes in textile treatment processes seems to be quite simple at first sight: just replace the traditional chemicals by enzymes. However, in practice, it was found that such an approach results in poor performance often leading to a reluctance to use modern enzyme technology in textile treatment processes (Bouwhuis et al. 2009). Because enzyme molecules are much bigger than the traditional chemical molecules such as sodium hydroxide, enzymatic treatment processes are often diffusion controlled, Nierstrasz and Warmoeskerken (2003). Moreover, the kinetics of enzymatic reactions is more complex than for traditional chemistry. This complex diffusion control of the transport of enzymes inside the pores of a fabric leads to a suboptimal adsorption of enzymes at the capillary surface. Therefore, the key for a good performance using enzymes in textile treatment processes is an optimal mechanical energy to prevent diffusion control. It could be argued that the enzyme amylase is successfully applied in the textile industry for the desizing process without transport limitations, but the reason for this is that the size is present only at the outer surface of the weft yarns and these surfaces can easily be accessed by enzymes without a limiting transfer process between the fibres of the yarns. 32 © Woodhead Publishing Limited, 2010

Developments in enzymatic textile treatments

33

There has been a great deal of research on the application of enzymes in wet textile processes. However, work in which the role of mechanical energy is also included is rather rare. There are some data on the role of mechanical energy in wet textile processes, but these are mainly related to fabric wash processes (Donk, Van der et al., 1998, Warmoeskerken and Boom 1999). Lenting and Warmoeskerken (2001) discussed the role of mechanical shear in enzymatic textile treatment. This work was focused on the removal of fibril pills during the treatment of cotton fabric with cellulose. Agrawal et al. (2008) were the first to show unambiguously that there is strong synergistic relation between the performance of enzymes in wet textile processes and the mechanical energy applied. They also showed that the mechanical action can be delivered by ultrasound resulting in a very effective and efficient enzymatic treatment process (Agrawal et al., 2007). Data on the suboptimal adsorption of enzymes at the surfaces of the textile capillaries is not available. In this chapter, the phenomena mentioned here are discussed and models that describe them are presented.

2.2

Current processes for the pre-treatment of cotton

In cotton pre-treatment, different types of installed base and processing are necessary to realize sufficient desized, scoured and bleached fabric for consecutive treatments such as dyeing, printing or finishing. The process steps for the pre-treatment of cotton are listed below. 1 2 3

Desizing: the removal of starch from the warp yarns. Scouring: the removal of waxes and pectin for sufficient hydrophilicity. Bleaching: the decolourization of the natural ‘grey’ colour of cotton.

If possible, one or more process steps are combined. For the pre-treatment, various types of processes can be distinguished.

2.2.1 Full-continuous pre-treatment The full-continuous process is done on two types of installed base. The first type is open width form. For the consecutive process steps, the fabric is impregnated with processing agents on an impregnation unit. This unit consists of rollers that guide the fabric into and out of the bulk. During the impregnation, the fabric is squeezed at least once. After impregnation, the fabric is incubated in a steamer where the fabric is guided over rollers. After incubation, the fabric is rinsed in washing compartments. A washing compartment consists of rollers to guide the fabric into and out of the bulk to


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rinse out the impurities. The process steps of desizing, scouring and bleaching are treated according to the same pattern (impregnation–incubation– rinsing). For each process step, different impregnation and incubation units can be distinguished depending on the time, temperature and mechanical energy necessary to achieve the desired result. In the second type of installed base for continuous treatment processes, the fabric is in rope form. Impregnation is done in a rope impregnation unit, followed by incubation in J-boxes and rinsing in J-boxes alternated with rope-squeezers. Characteristic of this process is the kneading of the fabric by filling the J-boxes completely with fabric in rope form and pushing the fabric through the J-box. The latter is only possible when large amounts of fabric are in the J-box and therewith long impregnation, incubation and rinsing times are achieved.

2.2.2 Pad batch–hot bleach process The pad batch–hot bleach process is an open width treatment. The fabric is impregnated in an impregnation bath in which the fabric is guided by rollers into and out of the bulk. After impregnation, the fabric is rolled on an A-frame and incubated for several hours. After incubation, the fabric is rinsed in washing compartments and consecutively impregnated with bleaching agents. For impregnation, high concentrations of bleaching agents (>100% pick-up) are added on the fabric. The fabric is incubated in a steamer and after incubation rinsed in washing compartments.

2.2.3 Discontinuous pre-treatment In a discontinuous installed base, the fabric is guided into the machine and transported/moved in the same compartment. The distinguished types of installed base are jigger, jet and winch. On a jigger, the fabric is moved from one roller, via the treatment bath containing the treatment agents, to the opposite roller (and vice versa). In a jet, the fabric is guided in rope form in a J-box. The fabric is made endless and guided out of the bulk through a pipe to the loading point of the J-box. In the J-box the fabric is kneaded and pushed forward for the next tour. The winch is the same type of installed base as the jet. The difference is in the transport of the endless fabric. The fabric is ‘stored’ in the bulk and taken out of the bulk by the winch. After passing the oval winch, the fabric is again laid down in the bulk to be pushed and kneaded again. The mechanical energy in the different process forms is represented in Table 2.1. The symbols used in the table are:



35

Table 2.1 Representation of mechanical energy in the various process forms (see text for key to symbols) Processing

Process step

Process [c] time

Impregnation Incubation Rinsing Full continous/ open width form

Desizing Scouring Bleaching

rr s rr s rs

rr rr rr

rr ss rr ss rr ss

t t t

cc cc cc

kk ss kk ss kk ss

kk kk kk

kk ss kk ss kk ss

tt tt tt

c c c

Pad batch–hot bleach

Desizing rr s Scouring/ rr ss bleaching

n rr

rr ss rr ss

ttt t

c ccc

Discontinuous

Jigger Jet Winch

ttt ttt ttt

c c c

Full continuous/ Desizing Scouring rope form Bleaching

r rr s ss k kk t c n

r k k

mechanical energy owing to rollers bending fabric once over a roller; mechanical energy owing to rollers bending fabric at least twice; squeezing once; squeezing, at least twice; kneading of the fabric; intensive kneading of the fabric; time consumption, an increasing number of ‘t’s indicates increasing time consumption; concentration of chemicals used, an increasing number of ‘c’s indicates increasing concentrations; and no mechanical energy available.

From Table 2.1, it is clear that there are many possible options for the pretreatment of fabrics and each option has its characteristic way to add mechanical energy to the process. Since the mechanical energy is a key factor in the performance of enzymes it is to be expected that enzymes perform differently on the different systems presented in the table. It is evident that there will be a strong relationship between the amount of mechanical energy applied to the process and the time required to achieve a good enzymatic performance. Therefore, a profound understanding of the mechanical energy in these systems and its effect on the enzymatic processes is a prerequisite for a successful introduction of enzymes in the textile industry.


36


2.3

Mass transfer limitations in textiles

To understand the role of mechanical energy in enzymatic textile processes, knowledge of the mass transfer phenomena in these systems is necessary (Nierstrasz and Warmoeskerken, 2003). Textile is a material with a dual porosity: pores between the yarns and pores between the fibres, Fig. 2.1. The former are called the inter-yarn pores and the latter the intra-yarn pores. For a woven cotton fabric the inter-yarn pores have a size of the order of 50 μm and the intra-yarn pores have a diameter in the order of 2 μm. This difference means that when water flows through the fabric, it flows through the inter-yarn pores rather than through the intra-yarn pores. The flow resistance in the small pores is higher than in the bigger pores. Therefore there will be a large area of the fabric where there is no water flow at all. A visual model for this has been developed by Warmoeskerken and Boom (1999) and is shown in Fig. 2.2. Figure 2.2 shows the cross section of a yarn and its fibres. The bulk of the water flows along the yarns through the inter-yarn pores. Depending on the water flow intensity, there is some flow of water between the fibres as indicated in the figure. Therefore, two zones are defined in the yarn: a stagnant core and a convective shell. In the stagnant core, there is no flow at all whereas, in the convective shell, the flow penetrates to some extent. It is clear that the mass transfer in the stagnant zone is completely determined by molecular diffusion whereas in the convective shell the mass transfer is controlled by convection. If we assume that a yarn is completely wetted so that all the intra-yarn pores are filled with water, and this yarn is put into Fabric Yarn

Inter-yarn pore

Intra-yarn pore

2.1 Inter- and intra-yarn pores in fabrics.



37

Stagnant core

Convective shell

2.2 Yarn model with a stagnant core and a convective shell.

a still standing enzyme solution so all the liquid in the yarn is stagnant, then the enzyme molecules will start to diffuse into the yarn. This process is described by the diffusion equation of Fick (Bird et al., 1960): ∂CE,Y D ∂ ⎛ ∂CE,Y ⎞ = ⎜ rY ⎟ ∂rY ⎠ ∂t rY ∂r ⎝

[2.1]

in which CE,Y is the enzyme concentration in the yarn (kg m−3), rY is the yarn radius (m), D is the diffusion coefficient of the enzymes (m2 s−1) and t is the time (s). This equation only describes the diffusion process in the radial direction of the yarn. The solution of this equation is often graphically given as in Fig. 2.3. The variable E on the vertical axis is the dimensionless mean concentration in the yarn: E=

CE,b − CE,Y CE,b − CE,Y,0

[2.2]

in which CE,b is the enzyme concentration in the enzyme solution (kg m−3), CE,Y is the mean enzyme concentration in the yarn (kg m−3) and CE,Y,0 is the mean initial enzyme concentration in the yarn (kg m−3). The variable on the horizontal axis is the Fourier number Fo and is a dimensionless expression for the diffusion time: Fo =

Dt dY2

[2.3]

in which dY is the yarn diameter (m).


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Concentration of enzyme in yarn, E

1

0.8

0.6

0.4

0.2

0 –4 10

10

–3

–2

10 Fourier number, Fo

10

–1

1

2.3 Graphical solution of equation [2.1].

Now we are able to calculate the diffusion time of the enzymes into the yarn. If the diameter of the yarn is 0.5 mm and the diffusion coefficient in the yarn D = 10−11 m2 s−1 based on an enzyme molecule of 25 kDa and a intra-yarn porosity of 0.4 (Nierstrasz and Warmoeskerken, 2003) and if E = 0.1, i.e. the mean enzyme concentration in the yarn is 90% of the enzyme concentration in the solution, assuming that the initial enzyme concentration in the yarn is zero, CE,Y,0 = 0, from Fig. 2.3, it follows that Fo ⯝ 0.1. From this value and equation 2.3, a diffusion time of 2500 s is calculated. Therefore, in this example, it takes approximately 40 min to achieve a mean enzyme concentration in the intra-yarn pores that is 90% of the final value. It is clear that this is too long for a normal treatment process. If we repeat these calculations for a small molecule such as NaOH, the diffusion time is in the order of 25 s which is much shorter than the value for big enzyme molecules. The mass transfer time for enzymes can only be reduced by applying mechanical energy that induces flow within the intra-yarn pores, consequently resulting in a smaller stagnant core and more convective transfer. The way to achieve this is by squeezing the fabric to force water to flow through the intra-yarn capillaries. This can be achieved when the textile is running over rollers or by passing between squeezing rollers. The mechanical energy delivered by these systems and their effect on the mass transfer inside the textile is unknown. To enable the squeezing action in textile treatment systems to be measured we have developed a model that relates the performance of wet textile equipment to a squeezing factor α.



2.4

39

The squeezing model

As previously stated, the mechanical energy in textile treatment equipment is aimed at deforming the porous fabric to create velocities in the pores of the textile. Figure 2.2 shows the yarn model with the stagnant core and a convective shell. In the stagnant core, the mass transfer is fully controlled by diffusion whereas in the convective shell this is based on convection, i.e. the flow of liquid. Here, we derive a model that describes this system. In this model, a squeezing factor α expresses the ratio between the stagnant and the convective regions. To make the model less complicated, the fabric is modelled as a porous slab with a stagnant centre layer and two convective outer layers as indicated in Fig. 2.4. In Fig. 2.4, df,S is the thickness of the stagnant middle layer and df is the thickness of the fabric. Thus, the thickness of the convective layers df,C is: df,C = df − df,S

[2.4]

The squeezing factor α is now defined as:

D=

volume of the convective layers in the fabric Vf,C = total volumee of the fabric Vf

[2.5]

If the surface area of the textile is A (m2):

α=

Adf,C df,C df,S = = 1− Adf df df

[2.6]

If α equals zero, df,S equals df, which is the case when there is no flow at all, so the liquid in the total volume of the fabric is stagnant. If α equals 1, df,S equals zero, which is the case if the water flows in all the pores of the fabric. If we relate the flow within the fabric to deformation we can also say that if α = 1, all the pores are deformed, i.e. maximum deformation effect of the mechanical energy and flow in all the pores. If α = 0, no pores are deformed, i.e. no deformation effect of the mechanical energy and no flow in the pores. It should be noted here that the value of the squeezing parameter α is a mean value. So when the fabric is going through a padding bath, the squeezing factor of this system is a mean value taken from the point where the

df

df,S

2.4 Model of a textile slab with a stagnant centre layer and two convective outer layers.


40


fabric enters the bath until the point where the fabric leaves the bath. The α-factor of the bath is an indication of how much of the fabric is stagnant and how much is convective in the padding bath. It must be noted that each system has its own value of the squeezing factor α. The value of α is related to the value of the forces or pressures that are exerted onto the load while passing over rollers or through squeezers. Figure 2.5 shows a schematical relationship between α and the pressure forces. The deformation parameter only gives information about the fraction of pores in which some flow can be generated as a result of the deformation. It does not give information about the magnitude of the flow in the pores. This value is dependent on the rate of deformation and not on the magnitude of deformation. The rate of deformation in represented by the Peclet number. This dimensionless number is defined as: Pe =

convective mass transfer vP LP = diffusive mass transfer D

[2.7]

in which vp is the mean liquid velocity in the pores (m s−1) and LP is the length of the pores (m). This parameter is related to the liquid velocities generated in the pores because these depend directly on the deformation rate. The higher the rate of deformation, the higher the induced pore velocities are. It can be argued that the pore liquid velocities that are in fact the liquid displacement velocities should be proportional to the pore deformation rate as drawn schematically in Fig. 2.6. If we now consider an enzyme padding processes with a particular value of the squeezing factor α and of Pe, then the curve that describes the concentration of enzymes in the fabric as a function of time looks as schematically drawn in Fig. 2.7.

a

1

0 Pressure force

2.5 Expected relation between the squeezing factor α and the pressure force exerted onto a textile.


41

Induced pore velocities


Pore deformation rate

2.6 Expected relationship between the induced pore velocities and the pore deformation rate.

1

Slow regime

E

Fast regime

0 Time

0

2.7 Expected enzyme concentration in the fabric as a function of time for a particular value of α and Pe.

In Fig. 2.7, the enzyme concentration is expressed by a factor E, which is the ratio between the actual enzyme concentration in the fabric and the concentration that is achievable after an infinitely long time: E=

CE,f,t CE,f,∞

[2.8]

where CE,f,t is the enzyme concentration in the fabric at time t and CE,f,∞ is the enzyme concentration in the fabric after an infinitely long time. Figure 2.7 shows two regimes: a fast and a slow increase of the enzyme concentration. The fast part of the curve represents the convective transfer process


42


whereas the slow part of the curve shows the relatively slow diffusion process. A value for the squeezing factor α can be obtained by a short cut method that approximates the two regimes by straight lines. as shown in Fig. 2.8. It can be argued that the E value at the cross point is also the value of α. It is the point where convective transport is changed into diffusion transport. The angle of the fast regime of the curve is proportional to the Pe number as indicated in Fig. 2.8 because Pe represents the rate of fabric deformation and thus the rate of convectional transport. To clarify the concepts of the squeezing factor α and the Pe number, Fig. 2.9 and 2.10 may be helpful. In Fig. 2.9, three curves have been drawn,

1

E

a

Pe 0 0

Time

2.8 Short-cut method to determine the value of the squeezing factor α graphically.

1

E

a1 = a2 = a 3

Pe3 Pe2 Pe1

0 Time

0

2.9 Three curves representing the transport of enzymes into a fabric as a function of time for different values of Pe and identical values of α.


Developments in enzymatic textile treatments 1

43

a3

E

a2

a1 Pe1 = Pe2 = Pe 3 0 Time

0

2.10 Three curves representing the transport of enzymes into a fabric as a function of time for different values of α and identical values of Pe.

all for a constant value of α and three different values of Pe. Because diffusion is identical for all three cases, the slow parts of the curves coincide at longer times. However, for high Pe numbers the diffusion regime starts earlier because the convective transport is completed faster. Figure 2.10 shows three curves for three different values of α and a constant value of the Pe number. The rate of convective transport is the same for all cases because the deformation rate is constant. However, for high values of α, the extent of deformation is higher, leading to a higher value of E at which the diffusion regime starts.

2.5

A mass transfer model

The mass transfer in a fabric can be modelled using the squeezing factor α. The model derived here is for rinsing mass from the fabrics to a rinsing bath. Because the impregnation of fabrics by enzymes from an enzymatic solution is just the other way around, the equations of the model can easily been applied for rinsing as well as for impregnation. The model is based on the squeezing principles leading to a stagnant core and a convective region. The release of enzymes is modelled by the principles of chemical reaction kinetics. The release ‘reaction’ can be written as: S C CE,f,S ⎯k⎯ → C E,f,C ⎯k⎯ → CE,B

[2.9]

The enzymes in the stagnant region of the fabrics CE,f,S become enzymes in the convective region in the fabric CE,f,C by diffusion. The enzymes in the convective region become enzymes in the bath CE,B by convective transport from the convective regions to the bath. The transfer coefficient for transport from the stagnant to the convective region is kS (s−1) and the transfer


44


coefficient for transport from the convective regions to the bulk is kC (s−1). During the release process, the enyme concentration in the bulk liquid CE,B increases. For an impregnation process, this bulk concentration decreases. The model describes the change of the bulk concentration as a function of time. The starting equation is a mass balance for the enzymes present in the total system. This balance is valid for all times. Vf CE,f,0 = Vf,SCE,S + Vf,CC E,C + VBC E,B

[2.10]

This balance assumes that there is initially a homogeneous enzyme concentration in the fabric CE,f,0. Vf, Vf,S and Vf,C are, respectively, the total fabric volume, the volume of the stagnant region and the volume of the convective region. CE,S, CE,C, and CE,B are, respectively, the enzyme concentration in the liquid bath, in the stagnant region and in the convective region as a function of time. Thus, from the mass balance an expression for CE,B can be derived: CE,B =

Vf CE,f,0 − Vf,SCE,S − Vf,CCE,C VB

[2.11]

The volumes Vf, Vf,S, Vf,C and VB can be expressed in α as defined earlier and the liquid to cloth ratio LCR: Vf,C =α Vf

[2.12]

Vf,S = 1−α Vf

[2.13]

LCR =

ρBVB ρf Vf

[2.14]

in which ρB is the density of the liquid in the bath (kg m−3) and ρf is the density of the fabric (kg m−3). Substitution of equations [2.12], [2.13] and [2.14] into equation [2.11] gives: CE,B =

ρB 1 [CE,f,0 − ( 1 − α ) CE,S − α CE,C ] ρ f LCR

[2.15]

Expressions for the enzyme concentration in the stagnant and convective regions CE,S and CE,C can be found with the following rate equations resulting from the release ‘reaction’ equation above. Vf,S

dCE,S = −Vf,S kSCE,S dt

[2.16]

Vf,C

dCE,C = Vf,S kSCE,S − Vf,C kCC E,C dt

[2.17]



45

This set of rate equations can be rewritten as: dCE,S = −kSCE,S dt

[2.18]

dCE,C 1 − α = kSCE,S − kCCE,C dt α

[2.19]

Equation [2.19] can be solved easily for the initial conditions: t = 0, CE,S = CE,C = CE,f,0 and CE,B,0 = 0. This means that the initial enzyme concentration in the fabric is everywhere CE,0 and that the initial enzyme concentration in the bath is zero. The solution of equation [2.19] now becomes: CE,S = CE,f,0 e− kSt

[2.20]

Substitution of this expression for CE,S in equation [2.19] gives: dCE,C 1 − α = kSCE,f,0 e− kS t − kCCE,C α dt

[2.21]

which is a standard first order differential equation of the type: dy = a + bekx + cy dx

[2.22]

The solution of this differential equation can be found in handbooks (Kreyszig, 1993). By applying the initial conditions t = 0, CE,C = CE,f,0, the solution of the standard equation for the enzyme concentration in the convective region gives: ⎡ k − D kC − kCt kS ( 1 − D ) − kSt ⎤ CE,C = CE,f,0 ⎢ S e − e ⎥ D ( kS − kC ) ⎣ D ( kS − kC ) ⎦

[2.23]

If we substitute now the expressions for CE,S and CE,C, equations [2.20] and 2.23], into equation [2.15] we finally obtain the expression for the enzyme concentration in the bath liquid: CE,B =

ρ B CE,f,0 ⎡ k − α kC − kCt kS ( 1 − α ) − kSt ⎤ 1 − ( 1 − α ) e − kS t − S e + e ⎥ ρ f LCR ⎢⎣ kS − kC kS − kC ⎦ [2.24]

For t = 0, this equation results in CE,B = 0, and for t = ∞, CE,B,∞ =

U B CE , f,0 U f LCR


[2.25]

46


Thus, equation [2.24] can also be written in a dimensionless form: CE,B k − α kC − kC t kS (1 − α ) − kS t = 1 − ( 1 − α ) e − kS t − S e + e CE,B,∞ kS − kC kS − kC

[2.26]

The dimensionless bulk concentration CE,B /CE,B,∞ has a value between 0 (for t = 0) and 1 (for t = ∞). Two limiting cases are also α = 0 and α = 1. If α = 0, there is no convection zone and the mass transfer is fully controlled by diffusion. From equation [2.26], it follows that: ⎛ CE,B ⎞ = 1 − e − kS t ⎜⎝ C ⎟ E,B,∞ ⎠ α = 0

[2.27]

If α = 1, the mass transfer is fully controlled by convection and equation [2.26] becomes: ⎛ CE,B ⎞ − kC t ⎜C ⎟ = 1− e ⎝ E,B,∞ ⎠D =1

[2.28]

Equation [2.26] can be applied to find the values for α, kS and kC. This can be done by measuring the enzyme concentration in the bath during a release experiment in time. Applying non-linear curve fitting then delivers the required values. We have also done some model calculations to show the effect of the squeezing factor α on the transfer of enzymes into the fabric. The values used for this example are: • • • • •

diffusion coefficient, D = 10−11 m2 s−1 fabric thickness, df = 0.5 mm liquid to cloth ratio, LCR = 5 liquid density, ρB = 1000 kg m−3 fabric density, ρf = 1200 kg m−3

Figure 2.11 shows the results of the calculations. In this case we used equation [2.26] to calculate the transport of enzymes from a bath to the fabric. For the mass transfer coefficient in the stagnant region, we have taken: kS =

D dS2

[2.29]

or by substituting the definition for α: kS =

D

[df (1 − α )]

2


[2.30]


47

Enzyme concentration, E

1.0 0.8 0.6 0.4 0.2 0.0

0

20

40

60 Time (s)

80

100

2.11 The results of model calculations using equation [2.26].

Furthermore, it has been assumed that the convective transfer coefficient is 100 times the stagnant transfer coefficient. The calculations have been done for five different values of the squeezing factor α as indicated in Fig. 2.11. These model calculations confirm that the squeezing factor has a determining effect on the rate with which enzymes are transferred from a padding bath to the fabric.

2.6

Adsorption limitation in textile pores

Another problem in enzymatic textile treatment processes results from the relatively high ratio of the surface area and the volume of the capillaries in the fabric. The enzymes in the capillary liquid adsorb at the capillary surface until an adsorption–desorption equilibrium is achieved between the surface enzyme concentration at the surface and the concentration in the capillary liquid. This adsorption–desorption equilibrium reads: kads ⎯⎯⎯ ⎯⎯ → ES S+E ← ⎯ kdes

[2.31]

where S is the number of sites at a substrate surface at which enzymes can be adsorbed, E is the enzyme concentration in the liquid, ES is the surface concentration of enzymes adsorbed at the surface, and kads and kdes are the adsorption rate constant and the desorption rate constant. To make the problem of suboptimal adsorption tangible, the next calculation example may be helpful. For a capillary in a fabric, filled with a solution of the enzyme pectinase, the characteristic properties of the capillary and the enzyme are: • capillary length, Lcap = 0.1 mm, i.e. the thickness of a yarn • capillary diameter, dcap = 2 μm, the intra-yarn pores • molecular weight of pectinase, ME = 50 kD = 50 kg mol−1


48


• specific weight of pectinase, ρE = 1300 kg m−3 and • pectinase concentration in the solution, CE = 5 g l−1, i.e. a frequently used concentration The surface area of the capillary Acap can be calculated by: Acap = π dcap Lcap

[2.32]

and the volume of the capillary by: Vcap =

1 2 π dcap Lcap 4

[2.33]

By using the values for the capillary length and capillary diameter as given above, Acap = 6.3 × 10−10 m2 and Vcap = 3.1 × 10−16 m3. If the enzyme is assumed to be a sphere, then its diameter can be calculated using: dE =

3

6ME NAV ρEπ

[2.34]

where NAV is the Avogadro number (= 6 × 1023 mol−1). From the values given above, the enzyme diameter is dE = 50 Å. This value is in accordance with values given in the literature. These enzymes adsorb at the surface of the capillary and it is suggested here that they adsorb in a simple rectangular array, which means that the maximum number of enzymes adsorbed at the surface nmax is given by: nmax =

A dE2

[2.35]

in which A is the surface area in m2. Thus, nmax = 4 × 1016 molecules per m2. The maximum surface concentration of enzymes, ΓE,max can be calculated by:

* E, max =

nmax NAV

[2.36]

For this case we calculated ΓE,max = 6.7 × 10−8 mol m−2. Therefore, the capillary surface area of 6.3 × 10−10 m2 can contain at maximum 4.2 × 10−17 mol pectinase. With an enzyme concentration CE of 5 g l−1 in the capillary liquid and a capillary liquid volume of 3.1 × 10−16 m3, the number of moles of pectinase in the liquid is calculated as 3.1 × 10−17. This example shows that the availability of pectinase in the capillary liquid is of the same order of magnitude as the number of moles of pectinase that can be adsorbed at the capillary wall. However, for enzymes, there is always an adsorption– desorption equilibrium. From the calculations made here, it is clear that the



49

enzyme concentration in the capillary liquid decreases immediately as a result of the adsorption process, leading to a lower equilibrium concentration at the surface. If the capillary liquid is not refreshed this results in a surface concentration that is lower than can be expected for particular enzyme concentrations in the liquid. There will be some transfer of enzymes from the inter-yarn liquid to the intra yarn liquid by diffusion. However, this is a relatively slow process as shown above and therefore it is expected that the adsorption process becomes controlled by diffusion. In most models describing the adsorption of enzymes at a surface, it is assumed that the enzyme concentration in the liquid bulk does not change as a result of adsorption. However, this assumption is only valid if the bulk liquid volume is much larger than the available surface. In capillary systems it is the other way around, the bulk liquid volume is much smaller than the available surface, and, therefore, the assumption of a constant enzyme concentration in the liquid can not be made. This phenomenon leads to a more complex model.

2.6.1 Modelling the adsorption limitation in textile pores The adsorption of enzymes at a substrate surface is shown schematically in Fig. 2.12. The rate equation for the enzyme concentration at the surface ΓES follows from the adsorption–desorption equilibrium, equation [2.31]: AC

d* ES = kadsVCCE AC * S − kdes AC * ES dt

[2.37]

where AC is the surface area of the capillary wall and VC is the volume of the capillary. It is assumed that this volume is totally filled with the enzymatic solution, so VC is also the volume of the enzymatic solution in the Enzymatic solution

E

E E

E

CE

G ES GS

E

E E

S

E

S

kads

S

S

S

E S

kdes

S

S

S

S

Substrate surface

2.12 A schematic view of the adsorption of enzymes at a substrate surface.


50


capillary. CE is the enzyme concentration in this liquid. ΓS and ΓES are the surface concentrations of the available sites for enzyme adsorption and the adsorbed enzymes, respectively. The mass balance for the total amount of enzymes in the system is: VCCE = VCCE,0 − ACΓES

[2.38]

where CE,0 is the initial concentration of enzymes in the solution. Thus, the amount of enzymes in the solution equals the initial amount of enzymes present in the solution minus the amount that has been adsorbed at the capillary surface. The number of available sites for enzyme adsorption can be related to the maximum amount of enzymes that can be adsorbed: ΓS = ΓES,max − ΓES

[2.39]

The number of vacancies S, varies from ACΓES,max for ΓES = 0 until 0 for ΓES = ΓES,max. In the latter instance, all places are occupied by enzymes. From equation [2.39], the equations [2.37] and [2.38] can be rewritten as: d* ES = kadsVCCE ( * ES,max − * ES ) − kdes * ES dt CE = CE,0 −

AC * ES VC

[2.40] [2.41]

Substitution of equation [2.41] into equation [2.40] and simplifying gives: d* ES 2 = kadsVCCE,0 * ES,max − ( kads AC * ES,max + kadsVCCE,0 + kdes ) * ES + kads AC * ES dt [2.42] This equation describes the dynamics of the adsorption–desorption process of the enzymes in a capillary pore of fabrics. The equation takes the form: dΓ ES 2 = γ + β Γ ES + αΓ ES dt

[2.43]

α = kads AC β = − ( kads AC Γ s, max + kadsVCCE, 0 + kdes ) γ = kadsVCCE, 0 Γ ES, max

[2.44]

where

Equation [2.43] is the differential equation of Ricatti. Although the equation looks rather simple, solving it is quite complex. Here we will only give the solution of the equation with the initial conditions t = 0, CE = CE,0 and ΓES = 0.



Γ ES = ψ −

2αψ + β ⎛β ⎞ −( 2αψ + β )t +α ⎜ +α ⎟e ⎝ψ ⎠

51

[2.45]

where

ψ=

−β + β 2 − 4αγ 2α

[2.46]

If the amount of adsorbed enzyme is small compared with the amount available in the liquid bulk, the model becomes simpler. In that case, it may be assumed that the enzyme concentration in the bulk is constant and equal to CE,0 so that equation [2.37] becomes: AC

d* ES = kadsVCCE,0 AC * E − kdes AC * ES dt

[2.47]

Substitution of equation [2.39] and simplification gives: d* ES = kadsVCCE,0 * ES,max − ( kadsVCCE,0 + kdes ) * ES dt

[2.48]

Equation [2.48] describes the dynamics of the adsorption process when the enzyme concentration in the capillary liquid does not change as a result of the adsorption process, which is the case when there is an excess of enzymes in the solution. This differential equation can be solved by separation of the variables. Its solution with the initial conditions t = 0, ΓES = 0 reads:

* ES =

kadsVCCE,0 * ES,max ⎡1 − e−( kadsVCCE,0 +kdes )t ⎤⎦ kadsVCCE,0 + kdes ⎣

[2.49]

For equations [2.45] and [2.49] we have performed some calculations to show clearly the effect of the decreasing enzyme concentration in the capillary liquid during adsorption of enzymes at the capillary wall. For the example we have used values calculated earlier: • • • •

AC = 6 × 10−10 m2 VC = 3 × 10−16 m3 CE,0 = 0.1 mol m−3 ΓES,max = 6.7 × 10−8 mol m−2

The values for the adsorption and desorption coefficients have been estimated to be: kads = 1015 mol−1 s−1 and kdes = 0.5 s−1. The results are expressed as the fraction of the adsorption sites that are occupied by enzyme molecules ΓES/ΓES,max as a function of time. Figure 2.13 shows the results


52

Advances in textile biotechnology 1.0

GES/GES,max

0.8 0.6 0.4 0.2 0.0 0

0.02 0.04 0.06 0.08

0.1 0.12 0.14 0.16 0.18 0.2 Time (s)

2.13 Model calculations for the adsorption dynamics of enzymes at a surface according to equation [2.49]. 0.06

GES/GES,max

0.05 0.04 0.03 0.02 0.01 0.00 0

1

2

3

4

5 6 Time (s)

7

8

9

10

2.14 Model calculations for the adsorption dynamics of enzymes at a surface according to equation [2.45].

if equation [2.49] is used. After 0.2 s, the adsorption is complete, which is the case when there is an excess of enzyme molecules in the capillary liquid. Figure 2.14 shows the results if equation [2.45] is used. In this case there is an equilibrium after 10 s at which the surface concentration is only 6% of the value that is theoretically achievable. So it can be concluded that a rapid refreshment of the capillary liquid during the adsorption is needed to prevent this undesired situation. This refreshment can be realized by squeezing using the mechanical energy. Therefore, the role of mechanical energy is to speed up the transfer of enzyme molecules to the pores of the fabrics and to prevent suboptimal adsorption because of exhaustion of the capillary liquid.

2.7

The application of ultrasound

It is known that ultrasound can accelerate wet textile processes to a large extent (Moholkar et al., 2004). This phenomenon is based on the mechanism


ΔPectin removal (%)


30

C

20

A, before pectinase incubation B, after pectinase incubation C, during pectinase incubation

53

B 10

A

0 0

2

4

6

8 10 Time (min)

12

14

16

2.15 Experimentally determined effect of ultrasound on the performance of enzymes.

of acoustic cavitation. The ultrasonic wave creates during its rarefaction period locally tiny bubbles that collapse during the compression phase. These collapses create micro liquid jets that can penetrate deeply into the pores of fabrics. Therefore, ultrasound can be seen as a source of mechanical energy to accelerate the mass transfer processes in the pores of fabrics. It has been found that ultrasound also accelerates the performance of enzymes in enzymatic textile treatment processes (Agrawal et al., 2007). Figure 2.15 shows the results of some experiments. To determine the effect of ultrasound on the removal of pectin by the enzyme pectinase, three experiments were performed: ultrasound applied before, after and during the enzymatic treatment. From Fig. 2.15, it is clear that applying ultrasound before the enzymatic treatment did not affect the enzymatic performance, whereas applying it after the enzymatic treatment led to a better performance. This is because of the more rapid removal of the enzymatic reaction products. Applying ultrasound during the enzymatic incubation resulted in a large effect on the performance. This can be explained by a fast transfer of the enzymes to the capillary liquid, a fast refreshment of the capillary liquid during the adsorption of enzymes at the surface and a fast removal of the reaction products. Therefore, ultrasound can be seen as a very effective and efficient way to apply mechanical energy to the enzymatic treatment of textiles and fabrics.

2.8

Future trends

It has been shown here that mechanical energy plays a dominant role in the performance of enzymes in textile treatment processes. Therefore, it is


54


not easy to translate the performance obtained on a laboratory scale to the performance on an industrial scale. This is because the deformation factor α is different in different systems. The best way would be to set up laboratory-scale experiments in such a way that they have an α-factor which is the same as its value on full-scale equipment. To prevent the phenomenon of enzyme exhaustion in the pores of the fabric, it is possible to enhance the enzyme concentration in the enzyme padding system. However, this will be very costly and therefore not a realistic option. The current installed base in textile treatment companies are based on traditional chemistry. It has been shown here that enzymes require another approach and, thus, it is expected that a breakthrough in the application of enzymes will only occur if machine manufacturers develop special systems in which the squeezing factor is as high as possible. Incorporation of ultrasound in such new systems could lead to the desired performance.

2.9

Acknowledgements

The authors acknowledge the support of the Foundation for Engineering of Fibrous Smart Materials and of the Saxion Universities of Applied Sciences.

2.10

References

agrawal p.b., nierstrasz v.a., warmoeskerken m.m.c.g. and veer van der, m. (2007) ‘Apparatus for enzymatic and ultrasound treatment of textiles and method of treatment thereof’, Patent GB 2432585. agrawal, p.b., nierstrasz, v.a. and warmoeskerken, m.m.c.g. (2008) ‘Role of mechanical action in low-temperature cotton scouring with F. solani pisi cutinase and pectate lyase’, Enzyme and Microbial Technology, 42(6), 473–482. andersson, k.m. and hovmöller, s. (2000) ‘The protein content in crystals and packing coefficients in different space groups’, Acta Crystallographica Section D: Biological Crystallography, 56(7), 789–790. bird, r.b., stewart, w.e. and lightfoot, e.n. (1960) Transport phenomena, Wiley & Sons. bouwhuis, g.h., dorgelo, b. and warmoeskerken, m.m.c.g. (2009) ‘Textile pretreatment process based on enzymes, catalyst and ultrasound’, Melliand International, 15(4), 150–151. kreyszig, e. (1993) Advanced engineering mathematics, Wiley & Sons. lenting h.b. and warmoeskerken, m.m.c.g. (2001) ‘Mechanism of interaction between cellulase action and applied shear force, an hypothesis’, Journal of Biotechnology, 89(2–3), 217–226. moholkar, v.s., warmoeskerken, m.m.c.g., ohl, c.d. and prosperetti, a. (2004) ‘Mechanism of mass-transfer enhancement in textiles by ultrasound’, AIChE Journal, 50(1), 58–64.



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nierstrasz, v.a. and warmoeskerken, m.m.c.g. (2003) ‘Process engineering and industrial enzyme applications’, Textile processing with enzymes, eds CavacoPaulo, A. and Gübitz, G.M., Woodhead Publishing, Cambridge, UK. van der donck, j.c.j., so, a. and frens, g. (1998) ‘The influence of stretching on salt release from porous yarns’, Tenside Surfactants Detergents, 35, 119–122. warmoeskerken m.m.c.g. and boom r.m. (1999) 90th AOCS Annual Meeting & Expo, May 9–12, Orlando, USA. warmoeskerken m. m. c. g., van der vlist p., moholkar v. s. and nierstrasz, v. a. (2002) ‘Laundry process intensification by ultrasound’, Colloids and Surfaces, A: Physicochemical and Engineering Aspects, 210(2–3), 277–285.


3 Smart textiles and biomaterials containing enzymes or enzyme substrates E. W E H R S C H Ü T Z - SIGL, A. H A S M A N N and G. M. GU E BI T Z, Graz University of Technology, Austria

Abstract: The high specificity of enzymes can be exploited for the design of smart materials. When immobilized on materials such as fabrics, enzymes can impart novel sophisticated functionalities ranging from antimicrobial effects to wound healing, self-cleaning or self-detoxifying properties. A variety of enzymes (in particular oxidoreductases and hydrolases) have been attached or incorporated into functional materials with distinct strategies which are reviewed here. In addition, enzymes can be used as triggers to impart bioresponsive properties to materials containing specific elements susceptible to modification by these biocatalysts. Thus the controlled release of functional molecules such as drugs, antimicrobial substances or perfumes from materials can be achieved. Key words: smart materials, enzymes, bioresponsive materials, controlled release.

3.1

Introduction

Today’s smart materials are expected to detect changes in their environment and respond with specific actions. Many ideas for such functional materials have been inspired by nature. The skin of a dolphin displays hydrolytic enzymes on the surface and is thus self-cleaning. In detail, a zymogel on the skin of Globicephala melas hydrolyses adhesive glycoconjugates secreted by biofouling micro-organisms thereby preventing their settlement (Baum et al., 2001). Mimicking this strategy, coatings containing proteases (i.e. subtilisin) have been developed for use in self-cleaning textiles (Tong et al., 2008) or antifouling membranes (Chen et al., 1992) whereas textiles self-decontaminating from warfare agents and pesticides or toxic microbes have already become reality (Edwards and Goheen, 2006). In several areas, smart materials are constructed such that they respond to triggers (e.g. enzymes) allowing a controlled release of active agents (such as drugs and perfumes). Many active agents used in pharmaceutics, food and agriculture require temporal stabilization and protection against degradation or oxidation (Hattori et al., 1995). In addition, the efficacy of such agents may be improved by increasing their solubility or by masking 56 © Woodhead Publishing Limited, 2010

Smart materials containing enzymes or enzyme substrates

57

unwanted properties, such as toxicity or bad taste, at least before the target environment is reached (Jain, 1989, Risch and Reineccius, 1995, Pothakamury and BarbosaCanovas, 1995, Poznansky and Juliano, 1984, Tomlinson, 1987, Vingerhoeds et al., 1994). Finally, a sustained or triggered release may be required. Therefore, a polymer, whether natural or synthetic, is judiciously combined with a drug or other active agent in such a way that the active agent is released from the material in a predesigned manner. Drug delivery and controlled release are of particular interest in cancer therapy and the treatment of many endocrinological disorders (Cox, 1993). For drug release, providing control over drug delivery can be the most important factor at times when traditional oral or injectable drug formulations cannot be used. A more effective therapy can be achieved while the risk for both under- and overdosing can be reduced. Other advantages of using controlled-delivery systems can include the maintenance of drug levels within a desired range, the need for fewer administrations, optimal use of the drug in question, and increased patient compliance. The ideal drug delivery system should be inert, biocompatible, mechanically strong, comfortable for the patient, capable of achieving high drug loading, safe from accidental release, simple to administer and remove, and easy to fabricate and sterilize. The field of controlled release, however, finds increasing application also in cosmetics and household materials (e.g. Joshi, 1996, Schaeffer and Brooks, 1992, Withenshaw, 1996), food (Risch and Reineccius, 1995, Yolles, 1973), agrochemicals/crop protectants (e.g. Allan and Neogi, 1971, Cohen et al., 1977, Knight et al., 1995, Kuderna and Saliman, 1974, Ventakraman, 2000), fertilizers (e.g., Fersch and Stearns, 1976, Knight et al., 1995) and industrial and other reactants. In recent years, controlled drug delivery formulations and the polymers used in these systems have become very sophisticated. Carrier systems have been developed including particulate systems like nano- and microcapsules, nano- and microspheres and liposomes (Amidon, 1997, BrannonPeppas, 1995, Cox, 1993, Felgner, 2009, Risch and Reineccius, 1995). Materials used for these systems normally consist of natural or synthetic biocompatible polymers. These can be proteins, such as gelatin, albumin, casein, or fibrin (Banga, 1995, Bissery et al., 1984, Chen et al., 2009, Gupta and Hung, 1989, Narayani and Rao, 1996, Senderoff et al., 1991, Wolkoff, 1987, Yu et al., 1996). The release of the active agent may be constant over a long period, it may be cyclic over a long period, or it may be triggered by the environment or other external events such as pH, ionic strength, ultrasound or by enzymes. In the following we will discuss the construction, function and application of smart materials based on the display of enzyme activity or response to trigger enzymes.


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3.2

Smart materials containing enzymes

3.2.1 Wound dressings Cotton-based wound dressings were functionalized by immobilization of various biomolecules. These include peptides acting as inhibitors of human neutrophil elastase or enzymes to impart haemostatic or antibacterial properties (Edwards and Goheen, 2006). Similarly, woven polyester fabrics (Dacron) were activated with ethylenediamine followed by surface immobilization of thrombin. Thrombin functionalized fabrics lead to thrombus formation after 1 min whereas on control fabrics there was only blood protein adsorption (Phaneuf et al., 2005).

3.2.2 Functional textiles Enzyme immobilization onto textiles was used to create smart materials with novel properties such as self-detoxifying or anti-microbial activity. For military purposes, organophosphorous hydrolase was covalently immobilized on cotton for detoxification of organophosphorus warfare agents such as sarin, soman, tabun (Edwards and Goheen, 2006). Cotton was first esterified with glycine followed by coupling of the enzymes with carbonyldiimidazole and glutaraldehyde. Organophosphorous hydrolase (EC 3.1.8.1), organophosphorous acid anhydrolase (EC 3.1.8.2), diisopropylfluorophosphatase (EC 3.1.8.2), phosphotriesterases (PTE) (EC 3.1.8) and combinations of these enzymes were suggested for immobilization on cotton textiles within polyelectrolyte multilayers allowing degradation of a wide range of toxic agents. Catalytic enzyme-modified textiles for active protection from toxins (USPTO Application #: 20070014838, 2009). Textiles with antimicrobial properties have been produced by immobilization of a variety of enzymes. Attachment of alkaline pectinase, alphaamylase or laccase lead to antimicrobial fabrics retaining full activity for at least ten consecutive wash cycles (Ibrahim et al., 2007). Lysozyme has also been used for the construction of antimicrobial textiles and packaging materials. This muramidase hydrolyzes the β-1,4 glucosidic linkages between N-acetylmuramic acid and N-acetylglucosamine of the cell wall of many micro-organisms. Lysozyme was covalently attached to cotton fabrics which were activated via esterification with glycine and a glycine dipeptide. Higher activity recovery was measured when the glycine dipeptide was used as a spacer. Consequently, in models of lysozymeattached through Asp-87, Glu-7, Asp-119, Asp-18, and COOH-terminal Leu-129 to cellopentaose-(3) gly-O-6-glycyl-glycine no steric hindrance of the active site was seen (Edwards et al., 2000). Lysozyme was also covalently immobilized on wool fabric activated with glutaraldehyde. Compared with control fabrics, antimicrobial activity of lysozyme-functionalized wool © Woodhead Publishing Limited, 2010


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fabrics was demonstrated against Staphylococcus aureus and 43% of the activity was retained after five cycles of use (Wang et al., 2009).

3.2.3 Functional packaging materials For applications in the food industry, lysozyme (registered as a food additive) was incorporated into chitosan films. Controlled release of the enzyme led to 3.8 log cycles reduction in S. faecalis and 2.7 log cycles reduction in E. coli (Park et al., 2004). Besides lysozyme, oxygen-consuming enzymes were immobilized in food-packaging materials to prevent microbial growth. Immobilization of glucose oxidase and alcohol oxidase on polypropylene (PP) and polyethylene (PE) has been demonstrated to improve the shelflife of food products by preventing growth of moulds, yeasts and aeraobic bacteria (Labuza and Breene, 1989, Vermeiren et al., 1999). Apart from antimicrobial properties, enzymes have been incorporated into packaging materials to control off-flavours. Naringinase, composed of α-rhamnosidase and β-glucosidase was incorporated into cellulose acetate (CA) in order to reduce the bitter flavour in grapefruit juice by hydrolysis of naringin to naringenin and pruning (Soares and Hotchkiss, 1998).

3.2.4 Enzyme immobilization on fabrics Fabrics are increasingly gaining importance as supports for enzyme immobilization (Table 3.1) and thus only selected examples will be discussed in the following. Horseradish peroxidase (HRP, EC 1.11.1.7) immobilized on non-woven polyester fabrics in the presence of glutaraldehyde as a crosslinking agent retained 85% of its activity after 4 weeks of storage at 4 °C compared with 90% activity loss of the free enzyme under the same conditions (Mohamed et al., 2008). HRP can be used for the production of electrically conductive polyaniline having a well defined thickness. Therefore, HRP was covalently immobilized on radio-frequency (RF) plasma-activated polyethylene and the synthesis of polyaniline was demonstrated (Alvarez et al., 2003). Catalase (EC 1.11.1.6) is widely used in applications ranging from textile bleaching and electronics to sterilization of liquid food products for conversion of residual hydrogen peroxide to oxygen and water. Apart from various inorganic carrier materials (Costa et al., 2001, Gudelj et al., 2001), fabrics have been used for catalase immobilization. Catalase was photochemically immobilized on polyester (PET) and polyamide 6.6 using diallylphthalate or cyclohexane-1,4-dimethanoldivinylether as cross-linking agents. The immobilized enzyme was highly stable with a 3.5 higher activity after 20 cycles when compared with the free enzyme (Opwis et al., 2004, 2005). Another strategy for catalase immobilization on cotton fabric was based on oxidation of cotton by sodium periodate followed by covalent attachment of the enzyme (Wang et al., 2008). © Woodhead Publishing Limited, 2010


Polyester (PET) Polyethylene

Polyester (PET), polyamide 6,6

Peroxidase (HRP)

Catalase

Polyamide 6,6

Silk fibroin, viscose rayon, PET, polyamide 6, polypropylene

Silk fibroin

Cotton

Laccase

Glucose oxidase

Alkaline phosphatase

Lysozyme

Cotton

Polyester (Dacron)

Organophosphate hydrolase

Thrombin

Wool

Silk fibroin Polyamide 6,6

Tyrosinase

Cotton

Material

Enzyme

Table 3.1 Strategies for enzyme immobilization on fabrics

Ethylenediamine

(1) Esterification with glycine (2) Carbonyldiimidazole/glutaraldehyde

(1) Esterification with glycine (2) Carbonyldiimidazole/glutaraldehyde Glutaraldehyde

Low-temperature plasma

Various activation strategies

(1) Enzymatic hydrolysis (2) Glutaraldehyde and spacer

Glutaraldehyde

(1) Photochemical (2) Diallylphthalate or cyclohexane-1,4dimethanol divinyl ether Oxidation by sodium periodate

Glutaraldehyde Plasma

Activation/binding

Phaneuf et al. (2005)

Edwards et al. (2000)

Wang et al. (2009)

Edwards et al. (2000)

Asakura et al. (1989)

Asakura et al. (1992)

Silva et al. (2007)

Acharya et al. (2008) Pialis et al. (1996)

Wang et al. (2008)

Opwis et al. (2005)

Mohamed et al. (2008) Alvarez et al. (2003)

Reference


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Tyrosinase (EC 1.14.18.1) has recently found applications for grafting and coating protein-based textile materials such as wool and silk (Anghileri et al., 2007, Freddi et al., 2006, Jus et al., 2008, 2009, Sampaio et al., 2005). In chemical synthesis, it is used for the production of l-DOPA (3,4dihydroxyphenyl-l-alanine) from tyrosin which is used for the treatment of Parkinson’s disease. Tyrosinase immobilization not only allows the design of continuous processes reusing the enzyme but also stabilization of l-DOPA by carrier autodegradation (Acharya et al., 2008). Using glutaraldehyde as crosslinking agent, tyrosinase has been immobilized on fibroin (Acharya et al., 2008) and on nylon 6,6 (Pialis and Saville, 1998, Pialis et al., 1996). Apart from glutaraldehyde attachment, other approaches were based on tyrosinase entrapment in alginate, polyacrylamide and gelatin (Munjal and Sawhney, 2002). Laccase (EC 1.10.3.2) is used for a variety of applications including the treatment of textile dyeing effluents (Kandelbauer et al., 2004), elimination of off-flavors in food products (Schroeder et al., 2008) and for lignocellulose processing (Kudanga et al., 2008). For numerous applications laccase has been immobilized on inorganic carrier materials such as alumina pellets (Abadulla et al., 2000). Also, laccase has been bound to organic polymers (i.e. PEG) to obtain water-soluble immobilized proteins with enlarged molecular weight and modified sorption properties. Such constructs have been used in detergents to prevent dye transfer (Schroeder et al., 2006). A novel approach for laccase immobilization on polyamide 6,6 involved limited surface hydrolysis of polyamide 6,6 by protease to introduce functional groups. Thereafter, first the spacer 1,6-hexanediamine followed by laccase were attached by glutaraldehye (Silva et al., 2007). Glucose oxidase (EC 1.1.3.4) is widely used in many applications including novel approaches for bleaching of textiles (Buschle-Diller, 2001, Tzanov et al., 2001). For example, glucose oxidase was immobilized on viscose rayon, polyethyleneterephthalate, nylon-6, polypropylene and non-woven fabrics of silk fibroin (Asakura et al., 1992). Urease (EC 3.5.1.5) was immobilized on cellulose fabrics to remove urea in an electrodialysis cell. Immobilization was achieved by biotinylation of both the enzyme and the cellulose fibres (Magne et al., 2002).

3.2.5 Enzyme immobilization strategies Various strategies have been developed for incorporation of enzymes into polymers including entrapment (Kim et al., 2001b), covalent attachment (Dessouki et al., 2001) and adsorptive binding (Akgol et al., 2001). Polydimethylsiloxanes have been used for covalent entrapment of enzymes including lipases and proteases (Gill and Ballesteros, 2000a, 2000b, Kim et al., 2001a). The biocatalytic films and paints can be applied to a wide


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range of materials such as metals and plastics. Pronase and alpha-chymotrypsin were successfully immobilized with this approach and thermostability was even enhanced. When attached to filters, antifouling properties were imparted. Since conventional antifouling paints contain toxic agents, such bioactive coatings offer great opportunities for food-processing equipment, ship hulls, medical implants, ultrafiltration membranes, and many other devices. Plasma treatment has been used for activation of polyethylene (Alvarez et al., 2003) whereas PET and PA were activated photochemically before cross-linking with diallylphthalate or cyclohexane-1,4-dimethanol divinyl ether (Opwis et al., 2005). Polypropylene-based ion-exchange textiles were constructed by UV-induced graft polymerization for immobilization of urease coupled with N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride, N-cyclohexyl-N′-[β-(N-methylmorpholino)ethyl]carbodiimide p-toluenesulfonate or glutaraldehyde (Yeon and Lueptow, 2006). Another strategy involves activation of polypropylene membranes with polyaniline using ammonium persulfate as the oxidizer. This pretreatment greatly facilitated both adsorptive and covalent immobilization of proteins such as HRP (Piletsky et al., 2003). Limited enzymatic hydrolysis of polyamide by protease has been used to insert anchor groups for covalent immobilization of laccase using glutaraldehyde together with 1,6-hexanediamine as a spacer (Fig. 3.1) (Silva et al., 2007). In general, enzymatic surface modification is a new strategy to activate the surface of synthetic polymers including polyamide (Heumann et al., 2009), polyalkyleneterephthalates (Brückner et al., 2008, Eberl et al., 2008) and polyacrylonitriles (FischerColbrie et al., 2007). Highly specific interaction of the glycoproteins avidin or streptavidin to biotin can be exploited for enzyme immobilization on surfaces. Biotin binds almost irreversibly to streptavidin with a Ka = 1 × 1015 M−1, which is similar to that of covalent bonds (Wilchek et al., 1988). Use of this interaction involves multiple attachment of biotin to the target proteins (i.e. enzymes) which consequently can bind several molecules of avidin (Fig. 3.2). These interactions are exploited in bioanalytics or affinity chromatography involving labeled or immobilized avidin, respectively. Alternatively, avidin is bound to biotinylated surfaces followed by immobilization of biotinylated enzymes (Janolino and Swaisgood, 2002). Usually this process can be carried out under mild conditions, which are beneficial for preservation of enzyme activity. Using this approach, urease was immobilized on cellulose fabrics (Magne et al., 2002). Similarly, trypsin was immobilized on cellulose beads biotinylated with sulfosuccinimidyl-6-(biotinamido) hexanoate (NHS-LC-biotin) resulting in a biotin content of 1.15 μmol g−1 (Janolino and Swaisgood, 2002). Specific binding modules of polymer-modifying enzymes also offer a potential for enzyme immobilization. Many (hemi)cellulolytic enzymes © Woodhead Publishing Limited, 2010


O HO

63

H

C

C

H N H n

N

O

Enzymatic surface hydrolysis NH2 NH2

O HO

C

C

OH

H2N

H N H n

O

NH2

NH

GA

NH

GA

NH

GA

GA

(a) Attachment of enzymes (b)

3.1 Immobilization of enzymes on polyamide activated by limited enzymatic surface hydrolysis. Enzymes are bound to the resulting amino groups by using glutaraldehyde (GA) via a spacer such as hexanediamine (a) or directly (b).

contain carbohydrate-binding modules (CBM) in addition to the catalytic module. These CBMs are responsible for controlled adsorption of the enzyme on the polysaccharides. Consequently, CBMs have been employed for targeting purposes (e.g. perfumes in detergents) and fused to enzymes of interest they provide another specific tool for enzyme immobilization on polysaccharide-based materials. (Kwan et al., 2005, Shoseyov et al., 2006). In a similar approach, polyhydroxyalkanoate (PHA) binding proteins have been fused to proteins of interest allowing their immobilization on ‘bioplastics’ and probably on other hydrophobic materials (Moldes et al., 2004).

3.3

Smart materials responding to enzymes as triggers

3.3.1 Controlled-release systems and mechanisms In most traditional systems, the control of release and the stabilization of the agent is based on encapsulation (Brownlee and Cerami, 1979, Cox, 1993, © Woodhead Publishing Limited, 2010

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Advances in textile biotechnology Polymer-NH2 Biotinylation

+ Avidin

+ Biotinylated enzymes

3.2 Immobilization of biotinylated enzymes.

Fischelghodsian et al., 1988, Hsieh et al., 1981, Kim et al., 1994, Kost et al., 1989, Pothakamury and BarbosaCanovas, 1995, Risch and Reineccius, 1995, Vingerhoeds et al., 1994). There are three primary mechanisms by which active agents can be released from this kind of delivery system: diffusion, swelling followed by diffusion and degradation. Many controlled-release systems are designed to create a sustained release by diffusion or swelling (Bagshawe et al., 1988, Cox, 1993, Merlin, 1991, Risch and Reineccius, 1995, Vingerhoeds et al., 1993). Diffusion occurs when a drug or other active agent passes from the polymer matrix into the external environment through the polymer that forms the controlled-release device. The only structure effectively limiting the release of the drug is the polymer layer surrounding the reservoir. Since this polymer coating is essentially uniform and of a non-changing thickness, the diffusion rate of the active agent can be kept fairly stable throughout the lifetime of the delivery system. The diffusion can occur on a macroscopic scale, as through pores in the polymer matrix, or on a molecular level, by passing between polymer chains. The diffusion controlled systems, which require hydrophilic polymers such as hydroxyethylcellulose and sodium carboxymethylcellulose, allow a sustained release of drugs, but do not permit a rigorous control because the release rate is not constant. Although the polymeric membranes or matrices do not change during the release process, uptake of solvent leads to polymer expansion and thus



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different release kinetics for swelling-controlled systems. For drug delivery systems, mostly hydrogels are used. Hydrogels are glassy in their dehydrated state. Drug release generally involves simultaneous absorption of water and desorption of drug via swelling controlled mechanism (Peschier et al., 1993, Rao and Devi, 1988). The presence of solvent in a glassy polymer causes the development of stresses that are accommodated by an increase in the radius of gyration and end-to-end distance of polymer molecules, which is seen macroscopically as swelling. The movement of solvent molecules into the dry (glassy) polymer matrix takes place with a well-defined velocity front and a simultaneous increase in the thickness of the swollen (rubbery) region with time in the opposite direction. The existence of a slow macromolecular relaxational process in the swollen region is believed to be responsible for the observed non-Fickian behaviour (Kim and Lee, 1992). However, most systems are designed to control the kinetics of release of the active agent in a site- and time-dependent manner. In this instance, the release of the active compounds can be triggered by the local conditions in the target environment (Albin et al., 1987, Brownlee and Cerami, 1979, Fischelghodsian et al., 1988, Kost et al., 1989, Hsieh et al., 1981, Jeong et al., 1985, Kim et al., 1994, Merlin, 1991, Pinnaduwage and Huang 2002) or by externally applied triggers (Amidon, 1997, Brannon-Peppas, 1995, Cox, 1993, Jain, 1989, Pothakamury and BarbosaCanovas, 1995, Vingerhoeds et al., 1994) (Fig. 3.3). Much effort is focused on creating biodegradable polymers for enzymatic drug-delivery systems that permit release of the entrapped drug only during degradation of the polymer matrix. For this to occur, the polymer matrix must be vulnerable to biodegradation by a component in the surrounding media. In a model system using dextran, enzymatic degradation of dextran by dextranase led to the release of insulin in a controlled manner (Moriyama and Yui, 1996). A similar system used polycaprolactone copolymerized with PEG to form enzymatically degradable gels which can be degraded by lipase (Rice et al., 2006). Another possibility is the preparation of films of different enzymatically degradable polymers such as chitosan (Yomota et al., 1990). Chemicals of interest were incorporated into chitosan films and degradation by lysozyme and release of the loaded chemicals were investigated. The enzymatic degradation rate of the films was dependent on the degree of deacetylation of chitosan used and decreased with an increase in deacetylation. The chemicals were released only in the presence of lysozyme as trigger, and their release rates were controlled by the degradation rate of the films (Yomota et al., 1990). Other natural materials such as gelatine, collagen, alginate or fibrin were described recently for various controlled-release applications (Patel and Mikos, 2004, Young et al., 2005). Apart from these polymers from natural


66

Advances in textile biotechnology pH release

Enzymes

Microcapsule

Drug release

Microcapsule

Magnetic release Moisture

3.3 Release of an active agent triggered by different external conditions.

sources, synthetic hydrogels incorporating defined biological moieties that can be degraded enzymatically are also under intensive investigation. One way to fabricate this type of hydrogel is to incorporate peptide substrates for enzymatic hydrogel formation and degradation (Hu and Messersmith, 2003, Lutolf et al., 2003a, 2003b).

3.3.2 Covalent attachment of enzyme substrates Beside encapsulation, covalent attachment of the active agent to the carrier system is often used (Albrecht et al., 1997, Dosio et al., 1997, Fiume et al., 1986, Franssen, 1993, Molema and Meijer, 1994, Narayani and Rao, 1996, Pirrung and Huang, 1996, Trouet et al., 1982). For binding the active agents, carriers with high affinity and selectivity like biotin-, lipid-, phosphate-, and sulfate-binding proteins, as well as bacterial periplasmic proteins, lectins, serum albumins, immunoglobulins, and inactivated enzymes are of great interest. Several systems are used for the specific targeting (homing) of the entrapped agents (Poznansky and Juliano, 1984, Tomlinson, 1987, Felgner, 2009, Vingerhoeds et al., 1994, Hirabayashi et al., 1996), whereas most systems are designed to control the kinetics of release of the active agent in a site- and time-dependent manner (Albin et al., 1987, Amidon, 1997, Bagshawe et al., 1988, Brannon-Peppas, 1995, Brownlee and Cerami, 1979, Cox, 1993, Fischelghodsian et al., 1988, Hsieh et al., 1981, Jain, 1989, Jeong et al., 1985, Merlin, 1991, Pothakamury and BarbosaCanovas, 1995, Risch and Reineccius, 1995, Vingerhoeds et al., 1993, 1994). The application of carrier components that selectively and reversibly bind the active agent with high affinity offers additional possibilities to stabilize the agent and improve the control of its release (Cohen et al., 1977, 1979, Hattori et al., 1995, Mitchell, 1986, Schaeffer and Brooks, 1992). A possible application for such release systems is chemotherapy, because of the severe side effects caused by the toxicity of traditional cytostatic agents towards normal tissues. Therefore, the site-specific activation of non-toxic prodrugs in tumours has been proposed in order to enhance their selectivity for the killing of cancer cells. Those prodrugs comprise trigger, linker and



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effector units releasing the effector unit after selective trigger activation via spontaneous chemical breakdown (Tranoy-Opalinski et al., 2008). A broad field of application is the linking of active agents on various surfaces such as wound dressings in medicine. A controlled-release system of antibiotics is described by Suzuki et al. (1997) to provide adequate local antibiotic concentration against an infection. A delivery system was created using a polyvinylalcohol (PVA) hydrogel carrying an enzymatically degradable peptide linker for binding gentamicin. In vitro experiments with this wound dressing showed that proteases from infected wounds cleaved the linker and gentamicin was released. The exudates from non-infected wounds had no hydrolytic activity towards the linker. The same group described a thrombin-sensitive peptide linker system using the elevation of thrombinlike activity in infected wound exudates as the key component of this system (Tanihara et al., 1998).

3.4

Conclusion and future trends

In summary, the high specificity of enzymes can be exploited for the design of smart materials in two ways. On the one hand, enzymes can impart novel sophisticated functionalities to materials ranging from antimicrobial effects to self-cleaning or self-detoxifying properties. On the other hand, enzymes can be used as triggers to impart bioresponsive properties to materials containing specific elements susceptible to modification by these biocatalysts. Both concepts have already been exploited for the construction of smart fabrics. An instructive good example are wound dressings containing enzymes to impart haemostatic or antibacterial properties or containing antibiotics released upon contact with enzymes from infected wound as triggers. As a result of advances in the discovery of novel enzymes and in genetic engineering, highly stable biocatalysts with increased efficiency for the target substrates are currently becoming available. Together with novel fabrication technologies this should lead to implementation of smart materials containing enzymes or enzyme substrates described here in various industrial sectors. Also, environmental legislation will favour materials that lead to a saving of resources by use of controlled release and biocatalytic reactions. Finally, customer/society benefits such as reduced resistances to antibiotics resulting from the controlled application of drugs or hygienic benefits owing to self-cleaning or antimicrobial properties will be a driver to bring these smart materials to the markets.

3.5

Acknowledgements

This work was financed by the SFG, the FFG, the city of Graz and the province of Styria within the MacroFun project and supported by the European COST868 program.


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3.6

References

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4 Enzymatic hydrolysis and modification of core polymer fibres for textile and other applications C. SI LVA and A. C AVAC O - PAU L O, University of Minho, Portugal; and V. A. N I E R S T R A SZ, Ghent University, Belgium

Abstract: Recent progress and new developments in enzymatic surface hydrolysis of the core textile polymers poly(ethylene terephthalate) (PET) and polyamide (PA) are discussed. The process produces modifications that yield charged functional groups at the surface of these synthetic materials. The use of cutinases, proteases and amidases for modification of the surface of fibres is described and possible applications for the bio-modified PET and PA are explored. Key words: biotechnology, textiles, enzymes, functionalization, surface hydrolysis, polymers.

4.1

Introduction

Synthetic fibres form an important part of the textile industry, the production of polyester alone surpassing that of cotton. In 2007, the global annual production of fibres and yarns was estimated to be 30.7 million tonne of polyester, 4.0 million tonne of polyamide, 2.4 million tonne of acrylics, and 26.1 million tonne of cellulose (Oerlikon, 2008). The production volume of polyester and polyamide fibres and yarns justifies research into effective production. A characteristic of synthetic fibres is their low hydrophilicity. The low hydrophilicity affects the processability of the fibres because the surfaces are not easily wetted, thus impeding the application of finishing compounds and a hydrophobic material hinders water from penetrating into the pores of fabric during production. Biotechnology has a vast potential in the industrial production of textile materials. The global market for industrial enzymes increased from US$ 1.5 billion in 2000 to US$ 2.25 billion in 2007, and it has been forecasted to grow to US$ 2.75 billion in 2012. Approximately 10% of industrial enzymes find their application in industrial textile processing. The increasing importance and potential of biotechnology and enzyme technology in textiles production has been assessed in the last few years (Araújo et al., 2008; Gübitz and Cavaco-Paulo, 2003; Lenting, 2004; Nierstrasz, 2009; Nierstrasz and Warmoeskerken, 2003). In industrial textile pre-treatment of natural 77 © Woodhead Publishing Limited, 2010

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fibres, such as depilling, scouring or ageing of cotton, enzyme technology has already proven to be very profitable. Enzyme technology contributes to the decoupling of economic growth and the use of natural resources leading to a more sustainable way of life; it enables us to produce more with less (Wu, 2008). Nevertheless, even though enzymes are produced by organisms to degrade or synthesize natural substrates, the use of enzymes is not limited to natural materials. Quite recently it has been demonstrated by several research groups that enzymes are also able to modify the surfaces of synthetic textile materials such as polyester and polyamide (for a review see for example Gübitz and Cavaco-Paulo, 2008). In this chapter, the focus is on novel promising enzyme applications for surface modification and hydrolysis of polyester and polyamide fibres. Classical methodologies to improve fibre hydrophilicity, like alkaline or acid hydrolysis, lead to the deterioration of fibre properties such as irreversible yellowing and loss of resistance (Cribbs and Ogale, 2003; Gübitz and Cavaco-Paulo, 2003; Miller and Wilmington, 1958; Shukla et al., 1997; Silva, 2002). These processes are based on an ‘all-or-nothing’ mechanism where the adsorption of high concentrations of chemicals can destroy the surface of the polymers and negatively affect the favourable bulk properties as well. Recent studies clearly indicate that the modification of synthetic polymers with enzymes is an environmentally benign method. Since enzymes are large molecules their action is restricted to the surface of the fibres maintaining their favourable bulk properties. The major advantages of enzymes in polymer modification, compared with chemical methods, are milder reaction conditions and highly specific non-destructive transformations, targeted to surfaces leading to less fibre damage (Gübitz and Cavaco-Paulo, 2008).

4.1.1 Polyester Polyesters are a category of polymers containing an ester group, and can be natural polyesters such as cutin in plants, but most commonly the name polyester refers to poly(ethylene terephthalate). Poly(ethylene terephthalate) (PET) is the most important synthetic fibre owing to its excellent fibre properties. Synthesis of polyester is achieved by a polycondensation reaction of a dicarboxylic acid and a diol, or a polycondensation of molecules containing both a carboxylic acid and an alcohol group. Linear polyester was first synthesized by Wallace Carothers (1928) at Dupont, who discovered that such polycondensation reactions could be used to produce fibres. However, the fibres had a low melting point and poor hydrolytic stability. Carothers turned to polyamide research. Advancing the early research of Carothers, John Rex Whinfield and James Tennant Dickson, at the Calico Printers Association, used aromatic polyesters and discovered PET in 1941.


Enzymatic hydrolysis and modification of core polymer fibres

79

The fibres had a high melting point (265 °C) and good hydrolytic stability. Commercial production of this fibre started in 1949 at ICI under the name Terylene, and was continued later by Dupont. The raw materials for PET production are terephthalic acid (TPA) or dimethyl terephthalate (DMT) and ethylene glycol. There are two production routes. TPA is directly esterified to bis-(2-hydroxyethyl)terephthalate (BHET) whilst forming water. Next, the polycondensation is performed under continuous release of ethylene glycol. Starting with DMT, the first step is a transesterification leading to (BHET), accompanied by the formation of methanol, after which the polycondensation is performed. Properties and applications of poly(ethylene terephthalate) PET is a strong and stiff material that is resistant to chemicals, with good abrasion resistance, a low water uptake (and consequently quick drying), good resistance to shrinking and stretching and low wrinkling properties. Depending on the processing and thermal history, PET appears as an amorphous or a semi-crystalline material (e.g. during drawing of the fibres the polymer chains become oriented resulting in more crystalline fibres). Amorphous PET is transparent and finds its applications in food packaging and soft drink bottles. Semi-crystalline PET has excellent thermal stability, and is used in, for example, housings for electrical appliances and recording tapes. The majority of global PET production is for synthetic fibers, with applications ranging from e.g. apparel, curtains, and carpets to e.g. sails, ropes, nets, tyre cords and fibre-reinforced materials.

4.1.2 Polyamide Polyamide (nylon) represents a family of synthetic polymers first produced in 1935 by Wallace Carothers at DuPont (Guillen, 1986; Stevens, 1999). The first product was a polyamide-bristled toothbrush (1938), followed more famously by women’s polyamide stockings (1940). Polyamide 6.6 was the first commercially successful polymer and it was intended to be a synthetic replacement for silk. This fibre was used for parachutes production after the USA entered World War II in 1941, making stockings hard to find until the war’s end (Burkinshaw, 1995; Guillen, 1986; Moncrieff, 1975). The US Federal Trade Commission defines nylon fibre as ‘a manufactured fibre in which the fibre-forming substance is a long-chain synthetic polyamide in which less than 85% of the amide linkages (—CO—NH—) are attached directly to two aliphatic groups’ (Yang, 1989). Polyamide (PA) is a polymer, which contains recurring amide groups (R—CO—NH—R′) as integral parts of the main polymer chain. Synthetic polyamides are produced by a condensation reaction between monomers,


80


Table 4.1 Examples of polyamide compositions (adapted from Yang, 1989) Nylon-6,10 Nylon-6 Nylon-11 Nylon-6,T mX,D,6 Nylon-6,6–6,10 (60 : 40)

—[—NH—(CH2)6—NH—CO—(CH2)8—CO—]— —[—NH—(CH2)5—CO—]— —[—NH—(CH2)10—CO—]— —[—NH—(CH2)6—OCO—(C6H4)—OCO—]— —[—NH—CH3(C6H3)—NH—CO—(CH2)4—CO—]— —[—NH—(CH2)6—NH—{—CO—(CH2)4—CO—}60={—CO— (CH2)8—CO}40—]—

in which the linkage of the molecules occurs through the formation of the amide groups. Polyamides may be made from a dicarboxylic acid and a diamine, or from an amino acid or its lactam that is able to undergo selfcondensation. One of the most common polyamides, nylon 6.6, is produced by the interaction of hexamethylenediamine and adipic acid. When εcaprolactam is the starting material, nylon 6, which consists more than 100 units of 6-aminohexanoic acids is obtained. Examples of polyamide compositions are presented in Table 4.1, but this chapter will emphasize the modification of nylon 6.6 fibres. Polyamide properties and applications The high strength, elasticity, lustre, abrasion resistance, dyeability and shape-holding characteristics of nylon make it suitable for many applications. In clothing and home furnishings, nylon is an important fibre, especially in socks, stockings, lingerie, stretch fabrics, sporting goods, carpets, luggage, and materials used in upholstering of furniture. Industrial uses of nylon fibre include automobile tyres, ropes, seat belts, parachutes and substrates for coated fabrics such as artificial leather, shower and fire hoses, and disposable clothes for the health-care industry. In addition, engineering plastics made of nylon exhibit high melting points; strength, toughness and chemical inertness, and are used as bearings, zippers, gears, automobile fan blades and pulleys (Yang, 1989).

4.2

Enzymatic hydrolysis and modification of poly(ethylene terephthalate) and polyamide: basic principles, methods and technologies

4.2.1 Modification of poly(ethylene terephthalate) fibres PET fibres have numerous favourable properties (see section 4.1.1) compared with other textile fibres, but there are considerable disadvantages, such as hydrophobicity, build-up of electrostatic charge, pilling tendency



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and difficult oil stain release. The hydrophobicity of the fibres hinders water penetration into the pores of fabric, resulting in bad comfort properties because perspiration is not absorbed by the material. From the consumer’s point of view improved hydrophilicity results in a number of improvements: improved washability, improved removal of oily stains, and enhanced comfort. In the textile industry various finishes, such as anti-static finishes, and colouring agents are applied. Owing to the crystallinity and hydrophobicity of the PET fibres, chemicals can hardly penetrate into the fibre or physically bind to the fibre surface. The absence of reactive groups at the PET surface hinders chemical reactions at the surface. Various techniques are available to increase hydrophilicity of PET. Copolymerization (Pang et al., 2006) is a suitable technology to adjust PET properties, though effects are not limited to the surface of the fibre, the bulk characteristics are modified as well. Another technique to make PET more hydrophilic is an alkaline treatment at high temperature. Polyester bonds are hydrolysed and, consequently, hydrophilicity is increased. However, because NaOH diffuses into the fibre, the favourable bulk properties of polyester, particularly the strength, are also affected. The alkaline treatment also results in pitting corrosion (Brueckner et al., 2008; Donelli et al., 2009; Kim and Song, 2006), not to mention increased energy consumption, the necessity to neutralize the pH of the liquid, and increased water consumption. Plasma treatments have been developed to modify the surface of polyesters (Höcker, 2002; Morent et al., 2007, 2008), but the technique is rather expensive and not yet generally accepted in the textile industry. The technique is non-specific, relatively difficult to control, and ageing can occur as a result of reorientation or migration. In the past decade new environmentally benign and specific processes have been developed based on modern enzyme technology. Synthetic materials, such as PET fibres, have generally been considered resistant to biological degradation. However, the application of enzymes is not limited to biological materials. Fairly recently it was established that enzymes are also able to act on synthetic textile materials. Various research groups assessed the potential of laccases, lipases, polyesterases (serine esterase) and cutinases in the oxidation or hydrolysis of PET textile materials with the aim of modifying and eventually functionalizing the material (e.g. Alisch-Mark et al., 2004, 2006; Araújo et al., 2007; Brueckner et al., 2008; Donelli et al., 2008, 2009; Feuerhack et al., 2008; Hsieh and Cram, 1998; Kim and Song, 2006, 2008; Kontkanen et al., 2006; Liebminger et al., 2007; Liu et al., 2008; Miettinen-Oinonen et al., 2002, 2004; Müller et al., 2005; Nimchua et al., 2006; O’Neill and Cavaco-Paulo, 2004; Silva and Cavaco-Paulo, 2008; Silva et al., 2005a; Vertommen et al., 2005; Wang et al., 2008; Yoon et al., 2002).


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Laccase Laccase (EC 1.10.3.2) is a copper-containing oxireductase found in white rot fungi. Laccases are non-specific enzymes, oxidizing primarily diphenol groups using molecular oxygen as an electron acceptor. The application of laccases has been extensively studied to increase strength of paper and board via lignin bonding, but it also has applications in the textile industry in the bleaching of denim and in wastewater decoloration. Laccases are often used in combination with a mediator such as 1-hydroxybenzotriazole (HBT) or 2,2′-azinobis-(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS). The purpose of the mediator is twofold: (i) owing to their size (65–70 kDa) laccases cannot penetrate deeply into fibres, smaller mediators do not necessarily have this limitation, and (ii) because of their low redox potential (0.5–0.8 V) laccases are not able to oxidize compounds with high redox potentials (larger than 1.5 V). Several studies (e.g. Miettinen-Oinonen et al., 2002, 2004) reported increased hydrophilicity of PET treated with laccases that caused surface oxidation of the material. However, Liu and co-workers hypothesised that the improved hydrophilicity when treating PET with a laccase from Streptomyces coelicolor in combination with a mediator (Denilite II Assist) and a non-ionic surfactant (Triton X-100) might to a large extent be caused by protein adsorption on the surface. After a protease treatment to hydrolyse adsorbed laccase, and a thorough rinsing processes to remove the proteins (laccase) from the surface, the hydrophilicity decreased dramatically (wetting time using a water drop test was 165 s for the laccase treated sample compared with 170 s for the sample treated with surfactant only, before protease treatment and rinsing the wetting time was 20 s) and almost lost the gain in hydrophilicity resulting from the treatment. The marginal improvement in hydrophilicity and the relatively high laccase concentration are significant hurdles in the actual application of laccases in modification of PET. Laccase is able to modify surfaces with aromatic groups; however, no real evidence for successful surface modification of PET has been presented so far.

Cutinase In contrast to laccases that oxidize the PET surface, lipases, polyesterases, and cutinases increase hydrophilicity by actual hydrolysis of PET (Fig. 4.1). Lipases, polyesterases, and cutinases are all carboxylic ester hydrolases. Cutinases from Fusarium solani pisi, Fusarium oxysporum and Thermobifida fusca are most frequently studied in enzymatic hydrolysis or modification of PET. The structure and properties of cutinases are well described (Carvalho et al., 1998, 1999). In contrast to lipases, cutinases do not require interfacial activation, the active site is accessible because it does not have


Enzymatic hydrolysis and modification of core polymer fibres O

O

O C

O

C O CH2CH2 O C

Cutinase Polyesterase Lipase

1

2

3

TPA

1

4

BHET

1

3

or

2

4

2

O

83

O

C O CH2 CH2 O C

3

4

MHET

4.1 Enzymatic hydrolysis of poly(ethylene terephthalate) by cutinase, polyesterase and lipase.

a lid, the oxyanion hole is preformed but considerably flexible in solution. Today, cutinases seem to have a large potential in the enzymatic surface modification of PET (Nierstrasz, 2009). The optimum pH and temperature for cutinase from Fusarium solani pisi are pH 8–8.5 at around 25 °C; above 35 °C the activity decreases rapidly. The relatively low reaction rate and limited temperature stability hinder industrial application of cutinases. Araújo et al. (2007) succeeded in increasing the reaction rate of cutinases on PET by modifying (enlarging) the active site through site-directed mutagenesis of cutinase from Fusarium solani pisi, whereas researchers from Novozymes improved the temperature stability to as high as 65–80 °C (Liu et al., 2008). An advantage of a cutinase or lipase treatment is that it does not result in pitting corrosion, as seen in alkaline treatments, but a more or less homogeneous surface treatment of PET (Brueckner et al., 2008; Donelli et al., 2009; Kim and Song, 2006, 2008). The homogeneity of the treatment depends on enzyme, enzyme concentration and incubation time. Using X-ray photoelectron spectrometry (XPS), Vertommen et al. (2005) demonstrated that cutinases strongly adsorb to the PET surface. This hinders proper determination of surface properties, such as wetting characteristics or formation of carboxyl or hydroxyl groups in the surface. Proper procedures have been developed to remove the adsorbed enzyme using proteases (Donelli et al., 2009; Liu et al., 2008) or using a thorough washing and extraction method (Brueckner et al., 2008). After protein (cutinase) removal from the surface it is possible to properly evaluate the surface modification. Donelli and co-workers (2009) measured water contact angles on PET films treated with cutinase, ∼75° for untreated PET and ∼58° for cutinase-treated PET. Brueckner et al. (2008) and Liu et al. (2008) measured wetting times


84


using the drop method; the results were 4 s for cutinase-treated fabrics, >180 s for untreated fabrics, and ∼120 s for cutinase-treated fabrics and >45 min for untreated fabrics. Results from the drop test are strongly affected by the porosity of the fabric; this makes it difficult to compare the results. Brueckner et al. (2008) demonstrated using XPS that, after enzymatic treatment, novel superficial carboxyl and hydroxyl groups are formed, whereas a hydroxide treatment does not result in the formation of new superficial groups. Donelli et al. (2008, 2009) assessed the surface changes using attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR). Donelli et al. (2008, 2009) successfully demonstrated that the free carboxylic surface groups generated by enzymatic hydrolysis of PET can be functionalized with functional molecules or groups. Müller et al. (2005) and Vertommen et al. (2005) demonstrated that cutinase displays higher activity towards amorphous PET and little activity towards highly crystalline PET. The amount and ratio of hydrolysis products formed/ released from the PET surface [viz. terephthalic acid (TPA), mono(2hydroxyethyl) terephthalate (MHET), and bis(2-hydroxyethyl) terephthalate (BHET), Fig. 4.1] depends on the enzyme, the source of the enzyme, the substrate, the enzyme–substrate ratio, the enzyme concentration and the progress of the reaction (Brueckner et al., 2008; Hooker et al., 2003; Vertommen et al., 2005). Despite the positive results obtained (increased hydrophilicity, significant depilling, increased reactivity with cationic dyes, and a small weight decrease) a better understanding of the action of cutinase on PET, the interaction of cutinase with PET and protein removal is required in order to make the potential of cutinase technology available for industry. Analytical techniques To monitor the changes resulting from the cutinase treatment a large variety of analytical techniques exist: techniques to assess the modifications of the PET itself and to determine the hydrolysis products released. When assessing surface properties, it is necessary to remove adsorbed cutinase thoroughly (Vertommen et al., 2005). Brueckner et al. (2008), Vertommen et al. (2005) and Yoon et al. (2002) used high-performance liquid chromatography (HPLC) to detect hydrolysis products formed (TPA, MHET and BHET), whereas O’Neil and CavacoPaulo (2004) used a spectroscopic assay to determine the release of TPA. Scanning electron microscopy (SEM) (Brueckner et al., 2008; Feuerhack et al., 2008; Kim et al., 2006, 2008), XPS (Brueckner et al., 2008; Vertommen et al., 2005) and ATR-FTIR (Donelli et al., 2008, 2009) have been used to analyse surface hydrolysis and surface properties. Donelli et al. (2008, 2009) and Brueckner et al. (2008) measured hydrophilicity using contact angle



85

measurements, Alisch-Mark et al. (2006) and Kim et al. (2008) measured the liquid uptake of fabric (rising height), whereas Brueckner et al. (2008) and Liu et al. (2008) used the drop method. Results from the liquid uptake test and the drop method are affected by porosity of the fabric. Alisch et al. (2004), O’Neil and Cavaco-Paulo (2004) and Brueckner et al. (2008) used a dyeing assay to determine the increase of hydroxilic groups.

4.2.2 Modification of polyamide fibres There is no doubt that polyamide polymers have unique properties, such as high uniformity, mechanical strength, and resistance against chemicals or abrasion. However, textile materials made of polyamide 6.6 are uncomfortable to wear because perspiration cannot penetrate the material and thus evaporate. This poor water permeability is directly related to the lower hydrophilicity of synthetic polymers, leading to static cling and stain retention during laundering process (Guebitz and Cavaco-Paulo, 2007). The reduced wettability and hydrophilicity also make these fibres unsuitable for specific finishing treatments such as coupling of flame retardants or covalent immobilization of proteins (Jia et al., 2006). Classical methodologies to improve hydrophilicity of the fibre, such as alkaline or acid hydrolysis, lead to the deterioration of polyamide fibres properties such as irreversible yellowing, resistance and weight loss (Cribbs and Ogale, 2003; Guebitz and Cavaco-Paulo, 2003; Miller and Wilmington, 1955; Shukla et al., 1997; Silva, 2002). These methodologies also require high amounts of energy and chemicals (binders, coupling agents, etc.), which are partially discharged to the environment. Furthermore, some of the substances used, owing to weak bonding with the fibre, are released from the end-products, causing serious health risks and reducing the technical lifetime of the products. Chemical finishers, based on hydrophilic carboxyl-containing polymers, are widely used to increase hydrophilicity of synthetic polymers, and their efficiency has continuously been improved, as evidenced by numerous patents filed (Soane et al., 2006). During the last two decades, functionalization of polymer surfaces has been recognized as a valuable tool to improve their adhesion properties. Functional groups like hydroxyl, amine and carboxylic acid groups have been introduced at polymers surface to adjust the surface free energy. Furthermore, these groups have been used to increase hydrogen binding and to facilitate chemical bonds between substrates and adhesives. Several studies have been carried out on the introduction of specific reactive groups at the surface of polyamide supports in order to reach higher levels of functionalization (Blencowe et al., 2006; Buchenska, 1996; de Gooijer et al., 2004; Herrera-Alonso et al., 2006; Jia et al., 2006; Makhlouf et al., 2007; Marcinčin, 2002; Saïhi et al., 2005; Tobiesen and Michielsen, 2002). Recently,


86


the covalent attachment of bioactive compounds to functionalize polymer surfaces including relevant techniques in polymer surface modification such as wet chemical, organosilanization, ionized gas treatments and UV radiation were described (Shearer et al., 2000; Goddard and Hotchki, 2007). Synthetic polymers coated with bioactive compounds include applications in textile manufacturing, microelectronics, bioprocessing and medical and food packaging (Shalaby et al., 2006). In order to reduce the use of chemicals and their environmental impact, new technologies have been developed to replace classical methodologies. Glow discharge (plasma) techniques are particularly useful for functionalization of surfaces. The surface modification of polyamide fibres and films by atmospheric plasmas has been described (Foerch and Hunter, 1992; Oktem et al., 2000; McCord et al., 2002; Pappas et al., 2006) and this technique has also been a valuable tool to improve the biocompatibility of fibres (Shmack et al., 2000). Surface modification with enzymes is considered to be a valuable tool to improve the quality and the processing properties of synthetic fibres. A biocatalytic method has the advantage of being performed under mild and environmentally friendly process conditions. Moreover, no complicated machinery is required, in contrast to plasma treatments, and few or no additional chemicals are needed. Furthermore, the direct control of the relative amount of specific groups formed is difficult to achieve by using physical techniques. Owing to the selectivity of enzyme reactions and their occurrence only at the surface of the fibres, enzymatic hydrolysis is desirable when compared with traditional chemical methods. Biocatalytic processes have been developed to modify polyamide surfaces enhancing hydrophilicity (Guebitz and Cavaco-Paulo, 2007). Enzymes that are able to hydrolyse polyamide surfaces are proteases, amidases and cutinases. Figure 4.2 shows the mechanism of action of these enzymes through polyamide structures. Heumann and co-workers have demonstrated that polyamide substrates can be efficiently functionalized by esterase, cutinase or protease action. A model substrate (adipic acid bishexylamide) was developed for screening polyamidase activity of a given enzyme. A protease from Beauveria sp., an amidase from Nocardia sp. and a cutinase from Fusarium solani were used to hydrolyze this model substrate (Heumann et al., 2006). The enzymatic hydrolysis of polyamide 6.6 fibres was extensively studied by Silva et al., and several process parameters were defined. For the first time, the action of a cutinase from Fusarium solani pisi towards polyamide fabrics was reported. The hydrolysis of nylon fabrics is based on the breakage of the amide linkages of the polymer surface resulting on amino and carboxylic groups yielding (Cavaco-Paulo et al., 2005; Silva and Cavaco-Paulo, 2004; Silva et al., 2005a).



R

O

H N

NH

87

R1

N H

O

O

Cutinase

Protease

Amidase

O R

OH

NH O

H2N

N H

R1 O

4.2 Enzymatic hydrolysis of polyamide 6.6 (amidase, protease and cutinase action) (adapted from Guebitz and Cavaco-Paulo, 2007).

The structure and function of cutinases are well studied and genetic engineering was previously used to improve their properties for several applications such as fat stain removal by detergents (Carvalho et al., 1999; Egmond and de Vlieg, 2000; Longhi and Cambillau, 1999). Site-directed mutagenesis of wild-type cutinase from Fusarium solani pisi was carried out to enlarge the active site in order to better accommodate the synthetic substrates. Several cutinase mutants with an enlarged active site were obtained showing higher activity towards polyamide substrates than wildtype cutinase (Araújo et al., 2007). In Fig. 4.3, the single exchanges in amino acids of a cutinase from Fusarium solani are shown. Despite the results attained, a better understanding of the enzyme–substrate interaction, regarding factors such as protein adsorption, mass transfer and binding modules, is required. The conjugation of all these factors will allow the development of enzymes with enhanced activity towards synthetic materials (Guebitz and Cavaco-Paulo, 2007). In addition to genetic engineering, reaction engineering (i.e. temperature, pH, type of agitation and additives) seems to play an important role in the enzymatic hydrolysis of PA (Silva et al., 2005b, 2007a). As esterases, amidases are also able to modify PA substrates. Enzymes and micro-organisms having amidase activity for hydrolyzing polyamides were patented in 2001 by Crouzet et al. (2001). Recently, an alkali stable polyamide was isolated from a new strain of Nocardia farcinica and its potential to hydrolyse water insoluble polyamide oligomers was studied (Heumann et al., 2009). Another polyamidase from Beauveria brongniartii was able to hydrolyse polyamide fibres (Almansa et al., 2008a).


88


L182A N84A L81A

V184A H188

OX

L189A

PA 6.6 model substrate-Tl

4.3 Detail of the active site x-ray structure of cutinase with the energy minimized structure of the tetrahedral intermediate (TI) PA 6.6 model substrate. The catalytic histidine (H188) and oxyanion-hole (OX) are shown. Residues mutated in this study are labelled as: L81A, N84A, L182A, V184A and L189A (Araújo et al., 2007).

In addition to hydrolytic enzymes, oxidases from lignolytic fungi have been shown to depolymerize polyamides. Some studies demonstrated that manganese peroxidase was able to modify the surface of PA 6.6 and PA 6 without reducing the fibre diameter (Friedrich et al., 2007; Klun et al., 2003). Nylon-degrading peroxidases attack methylene groups adjacent to the nitrogen atoms and the reaction then proceeds in an auto-oxidative manner (Deguchi et al., 1997, 1998). Fujisawa et al. (2001) have investigated a laccase-mediator system for its ability to degrade polyamide 6.6. Using 1-hydroxybenzotriazole (HBT) as a laccase mediator, they have shown that PA 6.6 membranes are disintegrated after 2 days of treatment. Other investigations where a laccasemediator system was used to increase the hydrophilicity of PA 6.6 have shown that the rising height as a parameter for hydrophilicity was enhanced from 1.8 (untreated) to 3.8 cm after a 300 s treatment with laccase and violuric acid as mediator, and to 5.5 cm with laccase alone (MiettinenOinonen et al., 2002, 2004). Despite all the developments in this area, the degradation of polyamide with oxidative enzymes seems to be difficult to



89

control, its effect being not only limited to surface degradation but also to other parts of the polymer, and therefore it has less potential for commercial application. Nylon oligomers degradation During polymerization reactions, some molecules fail to polymerize and remain as oligomers, and others undergo head-to-tail condensation to form cyclic oligomers. These nylon oligomers are the byproducts from the nylon factories, and, thereby, contribute to the increase in industrial waste materials. Biochemical studies revealed that hydrolases from Flavobacterium sp., namely 6-aminohexanoate-cyclic-dimer hydrolases are able to degrade 6-aminohexanoate oligomers (Fischer-Colbrie et al., 2006; Kakudo et al., 1993; Kato et al., 1994; Kinoshita et al., 1981; Negoro, 2000; Negoro et al., 1983, 1984). Negoro and co-workers have recently performed the x-ray crystallography of this enzyme as well its mutational analysis (Negoro et al., 2005, 2007; Ohki et al., 2006). Some work has been done on the isolation of a 6-aminohexanoate oligomer hydrolase from Agromyces sp. and Kocuria sp. (Yasuhira et al., 2007). Other reports describe oligomer degradation by Pseudomonas sp. (Prijambada et al., 1995). Methods to quantify enzymatic hydrolysis Enzymatic modification of polyamide materials involves the limited hydrolysis of amide bonds. The resulting changes in surface properties can be easily quantified by tensiometry or hydrophilicity measurements (Silva et al., 2007a). The corresponding chemical changes are measured in terms of formation of hydrophilic groups. The functional groups (e.g. amino groups) can be quantified at the surface of the treated materials by means of coloration with an α-bromoacrylamide reactive dye, which reacts specifically with amino groups (Silva and Cavaco-Paulo, 2004). The hydrolysis products released to the liquid treatment are quantified by colour change using 2,4,6-trinitrobenzenesulfonic acid (Silva and Cavaco-Paulo, 2004). The use of enzymes to modify polyamide materials requires limited hydrolysis at the surface of the polymer, leading to the production of new functional groups. High levels of hydrolysis can increase the amount of products released, therefore increasing materials degradation. A compromise between levels of hydrolysis and polymer degradation must be achieved (Silva and Cavaco-Paulo, 2008). When hydrolysis leads to the solubilization of the resulting smaller fragments, HPLC (high performance liquid chromatography) can be used to quantify them (Heumman et al., 2006). Attenuated total reflectance- Fourier transform infrared spectroscopy (ATR-FTIR) and scanning electron microscopy (SEM) have been used to analyse surface hydrolysis of polyamide


90


fabrics. X-ray diffraction can also be used to study the polymer changes in terms of crystallinity after enzymatic action (Silva et al., 2007a). The protein adsorption is an important factor to take into consideration during enzymatic treatment. Several washes and controls with inhibited enzymes are some of the techniques applied to eliminate the positive control attained when high amount of protein is used.

4.3

Applications

The enhancement of the hydrophilicity of synthetic polymers such as poly(ethylene terephthalate) and polyamide is a key requirement for many applications ranging from electronics to functional and technical textiles. The performance of PET can be improved by surface hydrolysis using cutinases, the enzyme affects the surface properties not the favourable bulk properties, especially when local modifications are required allowing specific functionalization. The functionalization of PET using fluorescent molecules after cutinase treatment was recently demonstrated by Donelli et al. (2009). Almansa et al. (2008b) increased the bonding strength of enzymatically modified PET with polyvinyl chloride (PVC). Polyamide performance can be improved by enzymatic hydrolysis resulting in an amine-enriched surface that can play an important role in processes such as the removal of heavy metal ions from aqueous solutions, removal of dyes from wastewater, biofouling prevention, covalent immobilization of proteins or coupling of flame retardants (Jia et al., 2006; Karimi and Amirshahi, 2000). The immobilization of a laccase from Trametes hirsuta onto an enzymatically modified nylon surface was reported by Silva et al. (2007b). The immobilization would protect laccase from denaturation by organic solvents and extend its half-life, at the same time allowing the enzyme to be reused in several reaction cycles (Brandi et al., 2006; D’Annibale et al., 2000; Durán et al., 2002).

4.4


The strong focus on the search for new sustainable processes, makes biocatalysis with enzymes a valuable alternative to traditional chemical methodologies, with advantages such as higher enantioselectivity, mild reaction conditions and lower energy and equipment requirements. Enzymes have proved to be environmentally friendly tools for hydrolysis of synthetic polymers, with a pronounced effect at the surface of the polymer, without compromising the favourable bulk properties of the polymer such as strength and crystallinity. The strength of enzymatic surface modification of PET and PA is to increase hydrophilicity locally, allowing specific functionalization and an important advantage of enzymes over conventional tech-



91

niques in surface modification of PET and PA is that the favourable bulk properties are not affected because the enzymes are too big to penetrate into the bulk phase of the material. Despite all the developments reported in this area, a better understanding of the interactions between enzyme and substrates is required. Factors such as protein adsorption, enzyme concentration, and level of agitation, mass transfer and the role of hydrophobins or binding modules will influence significantly the efficiency of hydrolysis. New strategies in terms of protein engineering and bioinformatics can provide new tools that will advance biocatalysis to a more competitive technology.

4.5


Despite some success already achieved, the potential benefits of biotechnological modification of synthetic fibres are far from being fully explored. The actual enzymatic modifications, described in this chapter, lead to the creation of hydrophilic groups at the surface of cheap materials like PET and PA. The new functionalized fibres can have a totally new range of applications such as filter media and smart, technical and high-performance materials. New methodologies have been developed to generate carboxylic, amino and hydroxyl groups over PET and PA. Immobilization of enzymes and other products such as chitosan at the surface of enzymatically modified fibres can be the future trends on this area. Already, new enzymes with improved activity towards synthetic polymers (Araújo et al., 2007) or better temperature stability have been produced (Liu et al., 2008). Future challenges are in the area of thermostable enzymes from extremophiles, molecular modelling, protein engineering and genetically modified enzymes. Novel strategies will need to be developed based on chemoenzymatic approaches to locally functionalize synthetic materials. Future trends will not only depend on enzyme technology but also on the technology to apply the enzyme to the material. Soft-lithography and inkjet technology allows application of chemicals and enzymes locally in a controlled way. This will enable specific surface modification and functionalization.

4.6

Acknowledgements

V. A. Nierstrasz acknowledges support of the European Commission, Marie Curie Grant, FP7-PEOPLE-2007-2-1-IEF, Grant Agreement Number PIEF-GA-2008-219665, BIOTIC, Biotechnical functionalization of (bio) polymeric textile surfaces. Carla Silva acknowledges support of the Fundação para a Ciência e a Tecnologia, Grant SFRH/BPD/46515/2008.


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4.7

References

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5 Enzymatic modification of polyacrylonitrile and cellulose acetate fibres for textile and other applications T. M ATA M Á and A. C AVAC O - PAU L O, University of Minho, Portugal

Abstract: The enzymatic reactions that lead to the creation of hydrophilic groups at the surface of polyacrylonitrile and cellulose acetate are reviewed and the main advantages and drawbacks are briefly discussed. New materials with distinct surface properties can be achieved using eco-friendly catalysts that preserve the desirable bulk properties of polyacrylonitrile and cellulose acetate. Key words: biomodification, polymer side chains, biotechnology, polyacrylonitrile, nitrilase, acetyl esterase, cellulose acetate.

5.1

Introduction

In the textile industry, the impact of biotechnology has been observed at three main levels: the introduction of enzymes in manufacturing wet processes and laundry detergents, the design of new and biodegradable fibres and the treatment of textile effluents. Biocatalysis is present in some of the oldest transformations known to humans: descriptions of various beer recipes were found in Sumerian writings (Ball, 2001). Studies on fermentation processes led to a ‘big bang’ in the knowledge of life’s chemistry. In the 19th century, Louis Pasteur came to the conclusion that the fermentation of sugar to alcohol by yeast was catalyzed by a vital force contained within the yeast cells called ‘ferments’, which were thought to function only within living organisms. Wilhelm Kühne was the first to use the term ‘enzyme’ and, years later, Emil Fisher proposed the ‘Lock and Key Model’ to visualize the substrate and enzyme interaction (Cabral et al., 2003). Enzymes are the subgroup of proteins that catalyse the chemistry of life, transforming both macromolecules and small molecules; they are the focus of current biocatalysis research (Walsh, 2001). The chiral nature of enzymes results in a remarkable chemical precision seen as different types of selectivity, even in the absence of functional-group protection (Rozzell, 1999; 98 © Woodhead Publishing Limited, 2010

Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 99 Koeller and Wong, 2001). Enzymes are environmental friendly catalysts not just because they are biodegradable themselves but also because of their mild operating conditions. They can result in processes that generate fewer waste disposal problems and that require lower energy input, leading to lower costs and lower emissions of greenhouse gases to the environment (Rozzell, 1999). The enzyme can be used as the sole catalyst in a reaction, in combination with other enzymes, or with inorganic reagents. Moreover, many enzymes accept unnatural substrates, and genetic, pathway and medium engineering can improve further their stability and specific activity as well as modulate their substrate specificity (Koeller and Wong, 2001). These qualities make enzymes remarkable catalysts. It is now widely recognized that enzyme-catalysed chemical transformations are convenient alternatives to traditional (non-biological) transformations. Biocatalysis is a tool of increasing importance for industries that aim at sustainable development (Schäfer et al., 2007; van Beilen and Li, 2002). Textile materials made from synthetic fibres are, in general, uncomfortable to wear because they are hydrophobic. This means that these materials can not absorb perspiration and water vapour can not easily be transported away from the body. The hydrophobic nature also leads to their characteristic static cling and stain retention during laundering (Gübitz and CavacoPaulo, 2008). Textile finishing is the final stage in the fabric manufacturing process and includes all the processes that modify the surface of fibres to add useful qualities to the fabric, ranging from interesting appearance and fashion aspects to high-performance properties for industrial needs (Schindler and Hauser, 2004). As the use of high performance textiles grows, the need for chemical finishes to provide the fabric properties required in these special applications has grown accordingly. Currently, more than 20 different types of chemical finishers exist for both natural and man-made fibres (Fig. 5.1) (Schindler and Hauser, 2004). There are several methods for the surface modification of synthetic polymers to increase hydrophilicity. The methods currently used are plasma treatments and chemical finishers, including alkaline and acid hydrolysis. In addition to the environmental issues, these methods are difficult to control, sometimes they are of reduced technical lifetime and in some instances severe fibre weight losses and yellowing occur (Gübitz and Cavaco-Paulo, 2003, 2008). On the other hand, enzymes are well-suited for targeted surface functionalization of polymers. In addition to the advantages referred to above, enzymes are macromolecules and for that reason their action is normally restricted to the most superficial layers of polymer fibres. The broad substrate specificity and catalytic promiscuity, exhibited by some enzymes, uncover a new range of possible biocatalytic transformations (Bornscheuer and Kazlauskas, 2004). The ability of enzymes to use


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Advances in textile biotechnology Softeners

0.01%

Repellents 14.0% 22.1%

1.4%

Flame retardants

2.3%

Products for coating and fibre/thread bonding Products for easy-care

0.3% 10.0%

4.1%

Hand builders Antimicrobial products

13.9%

Antistatic agents

13.5% Non-slip agents 18.4%

Anti-soiling products Others

5.1 Distribution of textile chemical finishers by amount in 2001 (adapted from Schindler and Hauser, 2004).

textile synthetic fibres as substrates, which are known for their stability and chemical inertia, under mild conditions, is evidence of the vast potential of these catalysts in industrial processes. There are several reports of successful enzymatic surface modifications of the major three synthetic textile fibres: polyester [poly(ethylene terephthalate), PET], polyamide (PA) and polyacrylonitrile (PAN). The targets for enzyme catalysis are the ester and amide bonds of the polymer backbone of PET and PA, respectively, which are the subject of chapter 4 of this book. In this chapter, the focus is on the biomodification of two man-made fibres: acrylic and cellulose acetate. For both fibres, enzymes can be used to accomplish the formation of reactive and/or hydrophilic groups at the surface by hydrolysis of their pendant groups, without affecting, in theory, the integrity of the main chain of the polymers.

5.2

Basic concepts and principles

5.2.1 Side chains from polyacrylonitrile as substrates for enzymes Acrylonitrile is the main monomer of synthetic fibres commonly called acrylics which are defined, according to the Federal Trade Commission of


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 101 the United States, as manufactured fibres in which the fibre-forming substance is any long-chain synthetic polymer composed of at least 85% by weight of acrylonitrile units (Guillen, 1987). In 2006, acrylic production was 6.8% of that of total chemical fibres (Yarns and Fibres Exchange, 2007). Acrylic fibres impart warm, natural-like aesthetics to most fabrics as opposed to the cold, plastic handle of polyester and nylon fibres (Lulay, 1995). According to a study performed to evaluate qualitatively the properties of several fibres, consumers considered acrylic as having moderate performance on highly desirable properties like abrasion, wrinkle and pill resistances, strength and wash-wear. Nylon and polyester were considered to have a better performance for those properties. Therefore, there is a continued effort to improve acrylic properties to compete with other synthetic fibres. Several methods have already been tested, including incorporation of comonomers and modification of the spinning process and/or finishing treatments (Frushour and Knorr, 1998). The term acrylic fibre covers a broad range of products, more diverse in composition than any other synthetic fibre (Masson, 1995). The major reason for this is that acrylonitrile can copolymerize with many different monomers with an ethylene unsaturated group, by free radical polymerization (Fig. 5.2). The comonomers are used in order to increase the solubility of the polymer in the spinning solvents and improve the rate of dye diffusion into the fibres (Frushour and Knorr, 1998). Acid and basic comonomers are also used to create additional sites for dye fixation and to provide a hydrophilic component in water-reversible crimp bicomponent fibres (Frushour and Knorr, 1998; Masson, 1995). Halogenated comonomers, usually vinylidene chloride, vinyl bromide and vinyl chloride, can be used to impart flame resistance to the acrylic textiles (Frushour and Knorr, 1998; Masson, 1995). The pendant group in PAN molecules is therefore the nitrile group. The distinguishing feature of the nitrile group is the large dipole moment, making it one of the most polar organic functional groups (Frushour, 1995). It is believed that these protuberant nitrile groups are responsible for the net attraction between adjacent PAN polymer chains and the ability of this vinyl polymer to form fibres with some degree of structural order, which is essential for its use as a textile fibre. Consequently, if the choice is to act on the nitrile group as a primary target in order to alter the acrylic properties or to add new functionalities, it is mandatory that these modifications happen at the fibre surface leaving intact the core structure that is essential for the resistance and integrity of the acrylic fibres. Current methods for the addition of functional groups or transformation of existent nitrile groups on the surface of acrylic fibres involve the action of chemical or physical agents, like hydrogen peroxide, concentrated acid and bases, or radiation and plasma (Battistel et al., 1995; Gübitz and Cavaco-Paulo, 2008). However,


102


R N

N

N

N

N

N

n

R = neutral comonomers CH3 CH2

CH2

CH

C

O

C

CH2

C

O

O O

O

O

CH

CH3

CH3

CH3 Methyl acrylate

C

Methyl methacrylate

Vinyl acetate

R = acid comonomers CH2

C

CH3

CH

CH2

CH O

CH2

OH

C CH2

SO3Na

Acrylic acid NaSO3

Sodium methallyl sulfonate

Sodium styrene sulfonate R = basic comonomers CH2

CH2

CH C NH2

Acrylamide

CH

O N 4-vinyl pyridine

5.2 The acrylic polymer structure and some examples of its common comonomers.

as mentioned above (see section 5.1) these methods present major disadvantages, leaving a very important gap that can be addressed with biotechnology. In nature, the formation and cleavage of carbon–nitrogen bonds are essential. Peptide bonds are the most obvious class of carbon–nitrogen bonds, but there are others whose metabolism is less well understood (Brenner, 2002). Such less obvious bonds include nitriles (R—C≡N), amides [R—C(=O)—NH2], secondary amides [R—C(=O)—NH—R′], N-carbamyl amides [R—NH—C(=O)—NH2]. They are hydrolytic substrates for 9 of 13 branches of the nitrilase superfamily (Brenner, 2002; Pace © Woodhead Publishing Limited, 2010

Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 103 and Brenner 2001). Nitrile-degrading activity is found in three of the 21 plant families and in a limited number of fungal genera, whereas it is relatively frequent in bacteria (Banerjee et al., 2002). There is a group of nitrile-metabolizing enzymes that can convert the nitrile groups into carboxylic or amide groups, depending on the type of enzyme system. Figure 5.3 illustrates the possible chemical transformations that may occur when treating copolymers of acrylonitrile with the various

R N

N

N

N

N

N

n

N

n

N

n

H2O

2 H2O Nitrilase

Nitrile hydratase

NH3

R HO

O

H2N N

O

N

N

H2O Amidase NH3

R HO

O

HO N

N

O N

5.3 Possible chemical transformations on polyacrylonitrile surface catalysed by nitrile-metabolizing enzymes.


104


classes of nitrile-metabolizing enzymes. Nitrilases (EC 3.5.5) and amidases (EC 3.5.1) are both hydrolases, whereas nitrile hydratase (EC 4.2.1) is a carbon–oxygen hydrolyase, meaning it catalyses the breakage of a carbon– oxygen bond by elimination of water. In the particular case of PAN biomodification the interest is on the reverse reaction. These enzymes are described in more detail in chapter 1.

5.2.2 Side chains from cellulose acetate as enzymatic substrates Acetate fibres are defined as manufactured fibres by the Federal Trade Commission of the United States, in which the fibre-forming substance is cellulose acetate (Needles, 1986). Cellulose acetates are classified as derivative cellulose fibres, as distinct from rayon and lyocell, which are regenerated cellulose fibres, the chemical composition of cellulose acetate being not cellulose but an ester of cellulose (Collier and Tortora, 2001). Each anhydroglucose repeating unit of cellulose (Fig. 5.4) has three hydroxyl groups located at the positions 2, 3 and 6 (La Nieve, 2007). These sites are available for acetylation to produce the acetate fibres. The degree of acetylation or substitution (DS) is the average number of acetylated positions per anhydroglucose unit. Commercial cellulose triacetate (CTA) has a DS of 2.91–2.96 whereas cellulose acetate has a DS of ≈2.4 (La Nieve, 2007). Therefore, the denomination of cellulose acetate fibres is used to refer to all commercial acetylated cellulose fibres, but it is also the common name for the cellulose acetate fibre with a DS ≈ 2.4. To avoid misinterpretations, throughout this text, the name cellulose acetate will be used to refer to both types of fibres whereas cellulose diacetate (CDA) will be used to refer the acetate with a DS of ≈2.4. Cellulose acetates are produced from high-quality cellulose, such as cotton linters and wood pulps, with an α-cellulose content above 95% (Saka and Matsumura, 2004). The most commonly used commercial acetylation process is the acetic acid system where acetic acid serves as solvent for the

6 CH2OH 5 4

O 1

OH

O

2 O

3 OH

n

5.4 Chemical structure of the anhydroglucose repeating unit of cellulose.


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 105 cellulose acetylation and acetic anhydride and sulfuric acid as catalysts (La Nieve, 2007). Acetate fibres are soft and cool, have silk-like aesthetics and good drape, and they can be easily blended with other fibres like silk, rayon, nylon, cotton and polyester (Law, 2004). The moisture regains for CDA and CTA are 6.5% and 3.5%, respectively (Steinmann, 1998; La Nieve, 2007). The CDA has a moisture regain close to the value 7% of natural cotton yarn, whereas the CTA has a lower value but still higher than the commercial synthetic fibres. Their unique attributes remain desirable and they are responsible for the survival of acetate production in the competitive market of man-made fibres. Another attribute that is gaining importance is the fact that cellulose acetate fibres are environmental friendly compared with the major synthetic fibres. Whereas cellulose, either from cotton linters or wood pulp, is highly crystalline, dry-spun cellulose acetates show very low crystalline order owing to the substitution of the hydroxyl groups by acetyl groups and consequent disruption of the original cellulose structure (La Nieve, 2007). In both CTA and CDA, hydrogen bonding between cellulose chains is substantially decreased and the bulky acetyl group prevents the close packing of cellulose chains (Needles, 1986). The van der Waals forces are the major associative forces between the polymer chains, and their lower magnitude is the reason for cellulose acetate being considerably weaker than cellulose fibres. Both CDA and CTA have a very low strength and their chemical stability is poor (Steinmann, 1998; Collier and Tortora, 2001). They are attacked by a number of organic solvents capable of dissolving esters, strong acids and bases, which result in saponification of acetyl groups. For these reasons, the physical and/or chemical modification of these fibres is of very limited use. Some methods were developed to improve the strength, abrasion resistance and dimension stability of acetate fibres, in particular of CDA (Steinmann, 1998). One approach was to apply polymer additives to the CDA spin dope. Several were tested but, unless their concentration was below 5%, the phase compatibility was poor (Steinmann, 1998). To improve the compatibility, some polymers were grafted onto CDA. In the case of acrylonitrile, the graft copolymer increased the compatibility of PAN and cellulose acetate and the resulting fibres had improved thermal and chemical stabilities (Steinmann, 1998). The effect of crosslinking agents was also investigated on CDA, though the improved properties were still not equal to those of heat-treated CTA (Steinmann, 1998). Work on the modification of cellulose acetate with enzymes has been done in the context of its biodegradation (Puls et al., 2004). Figure 5.5 summarizes the main reactions expected to occur during the biodegradation of cellulose acetate with special emphasis on the deacetylation reaction. The


106

Advances in textile biotechnology OCOCH3

OCOCH3 OCOCH3

O H3COCO

O H3COCO

O

O O H3COCO

O

H3COCO

CH3COOH

OCOCH3

OCOCH3 OH

O O HO

O

HO O

O

OH

H2O

OCOCH3

HO

O HO

O

O

O

OH

H3COCO

H3COCO

Endoglucanases

OCOCH3

OCOCH3 OH

O O HO

O

OCOCH3

H2O

Acetyl esterases (EC 3.1)

O

O

OCOCH3 H3COCO

OCOCH3

H3COCO

HO

OH HO

OH

O

O

O HO

HO O

OH

H3COCO

Exoglucanases

OCOCH3 HO HO

O

O HO

O O

H3COCO

H2O

OH

OCOCH3

O

HO OH

HO

OH

H3COCO

OCOCH3 O O

H3COCO

5.5 Main reactions occurring during biodegradation of cellulose acetate.

degradation of cellulose and hemicellulose is naturally carried out by microorganisms and requires the concerted action of many enzymes for their complete destruction. Among those carbohydrate-active enzymes, there is the group of carbohydrate esterases (EC 3.1) that hydrolyse the ester linkage of polysaccharides substitutents. This group is a potential supplier of biotechnological tools to hydrolyse the cellulose acetate fibres in a controlled manner, creating hydroxyl groups at the fibre surface that, besides imparting hydrophilicity, can be subsequently modified.

5.3

From theory to practice: current methodologies for the assessment of fibre biomodification

The modification of PAN and cellulose acetate with enzymes results in two types of products: soluble compounds and new chemical groups that remain attached to the insoluble substrate, the fibre. For PAN polymers, only nitrilase and amidase generate the soluble product ammonia (Fig. 5.3); nitrile hydratase generates amide groups as new side chains of the PAN


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 107 main chain. For cellulose acetate, the hydrolysis of its side chains releases acetic acid to the reaction media and the hydroxyl group is located on the polymer backbone (Fig. 5.5). The evaluation of enzyme activity on these textile substrates can be performed by measuring either the chemical changes at the solid substrate or the amount of the soluble product released into solution. Both ammonia and acetic acid can be easily quantified using enzymatic assays that are very specific, are simple to perform, and require only common laboratory equipment such as a UV/visible spectrophotometer. There are several diagnostic kits for both ammonia and acetic acid commercially available. By measuring the decrease or increase in absorbance at 340 nm, as the reduced nicotinamide adenine dinucleotide (NADH) is consumed or formed, respectively, the amount of ammonia and acetic acid are calculated, according to the stoichiometry of the reaction. Minor sample preparation is necessary and this consists of protein removal using strong acids and subsequent neutralization. Other methods based on chemical reactions exist for ammonia determination. The most common is the direct nesslerization method which involves the reaction of ammonia with an alkaline solution of mercury (II) iodide in potassium iodide–Nessler reagent to produce an orange–brown product (Greenberg et al., 1992, pp. 4.75–4.80). Banerjee et al. (2003) described a fluorimetric assay based on the reaction of ammonia with buffered o-phthaldialdehyde-2-mercaptoethanol solution (pH 7.4). The product is a fluorescent complex, whose intensity is measured with excitation and emission wavelengths of 412 and 467 nm, respectively. Acetic acid can also be determined by non-enzymatic assays. Gas chromatography is one of the most used methods to quantify acetic acid for which several columns are commercially available. A titration can also be used, but it is not specific for this acid. When assessing both soluble products (ammonia or acetic acid/ acetate), it is important to verify the stability of the solutions during treatment of the fibres and possible interferences, such as from the protein content, which can alter or mask the real concentration of these compounds. Controls with an inhibited/deactivated enzyme are important to eliminate background interferences; the use of biocide compounds during enzymatic treatment; or the acidification of aliquots from the treatment solutions containing ammonia (to prevent its volatilization) are some examples of what can be done to control as much as possible the quality of the measurements. For the evaluation of chemical changes on the solid material, analytical methods that have a good surface sensitivity and depth resolution should have an advantage because enzymes act mainly at the fibre surface. When comparing controls and samples from a particular enzymatic treatment, the


108


differences are diluted and the sensitivity of the method is weakened if the chosen analytical method takes into account both the bulk and the surface of the tested material. Nowadays, there are numerous analytical techniques available to study the surface of materials (Niemantsverdriet, 2007). The spectroscopic methods most used for the evaluation of chemical modifications in textile materials are Fourier transform infrared spectroscopy (FTIR) and x-ray photoelectron spectroscopy (XPS)/electron spectroscopy for chemical analysis (ESCA). FTIR provides specific information about chemical bonding. When analysing the surface, FTIR is used coupled to the attenuated total reflectance (ATR) technique or as diffuse reflectance infrared Fourier transform (DRIFT). XPS/ESCA is used to determine quantitative atomic composition; it is a surface analysis technique with a sampling volume that extends from the surface to a depth of approximately 50–70 Å. Knowing the chemical groups generated during biocatalysis on the surface of the fibres, it is possible to have a relative quantification using dyes that specifically react with them. When the PAN is treated with nitrilase, the formation of carboxylic groups can be evaluated by staining the fabric with a basic dye, which has a cationic group able to establish ionic bonds with anionic groups on the fibre. Similarly, when PAN is treated with nitrile hydratase, the formation of amide groups at the surface of the fibres can be evaluated by staining the fabric with an acid dye. Staining both controls and samples in the same dyeing bath (competitive assay) allows the increase in those particular chemical groups to be estimated as more dye can be absorbed into the biomodified fibre (seen as an increase in colour). The differences in colour strength are measured as K/S (the ratio between absorption K and scattering S) at the maximum absorption wavelength of the particular dye, a parameter proportional to the dye concentration in the fibre comparing the enzyme-treated and control samples (Kuehni, 1997). This staining methodology is a valuable and a very sensitive semiquantitative method because of the large molar absortivities of dye molecules (Matamá et al., 2006, 2007; O’Neill et al., 2007; Silva et al., 2005). The hydroxyl groups that result from the enzymatic hydrolysis of cellulose acetate can be evaluated using another class of dyes, the cotton reactive dyes, in particular, the warm brand vinylsulphone dyes. These dyes are able to covalently link to the hydroxyl group at low temperatures and relatively low alkaline pH, and the elimination of the protective group does not depend on the fibre to be dyed (Hunter and Renfrew, 1999). The moderate temperatures (below the glass transition temperature) are important to restrict the staining to the surface, stressing the differences between modified samples and controls (Burkinshaw, 1995). The pH is important in the particular case of cellulose acetate because this material is not chemically stable at high pH because of the chemical hydrolysis of the acetyl groups.


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 109 For both fibres, the enzyme introduces chemical groups that are more hydrophilic than the original ones; therefore it is also possible to indirectly assess the biomodification using methods that determine the hydrophilicity of materials. It may also be relevant to evaluate any changes that may occur in physical properties of the fibres such as mechanical performance (strength, elongation/elastic recovery, and resiliency) and electrical conductivity, using standard testing procedures. Valuable information can also be inferred from high-resolution surface images of treated materials. Scanning electron microscopy (SEM) is one of the most widely used analytical tools owing to the extremely detailed images it can provide. In addition, when it is coupled to an auxiliary energy dispersive x-ray spectroscopy (EDS) detector, this technique also offers elemental identification. Scanning transmission electron microscopy (STEM) has better spatial resolution and is capable of additional analytical measurements, such as crystallographic phase or orientation characterization (Niemantsverdriet, 2007). Whatever the method of choice, there is a factor that has to be carefully controlled: the amount of adsorbed protein that will interfere with the measurements. Efficient washing after enzymatic treatments is essential and, once again, good controls that use enzymes without activity (null mutants) but with the same adsorption profile are very important.

5.4

Polyacrylonitrile biomodification

5.4.1 Nitrile-metabolizing enzymes for the modification of polyacrylonitrile Tauber et al. (2000) reported the modification of PAN using crude enzyme preparations from the strain Rhodococcus rhodochrous NCIMB 11216: a nitrile conversion to the carboxylic acid with the release of ammonia, on granular PAN as substrate, and a nitrile conversion to the amide on acrylic fibres. In cell preparations, both nitrile hydratase and amidase were detected (at 30 °C and pH 6.5, the nitrile hydratase activity was 14.2 nkat mg−1 of cell dry weight using acrylonitrile as substrate and the amidase activity was 1.7 nkat mg−1 of cell dry weight using acrylamide as substrate). Two granular PAN standards were used, having different molecular weights (40 and 190 kDa). The enzymatic treatments of granular PAN were performed in 57 mM phosphate buffer pH 7, at 25 °C for a maximum period of 72 h, under vigorously shaking (300 rpm). The enzymatic system of nitrile hydratase/amidase was able to partially complete the hydrolysis of the nitrile to carboxylic groups, in both granular PAN standards, as indicated by the release of ammonia, although, the molecular weight of PAN negatively influenced the action of enzymes. When acrylic fabric samples (PAN copolymer containing 7% w/w of vinyl acetate) were treated at 30 °C for the


110


same period, under various media compositions, no ammonia could be detected. By means of XPS and dyeing assays with methylene blue (C.I. Basic Blue 9) and Coomassie brilliant blue (C.I. Acid Blue 90), we concluded that the superficial nitriles of acrylic fibres were converted to the corresponding amides that were not accessible to amidases. The best results in terms of increase in colour, measured as K/S for methylene blue, were obtained at pH 6.5 and in the presence of 0.5% v/v dimethylformamide (DMF). Owing to the reported enzymatic modifications the acrylic fabrics became more dyeable with methylene blue (an increase of 15% in K/S), which is somewhat contradictory to the conclusion that amide groups were formed and not the acidic groups, on the acrylic surface. The modification of nitrile surface groups into amide groups was also accomplished by the action of nitrile hydratases present in cell lysates from Brevibacterium imperiale and Corynebacterium nitrilophilus (Battistel et al., 2001). The PAN used in this study was a copolymer containing 10% of vinyl acetate in two formats, powder and fibre. The enzymatic treatments were performed in 100 mM phosphate buffer pH 7, at 20 °C for a maximum period of 48 h, with vigorous shaking (250 rpm). The newly formed amides were assessed by XPS. After 48 h, for both fibre and powder forms treated with the active cell lysate, the content of oxygen increased significantly with a concomitant decrease in the content of nitrogen. Further hydrolysis into carboxylic groups was ruled out because of the absence of the XPS peak characteristic of these groups in the spectrum of the modified PAN polymer. The oxygen content increased as a function of reaction time, mainly in the first 2 h, stabilizing after 10 h. This could be explained by the moderately fast deactivation of the enzyme during the treatment. The decrease in the contact angle for the modified PAN fibre was verified as well as the ability of this PAN to react with acid dyes that gave optimal colour yields for natural fibres such as wool and silk. The untreated controls were inert to the same dyes. The dye uptake behaviour during treatment correlated with the increase in the oxygen/carbon ratio of the modified polymer, as expected. The described enzymatic modification was a promising result for the improvement of the comfort properties of PAN and the possibility of dyeing simultaneously mixed yarns of acrylic and wool. A nitrile hydratase, isolated from the Arthrobacter sp. ECU1101, was used to selectively transform the nitrile into amide groups of acrylic fibres comprising 85% of acrylonitrile and 10% of methylacrylate (Wang et al., 2004). The enzymatic treatments were performed in 50 mM phosphate buffer pH 7, at 30 °C for a period of 48 h, with vigorous shaking (200 rpm). The addition to the media of 0.1% w/v of JFC (a non-ionic surfactant with fatty alcohol polyoxyethelene ether as the main ingredient) and 0.1% of DMF showed that the use of the non-ionic surfactant improved the enzyme activity whereas DMF did not (Tables 5.1–5.3). The modified acrylic fibre


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 111 Table 5.1 Working solution ingredients for the treatment of acrylic fibres with a nitrile hydratase preparation extracted from Arthrobacter sp. ECU1101 cells. Reproduced with permission from the original publisher (Wang et al., 2004) Samples

Enzyme (%, v/v)

JFC (%, w/v)

DMF (%, v/v)

Control 0 Control 1 Sample 1 Sample 2 Sample 3

0 5 5 5 5

0 0.1 0 0.1 0

0 0 0 0 0.1

(inactive) (active) (active) (active)

Table 5.2 Physical characteristics of samples and controls after treatment with a nitrile hydratase preparation extracted from Arthrobacter sp. ECU1101 cells. Reproduced with permission from the original publisher (Wang et al., 2004) Samples

Moisture regain (%)

Volume specific resistance (Ω cm)

Control 0 Control 1 Sample 1 Sample 2 Sample 3

2 2.1 2.7 3.1 2.6

1.65 9.86 1.25 7.29 2.08

× × × × ×

1012 1011 1010 109 1010

Water contact angle (°) 51 49 39 31 38

Table 5.3 K/S values of acrylic samples stained with acid dyes after treatment with a nitrile hydratase preparation extracted from Arthrobacter sp. ECU1101 cells. Reproduced with permission from the original publisher (Wang et al., 2004) Dyes

pH

K/S values Control 0 Control 1 Sample 1 Sample 2 Sample 3

C.I. C.I. C.I. C.I. C.I. C.I. C.I.

Acid Acid Acid Acid Acid Acid Acid

Black 24 Red 14 Blue 25 Red 249 Blue 225 Yellow 16 Red 1

4.5 2.0 2.0 4.5 4.5 2.5 2.0

1.76 0.05 3.03 0.77 0.16 0.82 0.55

1.89 0.08 3.12 0.87 0.27 0.95 0.64

2.16 0.12 3.49 1.13 0.43 1.12 0.92

2.73 0.25 3.96 2.22 0.57 1.56 1.12

2.12 0.10 3.41 1.42 0.41 1.20 0.89

became more hydrophilic as indicated by the results of moisture regain and water contact angle; the antistatic ability was also improved on treated acrylics as shown by the decrease in their volume specific resistance (Table 5.2). Several acid dyes were tested and the biomodified samples all showed


112


an increase in K/S (Table 5.3). The XPS analysis clearly demonstrated the conversion of nitrile groups into amides without further hydrolysis, despite the presence of amidases in the cell-free extracts, as also verified by Tauber et al. (2000). Another microorganism, Agrobacterium tumefaciens, was found to grow on PAN as a carbon source, converting it to polyacrylic acid (PAA) as shown by solid state nuclear magnetic resonance (NMR) (Fischer-Colbrie et al., 2006). The crude enzyme preparation isolated from the bacteria cells showed activities of both nitrile hydratase and amidase. PAN powder and fabrics were treated over 24 and 48 h, respectively, with crude enzyme preparations in a special stabilizing buffer, at 30 °C. The nitrile groups were converted either to amides or to carboxylic groups, depending on the reaction conditions (time of incubation and enzyme activity) and on the type of substrate. Once more, the amidase was less efficient in using acrylic fabric as a substrate, as found in other studies for amidases from other species. The ammonia was only detected after prolonged enzymatic treatment of PAN fabrics. Fischer-Colbrie et al. (2006) proposed the higher surface area of PAN powder compared with PAN fabrics as a cause of the higher yield of conversion to carboxylic groups in the powder. This work confirms that the nitrile hydratase is more active on PAN than amidase, resulting primarily in the formation of amides when such enzyme system is used. More recently, two reports were published referring to the complete biomodification of nitrile groups of PAN into carboxylic groups (FischerColbrie et al., 2007; Matamá et al., 2007). A membrane-bound nitrile hydrolysing enzyme was isolated from Micrococcus luteus BST20 by Fischer-Colbrie et al. (2007). Nitrile groups from both PAN powder and fabrics were converted to carboxylic acid groups, with the associated release of ammonia (5% of the theoretical possible amount for PAN powder). The fabrics treated with nitrilase showed higher K/S values when dyed with methylene blue than the fabrics treated with inactivated enzyme or without any enzyme at all. The results demonstrated that this micro-organism is an interesting source of enzymes for modifying the surface of PAN, producing acid groups as opposed to the previous nitrile-degrading enzymatic systems. Acrylic fabrics composed of a copolymer of PAN and 7% w/w vinyl acetate were treated with a commercial nitrilase (Matamá et al., 2007). The effect on nitrilase stability of the two most common PAN solvents, DMF and dimethylacetamide (DMA), and of polyalcohols was studied in order to establish a media composition that would favour the accessibility of PAN fabrics/fibres to nitrilase, without compromising the enzyme stability more than necessary. Treatments performed at 30 °C over 5 days, in the presence of 4% v/v DMA and 1M sorbitol, led to an improvement in the colour strength of treated fabrics (17% increase in K/S values compared with the


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 113 fabrics treated with nitrilase without additives). Further assays without additives were performed to study the progression of the modification of PAN catalysed by nitrilase over a maximum period of 36 h, at 40 °C and pH 7.8. The conversion of nitrile groups into carboxylic groups was followed by the release of ammonia and by improvement in the basic staining with methylene blue (Fig. 5.6). There was no significant ammonia release before 8 h of treatment; after this point a linear release of ammonia was verified which implies the absence of activity decay during the 36 h of treatment. The values of K/S, represented as a percentage of the initial value, demonstrated a varying behaviour with time, the maximum K/S value was observed for a treatment over eight hours and it corresponded to a relative K/S of 135%. The increase in carboxylic groups, evaluated by K/S was not steady during the 36 h of treatment. Instead, a succession of K/S values higher and lower than the controls, creating an oscillating pattern, was observed. The hypothesis proposed is that somehow the groups left the surface of the fabric. This was confirmed when PAA was determined in the treatment solutions (Fig. 5.6). A hypothetical model of surface erosion of acrylics was

Total nitrogen (mg l–1)

25

20

15

10

5

0 0

5

10

15

20 Time (h)

25

30

35

160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 40

100 90 80 70 60 50 40 30

Polyacrylic acid (mg l–1)

30

Relative K/S (lmax = 660 nm) (%)

Total nitrogen Relative K/S Polyacrylic acid

20 10 0

5.6 Total nitrogen and polyacrylic acid concentrations in solution, and relative K/S values for acrylic fabric samples as a function of reaction time. The acrylic samples were treated with 412 U of nitrilase per gram of fabric, at pH 7.8 and 40 °C. Samples and controls were competitively coloured at 70 °C with Basic Blue 9, C.I. 52015, and relative K/S values were calculated as K/Senzyme/K/Scontrol (%) (adapted from Matamá et al., 2007).


114


5.7 Surface erosion model for acrylic fibres treated with nitrilase at alkaline pH (for simplification the comonomers are not represented).

proposed to explain the results (Fig. 5.7). The oscillation of relative K/S values with time of treatment could be explained assuming that, above a certain degree of conversion of nitrile groups into carboxylic groups, the polymer chains are more stable in solution and therefore detach from the surface of the fibres. The modification of nitrile into carboxylic groups could create some instability in the arrangement of PAN chains at the surface of the fibres, mainly as a result of steric hindrance and pH-dependent charge repulsion. Increasing concentrations of PAA in treatment solutions were detected (Fig. 5.6). The removal of carboxylic groups from the surface of the fabric could explain the lower points on the K/S curve. When the PAA macromolecules leave the surface, the PAN chains underneath are exposed to further nitrilase catalysis. This would lead to increasing numbers of carboxylic groups, thus, to an increase of K/S, until the threshold value for chain solubilization would be again achieved or no more accessible nitrile groups would be available for nitrilase. The PAA release would be dependent on factors such as microstructural properties of PAN copolymers, molar mass of the polymer chains, enzyme adsorption, nitrilase deactivation, and mechanical stress during treatment and washing. The important consequence of these findings is that the nitrilase biomodification of acrylic


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 115 needs to be properly controlled in order to maximize the concentration of the surface acidic groups.

5.4.2 Advantages and limitations of polyacrylonitrile biomodification Table 5.4 summarizes the various reports of biomodification of polyacrylonitrile and its copolymers. Despite the different substrates, origin, and amount of enzyme used (not forgetting the different methods used to measure the initial activity applied in the treatments), it is obvious that, at moderate temperature and pH conditions, it is possible to specifically modify the nitriles of PAN into amides or carboxylic groups, with distinct chemical properties. This is an important outcome implying that the use of enzymes in the particular case of PAN, a non-natural fibre, is not an unobtainable concept but a feasible process. Several aspects, such as the staining properties and hydrophilicity were clearly improved for the polyacrylonitrile copolymers. A higher hydrophilicity is of great interest because it would improve wear comfort (by increasing moisture uptake capacity and reducing the static charge accumulation), dyeability, and fastness of some finishes. Above and beyond the textile perspective, modified PAN is also of interest to the filtration technology field where it is commonly applied in reverse osmosis gas separation, ion exchange, ultrafiltration and dialysis (Frushour and Knorr, 1998; Masson, 1995). The amide and/or carboxylic groups at the surface of PAN would facilitate and open new routes for its coating or functionalization in these particular technological areas (Huang et al., 2005). Huang et al. (2005) also inferred from their results that these transformations happened mainly at the surface of fibres. Tauber et al. (2000) compared several techniques (FTIR, Raman microspectroscopy and XPS) having different beam interaction depths inside the sample; only XPS (1–5 nm) allowed differences to be detected in the chemical composition between treated samples and controls. Wang et al. (2004) also analysed some samples and controls by FTIR that showed no significant chemical changes in the bulk fibre. In addition, they microscopically analysed cross-sections of treated samples after acid staining and verified that the fibres were ring dyed owing to the superficial action of nitrile hydratase. Matamá et al. (unpublished results) confirmed the superficial action of a commercial nitrilase conjugated with fluorescein isothiocyanate (FITC). Cross-sections of acrylic fibres were observed by fluorescence microscopy (Fig. 5.8) and the fluorescence signal was located mainly at the surface, the core did not emit fluorescence indicating that the conjugated enzyme did not penetrate inside the fibres.


Nitrile hydratase/ amidase

Enzyme system

Treatment conditions

Fibres

10% methyl acrylate

Arthrobacter sp. ECU1101

Powder

Fibres

Fibres

10% vinyl acetate

7% (w/w) vinyl acetate

7

25

327

8000– 60 000

7

7

30

20

19 and 6–7 30 96

Acrylonitrile 69 and Powder homopolymer 6900

48

48

72

—

Amide

XPS analysis; 0.1% v/v Enhanced acid DMF staining

0.1% (w/v) JFC

Detected (XPS analysis)

Detected (XPS analysis; acid 2% DMF staining)

Not detected

Not detected

Not detected

—

Carboxylic group

—

—

Not detected

Detected

Ammonia

Product detection

Detected (XPS 0.5–1.5% analysis; DMF acid staining)

—

Time Tested Co-monomer Format Enzyme/ pH T (°C) (maximum) additives composition substrate (h) ratio (U g−1)

PAN substrate

Corynebacterium nitrilophilus ATCC 21419

Brevibacterium imperiale CBS 49874

Rhodococcus rhodochrous NCIMB 11216

Source

Best enzyme performance at pH 6.5 and 0.5% DMF

—

Additive effect on catalysis

Antistatic ability

Hydrophilicity improvement JFC assisted (water the catalysis, contact angle; DMF did not moisture regain)

No improvement in catalysis Hydrophilicity by DMF improvement (water contact angle)

—

—

—

Other properties

Table 5.4 Summary of conditions and major results from published treatments with nitrile degrading enzymes performed on polyacrylonitrile

Wang et al. (2004)

Battistel et al. (2001)

Tauber et al. (2000)

References

Nitrilase

Nitrilase and/or Nitrile hydratase/ amidase

Commercial

Micrococcus luteus BST20

Agrobacterium tumefaciens BST05

7% (w/w) vinyl acetate

Fabric

Fabric

Powder

5% vinyl acetate

?

Fabric

Powder

?

5% vinyl acetate

412

24

?

72

0.9

8

7

7

40

30

30

30

36

116

24

48

24

—

4% DMA +1M sorbitol

—

—

—

Detected

—

—

—

—

Detected

Detected

Detected (basic staining dependent on the treatment duration)

Detected

Detected

Detected (basic staining)

Detected (basic staining)

—

Detected Not Not (ESCA; significantly significantly FTIR-ATR) detected detected

Detected ? (FTIRATR)

—

—

Secondary product formation (soluble polyacrylic acid)

—

Fabric weight Improvement loss in catalysis

—

—

Smoothing PAN surface (SEM)

Matama et al. (2007)

FischerColbrie et al. (2007)

FischerColbrie et al. (2006)

118

Advances in textile biotechnology (a)

0

(b)

μm

25

0

μm

25

5.8 Epifluorescent photographs of cross-sections from (a) acrylic controls and (b) treated samples. The samples were treated with FITC-conjugated nitrilase at pH 8 and 30 °C, for 15 h. Both images were acquired with a total magnification of ×1000.

Although the superficial action of nitrile degrading enzymes was demonstrated in the examples studied, the fact that the desirable bulk properties, such as the mechanical properties, of PAN fibres remained unaffected was not verified in practice. The study of mechanical properties is a pertinent subject, to ascertain the impact of the enzymatic modification on PAN materials, especially, when the nitrilase was used and the PAA was detected in solution as a secondary product. The modification of a solid substrate with enzymes constitutes heterogeneous biocatalysis because enzyme and substrate are in different phases. Therefore, it is necessary to consider two general steps in order to make the catalysed reaction occur: the physical adsorption of the enzyme (including the transport of enzyme molecules from the bulk solution to the substrate surface) and the formation of an active enzyme/substrate complex that will lead to the desired product and to the regenerated enzyme. This type of enzymatic catalysis is, in general, less efficient than in homogeneous systems because of steric constraints, mass transfer and partition effects. The use of certain additives served the main purpose of increasing the accessibility of PAN to enzymes. The organic solvents DMF and DMA were chosen because of their action on the fibre structure. They are known solvents of PAN and they are commonly used in the industrial production of acrylic fibre (Burkinshaw, 1995; Capone, 1995). Their plasticizer function disturbs the regular structure of the polymer, reducing the magnitude of interchain bonding, which should aid accessibility to the enzyme, improving its action on the fibre. The results showed that DMF had an effect dependent on its concentration that was best around 0.5% (v/v) for nitrile hydratase, not taking account of the different enzyme sources (Tauber et al., 2000; Battistel et al., 2001; Wang et al., 2004). The tested concentration of the nonionic surfactant JFC was beneficial as was a mixture of DMA and sorbitol


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 119 in the treatment with nitrilase (Matamá et al., 2007; Wang et al., 2004). The available data shows the possibility of improving the action of these nitriledegrading enzymes on PAN by manipulating the treatment media or, from another perspective, the possibility of their application in the PAN wet spinning process, which uses the already mentioned organic solvents (Capone, 1995). One important issue involving the application of enzymes to textile materials is the mass transfer of enzymes in textile wet processes as discussed by Nierstrasz and Warmoeskerken (2003). Mass transport across a porous material of large molecules like enzymes is a relatively slow process and dependent on the properties such as the porosity and density of fabrics. Unfortunately, it is not always clear in published works if PAN fibres refer to loose fibres or to fabrics, and when fabrics are clearly used their characteristics are not always provided. Different textile structures will have different outcomes in time and homogeneity of the biomodifications and it is important to know which ones are used to have a more realistic perspective. The enhancement of mass transfer during the enzymatic treatment is normally accomplished by mechanical agitation; in a lab scale, the horizontal agitation is the most frequent though vertical agitation (simulating the laundry washing machines) was also used in the treatment of PAN (Tauber et al., 2000; Matamá et al., 2007). The applied mechanical forces that imply a deformation of the porous matrix of the textile material (as it happens in general on industrial equipments) will lead to a higher flow rate in the intra-yarns pores and consequently to a faster and more homogeneous biocatalysis (Nierstrasz and Warmoeskerken, 2003). Other important aspects of biocatalysts are their stability under operational conditions and their ability to be reused in an industrial process. Nitrile-degrading enzymes, in particular purified nitrilases, have an intrinsically low stability (Harper, 1977 and 1985; Bandyopadhyay et al., 1986). The reported enzymes (Table 5.4) were used as cell lysate preparations, either centrifuged or not, in order to guarantee the highest activity and stability; it was observed that the nitrile hydratase activity was associated with cell membranes fraction (Battistel et al., 2001; Fischer-Colbrie et al., 2007). Matamá et al. (2007) observed that the adsorption of nitrilase on acrylic fabric led to stabilization of the enzyme (a linear increase in ammonia over 36 h of treatment at 40 °C, Fig. 4.6), similarly to the immobilization procedures employed to stabilize proteins. Even in the case of enzyme stabilization upon adsorption to PAN substrates, it would be interesting to look for naturally more stable nitrile-degrading enzymes, such as those found in extremophile micro-organisms (Cowan et al., 1998; Khandelwal et al., 2007; Liu et al., 2008 Mueller et al., 2006; Pereira et al., 1998), because the presence of high temperatures and organic solvents during the wet processing of acrylic fibres can have deleterious effects on their activities.


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The ability of enzyme recovery and recycling as well as a general improvement in the long term operational stability is commonly achieved by biocatalyst immobilization (Bornscheuer, 2005). When dealing with PAN substrates, the immobilization of the enzyme is not in theory a good option because it creates a barrier to mass transfer and diffusion phenomena. The industrial enzyme recovery or elimination requires other methods such as size exclusion chromatography and pH/temperature shifts (Lenting, 2003). The production of known nitrile-degrading enzymes that can be redesigned to use more efficiently PAN as a substrate is, beyond doubt, needed. The incubation times necessary to produce the desired surface modifications are too long for a large-scale industrial application. In order to achieve a proper control over the acrylic biomodification it is also important to have a better understanding of how nitrilase acts on such a substrate. Is the enzyme very sensitive to the crystallinity degree of the fibre? How does crystallinity affect the release of polyacrylic acid? How is the molecular size of the polymer related to the released amount of polyacrylic acid? Is there any difference in treating the fibre before and after orientation? Before any attempt to design a full-scale industrial process, the enzymes themselves need to be engineered and further optimized at a laboratory scale.

5.5

Cellulose acetate biomodification

5.5.1 Esterase for the modification of cellulose acetates Matamá et al. (2010) reported the superficial hydrolysis of acetate surface groups of cellulose diacetate (CDA) and cellulose triacetate (CTA) fabrics using cutinase (EC 3.1.1.74) from Fusarium solani pisi. Cutinase is a serine esterase from the superfamily of α/β-hydrolases (Longhi and Cambillau, 1999) and it also belongs to the family 5 of carbohydrate esterases, sharing a similar 3D-structure with two other members with known structure: the acetylxylan esterase (E.C. 3.1.1.72) from Trichoderma reesei and the acetylxylan esterase II from Penicillium purpurogenum (Ghosh et al., 2001; Hakulinen et al., 2000). However, the active centre F. solani pisi cutinase reflects a preference for hydrophobic substrates, a suitable and versatile feature to be chosen for surface modification of highly substituted cellulose acetates, together with the flexibility of cutinase in using soluble and insoluble substrates (Ghosh et al., 2001). To evaluate the effect of enzyme concentration, the release of acetic acid was determined for samples of CDA and CTA fabric incubated over 8 h with various esterase activity concentrations, at 30 °C and pH 8. The acetic acid production increased over the range of the tested enzyme concentrations. At the maximum enzyme concentration used, the acetyl esterase activity was 0.010 and 0.007 U which corresponded to a release of 0.54 and


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 121 0.36% of the acetyl groups from CDA and CTA, respectively. These results agreed with studies that describe a higher level of acetic acid released for CDA than for CTA, a more substituted cellulose acetate. Studies reported on the biodegradation of cellulose acetate, using micro-organisms or cellfree enzymes, provide evidence that the degree of substitution is inversely correlated with the degree of deacetylation (Altaner et al., 2001, 2003; Moriyoshi et al., 1999, 2002; Puls et al., 2004; Samios et al., 1997). It was proposed that steric hindrance and crystallinity are important factors in the effectiveness of the adsorbed enzyme to promote hydrolysis, thus favouring CDA over CTA (Lee and Fan, 1982). Altaner et al. (2001) reported that acetyl esterases from 13 different commercial origins could significantly use cellulose acetates with DS ≤ 1.4 as substrates. Among them only one enzyme from Humicola insolens was able to release a small number (10%) of acetyl groups from a cellulose acetate DS 1.8, after 220 h. Another report described the application of an acetyl esterase from Aspergillus niger that was able to hydrolyze 5% of acetyl groups on a cellulose acetate DS 1.8 after 140 h, but was unable to release acetic acid for a DS 2.3 (Altaner et al., 2003). Comparing these values with those obtained for an assay carried out for only 8 h, it was demonstrated that recombinant cutinase had an activity with as great a potential as acetyl esterase for CDA and CTA materials. Evidence of hydroxyl-group formation on the surface CDA and CTA fibres was also obtained by the improvement in the chemically specific coloration of the fabrics with a reactive dye (Remazol Brilliant Blue R, C.I. 61200) (Matamá et al., 2010). Increases in K/S of 25 and 317% were obtained for diacetate and triacetate, respectively, after treatment with cutinase for 24 h. The results showed that cutinase was able to modify the surface of the cellulose acetate fabrics, increasing the number of hydroxyl groups (both in CDA and CTA), and lowering the carbonyl groups (detected only in CTA by DRIFT). Cross-sections of fibres treated with cutinase conjugated with FITC were observed by fluorescence microscopy. The fluorescence signal was located mainly at the surface, the core of the fibres did not emit fluorescence indicating that the labelled protein did not penetrate inside the fibres, therefore confirming the superficial action of cutinase on CDA and CTA fibres. Matamá et al. (2010) presented a novel approach to increase cutinase adsorption on cellulose acetate fibres. Using molecular genetic tools, they constructed chimeric cutinases by fusing the catalytic domain with a carbohydrate-binding module (CBM). Four constructions were obtained using two distinct CBMs, fused independently to the C-terminal of cutinase, and varying the linker DNA sequence (Fig. 5.9). CBMs were selected on the basis of ligand affinity, because the two cellulose acetate fibres are structurally distinct from cellulose (the native ligand) and each other, with different


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Advances in textile biotechnology Cutinase

N

Cutinase

C 280aa

1

Cutinase

N

wt CBM

C

Cutinase-wtCBMCBHI

346aa

1

NPSGGNPPGGNPPGTTTTRRPATTTGSSPGP

Cutinase

N

s CBM

C

Cutinase-sCBMCBHI

330aa

1

NPSGGNPPGGSSPGP

CBM

Cutinase

N

C Cutinase-CBMN1 430aa

1

Cutinase

N

PT

C Cutinase-PTboxCBMN1

CBM

451aa

1

(PT)4T(PT)7

HSV tag

His6 tag

5.9 Schematic representation of the recombinant wild-type cutinase from F. solani pisi and its new chimeric proteins with the fungal carbohydrate-binding module of CBH I, from T. reesei, and the bacterial carbohydrate-binding module N1 of CenC, from C. fimi. The amino acid sequence of the linkers is specified in the figure using the one-letter code (Matamá et al., 2010).

overall crystallinities. CBM type A, of the cellobiohydrolase I (CBHI) from T. reesei belongs to the family CBM1 and has a preference for crystalline or microcrystalline regions of cellulose, whereas CBM type B, of the endoglucanase C (CenC) from Cellulomonas fimi, which belongs to the family CBM4, is able to bind to amorphous cellulose (Boraston et al., 2004). A more detailed description of these domains is given in chapter 1. Figure 5.10 shows the results obtained in competitive colouring assays, normalizing the K/S values for the enzyme activity concentration. These results showed a clear difference for CDA and CTA on the relative K/S obtained with the chimeric cutinases versus the native enzyme, with a more pronounced effect for the less substituted acetate, irrespective of the CBM used. The cutinase-wtCBMCBHI and cutinase-sCBMCBHI are the most efficient catalysts under the treatment conditions used for both fibres. On CDA (Fig. 5.10), the relative K/S was improved around 7- to 8-fold by cutinasewtCBMCBHI and cutinase-sCBMCBHI, respectively, compared with native cutinase. On CTA (Fig. 5.10), the increase in K/S was improved around 2-fold for cutinase-wtCBMCBHI and 3-fold for cutinase-sCBMCBHI. The results showed the presence of stronger steric constraints in the triacetate fibre, owing to the triacetate backbone having more acetyl groups than the diacetate. The better performance of chimeric cutinases with the fungal CBM could be explained by the difference in size of both CBMs, being more constrained by the bigger bacterial CBM than by the smaller fungal


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 123

Relative K/S (lmax = 595 nm)

10

Cellulose diacetate Cellulose triacetate

8

6

4

2

0 Cutinase

cut-N1

cut-PT-N1 cut-wtCBM cut-sCBM

5.10 Relative increase in K/S values for cellulose diacetate and triacetate, both treated with wild-type cutinase and chimeric cutinases. The samples were incubated independently with native cutinase (cutinase), cutinase-CBMN1 (cut-N1), cutinase-PTboxCBMN1 (cut-PT-N1), cutinase-wtCBMT.reesei (cut-wtCBM) and cutinase-sCBMT.reesei (cut-sCBM), at pH 8 and 30 °C. Samples and the control were competitively colored at 60 °C. Relative K/S was calculated as (K/S − K/Scontrol)/(K/Scutinase − K/ Scontrol), where K/S is the K/S value for a treated sample, K/Scontrol is the K/S value for the control and K/Scutinase is the K/S value for the sample treated with native cutinase (Matamá et al., 2010).

CBM. Another reason could be the inherent mechanism for ligand recognition displayed by each type of CBM. The results demonstrated that hydrolysis of surface acetyl groups from CDA and CTA with a chimeric cutinase-CBM constitutes a promising approach to increase reactivity and hydrophilicity in these fibres, exhibiting an enhanced reactive dye uptake of treated fabrics compared with the native cutinase. Taking inspiration from nature’s strategies, we designed and produced chimeric cutinases fused with a CBM that increased the performance of native cutinase.

5.5.2 Advantages and limitations of cellulose acetate biomodification Cellulose acetate and triacetate can be given more reactive surfaces using a lipolytic enzyme – cutinase; this is a new and important finding: the surface ‘regeneration’ of cellulose composition can impart these yarns with a bicomponent character that is potentially of great interest in the sportswear field. Because cellulose acetate is becoming the raw material of choice for a new generation of high-tech products in separations technology, the


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pharmaceutical industry, controlled release and the biopolymer fields, all these areas can also profit from a surface bio-regeneration of cellulose reactivity (Glasser, 2004). The affinity of cutinase for cellulose acetates can be improved by adding CBMs, resembling natural carbohydrate esterases. However, because the washing procedure after the enzymatic treatments was mild, in terms of pH and temperature, it would be important to include in future works controls like null mutated or inhibited cutinases for a better assessment of the interference in the fibre properties of adsorbed protein. In order to transform the reported biomodifications into cost competitive industrial processes, a considerable amount of work is needed. A better biochemical characterization of cutinase fusion proteins is required to understand their interaction with the cellulose acetates. The optimization of the expression system is also needed for higher yields of protein production, essential from an economics point of view. The properties of modified cellulose acetates have to be characterized in detail before any feasibility study is carried out into the upgrading of the optimized processes to an industrial scale and the application of such modified fibres for the production of textile products having speciality properties.

5.6

Future trends

The textile industry shows successful examples of enzyme applications in the processing of natural fibres. For cotton, enzymes are used in almost all stages of its wet processing, almost completely replacing conventional textile practices such as stone washing of denim. However, for man-made fibres, in particular the synthetic fibres, the scenario is very different. Nature did not design efficient ways to use and integrate these fibres as part of ecosystems and, therefore, they are not natural substrates for enzymes. Moreover, because of their chemical structure, man-made fibres require high amounts of energy and chemicals for their modification in order to achieve the desired properties. Taking into account that they have a market share of more than 60% (Engelhardt, 2008), it is imperative to address the environmental and safety issues brought by their processing and disposal. Finding eco-sustainable alternatives to traditional practices for manmade fibres in an extremely competitive market, such as the textile industry, is a hard challenge. It is essential to find, develop and improve methodologies for the enzymatic treatment of a given fibre (based on the target chemical bonds of the polymer molecules). Nature’s biodiversity provides us with a large collection of enzymes well suited for supporting life, however, they may not always be well suited for our technological interests (Arnold,


Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 125 2001). Therefore, the impact of enzymes is dependent on our ability to tailor their properties according to the demands of a particular technological process. Some important issues that often need to be addressed are: limited substrate range; limited stability to temperature, pH and solvents; limited enantioselectivity; and limited turnover number (Powell et al., 2001). Although screening naturally occurring enzymes continues to be an important approach to find better biocatalysts, the manipulation of the protein molecule itself and/or the reaction media are gaining increasing importance and becoming essential for the competitiveness of bioprocesses. Nevertheless, both maximizing those enzymes found through natural diversity and the optimization strategies are necessary to achieve the final goal of having a novel enzyme addressing the exact and specific demands of an industrial enzyme application. Although here we are concerned with textile biotechnology, PAN and cellulose acetate are materials of increasing interest for other areas where surface modification is important in their application. Owing to its excellent mechanical properties, stability and low cost, PAN is used in water treatment, protein immobilization, ultrafiltration and dialysis (Wang et al., 2007). Cellulose acetate also finds many non-textile applications such as desalting (by reverse osmosis), hemodialysis, drinking-water purification (by ultrafiltration), as filters for laboratory use, and as stationary phases in chromatography (Shibata, 2004). In membrane processes, protein fouling and biocompatibility (biomedical field) are major problems. The main driving forces for these undesirable adsorption processes are electrostatic, hydrophobic and entropic effects (Wang et al., 2007). To improve the performance of the membranes, the general approach is to make the surface hydrophilic. In some applications, biocompatibility is also an important and complex issue that can be addressed by the immobilization of certain molecules, to provide reactive functional groups at the surface (Wang et al., 2007). Materials made from PAN or cellulose acetate with no or few chemically reactive functional groups need a surface activation process to create sites for further grafting or immobilization. So far, the surface activation has been accomplished through chemical or physical processes (Deng and Bai, 2003, 2004; Huang et al., 2005; Jia and Yang, 2006; Lin et al., 2004; Wang et al., 2007; Yim and Sefton, 2009). Enzymes are the future of surface activation, and membrane technology will probably be a future application of biomodified PAN and cellulose acetate. For these two materials, enzyme engineering is still an unexplored research area or in the early stages of development, and it is urgent to address it for the simplification of modification processes and to give a wider range of PAN and cellulose acetate applications.


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5.7


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5.8

References

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Enzymatic modification of polyacrylonitrile and cellulose acetate fibres 131 wang z-g, wan l-s and xu z-k (2007), ‘Surface engineerings of polyacrylonitrilebased asymmetric membranes towards biomedical applications: an overview’, J Membr Sci, 304, 8–23. doi:10.1016/j.memsci.2007.05.012. yarns and fibres exchange, 2007. Highlights from the Acrylic Chain Report 2007. Available from: http://www.yarnsandfibres.com/revamp_ir/report_fullstory. php3?id=401&p_type=%2062&source_id=15&source=YarnsandFibres%20 Paid&story_type=F&BF=Special&reportshow=First [Accessed 18 March 2009]. yim e k f and sefton m v (2009), ‘Amidine surface modification of poly(acrylonitrileco-vinyl chloride) reduces platelet adhesion’, J Biomed Mater Res, 89A, 780–790. doi: 10.1002/jbm.a.32022.


6 Enzymatic treatment versus conventional chemical processing of cotton I. R. H A R DI N, The University of Georgia, USA

Abstract: The processes of conventional chemical preparation that include desizing, scouring and bleaching are compared with those of fully enzymatic processes that, to date, include desizing and scouring but not bleaching. Much of the driving force to replace conventional cotton preparation chemistry has been the belief that enzymatic treatments can save energy because they operate at lower temperatures, and that the effluent from enzymatic treatments will be much easier to treat and clean up. The processes are evaluated with regard to effectiveness on cotton properties, and environmental and energy impacts, and their costs and benefits are compared. Key words: chemical processing, cotton, enzymatic treatment, scouring, biopreparation, bioscouring.

6.1

Introduction

Over the past 15 years, there has been intense and continuing interest in the development of enzymatic treatment of cotton fibers and fabrics. The ultimate goals are to use enzymes to desize the fabric, remove surface materials and impurities so that water absorption is enhanced, and decolorize the cotton to provide either a white yarn or fabric or a consistent white base for dye application (Thiry, 2008). Much of the driving force to replace conventional cotton preparation chemistry has been the belief that enzymatic treatments can save energy because they operate at lower temperatures, and that the effluent from enzymatic treatments will be much easier to treat and clean up. In the past decade, an additional driving force has been the increasing interest from the consumer in purchasing fabric and clothing that can credibly be said to be produced by more environmentally friendly processes than conventional products. To be successful in convincing textile wet processors to switch from conventional to enzymatic processes the latter must be shown to be simple, of equal or lesser cost, it must be demonstrably less difficult to treat the effluent, and the same or better quality fabric must be produced (Kuildred, 2006). 132 © Woodhead Publishing Limited, 2010

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In this chapter the processes of conventional chemical preparation, including desizing, scouring and bleaching, are compared with those of fully enzymatic processes that, to date, include desizing and scouring but not bleaching. There have been research efforts regarding the latter, but no effective industrial methods. The processes will be evaluated with regard to effectiveness on cotton properties, their environmental and energy impacts, and their costs and benefits.

6.2

Chemistry and structure of the cotton fiber

Cotton is by far the most important natural textile fiber in use. Most accounts put the worldwide usage of cotton at about forty percent of all fiber consumed for textile purposes (Fiber Organon, 2005). Cotton comes from plants of the genus Gossypium, with only 4 of the 33 species being of major commercial importance (Chaudry and Guitchounts, 2003). For most of the world, G. hirsutum (American Upland) and G. barbadense (Egyptian, and other extra-long staple varieties) are the major cotton fibers. Two other coarse short staple cottons, G. herbaceum and G. arboretum, are grown in southeast Asia. Chemically, the cotton fiber is typically about 95% cellulose, the polymer of β-d-glucose in which the units are linked at the 1 and 4 carbons (Fig. 6.1). The other roughly 5% of the raw cotton fiber is a chemical mixture of proteins, amino acids, pectic substances, hemicelluloses, waxes (high molecular weight alcohols and esters), and various organic and inorganic salts. Many of these low-molecular-weight constituents can be removed by hot water, but the rest require stronger treatments, and these treatments have conventionally been dilute (0.5–4.0%) sodium hydroxide (NaOH) at or near the boil in the scouring process. The proteins, pectins and hemicelluloses are connected in complex ways to the cellulose microfibrils in the primary wall. Figure 6.2 gives a representation of how these constituents might be connected (Agrawal et al., 2007). A more detailed description of how these materials are joined to one another, and the implications for needs in enzymatic scouring are given in Fig. 6.3 (Agrawal, 2005).

OH OH O

HO

O

O

HO

O

OH OH

n

6.1 Cellulose structure.


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Advances in textile biotechnology Glycoprotein Ca2+ bridges between acidic pectins

Ca2+

Ca2+

Ca2+

Hemicellulose

Cellulose microfibril Acidic pectin

Esterified pectin

6.2 Connections between cellulose and non-cellulosic constituents of the cotton primary wall (Agrawal et al., 2007).

Cuticle

Primary wall

Secondary wall

Lumen

Waxes Pectin, protein, hemicelluloses and metal ions

Cellulose

6.3 Model of cotton structure (Agrawal, 2005, after Kim, 2001).

The outer layer of the cotton fiber, consisting of the chemical mixture mentioned previously, is the protective coating that all plant cells have evolved to protect themselves from environmental degradation. Cotton lacks the lignin that plant cells generally possess. The wax that is contained in the thin outer coating cuticle of the cotton fiber is a complex mixture of fatty acids, and high-molecular-weight alcohols and esters. This cuticle has been estimated to be approximately 12 nm thick (Ryser and Holloway, 1985). A number of studies have analyzed the mixture of compounds in the cuticle. Some references classify these as saponifiable and non-saponifiable,



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referring to whether or not they are a fat or oil (solid or liquid triglyceride) that can be hydrolyzed into the salt of a fatty acid and an alcohol. Of the non-saponifiable compounds, the high-molecular-weight alcohols and esters and fatty acids are more difficult to remove unless the materials they are attached to in the cuticle structure are excised. A comprehensive listing of these compounds is reproduced in Table 6.1 (Agrawal, 2005). Most of these compounds have melting points below the boiling point of water (100 °C) and high-temperature treatments can be effective in removal, or at least in redistribution on the cotton fiber surface. Removal or redistribution of the wax materials can be of great importance in allowing access to the primary wall non-cellulosic compounds.

6.3

Cotton seed coat fragments

Greige (undyed) cotton fabric coming off the loom contains, in addition to the starch size, a certain amount of cellulosic matter from plant fragments created during harvesting. These are more prevalent in mechanically harvested cotton than in that picked by hand. Opening, blending and subsequent operations in the yarn-spinning process remove most of this plant structure matter. More problematic are seed coat fragments that are not removed during the yarn spinning processes because the fibers attached to these become part of the yarn structure. These seed coat fragments are visible as small dark dots in the fabric. Figure 6.4 shows seed coat fragments in the fabric, and under magnification. Seed coat fragments are created during the ginning process when the cotton seeds are removed from the harvested cotton mass by the saw teeth on the rollers in the gin. The fragments result when various forces applied to separate fibers from the seed coat pull off pieces of the coat and cause

(a)

(b)

6.4 Seed coat fragments in fabric: (a) unmagnified, (b) ×100.



23–47

?

Resins

3–8

Phytosterol, sterol, glucoside and polyterpenes

Fatty acid (with even number of carbon atoms) in free or esterified form

7–13

Saturated and unsaturated hydrocarbons

Saponifiable part, 37–47%

40–52

High molecular weight alcohol C23 to C34

Unsaponifiable part, 52–62%

% Dry weight

Main constituents

Type

C27H56 C30H60 C31H64 C32H66 C20H50 C20H50 C35H60O6 C30H50O C21H43O20 C15H30O2 C16H32O2 C18H34O2 C18H36O2 C22H44O2

C30H61OH C30H30O8 C28H58O C28H58O C28—C30

Chemical formula

368.6 242.0 256.4 282.5 284.5 340.6

414.7 414.7

438.8 518.5 410.8 410.7 —

Molecular weight

Details of each constituent

n-Triacontanol Gossypol (colouring matter) Montanyl (1-octacosanol) Octacosanol Other alcohol Glycerol (low molecular weight alcohol, esterified with fatty acid) Heptacosane Triacontane Untriacontane Dotriacontane β-Sitosterol γ-Sitosterol Sitosterol glucoside (sitosteroline) α- and β-Amyrin n-Tetracosanic acid (Lignoseric acid) n-Pentadecanoic acid Hexadecanoic acid (palmatic) cis-9-Octadecanoic acid (oleic) Octadecanoic acid (steric) iso-Behenic acid Coloured resinous products

Constituent

Table 6.1 Detailed composition of cotton wax (Agrawal, 2005)

84 69.6 64 4 70 80

140 147

87 184–214 83 83 —

Melting point (°C)

Major Traces Major Traces Major Traces

Major Major Major Traces —

Remark

Enzymatic versus chemical processing of cotton –OOC

O

H3CO HO

O HO O

O

O O

O OH

O OH

O O

O

H3C

OH

H3CO

137

O HO O

O O

n

OH

HO

6.5 Structure of xylan with ferulic acid linked.

these to break into fragments. In turn, portions of these seed fragments remain attached to the fibers and are eventually embedded in the yarn during the spinning process. In the greige fabric they appear as black or dark brown colored material with fibers or linters attached to its surface. Because fibers are attached to the surface of a seed coat fragment, they can be difficult to eliminate during the normal cleaning and fiber spinning processes. Chemically, seed coat fragments are composed of 22–26% lignin, 5–10% pentosan (xylan, Fig. 6.5), 43–48% cellulose, and small amounts of wax, protein and inorganics (Reicher and Csiszar, 1992). Lignin forms a complex matrix around the other components, providing a protective shield, but also making the seed coat fragments difficult to break down for removal. Physically, the cotton seed coats consist of six layers, which are segregated into two main groups identified as internal and external integuments, depending on their positioning in the seed coat. The internal integument consists of an inner epidermis towards the center of seed followed by inner pigment layer and palisade cells. The external integument consists of an outer epidermis layer followed by an outer pigment layer and a colorless layer (Himmelsbach et al., 2003; Kim et al., 2001). Figure 6.6 shows this diagrammatically. Dhandapani has examined the weight loss in seed coat fragments artificially created, and subjected to 0.5% and 4.0% NaOH for 1, 5 and 10 h (Dhandapani et al., 2008). The results showed weight losses from the seed coat fragments ranging from 31–42% for the 0.5% treatments to 59–64% for the 4.0% treatments. These weight losses may be on the high side since some linters attached to the fragments are clearly removed after treatment. However, the results do seem to indicate that even with vigorous alkaline treatment the fragments are diminished and broken up, but are not eliminated from the fabric.


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Advances in textile biotechnology Fiber Outer pigment layer (diazonium, ruthenium red) Palisade A (ruthenium red)

Outer epidermis Colorless layer (acid phloroglucinol)

Palisade B Inner epidermis (ruthenium red)

Inner pigment layer (vanilline)

6.6 Diagram of cotton seed coat structure (Kim, 2001).

6.4

Conventional chemical treatments and conditions

Conventional preparation of cotton fabric for further chemical treatment usually consists of desizing, scouring and bleaching. There are two common textile chemical processes that use NaOH in their treatment of cotton fabric, mercerization and scouring. The former process uses a concentrated solution of NaOH (18–20%) to change both the crystal unit cell of cellulose and to change the physical and chemical properties of the fiber itself. Scouring (also referred to as boiling off), on the other hand, is most often done with dilute NaOH, usually in the range of 0.5 to 4.0%, and usually at temperatures approaching 90 °C or greater. Less frequently, calcium hydroxide or sodium carbonate (soda ash) may be used (Vigo, 1994, p. 16). Chelating agents such as ethylenediamine tetra-acetic acid (EDTA) are sometimes added if significant concentrations of calcium and magnesium are present in the cotton or the water source. Scouring is done with the purpose of removing some non-cellulosic impurities from the surface of the cotton fiber to give the cotton fabric adequate water absorbance (hydrophilicity) for subsequent dyeing and finishing. This requires at least partial removal of the cuticle waxes and the pectins and proteins attached the primary wall cellulose. Other introduced impurities that require removal before chemical processing are dirt, oils and even defoliants applied to the cotton plant before harvesting. Alkaline scouring, followed by hydrogen peroxide bleaching, is usually successful in both breaking up seed coat fragments and in decolorizing them. Even if they are not completely removed, they are no longer visible to the eye. The conditions used for scouring depend greatly on the fabrics being processed. Heavier fabrics may require higher concentrations of NaOH and surfactant to improve penetration of the alkali. The surfactant may also be needed to help emulsify those wax constituents that are not saponifiable



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and have not been made water soluble. Another variable that can affect the amount of alkali and surfactant used is the depth of shade intended for the fabric in subsequent dyeing; darker shades can actually require less vigorous scouring. If cotton/polyester fabrics are being scoured, the treatment may need to be milder and at a lower temperature, since polyester is subject to alkaline degradation. Although it is not often used, solvent scouring of cotton and cotton/polyester blends has been done with trichloroethylene (Hull, 1980). This will only remove the cotton wax and subsequent alkali treatment would be necessary if hydrophilicity needs improvement.

6.4.1 The effect of properties of cotton on conventional treatments Alkaline scouring is a process that has been used successfully for centuries. It generally does not unduly damage the cotton fiber and fabric unless the fabric is allowed to be in contact with air during the process, or if trace metals such as copper or iron are either in the scouring bath or come into contact with the fabric while it is at elevated temperature and under highly alkaline conditions. In that instance, oxidative damage can occur from the formation of oxycellulose. The highly alkaline conditions can cause depolymerization of the cellulose by reducing end elimination through the β-alkoxy-carbonyl mechanism. Loss of fabric strength can then result. Another question that has been raised with regard to conventional alkaline scouring is the non-specific nature of the removal of cotton fiber surface materials. It may well be that many alkaline scouring processes remove more wax from the surface than is necessary for achieving the necessary hydrophilicity, or for successful and problem-free spinning of cotton yarns. A side issue regarding the removal of wax during alkaline scouring is the creation of a harsh hand (feel of the fabric) as a result of complete wax removal. The weight losses for alkaline scouring are typically in the range of 5.6 (0.5% NaOH) to 6.5% (4.0%) when treated for 1 h at 95 °C with a 0.05% non-ionic surfactant (Dhandapani et al., 2009), indicating that almost all of the non-cellulosic materials are removed in most scouring operations.

6.4.2 Environmental problems resulting from conventional treatments The desizing operations and the use of NaOH in scouring and bleaching create high chemical (COD) and biological (BOD) oxygen demand of effluent from the starch removal process, and significant salt concentration, high alkalinity and high pH in effluents from scouring and bleaching operations.


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These then necessitate effective treatment of effluent before it can be discharged to the lake or river from which the water was sourced or before the effluent is sprayed on to land for remediation. Each of these types of effluent discharges requires permits and is governed by either state or national regulations, or both. Alkaline scouring of cotton requires large amounts of rinse water to lower the pH of the fabric before further chemicals can be introduced. Given the increasing concern about water usage, both in terms of cost and in environmental impact, reducing water consumption is a continuing industry goal. Energy consumption is also an environmental and fiscal concern in conventional alkaline scouring. Scouring is done near the boiling point of water (100 °C) and multiple rinses are also at elevated temperatures so that the energy consumption is substantial.

6.5

Enzymatic treatments of cotton

6.5.1 Desizing using enzymes Most cotton and cotton-polyester blend yarns are sized before weaving in order to strengthen the yarns and increase their abrasion resistance. The sizing materials are usually starches, modified starches or starch and cellulose derivatives (Vigo, 1994, p. 4). Synthetic polymers such as polyvinyl alcohol can also be used alone or in combination with starches. The fundamentals of these sizing materials have been extensively reviewed (Drexler and Tesoro, 1984). Once weaving is accomplished and the greige fabric comes off the loom, the first step in preparation for chemical dyeing and finishing is the removal of the size. If the size is not completely removed after weaving, the results may be uneven dyeing or printing, and poor reproducibility from batch to batch. For starch and starch derivatives, which constitute the vast majority of sizing for cotton and cotton/polyester fabrics, desizing may be done by enzymatic, oxidative or acid desizing. The latter two approaches are little used because of their potential to damage the cotton fiber during the process. Probably more than 75% of the sizing agents used worldwide are starches or modified starches and, therefore, the use of amylases for desizing is widespread. These enzymes break the starch polymer molecules down into small fragments that can either be washed away easily or that dissolve in hot water. Amylases are enzymes that hydrolyze starches. The major types of these enzymes are α- and β-amylase. The α-amylase type is capable of acting at random locations along the starch chain to break down long-chain carbohydrates, ultimately yielding maltose from amylose, or maltose and glucose from amylopectin. Because it can act anywhere on the substrate, α-amylase tends to be faster-acting than β-amylase and is therefore used for textile © Woodhead Publishing Limited, 2010


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desizing. Amylases as classical desizing agents date back to the early 1900s when malt extracts from barley containing the active enzymes were applied to greige fabrics. However, in 1917 bacterial amylases were isolated and ultimately bacteria became the source for industrial production. After World War II, the technique of growing bacteria under water in a fermentation tank allowed mass production of the needed amylases. The use of Bacillus subtilus by Novo Nordisk for production of amylase was initiated in the early 1950s (Diderichsen, 1995). Since then, genetic engineering has made possible the development of more heat stable amylases that can be used over a wider range of pH than conventional products. For example a regular amylase may be applied at pH 5.5–7.0 and at 25–55 °C. Amylases for use at medium temperatures can be used in the range of 50–95 °C, whereas high temperature amylases can be used successfully above 95 °C and also in a pad-steam process (Kuildred, 2006).

6.5.2 Bioscouring using enzymes Although amylases have been used in desizing for decades, using enzymes for scouring cotton fiber is a relatively new idea. German research in the early 1990s alluded to the possibility (Bach and Schollmeyer, 1992; Rossner, 1993; Stohr, 1995). Shortly thereafter, research on cellulases (Tyndall, 1992; Buschle-Diller and Zeronian, 1992a, 1992b, 1993, 1994; Buschle-Diller et al., 1994; Buschle-Diller, 1998), suggested that other enzymatic treatments for cotton fibers might be feasible. From 1996 to 2000, reports appeared that addressed the possibility of successful scouring of cotton fabrics with enzymes (Buchert and Pere, 2000; Hardin and Li, 1997; Hartzell and Hsieh, 1998; Lange, 1996; Lange et al., 1998; Li and Hardin, 1996, 1998; Sawada et al., 1998). Although several enzymes, including cellulases, proteases and lipases, were examined, pectinases proved to be the most effective. Though their work was not published at the time, Novo Nordisk (now Novozymes) had discovered an alkaline pectinase that was particularly effective in creating water absorbability in cotton fabrics, and they subsequently announced its commercial availability in 1999 (Lange et al., 1999). This product had two major advantages over the enzymes that had been examined previously. The pectate lyase, marketed as ‘BioPrep 3000L’ had very high specific activity and it had its maximum effectiveness at pH 8–10, which made its use compatible with subsequent hydrogen peroxide bleaching conditions. Over the next decade there have been numerous reports dedicated to what is now called ‘bioscouring’ (Csiszar et al., 2001b; Degani et al., 2002; Lenting and Zwier, 2002; Lin and Hsieh, 2001; Luis et al., 2006; Tzanov et al., 2001). As bioscouring was being explored by scientists and industry, it became necessary to determine the degree to which waxes and pectins needed to be removed in order to create the desired level of hydrophilicity in the fabric. A series of combinations of acidic and alkaline pectins and © Woodhead Publishing Limited, 2010

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cellulases, alone and in mixtures, were examined for their influence on wax and pectin removal (Ismal, 2008). The results showed that all treatments created fabrics giving less than 3 s in the AATCC 79 water drop test method (AATCC, 2009). All the enzymatic treatments gave better hand (soft feel of the fabric) than alkaline scouring, and the weight losses for the enzymatic treatments were all less than those in alkaline scouring. A comprehensive study by the group at the University of Twente in the Netherlands has also addressed the question of the essentials needed for enzymatic scouring (Agrawal et al., 2007). A TRI auto porosimeter (Miller and Tyomkin, 1994) was used to study changes in hydrophilicity. The porosimeter was said to have the advantage over the commonly used AATCC drop test and various absorption tests in that the results are independent of fabric density and structure. From these measurements they were able to calculate a structural contact angle. The research group used these calculations to benchmark the effects of enzymatic treatments against standard alkaline scouring. From this work they concluded that wax and pectin removal were the essentials for scouring, and that in the enzymatic processes the removal of the outermost layer of wax was foremost. Another report from this group introduced a cutinase–pectinase mixture, combined with added mechanical action, that allowed them to reduce the scouring temperature from 50 °C with pectinase to 30 °C with the mixture (Agrawal et al., 2008).

6.5.3 Cotton seed coat fragment removal by enzyme treatment The complex nature of the chemical and physical structures of cotton seed coat fragments ensures that finding an enzymatic route to removing and/or decolorizing these contaminants will not be easy. Research by Csiszar and colleagues has dealt with attempting to find viable enzymatic approaches to removing seed coat fragments in order to create a total enzymatic approach to the scouring of cotton fabric. They demonstrated that cellulases could be used to clip fibers from the fragments, enhancing their tendency to fall off, and that the cellulases could also help to open up the fragments to further chemical attack (Csiszar et al., 1998a, 1998b). Further work demonstrated that if a chelating agent such as EDTA is added to hemicellulase and xylanase enzymes, then the seed coat fragments are hydrolyzed much faster than the cotton fabric itself. In addition, it was shown that the removal of the calcium ions enhanced the removal of colored impurities in the alkaline process (Csiszar et al., 2001a; Losonczi et al., 2005). Additional research for enzymatic approaches to seed coat fragment removal involved the investigation of the degradation of the lignins in seed coat fragments by xylanase (Csiszar et al., 2006). Tavcer concluded that for



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effective removal of seed coat fragments, a treatment with a combination of cellulases and pectinases, followed by hydrogen peroxide, was needed (Tavcer, 2008). A unique approach was the application of a hydrolytic and oxidative enzyme mixture produced by solid-state fermentation on seed coat fragments (Csiszar et al., 2007). The seed coat fragments were used as a carbon source for the production of the enzymes, which were then applied to seed coat fragments in fabrics, with promising results. At present, despite the advances that have been made, a completely enzymatic process for removing and decolorizing seed coat fragments in woven fabric has not been achieved, though the solid state fermentation approach seems to generate the specific enzymes that will be needed for success.

6.5.4 Application of mechanical and sonic power to increase mass transfer The biggest drawback to the use of bioscouring in the textile industry is that enzyme actions are relatively slow, and though bioscouring lends itself to use in batch processing, its diffusion-controlled process means that using it in continuous processes is difficult. Yachmenev et al. (2001, 2003, 2007) described the use of ultrasonic energy to increase the rate of mass transfer of enzyme molecules to the fiber surface. Moholkar et al. (2003) looked in detail at the mechanisms involved in these processes and concluded that optimizing cavitation intensity is of paramount importance to achieve an efficient ultrasonic wet textile process. Another approach to improving the efficiency of the enzyme processes on cotton was that reported by Agrawal et al. (2008), where, in addition to creating a cutinase–pectinase mixture to remove wax and then pectin, they applied mechanical action using a wedge apparatus to flex the fabric. The net result was a significant improvement in both the time needed for the desired increase in hydrophilicity, and a dramatic decrease in the temperature at which the process could be operated, down to 30 °C (Agrawal et al., 2008).

6.5.5 Effects of bioscouring on cotton properties Alkaline scouring is done to improve the water absorbance of the cotton fiber. It accomplishes this by removing a mixture of materials from the surface of the cotton fibers. It is non-specific and works by hydrolyzing both the materials themselves and/or excising connecting compounds that are attached to the pectins, proteins and waxy compounds. Bioscouring done with enzymes is very much the opposite, with enzymes by nature being quite specific in how they act on various molecular entities. Bioscouring can create the needed hydrophilicity in the cotton fabric, but at the same time


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leave sufficient wax materials on the surface of the fabric to lessen the later addition of softeners for consumer attraction.

6.5.6 Comparisons of costs Very little work has been published regarding cost comparisons of alkaline versus enzymatic scouring, partly because the full enzymatic process for continuous processing is not complete. Durden et al. (2001) reported on an in-plant trial on combed ring spun, open-end spun and mercerized cotton yarns. The trials were done with the commercial alkaline pectinase then available, though no details of the processes were given in the article. Substantial savings were claimed but no figures were given. Novozymes has reported the following for their pectinase scouring process (Wu, 2008): • •

Enzyme use: 10 kg per tonne of yarn. Chemicals, energy and water saved: hydrogen peroxide, 40 kg; NaOH, 15 kg; acetic acid, 5 kg; yarn, 25 kg; steam, 2500 kg; electricity, 150 kWh; water, 20 m3.

A press release of the results of a plant trial collaboration between Novozymes and Esquel, the Hong Kong-based textile manufacturer, reported a preparation regime that included bioscouring with pectinase, a dyeing that included a cellulase treatment to remove fiber fuzz ends, and another enzyme treatment to remove excess dye (just-style.com, 2009). It was calculated that 70 000 l of water and 1000 kg of CO2 per tonne of knitted fabric can be saved in production of t-shirts. They also claimed a 20–25% reduction in process time. Ismal (2009) examined water, materials and energy consumption and costs in some detail for various preparation treatments based on mill and production conditions. The results showed that enzymatic processes save water, energy and time compared with conventional treatments. When activator agent-assisted hydrogen peroxide bleaching was combined with enzymatic scouring, there were no time and electricity savings, but water and steam consumption and cost was reduced. Though the enzymes had higher costs than conventional materials, the enzymatic processes gave significant water and energy savings, and a lower total cost than the alkaline scouring processes.

6.6

Future trends

Given the initial quantity of research regarding enzymatic scouring and bio-preparation in the mid to late 1990s, it is somewhat surprising that there are still significant problems to be solved in creating a totally enzymatic cotton preparation system. Desizing with enzymes is a long-established



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method, but progress on solving certain problems in bioscouring has been limited. Bleaching by enzymatic means has had attention but no industrial methods in this area have been put forward. For bioscouring, pectate lyases can definitely provide the required properties of water absorption to cotton yarns and fabrics, but the time necessary for the reactions has thus far prevented their use in continuous systems where fabrics move through the chemistry quickly. The use of pectate lyases for knitting yarns and fabric in exhaust systems has been demonstrated and is being used in Asia. There is also some success in using the pectinases for wovens in certain kinds of semi-continuous systems. Nevertheless, the lack of a method to remove and/or decolorize seed coat fragments limits the bioscouring systems to yarns and fabrics that will be dyed deeper colors. Despite the problems alluded to, there is reason for optimism regarding full development of enzymatic preparation systems. The research on combining cutinases with pectinases to optimize cotton wax removal in shorter processing times, the application of energy (both mechanical and ultrasonic) to significantly speed up the reaction time, and the work with mixtures of hydrolytic and oxidative enzymes derived from solid-state fermentation to attack the seed coat fragment removal problem all show promise for changing the perceived roadblocks to wider implementation of bio-preparation. To efforts of these research initiatives can be added the fact of increasing interest by the public in textiles that can credibly claim to have less environmental impact, both in effluent treatment needed and in energy use.

6.7

References

aatcc technical manual (2009), ‘Absorbency of Textiles’, Test Method 79-2007, 101–102. agrawal p b (2005), ‘The performance of cutinase and pectinase in cotton scouring’, PhD Dissertation, University of Twente, The Netherlands. agrawal p b, klug-satner v a, gubitz g m, lenting h b m and warmoeskerken m c g (2007), ‘Wax removal for accelerated cotton scouring with alkaline pectinase’, Biotech J, 2, 306–315. agrawal p b, nierstrasz v a and warmoeskerken m m c g (2008), ‘Role of mechanical action in low-temperature cotton scouring with F. solani pisi cutinase and pectate lyase’, Enzyme Microb Technol, 42, 473–482. bach e and schollmeyer e (1992), Textilveredlung, 27, 2. basra a s (1999), Cotton fibers, Development biology, quality improvement, and textile processing, Binghamton New York, Food Product Press, The Haworth Press. buchert j and pere j (2000), ‘Scouring of cotton with pectinases and lipases’, Textile Chem Color Am Dyestuff Rep, 35(5), 48–52. buschle-diller g (1998), ‘Effects of scouring with enzymes, organic solvents, and caustic soda on the properties of hydrogen peroxide bleached cotton yarn’, Textile Res J, 68(12), 920–929.


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buschle-diller g and zeronian s h (1992a), ‘Enzymatic hydrolysis of mercerized cotton cellulose’, 204th Natl Meeting of the American Chemical Society, 6. buschle-diller g and zeronian s h (1992b), ‘Enhancing the reactivity and strength of cotton fibers’, J Appl Polym Sci, 45(6), 967–979. buschle-diller g and zeronian s h (1993), ‘Photodegradation of oxidized cotton cellulose’, J Appl Polym Sci, 47, 1319–1328. buschle-diller g and zeronian s h (1994), ‘Enzymatic and acid-hydrolysis of cotton cellulose after slack and tension mercerization’, Text Chem Color, 26(4), 17–24. buschle-diller g, zeronian s h and pan n (1994), ‘Enzymatic-hydrolysis of cotton, linen, ramie, and viscose rayon fabrics’, Text Res J, 64(5), 270–279. chaudhry m r and guitchounts a (2003), Cotton facts, International Cotton Advisory Committee (ICAC), Washington DC. csiszar e, losonczi a, koczka b, szakacs g and pomlenyi a (2006), ‘Degradation of lignin-containing materials by xylanase in biopreparation of cotton’, Biotechnol Let, 28(10), 749–753. csiszar e, losonczi a, szakacs g, rusznak i, bezur l and reicher j (2001a), ‘Enzymes and chelating agent in cotton pretreatment’, J Biotechnol, 89, 271–279. csiszar e, szakacs g and koczka b (2007), ‘Biopreparation of cotton fabrics with enzymes produced by solid-state fermentation’, Enzyme Microb Technol, 40, 1765–1771. csiszar e, szakacs g and rusznak i (1998a), ‘Bioscouring of cotton fabrics with cellulase enzyme’, in Ericksson K E L and Cavaco-Paulo A, Enzyme applications in fiber processing ACS Symposium Series 687, Washington DC, American Chemical Society, ch. 17, 204–211. csiszar e, szakacs g and rusznak i (1998b), ‘Combining traditional cotton scouring with cellulase enzymatic treatment’, Text Res J, 68(3), 163–167. csiszar e, urbanszki k and szakacs g (2001b), ‘Biotreatment of desized cotton fabric by commercial cellulase and xylanase enzymes’, J Mol Catal B: Enzym, 11, 1065–1072. degani o, gepstein s and dosoretz c g (2002), ‘Potential use of cutinase in enzymatic scouring of cotton fiber cuticle’, Appl Biochem Biotechnol, 102–103, 277–289. dhandapani r, wilson s s and hardin i r (2008), unpublished data. dhandapani r, wilson s s and hardin i r (2009), unpublished data. diderichsen b (1995), ‘A new desizing enzyme shows the changing nature of enzyme production’, J Biotechnol, 38, 193–197. drexler p g and tesoro g c (1984), ‘Materials and processes for textile warp sizing’, in Lewin M and Sello S B, Handbook of fiber science and technology: vol 1, chemical processing of fibers and fabrics, fundamentals and preparation part B, New York, Marcel Dekker, 1–89. durden d k, etters j n, sarkar a k, henderson l a and hill j e (2001), ‘Advances in commercial biopreparation of cotton with alkaline pectinase’, AATCC Rev, 1(8), 28–31. etters j n, husain p a and lange n k (1999), ‘Alkaline pectinase: an eco friendly approach to cotton preparation’, Text Asia, 83–86. Fiber Organon (2005), Fiber Economic Bureau, Inc., Arlington, VA. hardin i r and li y (1997), ‘Enzymatic scouring of cotton: effects on structure and properties’, Text Chem Color, 29(8), 71–76.



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hartzell m n and hsieh y l (1997), ‘Removal of non-cellulosic cell wall components from cotton fibers for improved wettability’, 213th ACS National Meeting, San Francisco. hartzell m n and hsieh y l (1998), ‘Enzymatic scouring to improve cotton fabric wettability’, Text Res J, 68, 233–241. henriksson g, akin d e, rigsby l l, patel n and eriksson k e l (1997), ‘Influence of chelating agents and mechanical pretreatment on enzymatic retting of flax’, Text Res J, 67, 829–836. himmelsbach d s, akin d e, kim j and hardin i r (2003), ‘Chemical structural investigation of the cotton fiber base and associated seed coat: Fourier-Transform infrared mapping and histochemistry’, Text Res J, 73, 281–288. hull r b (1980), ‘Weighing the alternatives in scouring technology’, Am Text Rep J, 9(6), 41–43. ismal o e (2008), ‘Influence of wax and pectin removal on cotton absorbency’, AATCC Rev, 8(6), 37–42. ismal o e (2009), ‘Economic profile and evaluation of cotton preparation – water and energy consumption’, AATCC Rev, 9(4), 42–47. just-style.com (2009), ‘Denmark: Novozyme trials eco-fabric treatment at Esquel’, December 16. karmakar s r (1999), ‘Chemical technology in the pretreatment processes of textiles’, Textile Science and Technol Series, Amsterdam, Elsevier, 12. kerstiens g (1996), ‘Plant cuticles an integrated functional approach’, Environmental Plant Biol Series, Oxford, BIOS Science. kim j, choe e k, kim s y and nam s w (2006), ‘Optimization of the enzymatic scouring’, in Gubitz G M, Cavaco-Paulo A and Kozlowski R, Biotechnology in textile processing, New York, Haworth Press, 155–168. kim j, kim s h and hardin i r (2001), ‘Physical structure and chemical composition of cotton seed coat’, J Korean Fiber Soc, 38(12), 612–619. klug b g (2004), ‘A new pectate lyase from Bacillus pumilus, PhD thesis, Graz University of Technology, Graz, Germany. klug-santner b g, schnitzhofer w, visanska m and weber j (2006), ‘Purification and characterization of a new bioscouring pectate lyase from Bacillus pumilus BK2’, J Biotechnol, 121, 390–401. kolattukudy p e (2001), ‘Polyesters in higher plants’, Adv Biochem Eng Biotechnol, 71, 1–49. kuildred h (2006), ‘Combined desizing and bio-scouring’, AATCC ICE Proceedings, Charleston, SC. lange n k (1996), ‘Enzymatic preparation of cotton’, Book of papers, AATCC international conference and exhibition, 101–108. lange n k, liu j, husain p and condon b (1998), Book of papers, AATCC international conference and exhibition, 463–471. lange n k, liu j, husain p and condon b (1999), ‘Cotton biopreparation – a new enzymatic concept’, Enzyme Bus, 10, 1–12. lenting h b m and zwier e (2002), ‘Identifying important parameters for a continuous bioscouring process’, Text Res J, 72(9), 825–831. lewin m and sello s b (1983), Handbook of fiber science and technology: volume 1, chemical processing of fibers and fabrics, fundamentals & preparation part A, New York, Dekker.


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7 Engineering of plants for improved fibre qualities M. W RÓBE L - K W I AT KOWSK A, Medical University in Wrocław, Poland; J. SZ OPA, University of Wrocław, Poland; and S. H AW K I N S, Université de Lille, France

Abstract: In this chapter, the various types of plant fibres used in fabrication of paper, cardboard and textiles are defined and the link between genes, the structure of plant cell walls, and fibre quality is explained. Examples of how biotechnological engineering has been used to modify fibre properties in cotton and flax are given. Recent advances in genomics, proteomics and metabolomics are discussed in terms of how they are contributing to a greater understanding of fibre formation and development, thereby paving the way for more sophisticated genetic engineering in different plant species. Key words: cotton, flax, fibres, genomics, lignin, plant cell wall.

7.1

Introduction

Natural fibres from a number of different plant species have long played an extremely important role in human society. For example, remains of linen textiles made from flax fibres have been found in prehistoric sites (9000 bce) from Syria and Israel (Hillman, 1975, van Zeist and BakerHeeres, 1975) and recently 30 000 year-old wild flax fibres were found in Paleolithic caves in Georgia (Kvavadze et al., 2009). Today, plant fibres are mainly used in the fabrication of paper, cardboard and textiles. Approximately 190 million tonne of paper and paperboard were produced from cellulose wood fibres in 2009 (UNECE 2009) and the world market for textile fibres has been estimated at around 55 million tonne a year with an annual growth rate of 2.5% (Sampaio et al., 2005). Cotton is currently the world’s leading natural textile fibre with over 27 million tonne of cotton fibres used annually (Chen et al., 2007; Singh et al., 2009). The long bast fibres from flax and hemp are also being increasingly used in the automobile and construction industries (in ‘green’, environmentally-friendly composite materials) because the physicochemical and mechanical properties of these natural fibres are similar to those of glass fibres (Baley et al., 2006; Joshi et al., 2004). The importance of plant ‘fibres’ in human and animal nutrition (fruit, vegetables) is also becoming increasingly clear. Since the beginning of agriculture man has selected plants in order to improve the quality of 150 © Woodhead Publishing Limited, 2010

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the derived products, and plant fibres are no exception (Batista and Oliveira, 2009). The dramatic increase in the availability of genomic and biological information is now allowing scientists and plant breeders to utilise different biotechnological approaches to improve the quality of plant fibres.

7.2

Defining plant fibres

Despite such a long history of fibre use, a clear, simple definition of a fibre does not exist. In reality, this reflects the fact that fibres can be classified/ described according to their botanical origin, their chemical structure, or alternatively, in terms of their industrial use. For example, from a botanical point of view a single fibre (elementary fibre) corresponds to a single elongated cell, usually tapering with a narrow diameter and a lignified or nonlignified thick cell wall (Esau, 1977). Plant cells, unlike animal cells, are surrounded by a relatively rigid structure called the cell wall that is composed mainly of polysaccharide polymers such as cellulose. After harvest, more or less all that remains of the elementary fibre is the cell wall and it is this structure that enters into the fabrication of yarns for textiles. In contrast, from an industrial point of view, the ‘technical fibre’ (for textiles and/or composites) often consists of an agglomeration of individual cells (elementary fibres) giving rise to an elongated narrow structure (McDougall et al., 1993). For composite materials, such a technical fibre can then be granulated to facilitate subsequent production. In addition, it is clear that when nutritionists tell us to eat more fruit and vegetables because they are rich in fibres they are not necessarily thinking of the same type of cell. In this review we will use the term fibre to refer to a single elongated plant cell. Plant fibres can be classified into three main groups (for detailed review, see Ilvessalo-Pfafli, 1995, McDougall et al., 1993) according to their botanical origin: (1) (2) (3)

stem fibres, leaf fibres, seed and fruit fibres.

Stem fibres can be further divided into two main groups according to the chemical composition of their cell walls. Thus, one can distinguish lignified fibres, typical of the inner woody tissues (xylem) of angiosperm dicotyledons and gymnosperms (conifers), from the cellulose-rich fibres (bast fibres) that contain little or no lignin and are found in the outer non-woody stem tissues of plants such as flax (Linum usitatissimum L.), ramie (Boehmeria nivea), jute (Corchorus capsularis) and hemp (Cannabis sativa). It should be noted that the relative quantity of the phenolic polymer lignin has an extremely important impact on both the mechanical and chemical properties of plant fibres, and therefore represents a major target for engineering.


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The second group of fibres are derived from the leaves of monocotyledon plants such as yucca (Yucca elata), banana (Musa textilis) and Agave species (e.g. Agave sisalana, A. fourcroydes). The leaf fibres are mainly used for the production of ropes or thick cords that can be used in furniture and floor coverings. In addition, fibres can also be derived from the whole stems of monocotyledons such as sugarcane (Saccharum officinarum) and bamboo (Bambusa and Dendrocalamus species). The last group contains fibres obtained from seeds and seed pods of both monocotyledons and dicotyledons and includes the major fibre plant cotton (Gossypium sp.), coir (Cocos nucifera) and kapok (Ceiba pentandra).

7.2.1 Fibre improvement Despite recent controversies (mainly in Europe) over the use of genetic engineering techniques to improve plant growth and product quality, it should be emphasised that man, through the selection/utilization of seeds from ‘elite’ (high-quality) individual plants has practiced plant improvement since the beginning of settled agriculture (Batista and Oliveira, 2009). More recently, a better understanding of genetics and the development of molecular markers have allowed the development of more efficient plantbreeding programmes in order to improve plant/product quality (Moose and Mumm, 2008). Over the past few years there has been a veritable explosion in the amount of genomic information available, not only for model plants such as Arabidopsis thaliana and poplar, but also for economically important species such as rice, wheat and maize, as well as for cotton (Chen et al., 2007). Research programmes are also underway to provide similar information in bast fibre species such as flax and hemp. The current availability of such genomic data will allow us to improve our understanding of the biological bases underlying fibre formation thereby facilitating quality improvement. In the context of fibre production, plants can be ‘improved’ either by increasing/optimizing plant growth and biomass production (quantitative improvement) and/or by improving the basic physicochemical and mechanical properties of fibres (qualitative improvement). Quantitative improvements have been obtained in a wide variety of different plant species by targeting ‘agronomic’ parameters such as growth rate, photosynthesis, water and nutrient-use efficiency, disease resistance, and stress resistance (Yuan et al., 2009). For example, growth rate can be altered by introducing genes to modify the biosynthesis of key hormones involved in signal transduction in plants (Morinaka et al., 2006, Peng et al., 1999). Because abiotic stress and pathogen attacks limit biomass production, increased resistance to these factors also represents a key objective. In tobacco and potato, transformation with the gene encoding DREB1A (dehydration response element



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B1A) has led to increased resistance to cold and drought stress (Behnam et al., 2007, Kasuga et al., 2004). Interestingly, one of the most important commercially available engineered plant species in the world is Bacillus thuringiensis (Bt)-cotton. This fibre-plant has been modified to express the bacterial gene Cry1Ac (from Bacillus thuringiensis) that is responsible for the production of an endotoxin that protects cultures from insect pests (Mendelsohn et al., 2003). Various studies have shown that the use of Bt cotton leads to both an increase in harvest yield (and hence overall fibre production) and a reduction in the use of chemical insecticides (Downes et al., 2007; Vitale et al., 2008). Cotton has also been engineered to express another bacterial gene that encodes the enzyme 5-enolpyruvylshikimate-3phosphate synthase (CP4-EPSPS) thereby conferring resistance to treatment with the herbicide glyphosate and allowing efficient weed control during culture (Nida et al., 1996, Yasuor et al., 2006). It is therefore clear that a wide variety of different plant species can be modified in order to improve plant growth under both optimal and sub-optimal conditions. However, apart from cotton, such technology has not yet been used to improve the agricultural properties of fibre plants such as flax or hemp. Qualitative improvements of fibres depend upon targeting (modifying) those factors that specifically influence quality. Fibre quality is influenced by: (1) (2)

the architecture of the fibre (length, diameter, cell-wall thickness) and the composition/structure of the plant cell wall.

It is these two factors that determine the resulting physicochemical and mechanical properties (such as tensile strength, flexibility, hydrophobicity). Potential plant improvement programmes aimed at improving fibre quality should therefore target genes implicated in regulating these two parameters. In addition, another very important target concerns those components of the cell wall that potentially hinder fibre extraction from the plant and subsequent (elementary) fibre separation. In this chapter, we mainly address the use of engineering to improve fibre quality.

7.2.2 Fibre quality and genes However, before addressing the question of how to improve fibre quality, it is necessary to explain (for the non-biologists) the link between genes and fibre quality. As stated above, fibre quality depends upon both fibre architecture (length, diameter, cell wall thickness) and fibre cell wall composition. Fibres are biological structures (cells) and, as with individual human cells, the formation and development of these cells depend upon the biological information contained in the genes present in the DNA of the chromosomes. For example, genes encoding a pectin methyl esterase enzyme


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and genes associated with auxin metabolism have been shown to be involved in the control of fibre and vessel length in poplar wood (Nilsson et al., 2008; Siedlecka et al., 2008), and actin genes and ethylene-related genes have been shown to be involved in fibre elongation in cotton (Li et al., 2005; Shi et al., 2006). Similarly, plant cell walls are a complex assembly of various polymers (polysaccharides such as cellulose, hemicelluloses and pectins, proteins, and the phenolic polymer lignin) (Fincher, 2009; Weng et al., 2008). These polymers are present in different amounts (and types) depending upon the fibre (species, tissue) and it is the variability of cell wall composition that explains the different properties of different fibres (McDougall et al., 1993). For example, the secondary cell walls of flax fibres are unusual because they are only weakly lignified, containing an atypical angiosperm lignin, as well as relatively high amounts of non-cellulosic galactans (Day et al., 2005a; Gorshkova and Morvan, 2006). The synthesis and assembly of these different polymers into the structure that we call the cell wall are under the control of a complex network of several hundred genes (Fincher, 2009; Mellerowicz and Sundberg, 2008; Zhong and Ye, 2007). Up- or downregulating the expression (‘activity’) of various ‘cell wall genes’ provokes modifications in the composition/structure of the plant cell wall. For example, considerable research over the past two decades has demonstrated that cell wall lignin content and chemical structure can be engineered in both model plants and economically important species (Vanholme et al., 2008; Weng et al., 2008). The challenge, if we want to improve the quality of plant fibres via a gene-based engineering approach, is therefore to identify genes and to improve our knowledge about what genes are involved in fibre formation and development. Until relatively recently, identification of flax genes was performed on an individual basis based on the assumption that, for example, a flax ‘lignification gene’ would have a similar structure to the same gene from a model plant such as Arabidopsis. Following identification, a reverse genetics approach (in which the gene is inactivated) has been used by different teams to understand the role of key genes in flax fibre (and oil) formation (Day et al., 2009; Fofana et al., 2006; Lacoux et al., 2003; Vrinten et al., 2005; Wróbel-Kwiatkowska et al., 2007a). Although such studies have provided useful information on the role of the selected gene, they are not high-throughput approaches and do not provide large-scale genomic information (i.e. simultaneous information about hundreds/thousands of different genes) about these biological processes. As a result, other research teams are now starting to develop whole transcriptome/proteome approaches to study fibre and oil formation in flax and hemp (Day et al., 2005b; De Pauw et al., 2007; Hotte and Deyholos, 2008; Roach and Deyholos, 2007, 2008). Such powerful molecular tools are not only enabling



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the identification of large numbers of flax genes, but they are also helping us to understand the molecular bases underlying the formation of celluloserich bast fibers. For example, the use of flax cDNA microarrays revealed that the expressions of genes coding for lipid transfer proteins (LTPs) and arabinogalactan proteins (AGPs) were particularly well correlated with fibre-elongation and cell-wall thickening, respectively (Roach and Deyholos, 2007). Such approaches have also allowed the identification of cell-wall modifying enzymes such as β-xylosidases and β-galactosidases (Day et al., 2005b; Roach and Deyholos, 2008).

7.3

Plant engineering methods

Despite public concerns about the safety of engineered plant crops (also referred to as genetically modified organisms or GMOs), there is no scientific evidence to indicate that such crops are any more dangerous than other new plant varieties produced through conventional breeding techniques (Batista and Oliveira, 2009; Twyman et al., 2009). As indicated above, modifications in fibre structure can be induced by targeting genes associated with fibre development and/or cell-wall polymer biosynthesis. In both cases, the objective is to modify the activity (expression) of a given gene (or group of genes), either by stimulating its activity (up-regulation) or by inhibiting/ reducing its activity (down-regulation). Modifications are achieved by introducing a small piece of DNA (the ‘transgene’) into the targeted plant and a number of different mechanisms exist (Kohli et al., 2003). One of the most commonly used methods of introducing genes into plants takes advantage of a naturally occurring transformation system based upon a common soil bacteria (Agrobacterium tumefaciens). This micro-organism was first identified at the beginning of the 20th century as being the causative agent responsible for the formation of the crown gall on plants (Smith and Townsend, 1907). Since then, Agrobacterium has been widely used as a vector to introduce genes into plant DNA (Batista and Oliveira, 2009; Gelvin, 2003). In this method, the ‘gene’ to be introduced is firstly integrated into a loop of circular plasmid DNA (the binary vector) and then the binary vector is introduced into Agrobacterium for plant transformation. This method has been used to transform fibre plants such as flax (Mlynarova et al., 1994; Wróbel et al., 2004) and cotton (Zhao et al., 2006). Despite, the recent banning (2009–10) of Canadian flax imports into Europe because of concerns about the presence of extremely low quantities of GM flax in non-GM seed lots, recent studies suggest that engineered flax, as with other engineered plant species, does not represent a danger for the environment (Batista and Oliveira, 2009; Jhala et al., 2008, 2009). Although different attempts to engineer hemp have not proved successful (Ebskamp, 2002), the recent development of a transformation and regeneration system


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for ramie (Boehmeria nivea) raises the possibility of introducing desirable traits into another fibre plant (Wang et al., 2009b). Another powerful way of introducing genes into plants is the particle bombardment method in which metal particles (usually gold or tungsten) are coated with multiple copies of the desired gene and introduced directly into tissues or cells via a special apparatus (gene gun) (Christou et al., 1988; Klein et al., 1987; Sanford, 1990; Twyman and Christou, 2004). The main interest of this method is that it can be used to engineer economically important species (maize, rice, sugarcane, wheat, sorghum, papaya, spruce) that are currently recalcitrant to transformation by Agrobacterium. More recently (Naqvi et al., 2009), attention has been focused on introducing several genes (multigene transfer, MGT) in order to engineer complete metabolic pathways. Such an approach is interesting since single-gene transfer is often unable to modify the physiological pathway targeted.

7.4

Some examples of engineering in fibre species

7.4.1 Cotton As indicated above, the most widely cultivated fibre plant is cotton (Gossypium hirsutum L.) (Chen et al., 2007; Singh et al., 2009). Cotton fibres are single, long (30–40 mm), epidermal cells of the seed coat that undergo synchronous elongation during maturation (Li et al., 2002). The elongating fibre is highly vacuolated and its maturation is associated with secondary cell wall formation and intensive cellulose biosynthesis, followed by dehydration and mineral accumulation. The mature fibre consists mainly of cellulose (85–90%) with the rest being composed of hemicelluloses and pectins (4–6%), waxes and fats (0.5–1%) and proteins (0–1.5%). Cotton fibres contain no, or extremely low amounts of, lignin and other polyphenolic materials thereby explaining their white coloration (Ioelovich and Leykin, 2008). The chemical composition of these fibres, together with their form, as well as their relative accessibility post-harvesting, all explain the predominant position of cotton in the textile industry. Engineering has previously been used to increase the resistance of cotton plants to insect attack (Mendelsohn et al., 2003) and herbicides (Yasuor et al., 2006). More recently, a specific ‘cell-wall’-related gene has also been targeted (Wang et al., 2009a). In this study, the down-regulation of the GhADF1 gene that codes for an actin depolymerising factor was shown to improve cotton fibre properties. This study indicated that the length and strength of the transgenic fibres were higher (5.6 and 2–12%, respectively) when compared with wild-type fibres. The secondary cell walls of modified cotton fibres were also thicker and the deposition of cellulose was increased by 3–5% compared with control fibres. Actin is a cellular protein that makes



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up part of the ‘cytoskeleton’ associated with cellulose deposition in plant cell walls (Joshi and Mansfield, 2007). Recent work implicating the role of ethylene in cotton fibre elongation (Qin et al., 2007; Shi et al., 2006) suggests that genes involved in the metabolism of this plant hormone could also represent interesting targets for engineering. Similarly, an interesting study on the effects of apyrases on cultured cotton ovules raises the possibility of other interesting targets (Clark et al., 2010).

7.4.2 Flax Flax is an annual plant that is cultivated for its bast fibres as well as for linseed oil that is rich in unsaturated fatty acids (Bloeden et al., 2008; Day et al., 2005a; Lee, 2003). This species, as for the great majority of cultivated plants, can suffer extremely important yield losses (up to 100%) as a result of disease caused by attack by fungi such as Fusarium spp. (Rashid, 2003). In one attempt to find a possible solution to this problem, flax plants were engineered to produce higher quantities of the plant ‘defence’ protein β-1,3glucanase that belongs to the ‘pathogenesis related-2’ (PR-2) protein family (Wróbel-Kwiatkowska et al., 2004). Glucanases of this type are known to be able to hydrolyze fungal cell walls thereby producing oligosaccharides that activate plant defences. Analyses showed that the engineered plants were almost three times more resistant towards F. oxysporum and F. culmorum in comparison with non-modified plants. The flax fibres used in textiles and composites are derived from the procambium and develop in bundles located in the outer stem tissues surrounding the central vascular cylinder. At maturity, the flax fibre cell wall is made-up of cellulose (70%), hemicellulose (15%), pectin (3%) and lignin (2–4%) (Day et al., 2005a; Gorshkova et al. 1996). Although present in much lower quantities than in woody fibres (where the lignin level can reach 30% of the total cell wall polymers), the presence of lignin in the middle lamella zones that join individual flax fibres together is believed to have a negative impact on fibre flexibility, as well as on fibre separation (Day et al., 2005a; Girault et al., 2000; Sharma et al., 1999). Because thread fineness (and hence textile quality) is related to the capacity to separate elementary fibres, lignin has naturally been one of the first targets for fibre cell wall engineering in flax. In one study (Wróbel-Kwiatkowska et al., 2007a), flax plants were engineered to under-express the cinnamyl alcohol dehydrogenase (CAD) gene. The CAD gene encodes the enzyme catalyzing the final step in the biosynthesis of lignin monomers (hydroxycinnamyl alcohols or ‘monolignols’) and down-regulation of this enzyme has been shown to reduce/ modify lignin content in other plant species (Baucher et al., 2003; Kim et al., 2002; Vanholme et al., 2008). Analyses of different down-regulated flax plants indicated that stem lignin content was reduced by 16–40% depending


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upon the plant line analyzed, and that soluble phenolic content increased. In addition, GC–MS analysis of cell wall fractions indicated reductions in the amount of pectin and hemicellulose cell wall sugars. In contrast, no changes were observed in the overall cellulose content of stem cell walls. Interestingly, the reductions in lignin content were associated with modifications in whole stem mechanical properties (increased Young’s modulus and tensile stiffness) possibly as a result of the changed ratio between cellulose and lignin/pectins/hemicelluloses. Initial tests also showed that CAD down-regulated lines were approximately twice as sensitive as control plants to infection with the fungal pathogen F. oxysporum sp. lini. Although such an observation would tend to suggest that the CAD engineering leading to improvement in stem mechanical properties could be offset by increased sensitivity to plant pathogens, further investigations are necessary. In particular, it would be interesting to confirm that the improved mechanical properties of in vitro whole flax stems are equally associated with modifications in the mechanical properties of individual fibres. In another study (Day et al., 2009), flax plants were engineered to downregulate another lignin biosynthetic gene – caffeoyl coenzyme A O-methyl transferase (CCoAOMT). The CCoAOMT gene codes for a lignification enzyme involved in the control of both overall lignin monomer content (quantity), and lignin structure. Previous work has shown that flax lignin is poor in lignin syringyl units (S-units) and is therefore chemically closer to the lignin type found in conifers (Day et al., 2005a). Analyses of greenhousegrown plants showed that CCoAOMT down-regulation was associated with reductions (8–18%) in stem lignin content, as well as with slight modifications in the structure of the lignin polymer. Although the mechanical properties of the flax stems were not evaluated, microscopic examination of stem cross-sections revealed that CCoAOMT down-regulation was associated with decreased cell wall thickness and reduced mechanical strength as indicated by the irregular outline of certain xylem cells. In addition, CCoAOMT down-regulation also appeared to be associated with increased xylem cell size. However, despite careful examination of engineered plants, no evident modifications in the structure of bast fibres were observed. Other cell-wall associated genes such as those associated with pectin remodelling could also represent potential targets for engineering in flax (Al-Qsous et al., 2004). Another engineering strategy used in flax aimed to improve retting efficiency (Musialak et al., 2008). As indicated above, flax fibres occur as bundles in the outer stem tissue, and have to be isolated during industrial processing. The first step in this process is called retting in which plants are left in the field after harvesting (dew-retting) and are colonized by a variety of soil micro-organisms. During this process, bacteria and fungi produce hydrolytic enzymes that start to degrade the intercellular layers between the fibre



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bundles and surrounding stem cells, as well as between individual fibre cells. In an attempt to facilitate retting, flax plants were engineered to naturally produce fungal pectinases (polygalacturonase and rhamnogalacturonase) that were hypothesized to weaken the pectin-rich intercellular layers (Musialak et al., 2008). Subsequent analyses showed that engineered plants contained reduced amounts of pectin and showed a greater than two-fold increase in retting efficiency compared with non-modified plants. In contrast, no changes were observed in either lignin or cellulose content. Interestingly, modified plants also appeared to be more resistant towards F. culmorum and F. oxysporum, possibly in relation with the observed increase in soluble phenolic levels.

7.4.3 Introduction of novel properties In addition to modifying fibre quality by targeting native cell wall polymers such as lignin, fibre plants have also been engineered to produce polymers that are not normally produced by plants. For example, genes coding PHB (poly-β-hydroxybutyrate) synthesis were introduced into cotton plants by particle bombardment (John and Keller, 1996). PHB is a hydrophobic and thermoplastic agent displaying physical and chemical properties similar to those of polypropylene and is naturally produced by bacteria as a source of carbon and energy (Bohmert et al., 2000). In this study, eight engineered plants (Gossypium hirsutum L. cv DP50) were shown to contain both phbB and phbC genes, and the presence of PHB in the cytoplasm of transgenic cotton fibres was confirmed by electron microscopy and HPLC, GC and MS analysis. Although the amount of PHB in the cotton fibres was very low (0.34% fibre weight), it influenced the thermal properties of fibres. The rates of cooling and heat uptake were lower in engineered fibres than in unmodified cotton fibres. Interestingly, the degree of modification appeared to be correlated with the PHB content of the different plants examined, thereby suggesting that improved thermal properties (for example in winter clothing) could be obtained by increasing PHB content. A similar strategy has also been used to increase the elastic properties of flax fibres (Wróbel et al., 2004, Wróbel-Kwiatkowska et al., 2007b). In these studies, three bacterial genes (phbA, phbB, phbC) encoding different enzymes of the PHB biosynthetic pathway were introduced into fibre flax (L. usitatissimum L. cv Nike) by Agrobacterium. Analyses of modified plants showed that although PHB synthesis was not associated with major disturbances in growth or fertility as previously observed in Arabidopsis (Kourtz et al., 2007), it did give rise to a slight decrease in straw and fibre yield of engineered plants. Nevertheless, the engineered plants also showed increased retting efficiency and significant improvements in stem mechanical properties. In addition, technical analyses of fibres obtained from field-grown


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modified flax plants also indicated that various mechanical properties (elasticity, flexibility, tensile strength) were improved compared with the control plants. Stems from modified flax plants showed a reduction in lignin levels and in non-cellulosic sugars (xylose, galactose, rhamnose, galacturonic acid) potentially associated with the observed increase in retting efficiency. Other modifications included significant reductions in the amounts of primary metabolites such as glucose, starch, fatty acid and citric acid. It is possible that such reductions are associated with an increased production of acetyl coenzymeA (acetyl CoA) necessary for PHB synthesis. In addition, linoleate levels were increased in seeds as previously observed (Wróbel et al., 2004). Engineered plants were more than two-fold more resistant towards the flax fungal pathogen F. oxysporum sp. lini and from 10 to 200% more resistant towards F. culmorum. It is possible that the increased resistance towards pathogens is related to the increased amounts of soluble phenolics observed in engineered plants. Despite this interesting observation, care should be taken since increased levels of phenols in engineered plants can have negative effects on plant growth (Besseau et al., 2007). Although these results clearly suggest that the production of PHB in flax improves fibre qualities, recent studies (Wróbel-Kwiatkowska et al., 2009) indicate that the situation might be more complicated. For example, engineering improved the mechanical properties of whole flax stems, but not those of fibres (except in one modified line M13). Similarly, chemical analyses indicated that although whole stems from engineered plants showed potentially interesting cell wall modifications (increased cellulose content and decreased lignin, hemicellulose and pectin), such changes could not be detected in fibres. These results were also confirmed by spectroscopic analyses (FTIR). Interestingly, such an observation is similar to other studies (Day et al., 2009) indicating that CCoAOMT down-regulation in flax modified the amount of lignin in whole stem extracts and was associated with reduced mechanical resistance in xylem cells, but had no apparent effect on fibres. Nevertheless, the observation (Wróbel-Kwiatkowska et al., 2009) that the arrangement of the cellulose polymer in transgenic fibres differed from that of controls, together with the fact that the modified cellulose contained a significant increase in the number of hydrogen bonds clearly confirms that PHB-engineering modifies not only the cell walls of xylem cells, but also those of fibres. In another study (Szopa et al., 2009), fibres from PHB-engineered flax plants and polypropylene (PP) were used to produce composite materials. Scanning electron microscopy (SEM) showed that modified fibres exhibited enhanced adherence to the polypropylene matrix compared with control fibres. The corresponding composite material also showed better mechanical properties than composites prepared with control fibres. In addition, biocompatibility tests indicated that modified fibres provoked little/no



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platelet aggregation, thereby opening up the possibility of using such materials for the development of biomedical devices that are in regular contact with blood.

7.5

Future trends

As indicated above, an increasing body of experimental work has demonstrated that modifications in gene expression can change fibre cell wall structure and properties in various fibre species. Currently, gene targets in individual fibre species are chosen on the basis of what we have learned about cell wall formation in other model species (e.g. Arabidopsis, poplar, tobacco) and have, until fairly recently, concentrated mainly on genes encoding enzymes involved in the biosynthesis of different cell wall polymers. However, because the formation of cell walls involves not only biosynthesis, but also remodelling of cell wall polymers and their interactions during development, the genes encoding plant cell wall enzymes responsible for such developmental changes also represent potential targets for modification (Aspeborg et al., 2005; Irshad et al., 2008; Pelloux et al., 2007; Sloan et al., 2009). Interestingly, such genes have also been detected in flax (Day et al., 2005b; Roach and Deyholos, 2008) and hemp (De Pauw et al., 2007). More recently, information is also starting to emerge about the ‘developmental programmes’ associated with the formation of the secondary cell wall. For example, a number of studies have underlined the role of MYB and NAC transcription factors in controlling both lignin biosynthesis and secondary cell wall formation (Goicoechea et al., 2005; Ko et al., 2009; Zhong et al., 2008). These studies suggest that certain transcription factors are able to act as ‘master switches’ regulating the whole secondary cell wall developmental programme by activating genes controlling the biosynthesis of the main structural polymers (cellulose, lignin, hemicelluloses). In contrast, other transcription factors such as MYB58 and MYB63 appear to function as specific transcriptional activators of lignin biosynthesis (but not other secondary cell wall polymers) in Arabidopsis (Zhou et al., 2009). In addition, other studies have indicated that some transcription factors can also act as negative regulators of cell wall biosynthesis (Legay et al., 2007). These results clearly show that targeted genetic modifications are capable of modifying cell wall structure. The first challenge to successfully manipulating fibre quality in economically important species is therefore to obtain a better fundamental understanding of the various gene networks involved in controlling cell wall formation in plants. Secondly, it is necessary to obtain more complete data about the genetic make-up (genomic data) of different fibre species. This is important since cell wall structure is mainly under genetic control, and structural differences (e.g. lignified versus non-lignified) mainly reflect differences in the genetic


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regulation associated with fibre formation. In addition, a better understanding of the genomes of fibre species also allows us to integrate knowledge obtained from studies on model plants. Recent advances in whole genomic, transcriptomic and proteomic approaches have enabled us to gain a much better idea of the genes involved in a wide range of different biological processes in both animals and plants (Harismendy et al., 2009; Lister et al., 2009), and such strategies are now being adapted for fibre plants. For example, recent transcriptomic and proteomic analyses have been used to gain an overview of the genes and proteins potentially involved in fibre formation in cotton (Hovav et al., 2008; Shi et al., 2006), hemp (De Pauw et al., 2007; van den Broeck et al., 2008) and flax (Roach and Deyholos, 2007; 2008). For genomic data (i.e. sequencing the genome of a plant), fibre plants are no exceptions and an international consortium exists to sequence the cotton genome (Chen et al., 2007) and the sequencing of the flax genome is currently underway (Deyholos, personal communication). Having ‘catalogued’ the different genes present, the next challenge is to find out the role of these genes in the formation and development of fibres. Such information can be obtained by comparing what is known about the corresponding genes (orthologs) in other model plant species (Arabidopsis, poplar, tobacco). In addition both forward- and reverse-genetics can be used to determine the role of different genes. For example, high-throughput screening of chemical mutant populations has proved to be an extremely useful tool to identify plants showing modified phenotypes, thereby allowing identification of the gene associated with the observed modification (Martin et al., 2009; Perry et al., 2009). Although some natural and induced mutants in fibre plants are known (Bretagne-Sagnard et al., 1996; Roland, 1991; Sengupta and Palit, 2004), it is only recently that high-throughput screening of large-scale populations of fibre plants such as flax has been developed (Deyholos, personal communication; Hawkins, 2009). Interestingly, as already done in the case of bioenergy research (saccharification), mutant populations could also be screened against various fibre processing enzymes in order to identify plants showing increased sensitivity/resistance. Complete understanding of a given gene’s role in a particular biological process (such as fibre wall formation and development) will also require the development of a ‘systems biology’ approach in fibre species and involving the integration of transcriptomics, proteomics and metabolomics (Saito et al., 2007; Yuan et al., 2008). Finally, another extremely important aspect concerns the impact of environmental conditions on cell wall structure (and hence fibre quality). For example, in maize, it has been shown that water stress provokes changes in lignin synthesis, as well as in cell wall proteins associated with cell expansion (Fan et al., 2006, Muller et al., 2007). Similarly, soil pollutants such as the heavy metal cadmium have been shown to modify cell wall structure in



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flax hypocotyls (Douchiche et al., 2010). A better understanding of the genes involved in such stress responses could also provide novel targets for genetic improvements as well as leading to a better control of fibre formation in different conditions.

7.6

Conclusions

Although it is only 27 years since the production of the first transformed plants, it is now clear that genetic engineering is able to significantly improve a wide range of plant characteristics (Chua and Tingey, 2006). An increasing number of recent studies are also showing that this technology is able to modify fibre structure, either by acting on native polymers, or by inducing the production of novel polymers. The main challenge is to identify the best targets (genes) for modification and this ultimately depends on improving our basic understanding of the relationship between genes and fibre structure. Recent advances in whole genome approaches and systems biology are allowing us to rapidly increase our knowledge of fibre species genomes. It is therefore clear, at least from a biotechnological point of view, that the engineering of fibre plant species is possible. Certainly, as highlighted by the recent European embargo on the importation of Canadian linseed, the success of this approach also depends upon public acceptance of this technology.

7.7

Acknowledgements

This study was supported by grants NR 12 0009 06 and NN 302061834 from the Polish Ministry of Science and Education, and by the ‘Conseil Régional du Nord Pas-de-Calais’ of France (Project ARCir, Plant Teq 4).

7.8

References

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8 Enzymatic treatment of wool and silk fibres J. SH E N, De Montfort University, UK

Abstract: In this chapter, the major enzymes used as bio-catalysts for protein fibre processes in the textile industry are described and the protein structure of wool and silk fibres is explained. Recent developments are presented in the application of enzymes in wool scouring and finishing processes to achieve a variety of finishing effects including softness, reduction of prickle and shrink-resistance. Recent developments in the application of enzymes in silk degumming and finishing are also reviewed. Key words: enzymes, proteases, wool, bioscouring, textile finishing, shrink-resistance, silk fibroin and sericin, silk degumming.

8.1

Introduction

There is increasing demand for natural textile goods by consumers. Wool and silk are the major protein-based natural fibres. They are well recognised as luxury textile fibres for use in clothing, upholstery, rugs and bedding materials including wool or silk filled duvets. The world’s wool retail sale per annum is about $80 billion with more than 2.1 million tonnes of greasy wool produced every year (IWTO Roadmap, 2009). Silk is produced in smaller quantities: about 130 000 tonnes per year, but silk has remained as a speciality fibre for use in high value textiles. Interest in the environment and consumer demands will certainly increase the use of these natural fibres. Natural wool fibre processing involves extensive wet treatments to improve the function and properties of fibres and fabrics including softness, reduction of pilling and dimensional stability. These treatments bring about some environmental problems such as increasing the chemical oxygen demand (COD) of wastewater from wool scouring, the presence of absorbable organic halides (AOX) in the case of shrink-resist finishing of wool and the use of heavy metals to obtain certain colour fastness. The textile industries are being asked to lessen the environmental impact of textile wet processes, especially restricting AOX in effluent, which has become part of the legislation in EU countries. Owing to such environmental concerns, much research effort has gone into searching for environmentally friendly methods for wool processing. The environmental benefits gained by using enzymes as biocatalysts in wool processes to replace harsh chemicals, 171 © Woodhead Publishing Limited, 2010

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especially for wool shrink-resistance, and simultaneously to improve dyeability, handle and whiteness of wool are well recognised. Silk has excellent properties. Apart from uses in textiles, in recent years there has been increasing interest in the application of silk fibroin to the development of various biotechnological uses and biomedical devices. The structure of silk protein has therefore been extensively investigated through amino acids sequencing and other studies. Biotechnology has been used in the processing of raw silk: for example enzymatic degumming to produce lustrous silk fibres for textile uses and also for the extraction of silk proteins, especially highly crystalline fibroin protein for medical uses. Enzymatic treatments of protein fibres have been extensively investigated with the focus on wool and silk. This chapter provides information on the major enzymes used for protein fibre processes and a brief explaination of the protein structure of wool and silk fibres. Recent developments in the application of enzymes in wool scouring and finishing processes to achieve a variety of effects including softness, reduction of prickle and shrink-resistance are reviewed. The review also includes recent developments in the application of enzymes in silk degumming and finishing.

8.2

Enzymes used for processing of protein fibres

Enzymes are biological catalysts for specific chemical reactions and they require comparatively mild conditions. All enzymes are proteins and biodegradable. The precise reaction specificity of an enzyme can be used for specific or targeted textile finishing without causing undesirable effects. Enzymes are classified by the Enzyme Commission of the International Union of Biochemistry into six groups: oxidoreductases; transferases; hydrolases; lyases; isomerases and ligases. Most enzymes used in the textile industry belong to Group 3, hydrolases, and this group includes the amylases, cellulases, pectinases, catalyses and proteases that are used for various textile applications such as desizing, bioscouring, bio-polishing, bleach clean-up and imparting wool shrink-resistance. Proteases, proteolytic enzymes or peptidases are general terms for enzymes that catalyse the hydrolysis of certain peptide bonds in protein molecules (forming the group EC 3.4.X.X of hydrolases). These have been suggested for use in wool processing to improve scouring efficiency, and handle properties, and to impart shrink-resistance and low temperature dyeability. Proteases are also used for silk degumming processes. Proteolytic enzymes can be divided into exopeptidases (which hydrolyse terminal peptide bonds) and endopeptidases (which hydrolyse peptide bonds inside the substrate molecule). Proteolytic enzymes can be further grouped according to the chemical nature of the catalytic site. They are divided between 13 sub-subclasses (Table 8.1).


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Table 8.1 The Enzyme Commission’s system of classification of peptidases (NC-IUBMB, 2008) Sub-subclass

Type of peptidase

Number of entries

3.4.11 3.4.13 3.4.14

Aminopeptidases Dipeptidases Dipeptidyl-peptidases and tripeptidylpeptidases Peptidyl-dipeptidases Serine-type carboxypeptidases Metallocarboxypeptidases Cysteine-type carboxypeptidases Omega peptidases Serine endopeptidases Cysteine endopeptidases Aspartic endopeptidases Metalloendopeptidases Threonine endopeptidase

20 12 9

3.4.15 3.4.16 3.4.17 3.4.18 3.4.19 3.4.21 3.4.22 3.4.23 3.4.24 3.4.25

4 4 20 1 9 98 54 38 80 1

Four distinct families are the serine endopeptidases (such as chymotrypsin, trypsin and subtilisin), the cysteine endopeptidases (such as papain), the aspartic endopeptidases (such as pepsin) and the metalloendopeptidases (such as thermolysin). In particular, serine endopeptidases have been extensively studied. Within Group 3, hydrolases, other enzymes such as lipases and cutinases have been used in the treatment of wool fibres to try to remove surface lipid from wool to improve the wettability of the fibres and improve subsequent dyeing and finishing. Both lipase and cutinase belong to the ester hydrolases (EC 3.1.X.X) enzyme class, which are capable of catalysing the hydrolysis of ester bonds. Lipases (triacylglycerol lipase EC 3.1.1.3) catalyze the hydrolysis of lipid esters. Cutinases (EC 3.1.1.74) hydrolyze cutin, the wax-like, amorphous biopolyester in the outermost layer of higher plants, composed of hydroxyl and epoxy fatty acids, and predominantly, esters. Cutinases are specific for primary alcohol esters (Liu et al., 2008). They have been used in order to remove the lipid layer from the wool surface, resulting in more efficient subsequent enzymatic treatment with proteases. There is increasing interest in use of the protein-crosslinking enzymes transglutaminases (EC 2.3.1.13) for surface modification of wool fibres. Transglutaminases belong to the class of transferases and act by a different mechanism from hydrolases. These enzymes can catalyse the posttranslational modification of proteins by the formation of isopeptide bonds. This occurs either through protein cross-linking via ε-(γ-glutamyl)lysine bonds or through incorporation of primary amines at selected peptidebound glutamine residues (Griffin et al., 2002a). This crosslinking leads to


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increased protein stability and increased resistance to chemical and proteolytic degradation.

8.3

Application of enzymatic treatments to wool fibres

8.3.1 Wool fibres Raw wool sheared from sheep contains a high level of grease which contains valuable lanolin as well as dirt, dead skin, suint, and vegetable matter. A typical Merino fleece contains about 13.6% wool wax, 3.1% suint and 12.9% dirt (inorganic and organic) (Prins, 2009). The average yields of clean dry wool vary according to the breed, but generally Merino types yield 60–70% of clean wool after processing. Electron scanning microscopy shows that the wool fibre is covered by a thin sheath of overlapping scales like the slates on a roof (Fig. 8.1). These scales are responsible for the felting and shrinkage that occurs during laundering. This is due to the differential frictional effect (DFE) caused by the difference in friction between μa (friction coefficient when rubbing in the against-scale direction) and μw (friction coefficient when rubbing in the with-scale direction). Finishers can make use of this property in a process known as milling to achieve consolidation of the fabric and make it fuller and denser. On the other hand, felting shrinkage can be detrimental to the performance of wool fabrics. Shrink-resist finishing is required to give wool fabrics and garments machine washability.

10μm EHT = 12.00 kV 3μm

WD = 10 mm Photo No. = 1147

Mag = 1.75 K X Detector = SE1

8.1 Scanning electron micrograph of wool fibres.



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An understanding of the structure of the cuticlar cells in the surface scales of the fibres is necessary for the development of successful enzymatic processes for wool preparation and finishing including wool scouring and shrink-resist finishing. The wool cuticle is about 400 to 500 nm thick. The cuticlar cells comprise an endocuticle (120 to 180 nm thick), an exocuticle (150 to 200 nm thick) and an outermost epicuticle layer (5 to 7 nm thick) (Fig. 8.2). The epicuticle layer contains 18-methyleicosanoic acid covalently bonded to the surface of wool fibre via a thioester bond to form a hydrophobic barrier (a fatty acid monolayer called the F-layer). The exocuticle layer contains a high proportion of crosslinked disulfide and isopeptide bonds, resulting in resistance to attack by alkaline agents and proteolytic enzymes. The endocuticle, a layer lying below the exocuticle, has a relatively low crosslink density (3% half-cystine) and thus is easily permeable and more susceptible than the exocuticle to chemical attack (Feldtman et al., 1983; Mori and Inagaki, 2006).

8.3.2 Enzymatic scouring Raw wool fibres contain significant quantities of contaminants including wool greases, suint and soil particles which could be up to about 60%. Wool scouring is the major operation in the manufacturing process to remove contaminants from raw wool. There are several methods of scouring wool either with aqueous solution or with solvents. In the aqueous system, the scouring line consists of a number of bowls of hot liquor containing water, detergent and alkali. The traditional scouring process not only consumes large quantities of water and energy but also leads to effluent discharge with a high organic content, COD from 5000 to 150 000 mg L−1, and sludge (Pearson et al., 1976). The environmental impact of wool-scouring and the cost of the effluent treatment and sludge disposal increasingly concern the

Epicuticle (12% half-cystine) Exocuticle ‘A’ (35% half-cystine) Exocuticle ‘B’ (15% half-cystine)

Endocuticle (3% half-cystine)

Intercellular cement (1% half-cystine)

8.2 Schematic diagram of wool cuticle (Feldtman et al., 1983).


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wool scouring industry because of the tightening of environmental legislation. The use of enzymes in wool scouring might improve the efficiency of the removal of grease and natural impurities from raw wool and reduce water consumption and scouring effluents. However, in the literature there are only a few reports relating to the bioscouring of wool fibres (Christoe, 1984; Das and Ramaswamy, 2006; Gouveia et al., 2008; Kholiya et al., 2008; Sedelnik, 2003). As well as grease, a proteinaceous residue is found on the surface of raw wool fibres. The water-insoluble and water-soluble components of this protein have amino acid compositions similar to skin flakes and suint peptides, respectively; they can be present in significant quantities in raw wool (Anderson and Christoe, 1984; Christoe, 1984). Christoe (1984) studied the effect of adding a proteolytic enzyme in single-bowl scouring experiments. The enzyme caused a change in the distribution of grease between the emulsion, sludge, and middle phases in centrifuging. The results also varied with pH when a protease (Esperase from Novozymes) was used in the scouring experiments. Under alkaline conditions scouring effectiveness was improved marginally, but there were significant changes in terms of the characteristics of the scoured wool. Interference between enzymes, surfactants and buffer needs to be considered in the wool-scouring process. The presence of buffer and surfactants may affect the activities of enzymes. It is well known that proteolytic enzymes are compatible with non-ionic surfactants that are widely recommended to be used in enzymatic treatments. A recent study (Zhang et al., 2006) has shown that the activity of the protease Esperase towards wool can be promoted by an ethoxylated alkyl phosphate anionic surfactant. Therefore, this anionic surfactant can provide additional benefits if chosen instead of the widely used non-ionic surfactants. This could lead to the development of a more efficient enzymatic scouring process. Buffers are used to maintain the optimum pH for protease activity during enzyme treatment, but it is found that different buffer systems and their ionic strengths have different effects on the activity of proteolytic enzymes. This is because of the interference of buffer cations and anions with the conformation of enzymes and their biological reactions. Wool scouring can be divided into three stages. The first stage is to remove a substantial amount of the water-soluble contaminants and dirt. The second stage involves scouring in hot detergent solution to remove the easy-to-remove contaminants, which include the bulk of the wool wax and most of the remaining dirt and water-soluble matter. The third stage removes the remaining hard-to-remove contaminants such as oxidised wool wax, fine particulate dirt and strongly-adhering non-wool proteins (Christoe et al., 1994). The scouring process and surface modification of wool fibres could be integrated by adding enzymes to the later stage of wool scouring line.



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Smith et al. (unpublished work) investigated the use of proteases and a protease enlarged by chemical modification in the raw wool scouring process, which led to improved dyeing and shrink resistance. It was shown that protease improved the scouring process and modified the cuticlar layer. A reduction of lipid content was found and this led to an improvement in the dyeability. Wool fabrics may contain vegetable matter such as burrs, seeds and shives which may attach themselves to the fibre and can pass through to the final stage of weaving or knitting. To remove vegetable matter from wool fabrics, the wool may need to undergo an additional chemical carbonizing process. Alternatives using enzymes such as cellulases, pectinases, hemicellulases and xylanases to remove vegetable matter from wool have been investigated in the scouring process (Sedelnik, 2003; Das and Ramaswamy, 2006; Gouveia et al., 2008; Kholiya et al., 2008). The enzyme treatments may promote the degradation of vegetable matter, resulting in its easier removal. It was reported that enzymatic treatment might be able to lower the concentration of sulfuric acid needed for the carbonizing process (Sedelnik, 2003). Cleaning raw wool fibres using enzymes needs to be further studied and developed. A process combining a mixture of different enzymes might make scouring more efficient: saving water and energy.

8.3.3 Enzymatic finishing The application of enzymes in wool-finishing processes to achieve soft handle, shrink-resistance and improved dyeability has been extensively studied and developed in last decade, and has been recently reviewed by Heine and Höcker (1995 and 2001) and Shen (2009). Early studies and more recent work (Bishop et al., 1998; Moncrieff, 1953; Shen et al., 1999) have confirmed that the action of proteases on undamaged wool is slow. This is because of the protective nature of the hydrophobic epicuticle surface containing fatty acid molecules and highly crosslinked cuticle cell components. However, once some of the cystine disulfide crosslinks in the cuticle cells are broken, the rate of enzyme reaction is greatly increased. It was found that during treatment with proteolytic enzymes, enzyme attack occurred preferentially at the highly swellable cell membrane complex by the enzyme penetrating between cuticlar cells and then between cortical cells. Once the enzyme has diffused into the membranes between the cells, it can rapidly disrupt the cell membrane complex, and damage the fibre if the reaction is prolonged. A fluorescence microscopy study demonstrated the diffusion of fluorescently labelled enzymes between the cuticle scales, through the cell membrane complex and into the cortical cells (Heine, 1991; Fogorasi and Heine, 2006). Therefore, it is difficult to limit enzymatic degradation to the cuticle scales and to achieve machine-


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washable wool without significant fibre damage (Heine et al., 2000). Recently, scanning electron microscopy (SEM) has been used to observe the degradation of wool fibre cross-sections caused by proteases digestion. The SEM images confirmed that the proteases preferentially digest the cystine-poor endocuticle, cell membranes, nuclear remnants, and intermacrofibrillar materials on the wool cross-sections (Okada et al., 2008). During enzymatic treatment of wool using proteases on their own, an early study (Moncrieff, 1953) showed that the attack on wool fibres is variable. The enzymes appear to attack weathered fibres more rapidly and therefore attack the tips of fibres more than the root ends. Irregularity of damage of fibres by enzymes was clearly shown by SEM. Bishop et al. (1998) reported that carefully controlled treatments with proteolytic enzymes can reduce the buckling load and collapse energy of wool yarns. These treatments were shown to improve the softness and reduce the subjectively perceived prickle of wool fabric knitted from the treated yarns. The combination of chlorination and treatments with proteolytic enzyme was also reported to improve handle properties, especially for coarse wool and mohair fibres, as well as improving fibre whiteness, without causing any significant damage to the wool (Holme, 2006). Enzymatic treatments for lipid removal have been studied to make the wool fibre more hydrophilic, in order to make subsequent enzymatic treatment more efficient. Triacylglycerol lipase (EC 3.1.1.3) and lipoprotein lipase (EC 3.1.1.34) have been used. Heine (Heine, 1991; Nolte et al., 1996) investigated the removal of lipids from the outer surface layer of the wool fibre by treating grease-free wool with a lipoprotein lipase. It was found that the aliphatic hydrocarbon content of the cuticle surface was reduced by 20%. Mall et al. (2002) used ToF-SIMS (time-of-flight secondary ion mass spectrometry) surface analysis to monitor the loss of surface lipids caused by treatment of scoured wool with lipases. It was found that the anionic surfactants left on the surface of wool fibre from the scouring process might inhibit the enzyme. No change in the fibre wettability was observed, probably because the integrity of the hydrophobic surface layer was not sufficiently attacked by the enzymes. Recently, Wang et al. (2010) investigated the effect of lipase treatment on the fibre surface of wool fabrics cleaned by soxhlet extraction with chloroform/methanol 87 : 13(v/v). It was reported that the lipase treatment hardly affects the cuticle surface of the wool, resulting in no noticeable improvement of wettability. The use of cutinase to hydrolyse the outermost bound lipids in the wool surface has recently been investigated. Cutinase has hydrolytic activity towards a broad variety of aliphatic esters. Wang et al. (2009) reported that treatment by cutinase can improve the wettability of wool fabrics from a contact angle of 135° to 92° after 4 h incubation with cutinase activity at 10 U g−1 fabric. However, cutinase can not achieve the complete removal



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of the covalently bound lipids from the wool surface even after prolonged treatment time. It was believed to be associated with the accessibility of cutinase active sites toward the outmost hydrophobic bound lipids in the wool fibre surface. Treatment of wool fabrics with cutinase has the potential to improve the wettability and facilitate the shrink-resist finishing of wool fabric by the subsequent treatment with proteases. Other chemical methods of lipid removal have been studied to make the wool fibre more hydrophilic, which may make the subsequent enzyme shrinkproofing treatment more efficient. These chemical methods are: treatment with a cationic surfactant, preferably an alkylammonium halide such as cetyltrimethylammonium bromide (CTAB) or cetyltrimethylammonium chloride (CTAC), in alkaline conditions (Julie et al., 1985; Smith et al., 2010); treatment with hydroxylamine and a non-ionic surfactant at pH 7–8 (Meade et al., 2005, 2008); treatment with potassium tert–butoxide under anhydrous conditions (Leeder and Rippon, 1985; Meade et al., 2008). Pretreatment with cetyltrimethylammonium bromide (CTAB) in alkaline conditions can make the fibre become very hydrophilic because of the removal of surface lipid. It was found, however, that residual CTAB on the fibre significantly decreased activity of proteases. After CTAB treatment, thorough washing with anionic surfactant in acidic conditions was required to maintain the enzyme activity. It was found that the activity of a protease enlarged by chemical modification was improved on wool that had been treated with CTAB and washed with anionic surfactant, because chemical modification made the enzyme more resistant and stable (Smith et al., 2010). A two-step treatment consisting of a low-temperature plasma treatment (LPT) and subsequent enzymatic treatment by proteases has also been investigated to modify the cuticle cells to achieve wool shrink resistance (Dybdal et al., 2001; Jovancic et al., 2003). X-ray photoelectron spectroscopy (XPS) analysis revealed that the LPT/enzymatic treatment completely removed the outermost lipid layer (the F-layer) of the epicuticle. However, in order to avoid excessive damage to wool fibres, the subsequent enzymatic treatment was mostly used to remove fibre ends protruding from the surface of the fabric and thus increase its softness. In order to achieve machine-washable wool, severe treatments with proteases are required to degrade the cuticlar cells of wool fibres. It is difficult to avoid significant fibre damage caused by proteases. Scientists have made great efforts using different wool pre-treatments before enzyme treatment in order to limit enzymatic degradation to the cuticle scales. Various combinations of enzymes, oxidants/reductants and polymers have been proposed to improve resistance to felting and shrinking of wool (Cardamone et al., 2006; Jovancic et al., 1998; Levene and Shakkour, 1995; Levene et al., 1996). In some of the early enzyme-finishing processes, wool was pre-treated by chlorine or hydrogen peroxide before incubating the fibres with


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proteases. Oxidative treatments of wool can disrupt disulfide bonds and open up the wool fibre surface assisting enzymatic attack on the cuticle. In addition, oxidative pre-treatment probably causes a partial removal of the fatty acid barrier from the epicuticle, which confers hydrophilicity to wool (Cardamone et al., 2005). Consequently, the enzymatic attack on the cuticle can be selectively activated. It has been claimed that a combination of chlorination with chlorine gas or dichloroisocyanuric acid and subsequent enzyme treatments with a protease such as papain showed that preoxidation will limit enzymatic attack to the cuticular layer, resulting in the enzymatic descaling of wool fibres to enhance not only lustre but also shrink-resistance (Levene and Shakkour, 1995; Moncrieff, 1953). However, the process may cause fabric yellowing. Recently a two-step process which combines bleaching, shrinkage prevention and biopolishing was suggested as a way to make wool feel silky smooth. This involved a pretreatment using hydrogen peroxide enhanced by dicyandiamide and stabilised by gluconic acid allowing powerful oxidation. This was followed by treatment with proteases in the presence of sodium sulfite in triethanolamine buffer solution. Benefits claimed were a high level of whiteness, the removal of protruding fibre ends for fabric smoothness and shrink-resistance (Cardamone et al., 2004 and 2005). Other oxidising agents used as a pre-treatment for wool are peroxymonosulfuric acid, peracetic acid and potassium permanganate. Alkali and reducing agents such as bisulfite can alternatively be used to open disulfide bonds in the cuticle scales to make the fibre more susceptible to enzyme attack. These pre-treatments have been reported to enhance the activities of proteases and improve the efficiency of proteases in conferring anti-felting and shrink-resistance of wool (El-Sayed et al., 2001; Levene and Shakkour, 1995; Levene et al., 1996). However, these processes caused severe fibre damage when shrink resistance was reaching adequate levels for machine-washable wool. Recently, Lenting et al. (2005 and 2007) used the addition of a high concentration of sodium salt in a peroxide pre-treatment to restrict the oxidative reaction to the surface scale of wool fibres. This resulted in an improvement in the susceptibility of the protein layer in the outer surface to subsequent proteolytic hydrolysis. This agreed with an earlier study which suggested that the presence of high concentrations of salt lowered the swelling of the fibre and lowered the rate of diffusion of the oxidant through the cuticle to the cortex. This led to preferential attack on the cuticle (Maclaren and Milligan, 1981) and wool shrink resistance was claimed without substantial loss of fibre tensile properties. Although there is currently considerable interest in the use of enzymes to achieve shrink-resist finishing on wool, it is apparent that the results of enzyme treatments, especially with proteases, can be unpredictable and may sometimes lead to unacceptable degradation of the fibre.



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The EU funded PROTEX project (contract no.: G1RD-CT-2002-00695) has developed a new approach by increasing the size of proteases in order to limit the enzymatic degradation of wool fibre to its cuticlar scales (Cavaco-Paulo and Silva, 2003; Silva et al., 2004, 2005, 2006; Smith et al., 2008). The increase in the size of the proteases was achieved by covalently attaching them to soluble Eudragit polymer (an enteric copolymer of methacrylic acid–methyl methacrylate). This chemical modification of proteases with Eudragit improved the thermal stability of the enzymes as well as enzyme recycleability owing to soluble–insoluble reversibility of the Eudragit polymer attached to the enzyme. Modification of the enzyme was further developed through a genetic engineering approach to enlarge the size of proteases in a larger EU project ENZUP (contract No. 032877ENZUP) under the 6th EU Framework Programme. Detailed information on the genetic engineering of proteases is given in chapter 1 and recent publications (Araújo et al., 2009; Cavaco-Paulo et al., 2009). Shen et al. (2007) have published the results from bulk trials on wool fabrics carried out using the modified proteases. It was shown that the modification of the protease enabled the reaction of the enzyme with wool to be controlled, so that less degradation of the wool occurred compared with similar treatments with the unmodified protease. An anti-felting effect was achieved without any significant weight loss during the treatment. This novel enzymatic process leads to environmentally friendly production of machine-washable wool. Lenting et al. (2009) reported the industrial trials of wool fibres treated with modified proteases, and processed the treated wool as filling in the production of duvets. The machine-washability of these wool fibre-filled duvets was tested in a household washing machine using both the wool and normal wash programmes. The substantial improvement in shrink-resistance was achieved even at the high temperature of the normal washing programme. However, the treatment of the wool fibres with modified enzymes showed less impact on maintenance of the thickness of a fleece filling during machine washing. Alternatively, transglutaminases (EC 2.3.2.13) have been used for protein crosslinking within the fibres to compensate for the reduction of tensile strength and degradation of wool during treatment with an oxidative agent (e.g. chlorination for shrinkage prevention), a reducing agent (sodium sulfite) and/or a protease (Cortez et al., 2004; Du et al., 2007; Griffin et al., 2002b; McDevitt and Winkler, 1998). It was reported that treatment of wool with transglutaminases can slightly reduce the tendency to felting of wool without causing any negative effect of stiff and harsh handle. Hossain et al. (2008) used simultaneous protease and transglutaminase treatment of woven wool fabrics in a single bath. It was reported that the treatment reduced the felting tendency of woven wool fabrics by 9% with only 2% weight loss and tensile strength loss. Cardamone (2007) reported that the


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post-treatment with transglutaminase can regain 3–5% fabric strength from 10–18% strength loss during the peroxycarboximidic acid oxidation and protease treatment of wool. Transglutaminase has also been shown to graft protein polypeptides, or fluorescein cadaverine onto the surface of the wool fibre for special effects. It is reported that the crosslinking within wool fibres by transglutaminase can improve the resistance to enzyme attack, and increase the resistance of wool to chemical and mechanical disruption. This was claimed to be beneficial for protecting wool garments from detergent damage (Cortez et al., 2005, 2007). However, the amount of lysine and glutamine residues available for linkage by transglutaminase may be limited, especially on the surface of undamaged wool fibre, restricting the extent of enzyme reaction. Therefore, this may affect the wide application of transglutaminases on wool.

8.4

Application of enzymatic treatments to silk fibres

Silk is a natural protein fibre. The best-known type of silk is mulberry silk (Bombyx mori) obtained from cocoons produced by silkworm larvae, which are cultivated in provided habitats and fed with fresh mulberry leaves. There are other less cultivated silkworms, classified as wild silk, such as the Tussah silkworm, the Muga silkworm and the Eri silkworm, which feed on oak, polyanthus and castor oil leaves, respectively. Raw silk spun by the silkworm Bombyx mori is a continuous strand (about 300 to 900 m) of two fibroin filaments cemented together by a gummy substance called silk sericin or silk gum. Silk fibroin filament is the fibrous component that gives silk its unique physical and chemical properties. Sericin gum in raw silk provides protection during processing and so is usually retained until the yarn or fabric stage. Removing harsh and stiff sericin from silk fibroin leaves the silk fibre lustrous and with a soft handle. Silk is regarded as a luxury fibre and widely used for textile fabrics, garments and bedding materials including silk-filled duvets. Natural raw silk is composed of about 70–80% fibroin and 20–30% sericin on the weight of the raw silk, respectively, with the trace of other natural impurities such as wax, carbohydrates and mineral salts. Fibroin and sericin are two different types of protein which differ in their composition and structure. The difference between fibroin and sericin makes the removal of sericin gum from fibroin possible without causing considerable damage to the fibroin filaments. A manufacturing process for removing sericin gum from silk fibroin is known as the degumming process. This can be carried out in the stages of unreeled silk cocoons, yarns or fabrics. Some papers have comprehensively reviewed degumming processes (Gulrajani 1992; Mitra et al., 2009). Degumming processes have been developed based on extraction with boiling water and superheated water at about 115 °C,



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degumming with soap in alkaline or acidic solution, and enzymatic degumming. The most extensively used method is degumming with soap or detergents. Among them, the expensive Marseilles soap is outstanding for silk degumming. In alkaline conditions, soap makes sericin swell. It is then emulsified and removed from the fibroin filaments. Sericin gum can also be removed by using water only, but a high temperature, around 110–120 °C, and pressure is required to obtain complete degumming. In recent decades, research effort has gone into the development of enzymatic degumming of silk, which can be carried out under mild conditions and at low temperature, as an alternative to lessen the environmental impact of the chemical process (Arami et al., 2007; Gulrajani et al., 1996, 2000a; Freddi et al., 2003; Shukla et al., 1992; Sonthisombat 1993). An understanding of the difference between the protein structure of silk fibroin and sericin is necessary for the development of successful silk degumming. In recent years, there has been increasing interest in the use of silk fibroin in the development of various biotechnological and biomedical devices and applications. Fibroin protein has therefore been further investigated and attempts have been made to elucidate its structure (Asakura et al., 2002, 2005; Fossey et al., 1991; Lefevre et al., 2007; Mita et al., 1994; Takahashi et al., 1999). Fibroin has a predominance of five amino acids: 44.4% glycine, 30.2% alanine, 11.9% serine, 4.9% tyrosine, 2.1% valine and 6.5% other amino acids (Marsh et al., 1955a and 1955b). It consists of heavy-chain polypeptides (∼350 KDa) and light-chain polypeptides (∼25 KDa). Fibroin is about two-thirds crystalline and about one third amorphous. The primary structure of Bombyx mori fibroin may be approximately divided into repetitive region (R) and amorphous region (A) along the heavy fibroin chain (Fig. 8.3). Mita et al. (1994) and Zhou et al. (2000) predicted from the complete sequence of the Bombyx mori fibroin gene that the repetitive region was subdivided into three regions (Asakura et al., 2002): (a) (b)

(c)

highly repetitive -Gly-Ala-Gly-Ala-Gly-Ser- (GAGAGS) sequences constituting the crystalline region, relatively less repetitive -Gly-Ala-Gly-Ala-Gly-Tyr- (GAGAGY) and/or -Gly-Ala-Gly-Ala-Gly-Val-Gly-Tyr- (GAGAGVGY)sequences comprising semicrystalline regions containing mainly hydrophobic moieties, sequences similar to (a) plus an extension by AAS.

The crystalline blocks are composed of short-side chained, highly repetitive amino acid sequences, typically leading to the formation of antiparallel β-pleated sheets (Marsh et al., 1955a; Takahashi et al., 1999). In the crystalline region, these polypeptides chains, aligned along the fibre axis, are held together by a close network of interchain hydrogen bonds between the >CO and >NH groups within the β sheets while the β-pleated sheets are held


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Advances in textile biotechnology R02

R01

A01

R03 R04 R05 R06

A02

A03 A04

R07

A05 A06

R08 R09 R10

A07

R11

A08 A09 A10

R12

A11

a a a a b b b a a a a a a a a a a a a b b b b c

Repetitive crystalline regions a (GAGAGS)

b (GAGAGY) or (GAGAGVGY) c

(GAGAGSGAAS)

Amorphous regions consensus sequence: TGSSGFGPYVANGGYSGYEYAWSSESDFGT

8.3 Schematic representation of the organization of the B. mori fibroin heavy chain gene. One-letter standard amino acid abbreviations are used (Asakura et al., 2002).

Table 8.2 Properties and composition of sericin fractionated with hot water (Komatsu, 1979; Robson, 1985) Fraction

I

II

III

IV

Whole sericin

Content (%) Coefficient of dissolution velocity Crystallinity (%) Total of amino acids with polar side chains (Ap) Total of amino acids with nonpolar side chains (An)

41.0 5.33 3.0

38.6 1.76 18.2

17.6 0.70 32.5

3.1 0.22 37.6

100.0 — 15.06

76.30

75.29

70.69

58.97

74.61

23.74

24.70

29.27

41.03

25.68

together by hydrophobic bonds between the sheets. The amorphous regions contain negatively charged, polar, bulky hydrophobic, and aromatic residues. Sericin is distinctly different from fibroin. Bombyx mori sericin is rich in serine, glycine, and aspartic amino acids which together comprise about two-thirds of the total residues. There are approximately 75% of amino acid residues with polar side chains and 25% with nonpolar side chains. Komatsu (1979) has reported that the four stratified sericin fractions (I, II, III and IV) form different layers from the outside to the inside to cover the fibroin (Komatsu, 1979; Robson, 1985) (Table 8.2). These four layers constitute 41, 38.6, 17.6 and 3.1%, respectively, in content and differ in their rate of dissolution in hot water. Sericin I was shown to be random coil and is the most readily soluble outer fraction. The remaining fractions, which were increasingly more difficult to solubilize in hot water, showed a progressive increase



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in the proportion of their β-conformation and an increase in the degree of crystallinity. The crystallinity of the whole sericin is about 15% with the fractions I 3%, II 18.2%, III 32.5% and IV 37.6%. The hydrophilic nature of the sericin protein accounts for the relative ease with which it can be solublized during degumming and its susceptibility to proteolytic enzymes. As proteins, both silk fibroin and sericin are susceptible to degradation by proteolytic enzymes. Most proteolytic enzymes can hydrolyse the mainly amorphous sericin without much difficulty but not the highly crystalline fibroin. Proteolytic attack on silk fibroin is limited to its amorphous regions. Proteolytic enzymes such as chymotrypsin have been reported to degrade the amorphous regions of fibroins to obtain a highly crystalline fibroin fraction (Konishi et al., 1967; Tsukada, 1986). Successful removal of sericin from fibroin for silk degumming requires the selection of suitable proteases in order to limit enzymatic degradation to sericin to avoid damage to the silk fibre. Various proteolytic enzymes (alkaline, neutral, and acidic proteases) have been investigated for silk degumming (Freddi et al., 2003; Gulrajani et al., 1996). Among them, alkaline proteases performed better than neutral and acidic proteases. The degradation of sericin in terms of weight loss up to 24% has a linear relationship with the amount of alkaline protease used during the enzymatic degumming. Therefore the enzymatic process can be controlled through the enzyme dosage and treatment time. After enzyme treatment, the enzyme is inactivated at 80 °C for a short time and this enhances dissolution of the partially hydrolyzed sericin fractions from the silk. Enzymes have been used to remove sericin from fibroin in silk fabrics. For the crepe fabric, it was reported from morphological analysis that sericin was still found present in the highly twisted weft yarns whereas sericin was completely removed from the warp yarns of the fabric. Extended enzyme treatment might cause damage to the warp yarn fibres. A certain level of mechanical agitation during the degumming process might be needed to enhance enzyme penetration and to make sericin removal complete. Because of the small amount of wax existing on the raw silk and additional waxes applied to the yarns, Gulrajani et al. (2000b) incorporated lipase with the protease in the degumming process. It was reported that the combination of a lipase and a protease resulted in effective de-waxing and degumming and improved wettability of silk fibres and handle. Recently, low-frequency and high-power (20–100 KHz) ultrasound has been used to enhance enzymatic textile processing. The combination of enzyme and ultrasound treatments has two effects. One is the effect on the activity of enzymes. Very little is known about how ultrasound causes the inactivation or activation of enzymes. Another is that ultrasound induces vibration passing through a liquid and the resultant impact waves cause cavitation. Ultrasound was used to try to accelerate the degumming process


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(Krasowski et al., 1999; Mahmoodi et al., 2010). It was shown that the use of ultrasound in the degumming of raw silk with proteases can lead to a slight increase in the degumming effect. That might be due to the improvement in the movement of enzymes to the fibre surface, the increase in the swelling of sericin and/or the dispersion of the enzyme-degraded sericin. Degumming has also been attempted after pre-treatment of the silk with sodium thiosulfate or sodium hydrosulfite solution. The pre-treatment accelerates the removal of sericin by proteases. The fabric whiteness is also significantly improved. Novozymes recommend the pre-treatment of plain and crepe silk fabrics with sodium hydrosulfite at 95 °C for 5 min and 30–60 min, respectively, before to enzymatic treatment. Owing to the high temperature, some sericin is removed at this stage. The pre-treatment reduces the enzyme dosage and the reaction time needed. Subsequent enzyme treatment using alkaline protease (Alcalase from Novozymes) under mild conditions is able to remove the remaining sericin completely so that damage to the fibroin can be avoided (Novozymes technical sheet on Alcalase). Protease enzymes are also used in washing and finishing processes of silk. A biofinishing process, called ‘sand washing’, modifies the protein fibroin of degummed silk. It can achieve the effect of ‘peach skin’, a velvety look imparting a ‘worn’, ‘aged’, or ‘washed many-times’ look.

8.5

Future trends

Environmental issues have been an ongoing concern for the textile industry and will necessarily become of increasing importance. As a natural fibre, wool is perceived by consumers to be renewable and sustainable. Within the wool industry, a great effort has been made to develop green technology in order to replace the harmful chemicals traditionally used in wool processing. Biotechnology through the use of enzymes provides environmentally preferable methods in wet processing to produce wool products with improved softness, antipilling and easy-care properties, especially machinewashability. However, the reaction of conventional protease formulations on wool is difficult to control and may cause unacceptable damage owing to the unique surface structure of the wool fibre. Modification of enzymes or alteration of the absorption and penetration pattern of enzymes during treatment can confer specific finishing to wool including soft handle, shrinkresistance and low temperature dyeing without significant damage to the fibres. Integration of the use of enzymes with other modern technology might be able to expand the finishing process to enhance the wool performance for specific end uses. Natural silk has remained a luxury fibre. In the future silk materials will not only be widely used for textile fabrics, garments and bedding materials



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including silk-filled duvets but also used in the development of various biomedical devices and applications. Demand for silk materials and their production will increase continuously year by year. The textile industry will continue to explore the use of enzymes to improve the efficiency of silk degumming and the quality of the fibres. Research effort will be made to extract the silk proteins, sericin or fibroin and modify them to produce other textile fibres, or for biomedical uses. The design of processes using various available enzymes will be studied for raw wool scouring, silk degumming and their finishing. Integrated bioprocesses might be able to achieve multipurpose preparation and finishing of protein fibres. Innovation of enzyme-based processes needs to be brought into industrial production.

8.6

Acknowledgements

The author would like to thank Dr Peter Speakman for his valuable comments.

8.7

References

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cavaco-paulo a and silva c j s m (2003), ‘Treatment of animal hair fibres with modified proteases’, World Patent WO03097927. cavaco-paulo a, casal m, araújo r and silva c (2009), ‘Process for the treatment of animal hair fibres in order to increase the resistance to shrinkage thereof’, Portuguese Patent 104124. christoe j r (1984), ‘Scouring protein contaminants from raw wool, part I: single bowl laboratory studies’, Text Res J, 54(11), 713–721. christoe j r, pham d k and bateup b o (1994), ‘Raw wool scouring’, Text Asia, XXV(4), 45–47. cortez j, anghieri a, bonner p l r, griffin m and freddi g (2007), ‘Transglutaminase mediated grafting of silk proteins onto wool fabrics leading to improved physical and mechanical properties’, Enzyme Microb Technol, 40(7), 1698–1704. cortez j, bonner p l r and griffin m (2004), ‘Application of transglutaminases in the modification of wool textiles’, Enzyme Microb Technol, 34, 64–72. cortez j, bonner p l r and griffin m (2005), ‘Transglutaminase treatment of wool fabrics leads to resistance to detergent damage’, J Biotechnol, 116, 379–386. das t and ramaswamy g n (2006), ‘Enzyme treatment of wool and specialty hair fibers’, Text Res J, 76(2), 126–133. du g, cui l, zhu y and chen j (2007), ‘Improvement of shrink-resistance and tensile strength of wool fabric treated a novel microbial transglutaminase from Streptomyces hygroscopicus’, Enzyme Microb Technol, 40, 1753–1757. dybdal l, heine e and höcker h (2001), ‘Method for enzymatic treatment of wool’, US Patent 6258129. el-sayed h, kantouch a, heine e and höcker h (2001), ‘Developing a zero-AOX shrink-resist process for wool. Part I: preliminary results’, Color Technol, 177, 234–238. feldtman h d, leeder j d and rippon j a (1983), ‘The role of fibre structure in wool fibre and fabric performance’ in Postle R, Kawabata S and Niwa M, Objective evaluation of apparel fabrics, The Textile Machinery Society of Japan, Japan, 125–134. fogorasi m s and heine e (2006), ‘Spectroscopic methods of studying enzyme action in wool fibre’, Cent Eur J Chem, 4(4), 786–797. fossey s, nemethy g, gibson k and scheraga h (1991), ‘Conformational energy studies of β-sheets of model silk fibroin peptides I. Sheets of poly(Ala-Gly) chains’, Biopolymers, 31(13), 1529–1541. freddi g, mossotti r and innocenti r (2003), ‘Degumming of silk fabric with several proteases’, J Biotechnol, 106, 101–112. gouveia i c, fiadeiro j m and queiroz j a (2008), ‘Enzymatic wool treatment: preliminary evaluation of Trichoderma reesei cellulases and Aspergillus aculeatus pectinases and hemicellulases’, AATCC Rev, 8(10), 38–44. griffin m, casadio r and bergamini c m (2002a), ‘Transglutaminases: nature’s biological glues’, Biochem J, 368, 377–396. griffin m, cortez j m and bonner p (2002b), ‘A method for enzymatic treatment of textiles such as wool’, World Patent WO0204739. gulrajani m l (1992), ‘Degumming of silk’, Rev Prog Color, 22, 79–91. gulrajani m l, agarwal r and chand s (2000a), ‘Degumming of silk with a fungal protease’, Indian J Fiber Text Res, 25(June), 138–142. gulrajani m l, agarwal r, grover a and suri m (2000b), ‘Degumming of silk with lipase and protease’, Indian J Fibre Textile Res, 25(March), 69–74.



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gulrajani m l, gupta s v, gupta a and suri m (1996), ‘Degumming of silk with different protease enzymes’, Indian J Fibre Text Res, 21(December), 270–275. heine e (1991), Grundlegende untersuchungen zur einwirkung von enzymen auf wolle, PhD Thesis, Technischen Hochschule Aachen. heine e and höcker h (1995), ‘Enzyme treatments for wool and cotton’, Rev Prog Color, 25, 57–63. heine e and höcker h (2001), ‘Bioprocessing for smart textiles and clothing’, in Tao X, Smart fibres, fabrics and clothing, Woodhead Publishing Ltd, Cambridge, UK. pp. 254–263. heine e, hollfelder b, shen j and bishop d (2000) ‘Parameters affecting the morphology of wool during enzyme treatment’, The 10th International Wool Textile Research Conference, 26th Nov–1st Dec, Aachen, Germany. holme i (2006), ‘New frontiers in wool dyeing and finishing’, Int Dyer, 191(10), 9–11. hossain k h m g, juan a r and tzanov t (2008), ‘Simultaneous protease and transglutaminase treatment for shrink resistance of wool’, Biocatal Biotransform, 26(5), 405–411. iwto roadmap (2009), 100% natural renewable sustainable wool, change to a healthier and safer environment, International Wool Textile Organisation, Brussels, Belgium. jovancic p, jocic d and dumic j (1998), ‘The efficiency of an enzyme treatment in reducing wool shrinkage’, J Text Inst, 89 Part 1, No.2, 390–400. jovancic p, jocic d, molina r, julia m r and erra p (2003), ‘The combined lowtemperature plasma/enzyme wool shrink-resist treatment’, AATCC Rev, 3(2), 25–28. julie m r, erra p, dominguez j g and infante m r (1985), ‘The influence of a cationic surfactant on the shrink-resist treatment of wool with sodium sulphite in aqueous solution’, J Soc Dyers Color, 101(February), 66–71. kholiya r, khambra k, rose n and sangwan n (2008), ‘Optimization of process conditions for cellulase enzyme treatment of woollen fabric’, Colourage, January, 65–68. komatsu k (1979), ‘Chemistry and structure of silk’, Japan Agric Res Quart, 13(1), 64–72. konishi t, kondo m and kurokawa m (1967), ‘Study on the structure of silk fibroin under enzymatic hydrolysis’, Sen-i Gakkaishi, 23, 64–69. krasowski a, muller b, fohles j and hocker h (1999), ‘Degumming of silk in an ultrasonic field’, Melliand Textilberichte, 80, 543–545. leeder j d and rippon j a (1985), ‘Changes induced in the properties of wool by specific epicuticle modification’, J Soc Dyers Color, 101, 11–16. lefevre t, rousseau m and pezolet m (2007), ‘Protein secondary structure and orientation in silk as revealed by Raman spectromicroscopy’, Biophys J, 92, 2885– 2895. lenting h b m, broekman h, guebitz g m, kokol v and shen j (2009), ‘Industrial production of enzyme-modified wool fibers for machine-washable bed coverings’, Biotechnol J, 4, 1–9. lenting h b m, schroeder m, gübitz g m, cavaco-paulo a, and shen j (2005), ‘A new enzyme technology based method for obtaining machine washable wool’, The 11th International Wool Textile Research Conference, 4th–9th Sep, Leeds, UK.


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9 Functionalisation of wool and silk fibres using non-proteolytic enzymes G. F R E DDI, Stazione Sperimentale per la Seta, Italy

Abstract: Wool keratins, silk fibroin and sericin are potential substrates for various enzymatic reactions and proteases can be used in environmentally friendly processes for wool antifelting and softening, and for silk degumming. The use of non-proteolytic enzymes able to modify the chemistry of amino acid residues without breaking the peptide bond is reviewed. In particular, transglutaminases and tyrosinases have the capability to modify the properties of wool and silk fibres by targeting the reactive side groups of amino acid residues, thus resulting in activation, functionalisation, and crosslinking. Key words: wool, keratins, silk, fibroin, sericin, transglutaminase, tyrosinase.

9.1

Introduction

Wool and silk are protein-based textile fibres widely different in chemical and physical structure and morphology. Wool has a composite structure comprising two kinds of cells: cuticle and cortex cells (Zahn et al., 2003). Coarse fibres may also have a central medulla. Cuticle cells (also called ‘scales’) have a flat shape and overlap each other from root to tip forming a sheath around the cortex. The cortex forms the bulk of the fibre and consists of spindle-shaped interdigitated cells. Cortical cells are held together by the cell membrane complex (CMC), which also separates cortical cells from the cuticle. The CMC is a continuous region, containing slightly crosslinked proteins, that extends throughout the whole fibre. Being the only continuous phase in the fibre, it also provides a channel by which dyes and chemicals can diffuse in and out of wool. Keratin proteins are divided into four major classes: the low sulfur, the high sulfur, the ultra-high sulfur, and the high glycine–tyrosine proteins. They widely differ in amino acid composition molecular weight, and structure. In particular the content of cystine (Cys) and the degree of disulfide crosslinking are different. The raw silk fibres spun by the Bombyx mori silkworm are formed by two fibroin filaments surrounded by a cementing layer of sericin (Zahn and Krasowski, 2003). Fibroin and sericin account for about 75 and 25% of the 193 © Woodhead Publishing Limited, 2010

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fibre weight, respectively. Fibroin, the main constituent of silk fibres, consists of three polypeptides: the heavy (391.6 kDa) and light (27.7 kDa) fibroin chains, linked by one disulfide bridge, and a low molecular weight protein called P25 (25.2 kDa), also known as fibrohexamerin, which are present in a mass ratio of 93 : 6 : 1, respectively. The heavy fibroin chain, the real fibrous component, is rich in glycine (Gly), alanine (Ala), and serine (Ser) amino acids (about 85 mol%) in the molar ratio 3 : 2 : 1. The primary structure consists of eleven repetitive hydrophobic sequences (R) regularly interrupted by ten hydrophilic spacer sequences (A), in addition to two hydrophilic N- and C-terminus sequences (Zhou et al., 2000). The R sequences are characterised by repeating -(Gly-Ala)n- motifs which form the well known β-sheet crystalline regions present in the fibre as well as in regenerated silk fibroin materials like films and gels. Sericin, the silk gum holding together the fibroin filaments, is a complex mixture of 5–6 polypeptides widely differing in size (40–400 kDa), chemical composition, structure, and properties. The amino acid composition is characterised by an extremely high concentration of Ser, which ranges from 16 to 38 mol%. Sericin is usually removed during textile processing by a wet treatment called degumming. As proteins, keratins, fibroin, and sericin are potential substrates for various enzymatic reactions. The use of proteases as environmentally friendly processes for wool antifelting and softening or for silk degumming is reviewed in chapter 8. Here, the use of non-proteolytic enzymes able to modify the chemistry of amino acid residues without breaking the peptide bond is considered. In particular, transferases (e.g. transglutaminase) and oxidoreductases (e.g. tyrosinase, laccase, peroxidase) have the capability of modifying the properties of wool and silk protein fibres by targeting reactive side groups of amino acid residues, thus resulting in activation, functionalisation, or crosslinking. The specificity and selectivity of these enzymes may allow wool and silk to be processed under milder reaction conditions and their properties to be better tailored to end-use requirements, thus offering cleaner and safer alternatives to current processing practices which make use of harsh chemicals. The most recent studies on the biotechnological functionalisation of protein-based textile fibres with transglutaminase and tyrosinase are reviewed in sections 9.3 and 9.5, respectively. Biotechnology is typically a cross-sectoral technology able to drive product and process innovation in a range of textile and non-textile industrial sectors. In order to provide a wider overview of the great potentiality of these enzymes, not only textile, but also other biotechnological applications falling in different scientific and technological sectors are reviewed here. To outline the state of the art of non-proteolytic enzymes available for protein fibre modification, other enzymes having displayed some activity on protein fibre substrates, such as


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laccase and peroxidase, are briefly reviewed in section 9.6. Finally, emerging enzymatic activities, such as sulfydryl oxidase and protein disulfide isomerase, that are attracting the interest of scientists for their possible role in protein fibre functionalisation, are mentioned.

9.2

Transglutaminases

9.2.1 Biological properties and biotechnological utility of transglutaminases Transglutaminases (TGases; protein-glutamine γ-glutamyltransferase, EC 2.3.2.13) are involved in the post-translational modification of proteins by formation of isopeptide bonds (Griffin et al., 2002). TGase catalyses an acyl transfer reaction between the γ-carboxyamide group of a peptide-bound glutamine residue (Gln) and the amino group of lysine (Lys) (Fig. 9.1). Gln acts as the acyl donor, whereas Lys or other primary amines act as the acyl acceptor. In the absence of amine substrates, TGase catalyses the hydrolysis of the γ-carboxyamide group of Gln, resulting in deamidation and formation of glutamic acid (Glu). When the ε-amino group of a peptide-bound Lys is the substrate, peptide chains are covalently connected through ε-(γglutamyl)lysine bonds. The isopeptide bonds exhibit high resistance to proteolytic degradation. TGases have been found in micro-organisms (Ando et al., 1989; Tsai et al., 1996), plants (Serafini-Fracassini and Del Duca, 2008), invertebrates (Cariello et al., 1997), fish (An et al., 1996), birds (Weraarchakul-Boonmark et al., 1992), and mammals (Lorand and Graham, 2003). About eight calcium-regulated TGase activities have been identified in mammals at the

(a)

O

O TGase

NH2 + H2N (b)

O

O NH2 + H2N-R

(c)

+ NH3

N H

TGase N H

R

+ NH3

O

O NH2 + H2O

TGase

OH

+ NH3

9.1 Reactions catalysed by transglutaminase: (a) protein crosslinking by formation of a ε-(γ -glutamyl)lysine isopeptide bridge between the glutamine residue of one protein and the lysine residue of another; (b) incorporation of an amine into the glutamine residue of a protein with formation of an amine–isopeptidyl adduct; (c) deamination of a glutamine residue in a protein substrate with formation of glutamic acid.


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genomic level. All require the binding of Ca2+ for their activity. In general, mammal TGases display strict specificity in the recognition of glutamine protein substrates and lower specificity for the acyl acceptor amine group, which can either be the ε-amino group of peptidyl Lys or a low molecular mass primary amine. Ohtsuka et al. (2000) reported some differences in the substrate specificity of two TGases, one from guinea pig liver and one from Streptomyces mobaraensis. Accordingly, differences in the structure of the catalytic site between mammal and microbial TGases have been reported (Lorand and Graham, 2003). These features, together with the fact that microbial TGases are totally Ca2+ independent, confirm the high heterogeneity of this family of enzymes. TGases are present in most animal tissues and body fluids (Griffin et al., 2002; Lorand and Graham, 2003). These enzymes play a key role in physiology and development being involved in bone remodelling and ossification, skin maturation and integrity, development of heart, lung, salivary gland, and central and peripheral nervous system, hematopoiesis, spermatogenesis, egg-envelope formation following fertilisation, myofibrillogenesis, blood coagulation, and programmed cell death (apoptosis) (Della Mea et al., 2007; Ientile et al., 2007; Vanhoutte and Heymans, 2008; Bakker et al., 2008). One member of the TGase family (TGase 3) is involved in the progressive scaffolding of hair shaft through specific formation of isopeptide bonds between intermediate filaments and keratin-associated proteins (Thibaut et al., 2009). In general, the reactions catalysed by TGases endow the resulting supramolecular structure with extra rigidity and resistance against proteolytic degradation. Having knowledge of these enzymes is also essential for understanding the aetiologies of diverse hereditary diseases of blood and skin, and various autoimmune, inflammatory and degenerative conditions, a growing area of intensive TGase research (De Vivo and Gentile, 2008; De Vivo et al., 2008). The fact that TGase activity increases in various inflammatory and fibrotic conditions, such as rheumatoid arthritis, liver, renal and pulmonary fibrosis, erosive gastritis, leads to its implication in the pathogenesis of these disorders. Noteworthy, TGase activity is increased and/or crosslink formation is enhanced in all forms of mucosal inflammation (Crohn’s disease, ulcerative colitis, coeliac disease) and in mucosal epithelial lesions, as well as in fibrotic disorders (Stenberg et al., 2008; Wieser and Koehler, 2008). Last, but not least, TGases have been implicated in axonal growth and regeneration, as well as in various degenerative neurological diseases (for example: Huntington’s, Alzheimer’s, Parkinson’s and progressive suprabulbar palsy) which are characterised by the abnormal accumulation of insoluble protein aggregates that cause progressive neuronal death (Wang et al., 2008). The limited supply and the relatively high price have hindered for years the industrial utilisation of tissue TGases, commonly extracted from guinea pig liver, until the mass production of a microbial TGase from Streptomyces © Woodhead Publishing Limited, 2010


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mobaraensis has been achieved at low costs (Yokoyama et al., 2004). This TGase is commercialised by Ajinomoto under the Activa® brand name for food processing, and finds application at industrial scale for improving the texture of meat and fish or dairy products. The S. mobaraensis TGase is a secreted protein activated outside the cytoplasm, which participates in mycelial growth and has a role in morphological differentiation. Unlike many tissue TGases, it has a relatively low molecular weight (37 842 Da, based on its known primary structure). The isoelectric point is 9, the optimum pH for enzymatic activity is in the range 5–8, and the optimum temperature is 55 °C. The Ca2+-independency offers several advantages when biotechnological applications are sought. With respect to substrate specificity, the TGase from S. mobaraensis has the ability to crosslink most food proteins, such as legume globulins, wheat glutens, egg yolk and albumin proteins, actins, myosins, fibrins, milk caseins, α-lactalbumin, and β-lactoglobulin as efficiently as mammal TGases by formation of ε-(γglutamyl)lysine bonds. Protein solutions, such as soybean, milk, beef, and pork proteins, chicken and fish gelatine and myosins can be gelled. In a series of recent publications Lantto et al. (2005a, 2006, 2007a) reported results on the effect of S. mobaraensis TGase on the thermal, gel forming, textural, and water-holding properties of various meat systems, which further support the utility of this enzyme as a biotechnological food processing tool. In a recent study Kulik et al. (2009) investigated the reactivity of synthetic substitutes for Lys and Gln as substrates of microbial TGase from S. mobaraensis. The reactivity of ω-amino acids used as Lys substitutes increased with increasing the hydrocarbon chain length from C5 to C7. The conversion reached about 70% with 7-aminoheptanoic acid as substrate. With reference to Gln substitutes, high reactivity and conversion levels were achieved only with glutaric mono and diamine, i.e. with substrates having a C5 hydrocarbon chain similar to the natural Gln substrate. Substrates with longer hydrocarbon chains (i.e. C6, adipic diamine) displayed sensibly lower reactivity, indicating that restrictions for the Gln substitute are stronger than for the Lys substitute. Kulik et al. also succeeded in demonstrating the feasibility of a completely artificial microbial TGase-catalysed reaction by producing mono and diadducts of DNS-cadaverine and glutaric diamine, thus opening new opportunities for surface polymer modification by grafting functional compounds at distinct sites through an environmentally friendly reaction based on microbial TGase catalysis.

9.2.2 Applications of transglutaminases: advantages and limitations Although currently the main application sector of TGase is food processing, novel potential applications have emerged during the last years. These © Woodhead Publishing Limited, 2010

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non-food applications cover the areas of biological science, tissue engineering, material science, leather and textile processing (Zhu and Tramper, 2008). When the Activa® TGase available from Ajinomoto is used, care must be taken to consider the artefacts potentially caused by the presence of large amounts of maltodextrins, alone or together with sodium caseinate or lactose, in the commercial formulation. If undesired interferences are expected, these additives must be removed by applying suitable purification procedures (Lantto et al., 2005a). Tissue engineering TGase can facilitate the manufacture of tissue engineering scaffolds with structural and functional properties able to support healing in vivo. Scaffolds derived from collagen or gelatine are most commonly used for mammalian cell growth. Enzymatic crosslinking of collagen scaffolds represents a viable alternative to chemical or physical approaches that may negatively affect cell growth. Collagen scaffolds treated with tissue or microbial TGases displayed enhanced cell attachment, spreading and proliferation of human osteoblasts and human foreskin dermal fibroblasts, and also showed a greater resistance to cell-mediated endogenous protease degradation (Chau et al., 2005). Enzymatically crosslinked, composite collagen scaffolds containing hyaluronan, one of the principal extracellular matrix components of native intervertebral disc tissue, showed the greatest potential for developing an injectable cell-seeded scaffold for nucleus pulposus treatment in degenerated intervertebral discs (Halloran et al., 2008). Garcia et al. (2008) recently reported the development of enzymatic crosslinked collagen-based scaffolds for the production of dermal precursors with enhanced wound healing properties. Scaffolds crosslinked with microbial TGase provided an optimum substrate for cell migration, preventing wound contraction, stimulating epithelialisation and neoangiogenesis without inducing significant inflammatory reaction. Gelatine, a denatured derivative of collagen, is a versatile biomaterial that can be easily processed into scaffolds. However, one of gelatine drawbacks in tissue engineering is its solubility in aqueous medium, therefore gelatine-containing scaffolds for medium-to-long term applications need to be crosslinked. Enzymatic crosslinking may be a valid alternative to traditional strategies using potentially toxic chemical agents, such as glutaraldehyde. Moreover, the ability to control scaffold degradation in response to cell growth rates may represent an attractive and unique benefit of the enzymatically modulated scaffold structure. Increased thermal, mechanical and proteolytic stability and reduced citotoxicity as compared with chemically or physically crosslinked scaffolds have been reported (Barbetta et al., 2006).



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The enzyme technology was exploited by Chiono et al. (2008) to produce melt-extruded guides for peripheral nerve regeneration by blending poly(ε-caprolactone) (PCL) and gelatine. In one approach, gelatine was crosslinked with microbial TGase and then blended with PCL by solution mixing for the production of nerve guidance channels. In another approach, microbial TGase was used to graft poly-l-Lys, a biomimetic molecule able to confer to the channel guide a specific signalling for nerve cells attachment, proliferation and migration. Binding of poly-l-Lys to the exposed gelatine sequences in the inner surface of the nerve channel was proved by confocal microscopy, whereas in vitro tests with neuroblastoma cells showed that the nerve guides produced by melt extrusion were biocompatible and that the grafted signalling peptide greatly improved cell response. The same research group has recently reported the production of three-dimensional porous hydroxyapatite/collagen composite scaffolds for bone tissue regeneration characterised by enhanced stability and controllable swelling ratio by crosslinking via microbial TGase-catalysed reaction (Ciardelli et al., 2010). An alternative approach in the use of TGase for tissue engineering is to modify the surface of biomaterials by coating with the enzyme in order to exploit its ability to stimulate cell adhesion processes. Ball et al. (2009) used TGase in conjunction with fibronectin as a coating for tricalcium phosphate scaffolds and studied their effect on metabolic activity and other physiological parameters, including the rate and extent of mineralisation of osteoblast cells seeded on the scaffold surface. They found that fibronectin supported early cell differentiation, whereas increased matrix formation or stabilisation was induced by TGase, probably through nucleation of mineralisation promoted by the TGase crosslinked extracellular matrix. Site-specific modification of proteins Enzymatic approaches to site-specific modification of proteins have attracted a great deal of attention in biological research because this modification strategy enables protein manipulation without significant loss of function, it is highly selective and works under mild conditions compared with conventional chemical methods. Microbial TGase has been widely applied for site-specific modification, crosslinking, and conjugation of proteins with other biological molecules (e.g. DNA, carbohydrates). If protein substrates are not intrinsically reactive to microbial TGase because of reduced accessibility of target amino acid residues, the incorporation of specific peptide tags for enzyme recognition allows enhancing substrate specificity and reaction yield. By using an amino-derivative of poly(ethylene glycol) (PEG) as substrate for the enzymatic reaction with TGase, it has been possible to covalently bind the polymer to proteins of pharmaceutical interest thus making these protein drugs more water soluble,


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non-aggregating, non-immunogenic, and more stable to proteolytic degradation (Fontana et al., 2008; Mero et al., 2009). Interestingly, modification or PEGylation of protein-bound Gln residues occurred at specific sites. For example, it has been shown that in apomyoglobin only Gln 91 was modified and in human growth hormone only Gln 40 and Gln 141, despite these proteins having many more Gln residues. These studies shed light on the molecular features dictating the selective attack of protein-bound Gln residues by TGases and highlighted the close molecular similarities between TGase-mediated reactions and proteolysis. In fact, TGase can attack the same sites/regions of the polypeptide chain that suffer proteolysis, provided that a Gln residue is encompassed by that chain region. Enhanced chain flexibility, extended chain conformation, local unfolding/disorder of the protein secondary structure such as in connecting loops between hydrogenbonded structures (helices) are the key parameters driving the TGase attack to Gln residues. Thus, surface exposure alone does not seem a sufficient condition for TGase to use a Gln residue as substrate, as shown by the fact that not all surface exposed Gln residues in globular proteins are modified by the enzyme. Only those surface-exposed residues that are encompassed in a flexible or unfolded region can be attacked by TGase. These findings entail important consequences because it appears possible not only to explain but also to predict the sites of TGase-mediated modification of globular proteins of known sequence and structure, thus opening the prospect to properly design useful strategies for their modification. Biofabrication The integration of proteins with microfabricated devices is an attractive biofabrication approach for developing a range of applications, from proteomics to microfluidics, biosensors, medical diagnostics, microarrays, bioMEMS (biological microelectromechanical systems), and metabolic engineering. An elegant example of the enzyme-mediated controlled assembly of proteins is the stepwise assembly at electrode addresses of biological polymers, such as chitosan and proteins, by a combination of electrical stimuli and enzymatic reactions catalysed by tyrosinase and TGase (Yang et al., 2009). Chitosan is considered a good interface for assembling proteins at solid surfaces. The formation of a thin and resistant chitosan film onto solid electrode surfaces can be simply achieved by electrodeposition. Furthermore, proteins can be assembled onto the polysaccharide layer by combined enzymatic reactions that first exploit the nucleophilicity of the chitosan primary amine to bind a peptide tether via the tyrosinase-catalysed reaction, followed by the formation of a covalent bond between the tether and another protein via microbial TGase-catalysed conjugation. The latter protein could be an enzyme, a receptor, an antigen or an antibody, which



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confers on the device the desired functionality at specified addresses. This enzymatic assembly approach provides an alternative to existing chemical coupling methods. It positively responds to technological requirements, including spatial selectivity, orientational control, and mild treatment conditions, which allow the biological activity of proteins to be maintained during the multi-step assembling procedures because the enzymatic approach does not require reactive chemical agents or activated polymers. Leather processing Leather processing includes filling, i.e. the introduction of materials into the voids that exist between the fibres of the leather to smoothen any irregularity on the leather surface. Protein by-products such as gelatine and casein can be used as filling materials during leather processing. For example, glutaraldehyde crosslinked gelatines resulted in highly polymerised filling polymers that were able to fill the leather and, more importantly, also remained bound to the leather during washing steps (Taylor et al., 2006a). As a further development of this approach to leather filling, microbial TGase was used as crosslinking agent for various proteinaceous industrial by-products, specifically gelatine, whey, and whey protein isolate (Taylor et al., 2006b). The crosslinked biopolymers showed values of melting point, viscosity, and molecular weight distribution suitable for their use as fillers, suggesting that there is potential for these relatively inexpensive and sustainable resources to be recycled in leather processing. Filling experiments demonstrated that the enzymatically prepared filling materials could be effectively used as fillers, which were bound to the leather and could not easily be removed during further processing (Taylor et al., 2007).

9.3

Functionalisation of protein fibres using transglutaminases

Among protein-based textile fibres, wool has become an eligible substrate for the application of TGase enzymes. The content of Gln+Glu in wool is 1098 μmol g−1, corresponding to ∼12 mol% (Lindley, 1977), 40% of which can be attributed to Gln alone (Maclaren and Milligan, 1981). The amino acid analysis of the individual histological components of the wool fibre, i.e. cuticle, CMC, and cortex, has shown that Gln+Glu residues are present in roughly the same amount, i.e. 894 μmol g−1 in the cuticle (∼9 mol%), 968 μmol g−1 in the CMC (∼10 mol%), and 1081 μmol g−1 in the cortex (∼12 mol%) (Lindley, 1977). Considering that the relative amount of each histological component in fine wool fibres is about 10, 5, and 85 wt% for cuticle, CMC, and cortex, respectively (Bradbury and King, 1967), the content of Gln+Glu in the different morphological compartments of intact


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wool fibres can be estimated as follows: ∼95 μmol in the cuticle, ∼55 μmol in the CMC, and ∼950 μmol in the cortex. Endogenous TGases are implicated in the process of maturation of keratin fibres and about 15 μmol g−1 of ε-(γ-glutamyl)lysine isopeptide crosslinks have been identified in the enzymatic digest of untreated wool (Asquith et al., 1970). However, significant amounts of free Gln binding sites are still available in the wool fibre for further TGase-mediated crosslinking and/or incorporation of primary amines. The content of Lys, whose primary amino group may act as the acyl acceptor to form an isopeptide bond, is about 282 μmol g−1 (∼2.8 mol%) in untreated wool fibres. The pioneering work of Cortez et al. (2004) has demonstrated that TGase can use the Gln residues of wool as substrate and that the enzymatic approach to the modification of wool fibre properties is feasible. Both tissue and microbial TGases proved to be effective in: (i) (ii)

maintaining and/or improving wool fibre strength; reducing the propensity of wool fabrics to shrink under normal washing conditions; and (iii) incorporating functional amine compounds into wool fibres. When wool fibres previously treated with protease, or with reducing/oxidising agents to simulate adverse industrial processing conditions likely to lower fibre properties such as strength and handle, were exposed to the action of tissue or microbial TGases, the enzymatic treatment was able to restore the initial fibre properties, in particular tensile strength, thus remediating the damages induced by proteolytic and/or chemical degradation. TGases were able to improve strength of intact wool fibres as well, though to a lower extent. The higher the extent of fibre damage, the higher the beneficial effect of TGase, indicating that more extensive degradation made the fibre more accessible to the enzyme and facilitated the TGase-catalysed crosslinking via ε-(γ-glutamyl)lysine bridges. The potential of TGase to prevent/recover damages caused by the use of common household detergents containing proteolytic enzymes and to prolong the longevity of wool textiles was also demonstrated (Cortez et al., 2005). Incubation with microbial TGase before detergent washing led to a significant increase in tensile strength. After three washing cycles, the tensile properties of TGase-treated wool samples decreased but to a much lower extent than those not exposed to TGase. Treatment with TGase also had beneficial effects on colour fastness because it lowered the amount of dye released from the wool fabric into the washing liquor. All these results can be explained in terms of higher crosslinking density induced by the enzyme within the wool fibre matrix. The contribution of microbial TGase in improving yarn strength, decreasing area shrinkage and increasing wettability of wool materials damaged by chemical (oxidative) or enzymatic (protease)



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treatments has recently been highlighted by Ge et al. (2009). They also investigated the dyeing behaviour of wool and reported that treatment with microbial TGase resulted in an increase of initial dyeing rate and shortening of dye exhaustion time, which were attributed to the higher wettability of the wool fibres. The beneficial effects of microbial TGase on wool were further enhanced when primary amine ligands were grafted onto the fibres (Cortez et al., 2007). Silk-derived sericin peptides were used as bridging agents. Depending on reaction conditions, sericin peptides were grafted not only at the fibre surface but could penetrate inside the cuticle reaching the outer cortex layer. The increased crosslinking density improved not only wool fibre strength, but also resulted in better resistance to felting, which allowed TGase-treated wool to be classified ‘machine washable’ according to Woolmark standards. The perceived softness of wool fabrics, a property highly appreciated by end-users, was also significantly improved. In an attempt to overcome current limitations in the use of proteases as antifelting agents, a simultaneous one-batch protease-TGase treatment of wool fabrics was designed with the aim of exploiting the benefits of the individual enzymes (Gaffar Hossain et al., 2008). The protease is expected to erode the wool scales, which would bring about shrink resistance while opening the wool fibre texture for the penetration of TGase and formation of isopeptide crosslinks leading to improved dimensional stability and mechanical performance. The results reported seem to confirm the initial hypothesis. Higher level of shrink resistance, lower weight and strength losses, and less fibre damage were achieved in the combined enzyme treatment compared with a single protease treatment. The issue of recovering more or less extensive tensile strength losses caused by antifelting treatments was also addressed by Cardamone (2007). In an attempt to remediate the high strength losses caused by a two-step chemo-enzymatic shrink-proofing process, a third processing step based on the use of microbial TGase was performed. However, only a moderate recovery of tensile strength of about 3–5% was achieved by TGase treatment against an initial strength loss of 10–18% caused by the shrinkproofing treatment. In another study, wool knits were subjected to only one oxidative chemical treatment and then were treated with TGase in the presence of solubilised keratins with the aim of improving the dimensional stability (Cardamone and Phillips, 2007). Good results were obtained under optimised processing conditions in terms of shrink-proofing, which could be minimised to about 5%, whereas strength loss was reduced to 4.7%. According to SEM results, the solubilised keratins smoothed the fibre surface by filling the voids at the protruding scale edges. The mechanism of TGase-mediated grafting of solubilised keratin peptides onto wool fibres was reported in a subsequent paper (Cardamone, 2008).


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So far, most of the work on wool fibres has been done by using the microbial TGase from S. mobaraensis. A new stable form of microbial TGase from Streptomyces hygroscopicus WSH03-13 was recently obtained (Cui et al., 2006). This enzyme was used for wool treatment, the aim being to monitor its effect on shrink resistance, tensile strength (Du et al., 2007) and dyeing properties with natural dyes (Cui et al., 2008). In contrast to the S. mobaraensis TGase, this novel microbial TGase was almost ineffective towards intact wool fibres because properties such as tensile strength and area shrinkage were not influenced by enzyme treatment. This result was attributed to the fact that the external hydrophobic layer of intact wool fibres limited the accessibility of target reactive sites. On the other hand, oxidative, reductive, or proteolytic pretreatments opened up the fibres for the penetration of the enzyme, which could catalyse the formation of covalent crosslinks. As a result, tensile strength and area shrinkage were improved to different extents, depending on the kind of pre-treatment, thus confirming the ability of microbial TGase to repair chemical or biological damages of wool fibres. Crosslinking of gelatine onto the surface of wool fibres via microbial TGase-mediated reaction resulted in further improvement of mechanical and antifelting properties of wool (Cui et al., 2009). Good antimicrobial properties were also imparted on wool fibres by grafting ε-polylysine through microbial TGase-catalysed reaction (Jin et al., 2009). The good reactivity of TGases towards wool fibres can be explained in terms of amount and accessibility of Gln residues at the fibre surface (cuticle) and within the fibre texture (CMC and cortex). The microbial TGase from S. mobaraensis has a relatively small size (about 38 kDa), not too far from that of bacterial proteases, whose ability to diffuse inside wool fibres has been reported (Silva et al., 2005). The results of TGase-catalysed functionalisation of wool suggest that the enzyme is able to diffuse inside the fibre matrix and to reach the Gln residues, whose accessibility can be further enhanced by treatments able to lower the hydrophobic barrier of intact wool fibres. In fact, cuticle cells consist of lamellar components, whose thin outermost layer (epicuticle) is resistant to acids, alkalis, oxidising agents, and to enzymatic attack. The epicuticle is formed by heavily crosslinked proteins covalently bound to 18-methyleicosanoic acid through a thioester bond (Negri et al., 1993). This assembly makes the fibre surface strongly hydrophobic and hardly accessible to reagents and enzymes. Thus, accessibility of target amino acid residues to enzyme molecules sometimes might be a real challenge if suitable pre-treatment aimed at decreasing the hydrophobic barrier and opening the fibre surface are not performed. No results have yet been published on the modification and/or functionalisation of silk fibres with TGase. The total amount of Gln+Glu in silk fibroin is about 180 μmol g−1 (∼1.2 mol%) and the amount of Gln can be



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estimated to be about 70 μmol g−1 (∼0.5 mol%) based on the known amino acid sequence of the three fibroin polypeptides. Thus, the Gln content in silk is much lower than that of wool. The content of Lys is also low, accounting for about 40 μmol g−1 (∼0.3 mol%). To verify whether the Gln residues of silk fibroin could be used by TGase for transamidation reactions, an aqueous silk fibroin solution was incubated with microbial TGase from S. mobaraensis and then films were cast. When dissolved in aqueous solution, silk fibroin adopts a disordered random coil conformation with the more hydrophilic polypeptide sequences exposed to the aqueous phase (Canetti et al., 1989). This is considered the most favourable condition for the accessibility of charged amino acid residues like Gln and Lys. However, no detectable differences were observed in the chemical, physical, and mechanical properties of silk fibroin films upon incubation with TGase with respect to the control films cast without enzyme (Freddi et al., unpublished results). Noteworthy, another enzyme, i.e. the mushroom tyrosinase from Agaricus bisporus, under the same reaction conditions, had significant effects on silk fibroin properties (see section 9.4.2, Applications of tyrosinases: advantages and limitations). These results seem to indicate that the Gln residues of silk fibroin are hardly accessible to TGase, even under homogeneous reaction conditions. Based on these findings it is possible to assume that silk fibroin in the soluble state and, all the more so, in the form of intact fibres, might not be a good substrate for TGase-catalysed modification and/or functionalisation.

9.4

Tyrosinases

9.4.1 Biological properties and biotechnological utility of tyrosinases Tyrosinases (monophenol, o-diphenol: oxygen oxidoreductase, EC 1.14.18.1) belong to the type 3 copper proteins having a diamagnetic spin-coupled Cu pair in the active centre (Rosenzweig and Sazinsky, 2006; Solomon et al., 1996). Tyrosinases, which are often referred to as polyphenol oxidases (PPO), can catalyse both the hydroxylation of tyrosine (Tyr) into o-diphenols and the oxidation of o-diphenols into o-quinones (Fig. 9.2). In the first reaction, often referred to as the monooxygenase or cresolase activity, which is known to be the rate-determining step, a hydroxyl group is introduced into the ortho-position of the aromatic ring. In the second reaction, often referred to as the diphenolase or catecholase activity, the o-dihydroxy compound produced in the first step is oxidised to an o-quinone. Both reactions involve molecular oxygen as an electron acceptor which is reduced to water. Quinones are molecular species highly susceptible to non-enzymatic reactions leading to formation of brown or black pigments such as melanins.


206 (a)

Advances in textile biotechnology O

OH

OH

O

OH 1 2

O2

Tyrosine

Tyrosinase

+

1 2

O2

Tyrosinase

Dopa

+ H2O

Dopaquinone

(b) O HO

OH

[1]

O [2]

O

N

or HN HO

OH Phenol coupling

H2C

H2C Reaction with nucleophiles

9.2 Tyrosinase-catalysed oxidation of protein-bound tyrosine and subsequent non-enzymatic reactions: (a) hydroxylation of proteinbound tyrosine with formation of dopa by introduction of a hydroxyl group into the ortho position of the aromatic ring (monooxygenase activity), followed by oxidation of the o-dihydroxy compound to an o-quinone (diphenolase activity); (b) non-enzymatic reactions of o-quinones comprise phenol coupling, leading to protein–protein crosslinking, or reaction with nucleophiles, such as the free amine groups of proteins and other polymers, leading to protein–protein crosslinking or protein–polymer conjugation through Michael addition [1] or Schiff base [2] reaction mechanism.

The tyrosinase obtained from the edible mushroom Agaricus bisporus is one of the most studied fungal tyrosinases (Flurkey et al., 2008). Like other fungal tyrosinases, it is intracellular. The enzyme is a heterotetramer with an apparent molecular mass of about 120 kDa, comprising two heavy (about 43 kDa) and two light (about 14 kDa) polypeptide chains. The isoelectric point is 4.7–5, and the optimum pH is in the neutral-slightly acidic range (6–7). Like other microbial tyrosinases, it is generally reported not to be a very thermostable enzyme. The active site of A. bisporus tyrosinase contains two Cu binding sites, CuA and CuB, each one coordinated by three histidine residues, which interact with molecular oxygen. Depending on the Cu-ion valence and the linking with molecular oxygen, the active site exists in three intermediate states: the deoxy, oxy, and met-states. These three states determine the ability of tyrosinase to bind to its substrates and therefore determine the reaction kinetics. The met-form (Cu2+-OH−-Cu2+) of the enzyme is the resting state predominant in vivo. This form can bind diphenols and,



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during the subsequent reaction in which diphenols are oxidised and o-quinones are released, is converted into the deoxy form (Cu+-Cu+) able to bind reversibly with molecular oxygen, producing the oxy form (Cu2+-O2-Cu2+), which can act on both monophenols and diphenols. Selinheimo et al. (2006) purified and characterised a novel extracellular tyrosinase from the filamentous fungus Trichoderma reesei, whose substrate specificity, stereospecificity, inhibition, and ability to crosslink model protein substrates was compared with that of other fungal and plant tyrosinases (Selinheimo et al., 2007). Mattinen et al. (2008) reported that the T. reesei tyrosinase displays better ability to oxidise Tyr-containing model peptides and to crosslink random coil protein substrates such as α-casein than A. bisporus tyrosinase. The substrate specificity of T. reesei and A. bisporus tyrosinases was further compared by using mono- and diphenolic compounds bearing different substituents and tripeptides containing Tyr in the amine, middle, or carboxyl terminal position (Selinheimo et al. 2009). The presence of an amine group in the structure of the substrate was found to have negative effects on T. reesei tyrosinase, and phenol substrates with a carboxylic group were not effectively oxidised by A. bisporus tyrosinase. Differences in the structure of the catalytic site were hypothesised to explain these results. With reference to the ability to oxidise tripeptides, T. reesei tyrosinase showed higher Km and Vmax values but the oxidation products did not differ significantly between the two enzymes with the same substrate. The catalytic activity of tyrosinase from three different sources, including one from A. bisporus, was also investigated by Martin et al. (2008). The results showed that the catalytic efficiency towards each substituted phenol substrate is different for each enzyme source and that large differences in affinity and turnover are observed for different enzyme sources towards the same substrate. These results demonstrate how important it is to understand the substrate specificity of tyrosinases as well as their capability to oxidise different substrates in view of selecting the best enzyme for the targeted application. Tyrosinases are widely distributed enzymes in nature. They are found in mammals, invertebrates, plants, and in prokaryotic and eukaryotic microbes (Claus and Decker, 2006; van Gelder et al., 1997; Garcia-Borron and Solano, 2002). Most of the reported tyrosinases are intracellular enzymes, whereas bacterial tyrosinases, like the one found in Streptomyces, are secreted. Tyrosinases are involved in several biological functions. In mammals, pigmentation of skin, eye, and hair is a tyrosinase-mediated process called melanogenesis which contributes to protection against absorption of UV radiation (Ito and Wakamatsu, 2008). Abnormal increase or decrease in tyrosinase activity is the cause of hyperpigmentation and albinism, respectively. The control of abnormal skin pigmentation by inhibiting tyrosinase


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activity has recently become one of the most popular methods in cosmetics. Tyrosinases are also thought to play a key role in skin diseases like melanoma and in the damage of neurons related to Parkinson’s disease (Sato and Toriyama, 2009). In invertebrates, the tyrosinase-catalysed melanogenesis is related to defence reactions and sclerotisation. In plants, tyrosinases have been suggested to participate in wound healing and defence reactions mediated by quinones which can create a toxic environment, reduce the bioavailability of proteins, or contribute to the formation of barriers by polymerisation reactions. The undesired browning reaction in fruits and vegetables is related to tyrosinase activity, and methods for controlling it are constantly being searched in the food industry. In fungi, tyrosinases have been proposed to participate in spore formation, defence reactions and pigmentation. In fact, melanogenesis has a role in the formation of reproductive organs and spores and in cell wall protection after physical damage. All the most recent reports underline the very promising properties of tyrosinases for biotechnological applications. Fungal tyrosinases appear to be the most suitable candidates for the establishment of industrial processes, because they have been widely used in laboratory studies. Current limitations to establish commercial sources of tyrosinase enzymes stem from the fact that native fungal tyrosinases are generally intracellular (with only few exceptions, such as T. reesei tyrosinase) and are produced in low quantity, with poorly reproducible biological characteristics. Flurkey et al. (2008) recently examined the characteristics of various commercial preparations of mushroom tyrosinase from A. bisporus and found that they contain variable amounts of other proteins, enzymes (e.g. laccase, β-glucosidase, cellulase, xylanase), carbohydrates, and phenolic compounds. Indeed, the presence of contaminants in tyrosinase preparations, if not carefully evaluated, might affect the results and their interpretation. Current problems to be overcome for a wider biotechnological use of tyrosinases have been outlined by Halaouli et al. (2006) and by de Faria et al. (2007). One problem is to optimise the production techniques in mycelial culture, either by submerged culture or solid-state fermentation. When tyrosinase is produced from the fungal mycelium, it is necessary to use standardised purification protocols in order to remove the melanin pigments, which often remain bound to the protein after extraction, and other contaminants. The generally low stability of tyrosinase sometimes hampers its use and immobilisation techniques have been proposed as a tool to enhance enzyme stability and to allow repeated use. However, the largest and still unsolved problem seems to be related to the lack of suitable expression systems for large-scale heterologous production of tyrosinases, possibly in the extracellular medium. More research efforts in this direction are needed to design suitable expression constructs for extracellular production and to improve productivity.



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9.4.2 Applications of tyrosinases: advantages and limitations Tyrosinases are of great interest for many biotechnological applications in industry (food processing, textile, pulp and paper industry), medicine (production of l-dopa), biosensing (detection and quantification of phenolic compounds), the environment (removal of phenolic compounds from wastewater), and for the production of various kinds of proteins and/or mixed polymer crosslinked networks owing to their ability to oxidise phenolic compounds and to the high reactivity of these primary oxidation products. However, despite the active research and promising results, commercial tyrosinases suitable for industrial applications are not yet available on the market. Production of L-dopa The ability of tyrosinases to convert monophenols into diphenols can be exploited for the production of antioxidants with beneficial properties as food additives or pharmaceutical drugs (Ullrich and Hofrichter, 2007). The biosynthesis of l-dopa, the preferred drug for treatment of Parkinson’s and myocardium diseases, has been attempted using Tyr as substrate (Acharya et al., 2008). The productivity so far achieved by using mushroom tyrosinase in cell-free systems is still relatively low owing to incomplete conversion of Tyr and to side reactions leading to formation of dopaquinone, leukodopachrome, and then dopachrome molecules, which spontaneously polymerise into melanins (de Faria et al., 2007). The use of tyrosinase immobilised in Cu-alginate gels forming continuous-packed-bed systems allowed increasing productivity, though to a level still not sufficiently high to be competitive with current chemical processes or with a novel enzymatic process involving tyrosine phenol-lyase enzyme (Lee et al., 1999). Biosensing and bioremediation Enzymes share nanoscale dimensions comparable with those of metal or semiconductor nanoparticles. This paves the way for the combination of the unique electronic and photonic properties of nanoparticles with the catalytic functions of biomolecules leading to assembly of novel biosensors, nanoscale biocircuits, and bioelectronic systems (Willner et al., 2007). The tyrosinase-stimulated synthesis and growth of gold nanoparticles has been exploited for developing an optical biosensor applicable to the detection of tyrosinase activity as a diagnostic system for detecting melanoma cancer cells and Parkinson’s disease (Baron et al., 2005). The analytical arrays are relatively rapid and highly sensitive. The tyrosinase-catalysed oxidation of


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Tyr generates l-dopa. Similarly to other neurotransmitters, the latter product stimulates the formation of gold nanoparticles, which are detected by optical methods. The sensitivity of the detection of tyrosinase activity was as low as 10 units. Since tyrosinase is specifically expressed in melanocytes and melanoma cells and is viewed as a specific marker of these cells, this analytical protocol may have positive outcomes in clinical diagnostics. Increasingly strict environmental regulations require the development of more efficient analytical techniques for monitoring wastewater and air pollutants such as the phenolic derivatives generated by textile, coal, chemical, petrochemical, and mining industries. To achieve this, biosensors based on immobilised mushroom tyrosinase are currently being developed (Abdullah et al., 2006; Gu et al., 2009; Perez Lopez and Merkoci, 2009). Various measuring principles, such as detection of oxygen consumption, direct reduction of liberated o-quinones, or reduction of the o-quinones by a redox mediator, can be utilised in these biosensors. However, the previously outlined relatively poor stability of mushroom tyrosinase under storage and measuring conditions still represents a limitation for the commercial exploitation of these devices. Bioremediation, i.e. removal of phenolic compounds from wastewater, is another forefront application for which an enzymatic process could provide advantages compared with the traditional chemical and physical methods. Mushroom tyrosinase has been used as a biocatalyst confined in a flat membrane bioreactor to oxidise polyphenols present in agro-food wastewaters, and the results reported confirmed the validity of the membrane bioreactor system (Calabro et al., 2009). Wu et al. (2001) studied the ability of volatile phenols to react with tyrosinase and to couple by chemisorption onto a chitosan support. The p-cresol could be removed from vapours if contacted with tyrosinase-coated chitosan films. Mushroom tyrosinase immobilised onto chitosan provided optimum performance in terms of phenol removal efficiency, even at low phenol concentration, and for the segregation of oxidation products. Food processing Texture plays a major role in food product quality. Besides a number of hydrophobic, hydrogen bonding, and electrostatic bonds present in a food matrix, the covalent linkages in and between the food biopolymers, i.e. proteins, carbohydrates and lipids, make a major contribution to the textural characteristics of food. Enzymes are interesting tools for altering the number of covalent bonds and tailoring the properties of the end-product (Buchert et al., 2007). As previously discussed for TGases, other crosslinking enzymes such as tyrosinases can be exploited for improving various food properties. The effects of T. reesei tyrosinase-catalysed crosslinking on the



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thermal stability and gel formation ability of isolated chicken breast myofibril proteins, and the potential of tyrosinase to modify the firmness, waterholding capacity, and microstructure of cooked chicken breast meat homogenate gels were recently investigated (Lantto et al., 2007b). The results showed that tyrosinase was effective in crosslinking myofibrillar proteins. Crosslinking resulted in enhanced thermal properties, improved gel-forming ability and gel properties of the meat protein systems. Particularly interesting is the contribution of tyrosinase to improve water-holding capacity, a target that could not be achieved by TGase-aided crosslinking alone. In an attempt to tailor the textural and nutritional properties of food polymers and to develop novel food formulations, Selinheimo et al. (2008) studied the ability of two oxidative enzymes, the tyrosinase from T. reesei and the laccase from Trametes hirsuta, to catalyse the conjugation between proteins and carbohydrates. The results reported indicate that both enzymes were able to catalyse the conjugation between the protein substrate casein and the phenolic acids of hydrolysed oat spelt xylan, although to a different extent. The higher efficiency of tyrosinase to crosslink casein was confirmed, but the possible inhibitory effect of phenolic acids added to the reaction system was also highlighted. To fully exploit the potential of these enzymes for the conjugation of food polymers, further optimisation of the reaction system is needed. Biopolymer conjugation The ability of fungal tyrosinases to crosslink biopolymers has been extensively investigated using several types of substrates such as proteins and polysaccharides. Tyrosinases produce quinones, which are prone to react non-enzymatically with nucleophilic moieties, such as lysyl, tyrosyl, cysteinyl and histidinyl residues of proteins and primary amino groups of other biopolymers (for example chitosan) resulting in formation of crosslinks in the protein structures or formation of polymer conjugates. Chitosan has been widely used as model substrate to demonstrate the potential of tyrosinase-catalysed polymer modification. Payne et al. (1996) devised an enzymatic strategy to modify this natural polymer by binding tyrosinase-oxidised phenolic reactants to the polysaccharide chains, thus obtaining a polymer with new properties. Interestingly, the enzymatic approach was extended to the synthetic polymer poly(4-hydroxystyrene), whose tyrosinase-oxidised phenolic moieties were grafted with aniline or chitosan, thus proving the validity of the concept of enzymatic polymer grafting (Shao et al., 1999). Tyrosinase-aided grafting of chlorogenic acid onto chitosan under homogeneous conditions allowed widening of the water solubility window of the polymer under alkaline conditions (Kumar et al., 1999). Gelation of chitosan was enhanced by reaction with small


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molecular weight freely diffusible reactive o-quinone species obtained by tyrosinase oxidation of p-cresol (Kumar et al., 2000). The enzymatic grafting of phenolic compounds containing carboxyl groups allowed conferring on chitosan the ability to absorb cationic dyestuffs from wastewaters (Chao et al., 2004). By using dopamine as tyrosinase substrate, water-resistant adhesive properties were imparted on chitosan (Yamada et al., 2000 and 2008). Novel bio-based functional coatings for packaging materials for foodstuffs were developed by tyrosinase-catalysed grafting of octyl or dodecyl gallate onto chitosan and the coated polypropylene or paper sheets resulted in strong antimicrobial activity and effective barrier effect against oxygen transmission under dry conditions (Vartiainen et al., 2008). Chen et al. (2001) tried to disclose details of the quinone-driven conjugation of biopolymers by mimicking quinone-based natural processes, such as the transformation of animal secretions into strong adhesives (e.g. setting of mussel glue) or the sclerotisation of cuticle in insects. Various biologically active proteins were coupled to chitosan by means of a series of phenolic coupling precursors. In most cases, the protein–polysaccharide coupling was favoured by the phenolic precursor indicating that the accessibility of surface Tyr residues in globular proteins might be a limiting factor for the yield of the reaction, although it might play a key role in terms of selectivity and specificity of the reaction. All the proteins coupled with chitosan maintained their biological activity. Owing to the more open structure of gelatine compared with the globular proteins previously investigated, tyrosinase was able to oxidise the protein-bound Tyr residues and the quinone species thus formed could bind to the amino group of chitosan leading to formation of gelatine–chitosan conjugates (Chen et al., 2002). Rheological measurements gave evidence of the conjugation. Properties of these chitosan–gelatine gels and their potential use for in situ applications were further investigated by Chen et al. (2003a). This approach was further extended to grafting casein-derived peptides onto chitosan with the aim of exploiting food processing by-products for the production of environmentally friendly polymers with useful functional properties (Aberg et al., 2004). To overcome the difficulties related to accessibility of Tyr residues in globular protein the fusion proteins technology was exploited to produce a five Tyr residues tagged green fluorescent protein (Chen et al., 2003b). The Tyr tail was designed to facilitate the tyrosinase-catalysed activation, a prerequisite for the effective formation of protein–chitosan conjugates. Because of the pH-responsive properties and the electrostatic nature of chitosan, these conjugates could be selectively deposited onto micropatterned surfaces in response to an applied voltage, thus paving the way to the production of bioactive microdevices. The protein tagging technique was further exploited by Lewandowski et al. (2006) for a non-chromatographic, fully enzymatic approach aiming at recovering proteins from cell



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lysates. In a first enzymatic step, the penta-Tyr tagged green fluorescent protein was bound to the stimuli-responsive polysaccharide chitosan via tyrosinase-catalysed conjugation, thus enabling multiple washing steps for protein purification. Then, the protein of interest could be released by another enzymatic step involving either an enterokinase or a chitosanase. More interestingly, Lewandowski et al. (2008) further developed the concept of enzymatic coupling of proteins onto micropatterned surfaces upon tyrosinase activation of Tyr-tagged proteins, leading to the production of biofunctional microfabricated devices (see section 9.2.2). Binding proteins without loss of biological activity (e.g. growth factors or enzymes) onto the surface of drug delivery devices or tissue engineering scaffolds is still a field of intensive research. Some authors have recently developed a novel approach based on a highly specific coiled-coil capture/ release system to functionalise the surface of chitosan scaffolds with tethered epidermal growth factor (EGF) (Demolliens et al., 2008). In this approach, one of the coil partners (Kcoil) is coupled to chitosan via tyrosinase-catalysed oxidation of a Tyr residue of the peptide sequence. Then the EGF coil-tagged (Ecoil) protein is captured onto the surface of the chitosan scaffold by formation of a stable K/E coiled-coil structure. This versatile strategy for capturing proteins onto the surface of biomaterials, which exploits the chemoselectivity of tyrosinase in one of the key steps of the reaction protocol, seems very promising for the preparation of multifunctional devices applicable in regenerative medicine.

9.5

Functionalisation of protein fibres using tyrosinases

The quinone-mediated crosslinking of biopolymers is the basis of many biological processes dealing with the production of strong composite materials such as the exoskeleton of insects and the byssus threads of marine mussels. Their astonishing chemical, physical, and mechanical properties have stimulated researchers to exploit the quinone chemistry for the production of biomimetic materials such as adhesives (Yamada et al., 2000 and 2008) and novel hybrid fibres (Kuboe et al., 2004; Tonegawa et al., 2004). Chitosan and poly-l-Lys N-substituted with the tetrapeptides ‘Lys-Gly-TyrGly’ were synthesised and blended with the anionic polysaccharide gellan, and then fibres were spun with addition of tyrosinase to the spinning dope. In this way, the polycation–polyanion hybrid fibres formed by polyionic complexation could be further reinforced by enzymatic-assisted crosslinking, as confirmed by the results of tensile measurements. Mushroom tyrosinase from A. bisporus has been widely used to activate/ functionalise/crosslink protein fibres, e.g. wool and silk, as well as protein fractions extracted from them (keratin hydrolysate, fibroin, and sericin


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polypeptides). The amount of Tyr in whole wool fibres is 366 μmol g−1 (∼4 mol%). The Tyr content of cuticle, CMC, and cortex is 292 μmol g−1 (∼2.9 mol%), 571 μmol g−1 (∼6.2 mol%), and 389 μmol g−1 (∼4.2 mol%), respectively (Lindley, 1977). The relatively higher content of Tyr in CMC is mainly caused by the presence of the high glycine–tyrosine keratin fraction. Considering the relative proportion of cuticle, CMC, and cortex in the wool fibre, it can be calculated that about 30 μmol of Tyr are present in the cuticle, and almost the same amount is contained in the CMC. Cuticle can be reasonably considered to be the morphological fraction more exposed to enzymatic attack, but also CMC is a potential path for enzyme diffusion because it is a lightly crosslinked continuous phase that provides channels by which various molecular species can diffuse into and within wool fibres (Brady, 1992). The remaining Tyr residues (about 300 μmol) are contained in the cortical cells. The amount of Tyr residues does not seem to be a limitation for the reactivity of tyrosinase towards wool, because when wool hydrolysates were incubated with tyrosinase under homogeneous reaction conditions, the tyrosinase-catalysed oxidation proceeded steadily (Jus et al., 2009; Lantto et al., 2005b). However, under heterogeneous conditions, by using wool fibres as substrate, a range of chemical, morphological, and structural factors are likely to impose restrictions to the effective mass transfer of the enzyme from solution to fibres and to its diffusion inside the fibre matrix. Provided that adsorption and diffusion occur, the target keratin-bound Tyr residues must be located in exposed chain segments endowed with sufficient flexibility to properly fit the enzyme’s active site. For wool, enzymes should first overcome the external hydrophobic barrier characteristic of intact fibres (epicuticle) before reaching the cuticle- and/or CMC-bound Tyr residues. The possibility of modifying the Tyr residue in untreated wool fibres with tyrosinase and other oxidative enzymes was first investigated by Lantto et al. (2005b). Upon incubation with tyrosinase, no detectable activation of fibres was observed by oxygen consumption measurements. However, XPS measurements allowed small chemical changes to be detected at the fibre surface attributable to the formation of oxidised amino acid species. Intact wool fibres were used in this work and their slight reactivity under the reaction conditions used was attributed to limited accessibility of Tyr residues. Jus et al. (2008) reported the tyrosinase-catalysed grafting of antioxidant compounds (caffeic acid, chlorogenic acid) onto wool fibres. Interestingly, wool fibres incubated with A. bisporus tyrosinase displayed a continuous decrease of oxygen concentration in the reaction system over 24 h of the reaction, in apparent contrast to the results reported by Lantto et al. (2005b). However, it is not reported whether intact or surface pre-treated wool fibres were used. Wool fibres enzymatically grafted with caffeic or chlorogenic acid showed antioxidant activity, which was detectable even after a strong



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washing cycle. This finding led to the suggestion that the antioxidant molecule was effectively grafted onto the fibres. The same enzymatic approach was further extended to grafting molecules of biological interest (collagen, gelatine, elastin) onto the surface of wool fibres (Jus et al., 2009). The use of FITC-labelled collagen confirmed that the protein was grafted onto the surface of wool fibres and that it was resistant to severe washing. Mattinen et al. (2008) studied the reactivity of A. bisporus tyrosinase and of the novel T. reesei tyrosinase towards wool fibres. Because the close packing of keratins into hierarchical three dimensional fibrous structures is likely to restrict the chain mobility and to make direct crosslinking of enzyme-activated species difficult, small molecular weight freely diffusible mono- and diphenols were used as coupling agents. Moreover, wool fibres were pre-treated with an oxidising agent in order to facilitate the penetration of the enzyme into the wool fibres and to enhance the accessibility of wool-bound Tyr residues. The T. reesei tyrosinase/l-dopa system was the best performing one in terms of high fluorescence intensity of wool fibres, whereas A. bisporus tyrosinase was almost unable to activate wool fibres under the same reaction conditions. The silk proteins, sericin and fibroin, have been extensively investigated as substrates for the tyrosinase-catalysed oxidation of tyrosyl residues and subsequent grafting of chitosan under both homogeneous and heterogeneous reaction conditions (Anghileri et al., 2007; Freddi et al., 2006; Kang et al., 2004; Sampaio et al., 2005). The amount of Tyr residues in fibroin and sericin is about 5.1 and 3.7 mol%, respectively. Sericin and fibroin polypeptides dissolved in aqueous solution were effectively oxidised by A. bisporus tyrosinase. An average amount of 56–58% of the available Tyr residues were oxidised as determined by oxygen consumption measurements (Anghileri et al., 2007). The behaviour of the FTIR and Raman bands characteristic of Tyr confirmed that only a fraction of the tyrosyl residues was oxidised by the enzyme (Taddei et al., 2007). The random coil conformation taken by both sericin and fibroin in aqueous solution allowed sufficient chain flexibility for the Tyr residues belonging to the more hydrophilic polypeptide sequences exposed to the solvent to be accommodated into the tyrosinase active site and to be oxidised. On the other hand, the tyrosyl residues buried into the apolar micro-environments from which the solvent was excluded were less or not accessible to the enzyme for steric reasons. The yield of oxidation decreased significantly when the enzymatic reaction was carried out under heterogeneous condition, with fibroin in the form of gel. About 10–12% of the tyrosyl residues were oxidised based on the amino acid analysis (Freddi et al., 2006). This result can be easily explained if the primary and higher order structures of silk fibroin are considered. About 80% of the Tyr residues belong to the repetitive


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-(Gly-Ala)n- sequences (R) responsible for the formation of the more ordered and crystalline regions in both regenerated silk fibroin materials and fibres. It is reasonable to assume that these residues were hardly accessible to A. bisporus tyrosinase owing to steric hindrance. The remaining Tyr residues are located in the amorphous (A) and in the N- and C-terminus sequences, which are characterised by a less regular amino acid sequence and by a loose arrangement of the chain segments. When regenerated fibroin materials such as gels are immersed in aqueous solution the A-rich sequences can swell quite easily thus favouring the diffusion of the enzyme towards the Tyr residues. About a half of these theoretically accessible tyrosyl residues of fibroin gels were oxidised by tyrosinase. When fibroin substrates in powder (mechanically generated from fibres) or fibre form were incubated with mushroom tyrosinase, the yield of oxidation was very low or undetectable. In general, silk fibres display a high degree of inertness towards enzymes owing to a combination of morphological and structural factors. In practice, after removal of sericin, the surface of silk fibres appears very smooth, without any gap to the inside, which makes it very difficult for the enzyme molecules to find paths for diffusing into the fibre matrix. In addition to morphology, structural features of silk fibres, such as the close packing of the fibroin chains and their high degree of order, orientation, and crystallinity, also hinder penetration and diffusion inside the fibre of relatively large molecules like enzymes. In contrast to wool, where the scaled surface structure and the continuous, poorly crosslinked CMC phase allow paths for enzymes to penetrate inside the fibre and to reach the target amino acids, only amino acid residues available at the surface of silk fibres can be reasonably considered to be accessible to enzymes. The behaviour of wool and silk with proteases supports this assumption. Both fibres are susceptible to proteolytic degradation, but proteases hydrolyse more readily wool than silk. Large decreases in strength and weight are suffered by wool fibres upon exposure to proteases for relatively short times (Shen et al., 1999), whereas it takes days to detect extensive fibre degradation of silk (Arai et al., 2004). Accordingly, A. bisporus tyrosinase was able to oxidise wool fibres (Jus et al., 2008), but failed to oxidise tyrosyl residues of silk fibres (Freddi et al., 2006). A protease pretreatment lasting up to 14 days was slightly effective in opening the fibre texture and enhancing the accessibility of surface-available Tyr residues (Freddi et al., unpublished results). Hence, these results demonstrate that the accessibility of reactive amino acid residues plays a key role in determining the outcome of enzymatic reactions on protein fibres. FTIR and Raman measurements confirmed that tyrosinase oxidised the accessible Tyr residues of silk fibroin and sericin and converted them into reactive quinones and corresponding tautomers (Taddei et al., 2007). The intensity of the Tyr Raman bands decreased upon oxidation. The Raman



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band at 1615 cm−1, assigned to νC=C modes (where ν indicates stretching) of both aromatic amino acids and conjugated dienes, increased in intensity, confirming the formation of quinone species and their tautomers. The trend of the Raman I852/I830 intensity ratio, which increased upon oxidation, indicated a shift towards a more hydrophobic environment for non-oxidised Tyr residues, confirming that only those more exposed to the solvent and more accessible to the enzyme were oxidised. Broadening of FTIR band at 1650 cm−1 (νC=O mode) confirmed the presence of Tyr oxidation products. Moreover, the appearance of a band at 1045 cm−1 (νCO mode of conjugated secondary alcohols) disclosed the formation of a certain amount of the quinone methide tautomer. In a second stage of the study, the efficiency of the tyrosinase-mediated grafting of chitosan onto silk proteins was investigated (Anghileri et al., 2007; Freddi et al., 2006; Sampaio et al., 2005). pH had a strong effect on the yield of grafting. A pH value close to 7 (i.e. more favourable to tyrosinase activity) allowed a higher grafting yield, confirming that the determining step of the whole grafting reaction is the formation of the o-quinone species. FTIR spectra of chitosan-grafted silk fibroin and sericin showed a significant increase in intensity of the bands near 1400 and 1000 cm−1, where chitosan absorptions fall. Accordingly, the Raman spectra also showed several bands ascribable to chitosan. Although the mechanism of the nonenzymatic grafting step is still poorly understood, some spectroscopic data seem to support the occurrence of a Schiff base reaction. In fact, the presence of a shoulder at about 1646 and 1650 cm−1 in Raman and FTIR spectra, respectively, may be attributed to the imine group, whose νC=N mode falls in this region. However, the Michael addition-type mechanism cannot be completely excluded, as indicated by the trend of the νNH FTIR region, where the grafted samples showed higher wavenumbers broadening and the presence of a single νNH band. These spectral features are consistent with the conversion of some primary amino groups of chitosan into secondary amino groups (according to the Michael addition-type pathway). The yield of grafting was dependent on the extent of oxidation and followed the order: sericin/fibroin aqueous solution >> hydrated gel > dried gel >> mechanically generated fibroin powder. Grafting onto silk fibres did not occur, or occurred at an undetectable level. The results reported on the silk proteins/chitosan model system demonstrate the potential of the tyrosinase-initiated grafting for the formation of novel biopolymer conjugates suitable for a range of applications (e.g. textile, coating, packaging, biomedical). Grafting chitosan onto sericin or fibroin polypeptides may complement the outstanding properties of the polysaccharide (antimicrobial activity) with those of the silk proteins (antioxidant, UV-resistant, moisturising properties of sericin; strength, biocompatibility, biodegradability of fibroin), resulting in the production of valuable bio-based polymers under


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the mild conditions assured by the specificity and selectivity of enzymes. With reference to silk fibres, the accessibility and reactivity of surfaceavailable tyrosyl residues still remain to be proved. Overcoming of the current limitations will allow development of innovative techniques for surface functionalisation of textile fibres.

9.6

Other enzymes for protein fibre functionalisation

In addition to transglutaminases and tyrosinases, other enzymes have attracted some interest for their ability to interact with protein fibre substrates. Such is the case for laccases and peroxidases, both belonging to the group of oxidoreductases. Laccases (p-diphenol : dioxygen oxidoreductase, EC 1.10.3.2) are Cu-containing enzymes, having four Cu atoms in their reactive site, which are able to oxidise their substrates by removing one electron and using oxygen as terminal electron acceptor (Riva, 2006; Widsten and Kandelbauer, 2008) (Fig. 9.3). The radical species thus formed undergo further non-enzymatic reactions. Despite intensive studies, the detailed mechanism of laccase-catalysed reactions is still poorly understood. A typical feature of these enzymes, which can be found in plants, fungi, and insects, is their broad substrate specificity, ranging from the oxidation of o- and p-diphenols, aminophenols, polyphenols, polyamines,

(a)

(b)

H2 O

LaccaseOX

SubstrateRED

O2

LaccaseRED

SubstrateOX

H2O

LaccaseOX

MediatorRED

SubstrateOX

O2

LaccaseRED

MediatorOX

SubstrateRED

(c)

O•

OH

OH

n/4 O2 n/2 H2O

n

LACCASE

R

n

*

R

*

R

n

9.3 Redox cycle of laccase in (a) the absence or (b) the presence of a chemical mediator. (c) Laccase-catalysed oxidation of phenolic substrates. Intermediate radical species can undergo polymerisation by radical–radical coupling.



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lignins and aryl diamines as well as some inorganic ions. In the presence of small molecules capable of acting as electron transfer mediators, laccases are also able to oxidise non-phenolic structures or substrates otherwise not accessible to the enzyme, thus expanding the range of compounds that can be oxidised by these enzymes. Laccases have received much attention for their application in several biotechnological processes. Such applications include the detoxification of industrial effluents, bioremediation of soil, medical diagnostics, manufacture of anti-cancer drugs, and as ingredients in cosmetics (Rodríguez Couto and Toca Herrera, 2006). Peroxidases (hydrogen-peroxide oxidoreductase, EC 1.11.1.7) have been discovered in various plants, microbes, and mammals (Floché and Ursini, 2008). The plant peroxidase from horseradish (HRP) is one of the best characterised enzymes in this group of oxidoreductases, which has found a wide range of biosensing and biotechnological applications, including the peroxidase-catalysed polyphenol synthesis (Ayyagari et al., 1996). HRP is a hemoprotein of about 44 kDa containing an atom of iron in the protoporphyrin IX prosthetic group. It uses hydrogen peroxide (H2O2) as electron acceptor for the oxidation of a wide range of substrates. The generally accepted reaction mechanism of HRP involves the formation of two oxidation states of the Fe-containing prosthetic group, i.e. compound I and II (Fig. 9.4). The catalytic cycle is initiated by the rapid oxygen transfer from a peroxide to the resting ferric state of the enzyme (two electron oxidation), resulting in the formation of compound I intermediate state. The enzyme regains its resting state by passing through two consecutive one-electron reduction steps, with the formation of the intermediate compound II (one electron reduction of compound I). In the presence of excess H2O2, HRP is reversibly transformed into compound III, an inactive form of the enzyme (Bagger and Williams, 1971). Electrons are extracted from aromatic substrates, which form reactive radical species that usually undergo nonenzymatic radical–radical coupling. Formation of dityrosine by in vitro HRP-catalysed oxidation of free Tyr and Tyr-containing peptides and proteins in the presence of H2O2, has been reported (Michon et al., 1997). Current applications of laccases and peroxidases in textile processing mostly refer to laccase-mediated indigo decolorisation in denim finishing and to removal of reactive dyes by means of a chemo-enzymatic peroxidase-based technology (Schäfer et al., 2007). Both enzymatic systems are commercially available. Several enzymatic techniques for dyeing based on the use of laccase and peroxidase redox enzymes have been tested, but not yet commercialised. Worth mentioning are the studies on decolorisation of textile and other industrially important dyes from wastewater as an alternative strategy to conventional chemical, physical and biological treatments (Husain, 2006). The potential of peroxidases, laccases, and other oxidases is under investigation. The addition of certain redox mediators enhances the


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Advances in textile biotechnology H2O2

(a)

H2O

HRP

Compound I

R•

RH Compound II

RH

•

R

(b) nH2O2 2nH2O

n RH

n R•

HRP

*

RH

* n

(c)

O·

OH

2

HRP/H2O2

OH

2 OH

9.4 (a) Redox cycle of horseradish peroxidase (HRP). (b) Simplified scheme of the HRP-catalysed oxidation of a substrate. Intermediate radical species can undergo polymerisation by radical–radical coupling. (c) HRP-catalysed phenolic coupling of tyrosine residues in proteins with formation of dityrosine.

range of substrates and efficiency of degradation of the recalcitrant dyes. Because soluble enzymes cannot be exploited on a large scale owing to limitations such as stability and reusability, immobilised enzyme systems are used. Despite great efforts in this field of study, treatment of wastewater on a large scale has not yet been implemented. The utility of laccases not only for dye degradation but also for the biosynthesis of new dyes suitable for textile and leather coloration has recently been demonstrated (Enaud et al., 2010). A novel azoanthraquinone dye called Laccase Acid Red 1 has been synthesised by means of an immobilised laccase system. The resulting dye preparation was neither toxic nor mutagenic, and displayed good dyeing properties on polyamide fibres. The enzyme-catalysed synthesis of dyes represents a promising safer alternative to chemical synthesis, which still poses health and environmental problems. There have been few studies on the application of laccases and peroxidases to the functionalisation of protein fibres. Lantto et al. (2004) reported the laccase-mediated oxidation of wool and model compounds (Cys and Tyr) with and without the mediators violuric acid (VA) or 1-hydroxybenzotriazole (HBT). Without a mediator, laccase was unable to oxidise wool or the amino acids. In the presence of mediators, oxygen consumption was



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observed, indicating oxidation of the secondary substrates of laccase, i.e., wool, Cys, or Tyr. Lantto et al. calculated that with the laccase/HBT system about 5% Cys and 20% Tyr residues present in the cuticle were oxidised. However, only very low amounts of oxidised species were detected by XPS analysis of the wool surface, probably because oxidised mediator molecules that were absorbed or covalently bound to the fibre surface interfered with measurements. When wool fibres were exposed to the action of two peroxidases, HRP and a peroxidase of fungal origin (NS51004), about 35–40% of the Tyr residues located in the cuticle were oxidised based on H2O2 consumption (Lantto et al., 2005b). XPS and FTIR analyses could detect only slight changes in surface chemistry of wool fibres incubated with the fungal peroxidase. The reaction of silk fibres with HRP and H2O2 was investigated by studying the intermediates enzymatically generated by means of electron paramagnetic resonance (EPR) and UV/vis spectrophotometry (Oliva et al., 2003). The time dependence of the EPR spectrum indicated that HRP compound II was still present after 5 min of silk fibre oxidation, and this was confirmed by spectrophotometric results. On the contrary, after about 10 min, only the presence of HRP compound I was detectable. The latter species became weaker within a few weeks with sample ageing. Furthermore, both HRP compounds I and II were observed to decay in the presence of silk, the latter more quickly than the former. By contrast, these two oxidised forms of HRP remained stable (compound I) or decayed more slowly (compound II) without silk. This suggests that silk took part in the reaction as an electron donor probably by oxidation of the Tyr side chains of silk fibroin by HRP. The peroxidase- and laccase-catalysed coloration of wool fibres has been reported by Shin et al. (2001). In situ production of oligomers and polymers from different phenolic compounds by HRP/H2O2 allowed different colours to be obtained on wool fibres, from black to brown, grey, and yellow. To avoid the use of H2O2, laccase was used to oxidise hydroquinone or ferulic acid for the enzymatic dyeing of wool. Colour depth was sensitive to both enzyme and phenol concentration. A mordanting treatment with chromium compounds enhanced the fixation of the phenolic colouring matters onto wool fibres. The enzymatic coloration of wool fibres was further investigated by Tzanov et al. (2003), who applied a factorial experimental design to understand the effect of different reaction parameters in an enzymatic system comprising laccase, a dye precursor, and catechol or resorcinol as dye modifiers. Different hues and depths of shade were obtained by varying the concentration of the modifiers and the time of laccase treatment. By coupling ultrasonic irradiation with cyclic voltammetry for the laccasemediated oxidation of 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonate) (ABTS) in the presence of wool fabrics, a high depth of wool coloration


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could be achieved owing to the high affinity of ABTS oxidation products for keratins and to improved mass transfer within the reaction system (Munteanu et al., 2007). These results show the potential of the enzymatic approach to wool dyeing and how the process can be intensified by suitable technological developments. However, the range of colours, hues, and depth of shades so far obtained is too limited for enzymatic dyeing to be an attractive alternative to conventional chemical dyeing. Their technological performance (reproducibility, colour fastness) is still a challenge. Further studies are needed to explore a wider range of enzyme, phenols, and modifiers/additives combinations in order to widen the range of colours, to enhance their performance, and to achieve industrially acceptable standards. This overview of non-proteolytic enzymes of textile interest cannot be considered complete without mentioning emerging enzymatic activities that are likely to open new perspectives for the functionalisation of protein fibres. These novel enzymes are sulfydryl oxidases (SOXs) and protein disulfide isomerases (PDIs). In living organisms, these enzymes play a key role in the post-translational modification and folding of newly synthesised proteins leading to stabilisation and regulation of protein structure and function. In particular, these enzymes are involved in the formation and rearrangement of protein disulfide bonds (Appenzeller-Herzog and Ellgaard, 2008; Fass, 2008). Disulfides are covalent bonds produced by oxidation of two free thiols of cysteine residues, both as intramolecular or intermolecular bonds, to stabilise the protein structure, but they also have the capacity to work as regulatory switches in redox signalling in cells. SOXs have been identified in animal, plant, and fungal species, as well as in the genomes of viruses. The eukaryotic SOXs characterised to date are classified into two families: Ero1 and Erv. All SOXs contain a flavin adenine dinucleotide (FAD) bound to the protein. SOXs promote cysteine pairing by transfer of electrons from thiol groups to molecular oxygen. Common to both eukaryotic families of SOXs is a CXXC motif (C is cysteine) adjacent to the FAD that participates in the two-electron transfer. The redox activity of PDI enzymes is also governed by the CXXC active site. When in the oxidised state, the disulfide can be transferred to the substrate to catalyse its oxidation whereby the active site itself becomes reduced. When in the reduced state, substrate disulfides can be reduced and the active site ends up in the oxidised state. These thiol–disulfide exchange reactions proceed through the formation of a transient mixed disulfide between enzyme and substrate. Therefore, catalytic PDIs harboring a CXXC active site sequence are specialised disulfide carrier proteins that act both as disulfide-donor and -acceptor enzymes, i.e. catalysing both cysteine oxidation and disulfide reduction in their substrates. Disulfide isomerisation is a crucial reaction in biological systems not only for its key role in driving



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protein folding but also because PDIs can catalyse the rearrangement of wrongly formed disulfides, which often occur during folding and prevent the formation of the native structure. The biotechnological potential of these enzymes is still under scrutiny. Research efforts dealing with screening of micro-organisms for the identification, isolation, and purification of SOX and PDI activities suitable for laboratory-scale trials are in progress. Approaches aimed at inducing overproduction or heterologous production of these enzymes are also being investigated. No results have been reported yet on their application on protein fibres. Preliminary studies on model peptides and proteins are needed to disclose the real potential of these enzymes for the functionalisation of wool and silk fibres.

9.7

Conclusions and future trends

Enzymatic reactions on protein fibres typically occur under heterogeneous conditions, with fibres in the form of solid substrate. Most of the studies reviewed in this chapter, as well as others not considered here, demonstrate how complicated these systems are and how many chemical, physical, structural, and morphological parameters can influence the outcome of the enzymatic reactions. The experimental results obtained do not always comply with the initial hypotheses. A deeper understanding of the enzyme/ substrate interactions, as well as a more detailed knowledge of the effects of various experimental variables on reaction kinetics and yield would be very helpful for devising proper strategies for the enzyme-mediated modification of protein fibres. The individual polypeptide chains forming wool and silk fibres are held together by a range of weak-to-strong molecular interactions (from hydrophobic interactions to electrostatic, hydrogen, and covalent bonds) which result in the formation of compact and tough three-dimensional fibrous structures, whose main primary function in nature is to resist external chemical, physical, mechanical, and biological stresses. When immersed in aqueous solution, these fibres may undergo more or less extensive swelling thus allowing penetration and diffusion of reagents inside the fibre matrix. Owing to morphological, chemical, and structural factors, enzymes can penetrate more easily inside wool than silk fibres. This suggests that only amino acid residues located at the surface of the fibres can be potentially exploited for the enzymatic functionalisation of silk, whereas for wool, enzymes can also use a certain amount of the amino acid residues confined in the fibre matrix. Although the total amount of potentially reactive amino acid residues in a fibre substrate is important, the effective accessibility of these amino acid residues is much more important for a reaction to occur. Enzymes must not


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only be able to reach the polypeptide chain segments where the target amino acidic groups are located, but must also be able to bind with them. As demonstrated by the results reported for the TGase-catalysed modification of globular proteins in aqueous solution, even exposed Gln residues are not attacked by the enzyme if other requirements are not fulfilled. In particular, the pattern of surrounding amino acidic sequences, as well as the local structure of the polypeptide chain segments where the amino acidic groups are located must be endowed with sufficiently high mobility and flexibility to allow the active site of the enzymes to come into close contact with the substrate and to bind to it. Consequently, it is reasonable to suppose that a certain number of potentially reactive amino acid residues of wool and silk, either at the surface or inside the fibre matrix, do not match the above requirements, being involved in hydrogen and/or covalently bonded secondary and higher order structures that are not flexible enough to interact dynamically with the enzyme molecules. However, various enzymatic or chemical treatments can modify protein fibres not only at the surface level but also in the bulk, thus enhancing the accessibility of amino acid residues by removing barrier elements, by changing their chemical environment, or by inducing conformational transitions of the backbone peptide chains where they are located. Far from being considered a limitation, the above morphological, molecular, and structural features must be viewed as a way to address selectivity and specificity in the enzymatic modification of wool and silk fibres. To fully exploit the great potential of enzymatic tools, it is important to gain wider understanding of the accessibility and reactivity of the target amino acid residues in wool and silk. Building a database of accessible amino acid residues in protein fibres is still a challenge for textile biotechnology scientists. However, unravelling fundamental aspects of the enzyme/substrate interactions would make it possible to design suitable enzymatic functionalisation strategies able to address specific physicochemical and functional properties of wool and silk fibres. TGases and tyrosinases are both protein crosslinking enzymes. As described in the previous paragraphs, inter- or intramolecular crosslinking may directly result from the TGase-catalysed reaction between Gln and Lys or may be the outcome of a non-enzymatic reaction step between tyrosinase-activated chemical species, typically quinine–quinone or quinone– nucleophile coupling. To allow crosslinking, the amino acid groups involved should be in the right position to meet the steric requirements of the enzyme active site or must be close enough to reach the reactive counterparts. It is reasonable to suppose that these conditions are not always fulfilled when the amino acid residues belong to polypeptide chains that are blocked into closely packed three-dimensional fibrous structures such as wool and silk. Crosslinking of wool fibres by TGase-catalysed reaction has



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often been claimed, but it has never been directly proved by determination of the newly formed ε-(γ-glutamyl)lysine bonds. Tensile measurements have always been used as an indirect method to assess crosslinking. However, single-fibre measurements have never been reported. This does not mean that TGase-catalysed crosslinking did not occur within wool fibres, but it is very difficult to understand to what extent the observed changes in fibre strength were the result of crosslinking alone or if other unknown experimental factors played some role. To overcome some limitations imposed by the accessibility of amino acid residues and to effectively functionalise protein fibres, TGase and tyrosinase can be used in combination with suitable freely diffusible small molecular weight compounds able to take part in the reaction. Various primary amine reagents and a wide range of phenolic compounds are TGase and tyrosinase substrates, respectively. These compounds covalently bound to the surface or in the bulk of protein fibres by enzyme-mediated reaction will impart on silk or wool the specific functionality carried by their pendant group. Another approach would be the preliminary induction of suitable changes in surface chemistry by plasma or electron beam irradiation or grafting treatments in order to increase the number of potentially reactive sites available for subsequent enzymatic functionalisation. The combination of emerging nano- and biotechnologies represents an attractive field of intensive research likely to bring real innovation in textile processing in the near future. With reference to the different sources of TGase and tyrosinase enzymes, a careful screening and selection of the enzymatic activities better fitting the needs of the intended application is an important prerequisite for the success of any laboratory-scale or pilot trial. The possible interference of additives or contaminants (especially the presence of other enzyme activities) must be firstly evaluated. More importantly, enzymes must be selected with a special attention to their intrinsic properties and performance, as it has been shown that different sources of the same enzyme, such as the tyrosinases from T. reesei and from A. bisporus, display radically different properties in terms of reaction kinetics and substrate specificity. Finally, one of the most critical issues for the industrial exploitation of enzymes is their mass production and supply at a reasonable price. In recent years, TGases and tyrosinases have attracted much interest from scientists and technologists from various sectors, including textiles. Despite the high attractiveness of enzymatic functionalisation and crosslinking of protein fibres, the development of large-scale industrial applications is still blocked because tyrosinase is currently not commercially available in sufficiently large quantities and at a reasonable cost to support process scale up, whereas TGase is commercially available but its formulation does not fit the requirements of textile processing. TGases and tyrosinases have demonstrated that


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they work reasonably well in laboratory-scale trials and meet the needs of a broad range of applications, from biology, to medicine, pharmacology, biosensing, tissue engineering, food, leather, and textile processing. Although there appears to be a lack of enzyme robustness, the results so far obtained offer many options for further research efforts leading to more consistent improvements in terms of reaction engineering. Taking advantage of the originality and the effectiveness of some biotechnological approaches devised in non-textile sectors, textile biotechnologists should make further research efforts aimed at developing novel enzymatic strategies able to fully exploit the potential of TGases and tyrosinases, as well as of other nonproteolytic crosslinking enzymes, for protein fibres functionalisation. Future results, if positive, will probably create the conditions for raising a large market demand for industrial TGases and tyrosinases, which, in turn, will stimulate biotechnology companies to invest in the development of these enzymes, thus paving the way for the establishment of commercial processes. Not only will the textile industry value this, but consumers will also be able to take advantage of the tangible benefits offered by the biotechnological functionalisation of protein fibres.

9.8

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10 Developments in recombinant silk and other elastic protein fibers for textile and other applications J. C. RODR ÍGU E Z - C A BE L L O, C. GA RC Í A-A R É VA L O, L. M A RT Í N, M. SA N T O S and V. R E BO T O, University of Valladolid, Spain

Abstract: The ability to manipulate genes and their products by recombinant DNA has signaled a number of new possibilities for the production of modified or new fibrous biopolymers or protein-based polymers with a combination of strength and elasticity similar or even superior to that of synthetic high-tech fibers. Biotechnological approaches offer the opportunity to replace existing chemical or mechanical processes for a cleaner production technology than conventional procedures, which cause severe pollution problems from textile effluents. Key words: recombinant DNA, fibers, silks, collagens, elastin-like polymers, resilin.

10.1

Introduction

Nature is replete with structural materials in the form of fibers and biocomposites that have attained remarkable levels of efficiency and performance through eons of evolutionary selection (O’Brien et al., 1998). Although these natural polymers remain, in some cases, commercially important because of their cost, functionality and consumer preferences, those shortcomings associated with quality variations along with their hydrophilic nature and low thermal stability have led to them being replaced by synthetic polymers with more desirable properties (Kalia et al., 2009). With the advent of organic and petroleum-based chemistry in the early 20th century, natural polymers have been increasingly substituted by synthetic polymers and fiber development that, years ago, resulted in a family of new products, such as nylon, polyester, acrylic, aramid, spandex, olefin resins and fibers, with superior tensile strength and stress–strain behavior (O’Brien et al., 1998). A new class of ‘engineered’ peptide-based biopolymers that has attracted much attention, consists of materials deriving from two scientific developments: the increasing understanding of protein structure–function, which provides peptide motifs that are useful for the design of repetitive, 235 © Woodhead Publishing Limited, 2010

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polypeptide-based materials, and the maturation of recombinant DNA technologies, which allows these materials to be synthesized in large yields with precise control over the chain-length, stereochemistry and monodispersity (Chow et al., 2008). Protein polymers with amino-acid sequences based on the consensus repeat sequences of silk, collagen, elastin glutenin, and resilin have been a primary focus in the design of fibrous protein based polymeric materials (Kiick, 2007).

10.2

Principles of recombinant DNA methods applied in the production of recombinant proteins

The ability to manipulate genes and their products by recombinant DNA technology in microbial, animal and plant systems has signaled a number of new possibilities for the production of modified or new fibrous biopolymers or protein-based polymer as well as modified or new enzymes with better properties (more temperature stability, increased activity and improved performance) by ‘splicing’ relevant enzymatic genes into another organism which has improved pre-existing treatments such as many enzymatic transformation processes of textile processing and after care. Indeed, biotechnological approaches offer the opportunity of replacing existing chemical or mechanical processes for a cleaner production technology than conventional procedures which cause severe pollution problems from textile effluents (Chen et al., 2007, Rogers, 2000). The control of biosynthetic methods for specifying polymer sequence, molecular weight and consequently structure, arises from the high fidelity (error rates less than one in 104) with which a gene encoding a target protein is converted to its corresponding protein sequence at the ribosome. The two general steps by which this conversion occurs are: first, a specified DNA sequence is copied into a corresponding mRNA sequence (transcription), and second, these mRNA fragments are used as templates to direct the synthesis of proteins (translation). In the translation of the mRNA template into a protein, transfer RNA (tRNA), which acts as an adaptator between the mRNA and the amino-acids of the proteins, and the enzymes aminoacyl-tRNA synthetases, which control the attachment of amino acids to their cognates tRNAs, are involved. Because there are multiple approaches for assembling target genes, this strategy has allowed the synthesis of many types of protein polymers and block co-polymers (Kiick, 2007, Sankaranarayanan and Moras, 2001). Frequently, discrete domains of proteins support a certain structural, biochemical or biological activity and thus it is often not necessary to express the full-length protein to address a particular biological question (Hallberg, 2008). Known fibrous proteins such as silks, elastin and collagens are characterized by a highly repetitive primary sequence (mainly composed of tandem of repeated blocks of amino-acids sequences), which leads © Woodhead Publishing Limited, 2010

Developments in recombinant silk and other elastic protein fibers

Unfolded

α-Helix

310-Helix

β-Sheet

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β-Turn

10.1 Common secondary structural motifs in proteins.

to significant homogeneity in secondary structure (Fig. 10.1) and usually exhibit important mechanical properties (Altman et al., 2003). By combining polypeptide sequences derived from these proteins in different ways, a new protein polymer can be biosynthesized using DNA techniques with unique physical, mechanical and biological features, such as variations of folding, chain interactions within the synthetic protein structure, temperature or pH responsiveness or others (Qiu et al., 2009). Devising an appropriate strategy at the nucleotide level is essential for the efficient synthesis of the protein encoding sequence and for producing a uniform protein product with an optimal quality and yield of the final protein product. The biosynthesis of any artificial protein generally includes: (1) constructing a synthetic gene encoding the protein of interest in a plasmid with tight transcription control; (2) cloning of the recombinant gene with the necessary transcriptional regulatory elements to competent cells; (3) screening plasmid containing cells for ones containing the desired clones and verifying the DNA sequence; (4) transforming the chosen plasmids into expression competent host cells; (5) growing appropriate volumes of host cells and inducing protein expression; (6) purifying the protein of interest from cell lysates (Mi, 2006).

10.3

Biomimetic design of recombinant proteins

In designing genes encoding repetitive protein-based polymers, the techniques of molecular biology are typically employed to self-ligate monomer DNA fragments in a process of oligomerization that relies on restriction enzyme-based approaches. In this instance, the monomer fragments must be oligomerized in a ‘head-to-tail’ orientation, and can be seamless in sequence or can contain intervening linkers between the desired repeats. The approaches to the oligomerization can be classified as: 1

Iterative technique, where a DNA segment is oligomerized in a series of single, uniform steps, each step growing the oligomer by one unit length of the monomer gene. © Woodhead Publishing Limited, 2010

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Random method or ‘concatemerization’, where an uncontrolled number of monomer DNA segments are oligomerized in a single step, creating a population of oligomerized clones of different lengths. This method has the advantage of creating, in a simple ligation step, a library of genes of different lengths that encode oligomeric polypeptides with the same repeat sequence. However, it sacrifices precise control over the oligomerization process, because it is a statistical process and, therefore, it is difficult to, a priori, design a polypeptide of a certain size and difficult to synthesize genes encoding large polypeptides (>100 kDa). Recursive directional ligation (RDL), where DNA segments with two different restriction sites flanking the insert are joined in sequential steps with the length of the ligated segments growing geometrically in each step. This approach is applicable for the synthesis of repetitive polypeptides of specific and predetermined chain length because this process seamlessly joins the two monomeric inserts and also eliminates the restriction site at the junction, so that the two unique restriction sites are now on either end of the dimerized gene (Fig. 10.2) (Chilkoti et al., 2002, Rodríguez-Cabello et al., 2008).

3

E E

E E

E

E

E

E

E

E

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E E

E E

E

E E

E E Random

E E

E E

E

E

E

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10.2 Schematic representation of three DNA oligomerization techniques (adapted from Rodríguez-Cabello et al., 2008 and Chilkoti et al., 2002). ‘E’ represents the targets for restriction enzymes flanking the inserts that are joined in the construction of the repetitive polypeptides.



10.4

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Expression systems

When recombinant DNA techniques are used to express certain genes outside their natural expression habitat, the process is described as heterologous gene expression. In order to express a target protein heterologously in a host, the target DNA-coding sequence has to be known and a suitable transformation and selection strategy has to exist (Mahmoud, 2007). When a designed gene is introduced into a suitable vehicle (called a vector; usually on a plasmid), and then into a selected host, the progeny of such a hosting cell (called a clone), produce the recombinant protein specified by the codified gene (Jackel et al., 2008). For DNA delivery and integration non-viral gene transfer remains the preferred approach to generate stable cell lines for manufacturing purposes. Calcium phosphate transfection, electroporation, lipofection and biolisticand polymer-mediated gene transfer, are routinely used and are all reasonably efficient and reliable procedures for transferency (Godbey and Mikos, 2001, Jo and Tabata, 2008). Expression constructs are chimerical structures in which the transgene is bracketed by various regulatory elements. To achieve high yields, the design of expression constructs must optimize all stages of gene expression, from transcription to protein stability. The central elements essential in the design of any recombinant expression systems must include: 1

2

3

4

5

origin of replication (ori), which is a particular sequence at which replication is initiated and that allows the autonomous replication of the vector within the cell; selection markers are sequences encoding a selectable marker that confers particular features to the transformed hosts for subsequent selection and assures maintenance of the vector into it; a promoter that controls the level of gene expression. Expression vectors generally use a strong viral or cellular promoter/enhancer to drive the expression of the recombinant gene; a terminator, which is a sequence for ensuring that the RNA polymerase disengages and does not continue to transcribe downstream genes. A strong transcriptional terminator should be used with a strong promoter; polylinkers or cloning sites for cutting and pasting of DNA fragments to simplify the insertion of the heterologous gene in the correct orientation within the cell (Rai and Padh, 2001).

A variety of surrogate hosts are available for in vivo heterologous recombinant protein production. These actually encompass five groups of organisms: bacteria, fungi, plants, insects and mammals (Makrides, 1996). The optimal expression system for generating sufficient amounts of one protein must be selected based on protein yield, purity, structure, final application


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and economics of scale-up (Huang et al., 2007). These factors depend, in turn, upon the nature of the target protein encoded, their chemical properties such as size, iso-electric point, stability over various pH and temperature ranges, and proteolitic resistance as well as the post-translational modifications required, such as proper-folding, glycosylation, phosphorylation, formation of disulfide bridges (Rai and Padh, 2001). Prokaryotic systems are generally easier to handle and are satisfactory for most purposes. Many microbial species have a strong natural capability of expressing proteins, which makes them very attractive candidates for the heterologous production of recombinant proteins. The most successful and routinely used microbial recombinant protein expression systems mainly belong to bacterial and fungal species.

10.4.1 Bacteria Escherichia coli offers a means for the rapid and economical production of recombinant proteins and is, by far, the most widely employed host for many reasons: it is well characterized, its genome sequence is known, many of its biological processes and metabolic pathways are understood and there are many genetic tools readily available for its manipulation. Indeed, high growth rates combined with its ability to express high levels of heterologous proteins (i.e. strains producing up to 30% of their total protein as the expressed gene product), result in high volumetric productivity. Furthermore, it can grow rapidly to high densities in simple and inexpensive media. However, there are various limitations for E. coli as an expression system. Many proteins are complex, containing multiple subunits, cofactors/ prosthetic groups, disulfide bonds, and post-translational modifications, including glycosylation, that are essential for their function. Producing such complex proteins in E. coli may be quite challenging. In addition, high-level gene expression levels can make these cells physiologically ill and stresssensitive, thus creating a metabolic burden that limits nutrient and oxygen availability, and the recombinant product can induce various levels of toxicity. For this reason, a compromise in balancing the levels of gene expression and cell growth needs to be reached to maximize the volumetric recombinant protein productivity. Indeed, bacterial lysis to recover the cytoplasmic proteins often results in the release of endotoxins, which must be removed from the final product (Chou, 2007). Recombinant expressed proteins in E. coli can, in principle, be directed to three different locations, namely the cytoplasm, the periplasm or the cultivation medium. Expression in the cytoplasm is normally preferable because production yields are higher. However, protein degradation is more likely to occur in the cytoplasm of E. coli than in other compartments (Makrides, 1996). Proteins expressed in large amounts in the cytoplasm



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often accumulate as soluble aggregates, called inclusion bodies, because of hydrophobic interactions between exposed patches on unfolded chains or other unknown specific clustering mechanisms and to inadequate protection from chaperones. These inclusion bodies are a set of structurally complex aggregates that can be isolated by differential centrifugation and provide a useful concentration and purification step in simple plasmid constructs and protect target proteins from protease degradation (Swartz, 2001). Extensive efforts have been made in order to improve the recombinant expression of heterologous proteins in E. coli and many strategies have been employed for achieving the high yields demanded: 1

strains deficient in the most harmful natural proteases belonging to the genetic bacterial background, such as E. coli BL21 and derivative strains; 2 strains with mutations inhibiting mRNA proteases (such as RNase E, RNase K and RNase III) that stabilize mRNA and increase the frequency of mRNA translation; 3 strains modified by site-directed mutagenesis or co-expressing rare tRNAs for avoiding codon bias; 4 the development of protein fusion partners in order to simplify the purification and expression of recombinant proteins such as His-tag (polyhistidine tag) or glutathione S-transferase (GST) tag, that can protect target proteins in vivo from intracellular proteolysis, enhance solubility, or be used as specific expression reporters, as well as increasing expression levels as result of mRNA stabilization (Swartz, 2001, Sørensen and Mortensen, 2005).

10.4.2 Fungi Many advantages have made yeast a favored alternative to bacterial systems for recombinant expression of foreign proteins: these lower eukaryotic systems include a rich diversity of species and metabolic routes able to produce high yields in recombinant target proteins secreted to the media, the availability of the complete genomic sequence and the possibility to manipulate it, provide a good compromise between bacteria on one side and mammalian cell lines on the other (Rai and Padh, 2001). However, only a few, mainly yeast and moulds, have been extensively exploited for their heterologous protein production skills. Yeast species such as Saccharomyces, Pichia, and Hansenula are well known as efficient gene expression hosts with good fermentation properties (Pichia pastoris until 65 g L−1). Similarly, mould species such as Trichoderma and the Aspergilli, are distinguished by their high enzyme secretion levels that can reach 20 g L−1. Some disadvantages of these expression systems include hypermannosylation, hindering


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proper folding and therefore the activity of some proteins and hence lower efficiency in introducing the foreign DNA into the host cells (Mahmoud, 2007).

10.4.3 Plants Recombinant production of proteins in plants has many advantages compared with other expression systems. These include the potential for rapid agricultural scale production at an extremely competitive cost, the availability of established practices for their efficient harvesting, transporting, sorting and processing, the absence of human pathogens, the ability to fold and assemble complex proteins accurately and the possibility of governing sub-cellular targeting (seeds or endoplasmic reticulum) instead of the secretory pathway, thus affecting the interlinked processes of folding, assembly post-translational modification and long-term storage. Plants can also be used in large-scale microbial-like fermentation production facilities by using plant-cell suspensions. The range of recombinant proteins made in plants has extended to include industrial enzymes, technical proteins that are used in research, milk proteins that are a suitable nutritional supplement, and new protein polymers with both medical and industrial uses. There is little doubt that recombinant antibody production in plants is likely to have a great impact on human health. Protein polymers that have been produced in transgenic plants include human collagen, a synthetic spider silk and bovine elastin, the latter being produced at low yields (Ma et al., 2003). Several challenges remain to be met in plants in terms of increasing yields, lack of sufficient information on post-translational events, downstream processing and addressing biosafety (the main perceived risks are transgene spread by pollen dispersal and horizontal gene transfer, and the effects of potentially toxic recombinant proteins on herbivores, pollinating insects and micro-organisms in the rhizosphere. There is also concern that plant material that contains recombinant proteins could inadvertently enter the food chain) and acceptability issues, as well as industry inertia (Ma et al., 2003).

10.4.4 Insects The use of recombinant baculoviruses such as the nuclear polyhedrosis virus of the ‘fall armyworm’ Autographa californica (AcNPV), of the silkworm Bombyx mori (BmNPV) or of the gypsy moth Lamantria dispar (LdNPV) and insect tissue-culture cells have demonstrated considerable advantages. These include: high protein yields up to 30–500 mg L−1, large scale production, and the option of expressing proteins with posttranslational modifications and others that would otherwise be toxic at high



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levels in mammalian cells (Griffiths and Page, 1997). Insect cell cultures provide an environment in which the process parameters can be carefully controlled and monitored allowing the use of a single cell line with more uniform post-translational modifications. The main drawbacks are that the process requires a significant set-up and running cost for the sterile growth facilities. The employment of larvae is more advantageous because the cost is lower than insect cell culture and some post-translational processing events such as glycosylation and secretion may be more efficient in larvae, with final protein yield approximately equal to 1 L of cultured cells (Mahmoud, 2007).

10.4.5 Mammalian cells There are two approaches for mammalian host transformation, i.e. by using mammalian cell lines or by producing transgenic animals delivering the genetic material to embryonic cells. Cultivated mammalian cells have become the dominant system for the production of recombinant proteins for clinical applications because of their capacity for proper folding, assembly and post-translational modifications (today about 60–70% of all pharmaceuticals based on recombinant proteins are produced in mammalian cells). Many of these proteins are expressed in immortalized Chinese hamster ovary (CHO cells), but other cell lines, such as those derived from mouse myeloma (NS0), baby hamster kidney (BHK), human embryo kidney (HEK-293) and human retinal cells have gained regulatory approval for recombinant protein production. All mammalian recombinant therapeutics are naturally secreted proteins or have been developed from gene constructs that mediate protein secretion. On the other hand, transgenic animal technology has enabled targeting of recombinant proteins to specific animal tissues or secretions allowing their production in living whole animals (Wurm, 2004). Mammalian expression techniques, however, have several drawbacks. They are time-consuming and technically difficult, with complex nutrient requirement and low product concentration, and are much more difficult to perform on a large scale, all of which means that the end product is extremely expensive. For transgenic animal technology, the process requires thousands of fertilized oocyte microinjections to generate a single transgenic animal and there are also ethical issues to consider (Rai and Padh, 2001, Wurm, 2004).

10.4.6 In vitro systems Cell-free protein synthesis systems are based on extracts from prokaryotic or eukaryotic organisms known for their good protein synthesis capabilities.


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These extracts essentially lack both the genomic material and the cellular boundary system but they contain the cellular components required for transcription and/or translation of genes. Cell-free translation systems, provide a useful alternative to the cellular-expression systems as they can synthesize large proteins with a speed and accuracy that approach those of in vivo translation and they can express proteins that would otherwise interfere with host cell physiology such as protein aggregation in inclusion bodies or proteolysis owing to endogenous proteases which are encountered with the cellular host. It is also a very useful system for isotope labeling of proteins and synthesis of artificial proteins that contain unnatural amino acids (Sawasaki et al., 2002).

10.5

Purification of recombinant proteins

Unless recombinant proteins are being secreted to the extracellular space, the high-level gene expression in the cytosolic compartment usually requires either cell lysis by high-pressure homogenization or sonication or lysis by freeze–thaw procedures with lysozyme to rescue the protein of interest (Hallberg, 2008). Most protein purification techniques are based on the intrinsic differences in the physicochemical properties of proteins, such as solubility, size, charge, hydrophobicity and shape. Commonly used methods that exploit these physicochemical properties of proteins include precipitation, dialysis, electrophoresis and chromatography, the latter being, by far, the most widely used (Chow et al., 2008). Recent reports have included downstream processing strategies for recovering recombinant proteins, by the inclusion of affinity tags for purification purposes. The use of affinity tags enables various proteins to be purified using conventional chromatographic purification techniques, which rarely have adverse affects on biological or biochemical activity. Affinity tags can be typically defined as exogenous amino acid sequences with a high affinity for a specific biological or chemical ligand. His-tags are the most widely used affinity tags and their purification is based on the use of chelated metal ions as affinity ligands. Other groups include tags that use antibody based purification (FLAG, Softag1, Softag3, Streptag II) or antibody purification using protein A affinity chromatography. The choice of a suitable affinity tag depends both on the type of application for the protein of interest and the stage of development of the protein but it is additionally conditioned by the cost of the chromatographic support and the scalability of the process. Once affinity tags have served for extracting the target protein from the crude extract, a tag-removal step is usually developed for obtaining the pure detagged protein. Most of the available methods for affinity tag removal include enzymatic cleavage of the tag followed by specific removal of the



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process enzyme(s) by affinity chromatography. Only in a few specific cases can the affinity tag be removed by harsh chemical treatment (Arnau et al., 2006). Immobilized metal affinity chromatography (IMAC) is the most widely used technique for single-step purification of recombinant proteins containing specific amino acid side chains, particularly those of histidine, cysteine, and tryptophan as fusion tags from bacterial cell lysates (BolanosGarcia and Davies, 2006). Novel fusion tags are being developed for increasing solubility such as NuscA and T7 protein kinase or a small chaperone, Skp; and for aid in protein folding such as the ubiquitin-based tag SUMO. Others, such as inteins, are self-cleavable proteases by using either a thiol reagent or pH and temperature shift to yield intein cleavage and elution of the target protein (Arnau et al., 2006). Elastin-like polymers (ELPs) can be used as fusion proteins for easy purification by two strategies, either engineering the ELP tag into a recombinant protein (direct ELP tagging) or recombinantly expressing or chemically attaching to a capture partner that binds specifically to the target protein in solution (ELP-mediated affinity capture). The ELP imparts its phase transition behavior to the target protein and this can be purified by inverse transition cycling (ITC). This strategy has several advantages over conventional chromatography making it a universal method for soluble recombinant proteins; the ELP acts as a capture mechanism, so no chromatography should be needed and the method does not require a concentration step to recover the final product. In the second approach, ELP-mediated affinity capture does not require enzymatic or chemical cleavage or the ELP tag (Chow et al., 2008).

10.6

Experimental characterization of recombinant proteins

Depending on the individual applications of the obtained biomaterials, and in order to fully determine the functionality after protein purification when necessary, several characterization techniques may be required. Many of them, originally developed for the characterization of proteins, are also useful for the biophysical characterization of repetitive peptides. Biophysical characterization can include: the determination of protein concentration (spectrophotometrically or colorimetrically); the molecular weight of recombinant proteins can be measured by electrophoresis-based methods such as SDS-PAGE, mass spectrometry methods such as matrixassisted laser desorption ionization (MALDI-MS) or electrospray mass spectrometry or analytical ultracentrifugation based on sediment coefficients. Surface plasmon resonance spectroscopy (SPR) provides a way to determine the binding behavior of peptide-based biomaterials, and binding


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constants such as ka, kd, and Kd are calculated based on the binding curves of the peptides. Isothermal titration calorimetry (ITC) can also be used to study the binding thermodynamics of peptides with their binding partners and thermodynamic parameters such as the binding enthalpy, entropy and free energy, and binding constant can be calculated from calorimetric data. (Chow et al., 2008) For ELPs, the inverse temperature transition is often determined by turbidity (optical density at 350 nm) as a function of temperature or by differential scanning calorimetry (DSC) (Reguera et al., 2003). Nuclear magnetic resonance (NMR) and x-ray crystallography are the methods of choice for obtaining comprehensive structural information for proteins. In addition, protein folding can be studied using circular dichroism (CD), which defines the unfolding and folding transitions of peptides. Differential scanning calorimetry (DSC) can be used to elucidate the folding and refolding properties of peptides during cooling and heating. Dynamic light scattering (DLS) also named photon correlation spectroscopy, can be used to determine the hydrodynamic radius (Rh) of polypeptides in aqueous solution, especially useful for examining the formation of self-assembled polypeptide micelles (Chow et al., 2008). Fluorescence spectroscopy combined with previous techniques can provide additional information about the structural state of the proteins (Woestenenk et al., 2003). Rheological properties: the relationship between the molecular structure of a polypeptide and is rheological properties can be estimated by measuring G′ elastic (storage) modulus, which represents the solid-like component of a material, G″, viscous (loss) modulus, that represents the liquid-like component, η*, dynamic (complex) viscosity, and δ, loss angle, that is a measure of the dissipation of energy inherent in the material and is a useful parameter for quantifying the viscoelasticity of a material, as a function of strain, frequency, temperature, time and other parameters (Chow et al., 2008). The sol–gel transition for a given polymer solution or gelation point can be easily detected by a measurement of G′ and G″ and is defined as the crossover between G′ and G″ as a function of the previous parameters cited (Nagapudi et al., 2005). The mechanical properties of fiber proteins are determined by stretching them at a particular strain rate d(l/l0)/dt, and measuring the force required to extend the fiber a certain length, defined as dl. The strain (ε) represents the normalized deformation (elasticity), which is defined as the ratio of change in length (dl) to the initial length (l0) or (dl/dl0). The stress (σ) is defined as the force (F) divided by the cross-sectional area (A) of the fiber (σ = F/A). Young’s modulus (E), which can be obtained from the slope of the stress–strain curve, is a measure of the stiffness of the fiber. If the fiber breaks during extension, the area under the curve is a measure of the toughness of the fiber, a parameter that indicates the amount of energy absorbed by the fiber. During extension, stress–strain profiles can often display



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sudden slope changes, indicative of major structural transitions of the material that are known as ‘yield points’ (Hu et al., 2006). When post-translational modifications are essential for protein function, such as protein activity, stability, localization, turn-over, and protein interactions, the study of phosphorylation, glycosylation and sulfation as well as many other modifications is extremely important. Two-dimensional gels, 32P labeling, western-blot, immunological methods, protein chips and other generic strategies based mainly on affinity methods, depending on the event that it is studied, can be applied for this purpose (Pandey and Mann, 2000).

10.7

Examples and applications of recombinant protein fibers (silk, elastin, collagen, resilin)

Elastomeric proteins have been identified in a range of biological systems. The mechanical and biochemical properties of some have been extensively studied, especially elastin, abductin and flagelliform spider silks and their potential as biomaterials for industrial and biomedical applications are well documented (Lyons et al., 2007). From all the biopolymers that commonly occur as structural elements of biological systems some protein-based biopolymers have been employed to mimic and/or improve the structural and mechanical properties found in their natural sources, in order to be useful in the textile industry. Biotechnological production of fibers could allow the preparation of a new generation of high-performance fibers having even more built-in properties for a variety of textiles-related applications. With traditional fibers in the textile industry, it is known that only when continuous single nanofibers or uniaxial fiber bundles are obtained, can they be employed in numerous applications. Thus, one of the main targets in obtaining recombinant protein-based polymers for applications in textile industry is to achieve aligned nanofibers. Several fabrication techniques such as electrospinning (Huang et al., 2003, Li and Xia, 2004), phase separation (Yang et al., 2004) and self-assembly (Hartgerink et al., 2001) amongst others have been employed to produce polymer nanofibers for different applications. Most of these techniques are time-consuming or have prerequisites regarding the viscoelasticity or the cohesivity of the material employed. Therefore, the most popular technique used is electrospinning, which provides a convenient method for many materials and produces continuous nanofibers that can be developed for large-scale production (Nagapudi et al., 2005). Electrospinning utilizes a high-voltage source to inject charge of a certain polarity into a polymer solution, which then dries to leave a polymer nanofiber mesh. The fibers produced by this process usually have diameters on the order of a few micrometers down to less than a hundred nanometers and their structural properties depend on processing parameters such as


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Advances in textile biotechnology Medical implants Molecular filtrations

Drug release

Scaffolds for tissue engineering

Applications of polymer nanofibers

Dressings for wound healing

Dental applications

Biosensors Preservation of biological agents

10.3 Potential applications of polymer nanofibers.

polymer concentration and viscosity, flow rate, and applied voltage, amongst others (Sill and von Recum, 2008). The ability to vary fiber size in the nanometer range has opened up the possibility of using polymer nanofibers for biomedical and nanotechnological applications. Recent uses of these materials have included tissue engineering, medical implants, biosensors and drug release amongst others (Fig. 10.3) (Huang et al., 2003). Thus, some examples of fibers made from proteins such as fibrinogen (Wnek et al., 2003), gelatin (Li et al., 2005), collagen-elastin mixtures (Buttafoco et al., 2006), or silk-like proteins (Jin et al., 2004) have been obtained. Some of the most important natural protein-based fibers with potential for replacing synthetic fibers owing to their outstanding properties in terms of strength, elasticity and the attempts for their biotechnological production are described in the following subsections.

10.7.1 Silks Silks are protein-based fibers produced in specialized glands of arthropods for a variety of task-specific applications. Of all natural silk-producing animals, mulberry silkworms (Bombix mori) are of the most economic importance. B. mori silk fibers have been used for the production of textile goods for centuries owing to their characteristic luster, moisture absorbance and strength. Indeed, the variety of reactive amino-acid residues displayed on the backbone of this kind of silk protein has mainly led the modification of their affinity for disperse dyes and water and their wrinkle recovery (Hardy et al., 2008). Orb-web spinning spiders are able to produce a variety of task-specific silks, some of which have mechanical properties unmatched in the natural world and similar to the very best synthetic fibers produced by modern © Woodhead Publishing Limited, 2010


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technology (Gosline et al., 1999). Spider silks are excellent biomaterials, being lightweight and exceptionally strong, and having elastic properties, including high impact resistance. The molecular structure of spider silks consists of regions of protein crystals separated by less organized protein chains. The primary structural modules give rise to diverse secondary structures that, in their turn, direct functions of different silks (Kluge et al., 2008) (Table 10.1). Biomaterials based on silks have applications ranging from medical (micro-sutures, artificial ligaments, coatings), military (body armor, lightweight gear) to textiles fields. Although variable in their primary structure, silk proteins in general reveal a high amphiphilicity and a highly repetitive nature. The key properties of web silks involve a balance between strength and extensibility, giving a high degree of toughness and internal molecular friction. Spider silks are able to absorb a large quantity of kinetic energy with a minimal volume of silk. The viscoelastic nature of the spider silks transforms this energy into heat instead of being available through elastic recoil (Gosline et al., 1999). The combination of strength and elasticity is judged to be similar or even superior to that of synthetic high-tech fibers made of polyamide or polyester. Indeed, compared with Kevlar, which must be spun from almost boiling sulfuric acid, the spider silks are produced in aqueous solutions at ambient temperature. Spider silks exhibit mechanical properties similar to the best synthetic fibers produced by modern technology, commonly employed to transmit and support tensile forces (Gührs et al., 2000) (Table 10.2). All silks contain protein crystals, and the majority of these silks contain β-pleated sheet crystals that form from tandem repeated amino acid sequences rich in small amino acid residues. The β-sheets contribute to the high tensile strength of silk fibers and form through natural physical crosslinking of amino-acid sequences, which on spider and silkworm silk consist of multiple repeats of mainly alanine: poly(A)n, glycine–alanine (GA)n or glycine–alanine–serine (GAGAGS). The non-crystalline regions of silk are commonly made up of: β-spirals similar to a β-turn composed of GPGXX repeats (where X is mostly glutamine) and helical structures composed of GGX. These semi-amorphous regions provide silk with elasticity. In addition to elastic fiber and semi-amorphous regions, non-repetitive regions are present at the amino- and carboxyl termini of the proteins and they probably play a role in the controlled assembly of silk proteins (Gosline et al., 1999, Kluge et al., 2008). Although spiders produce seven different types of silks, only the major ampullate gland silk and flagelliform silk have been produced as heterologous proteins using genetic engineering techniques. Heterologous expression of spider-silk proteins has been achieved, as well as the formation of new materials from spider-silk proteins derived recombinant DNA. © Woodhead Publishing Limited, 2010


Web reinforcement and temporary capture silk

Capture spiral

Minor ampullate (MI)

Flagelliform

Flag

MiSp2

(GPGXX)n, GGX and spacer

GGX, (GA)n, poly(A) blocks and spacer GGX, (GA)n, poly(A) blocks and spacer

MiSp1

Poly(A) blocks, (GA)n and GGX

GGX, GPGXX, (GA)n and poly(A) blocks

MaSp1

Web frame, safety line

Major ampullate dragline (MA)

Ensemble repeats

MaSp2

Core fiber proteins

Location

Silk

• Repetitive regions consist mainly of alanine and glycine, with lower levels of tyrosine, glutamine and arginine • There is no proline present in MI silks • MI silk has similar mechanical properties to MA silk but has lower elasticity

• Dominant α-helical nature with reduced β-sheet structure • Conformations of the alanine residues are more heterogeneous in nature with a large fraction present in a non-β-sheet conformation • Repetitive regions are interrupted by a highly conserved serine-rich 130 amino-acid non-repetitive sequence that acts as a spacer • Concatenated repeats of the pentapeptide GPGXX, which can appear up to 63 times in tandem arrays, have been proposed to form a β-spiral structure which provides fiber elasticity/flexibility

• The tripeptide GGX occurs tenfold fewer times than GPGXX • Flag silks are the most extensible, but have lower tensile strength relative to dragline silk

• X is typically tyrosine, leucine or glutamine • MaSp1 levels are higher relative to MaSp2 • Repetitive structural modules are proline free Iterated sequences contain 15% proline

Comments

• Alanine-rich blocks form β-sheet stacks that are responsible for the high tensile strength of MA silks • Blocks of GGX presumably form 310-helices • Blocks of GPGXX form β-turn spirals imparting elasticity/ flexibility to the proteins

Secondary structures

Table 10.1 Spider silks, natural functions and common modules and secondary structures within silk proteins (adapted from Hardy et al., 2008, Hu et al., 2006)


Large-diameter egg case silk fiber

Glue coating for capture spiral Attachment disc and joining fiber

Tubuliform

Aggregate

Pyriform

Wrapping silk, small-diameter egg case fiber

Aciniform

Unknown

Unknown

The precise molecular identities of the proteins involved remain to be determined

Short poly(A) blocks and GA repeats Short poly(A) blocks and GA repeats

ECP-1

ECP-2

Poly(S), GX, AAQAASAA, AAAQA, AASQAA and SQn

Poly(S) blocks and GGX

TuSp1

AcSp1

These sequences are derived from the C-termini and do not reflect ensemble repeat unit

• Large ensemble repeats that are 184 amino acids in length, rich in serine but low in glycine • Blocks of alanine and GGX yield protein structures rich in β-sheet stacks

GGX domains are notably different from other types of silk proteins

• Blocks of GGX assumed to form 310-helices • The spacer region is a highly conserved and glycine-poor sequence of 28 amino acids

These two silks contain very polar amino acids, which include Ser, Asp, Glu, Thr, Lys and Arg

ECPs may function as intermolecular crosslinkers, with potential structural roles in the egg case fiber

• Low percentages of glycine and alanine • Low tensile strength relative to drag line silks but higher extensibility and toughness


Dragline spider silk Flagelliform spider silk Bombix mori silk Elastin Mussel byssus (distal part) Tendon collagen Resilin Rubber Nylon Kevlar 49 fiber

Material

1.5 0.002 0.001 5 130

10 0.003 7 0.001

Stiffness, Einit (slope of stress– strain curve) (GPa) 1.1 0.5 0.6 0.002 0.075 0.15 0.003 0.05 0.95 3.6

Strength, σmax (yield point) (GPa) 30 270 18 150 109 12 190 850 18 2.7

Elasticity, εmax (elongation at break) (%) 160 150 70 2 45 7.5 4 100 80 50

Toughness (work to fracture) (MJ m−3)

7 6

10

65 65

Hysteresis (%)

Table 10.2 Mechanical properties of fibrous materials (adapted from Gosline et al., 1999, Vendrely and Scheibel, 2007)


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Biotechnological strategies concerning silk-protein production have to take into account several aspects: silk genes are large (up to 15 Mbp) and the use of bacterial host, such as E. coli, as industrial production host is hampered by the fact that the size of expressible genes is limited and bacterial host have a distinct codon usage different to that of spiders. Successful expression of spider silks have been achieved in bovine mammary epithelial and hamster kidney cells, in the milk of transgenic goats and in baculovirus expression systems with their genome modified with silk genes and selectively infecting insect cells. In addition to bacterial hosts, the yeast Pichia pastoris and plants are currently under investigation as a cheap alternative to bacterial host for silk protein production like tobacco, potato and Arabidopsis thaliana (Vendrely and Scheibel, 2007). Silk proteins have been shown to solubilize in water, organic solvents and ionic liquids, indicating the versatile options available, and they can then be processed into new materials including fibers, films, gels, porous sponges and other related systems, thus making spider silks potentially promising for future materials. Silk-like proteins (SLP) based on the repeating motif (GAGAGS) produce crystalline structures similar to the β-sheet structures of native silk proteins (Cappello et al., 1990). Several methods to increase the water solubility have been developed such as the addition of sterical crystallization triggers such as methionine residues or charged phosphate groups (Winkler et al., 1999), which interrupted the β-sheet formation preventing insolubility. Subsequent modifications of spider silk sequences have been performed in an attempt to control the mechanical properties showing that the nature of the fifth and tenth amino acid (X) in the (GPGGX)n elastic motif and temperature have a large impact on the selforganization process and in the level of elasticity of the proteins in the film or fibers (Teule et al., 2007). Recently, mechanical properties of a recombinant miniature spidroin have been increased by genetic engineering with disulfides increasing the stiffness and tensile strength without changes in the ability to form fibers, or in fiber morphology (Grip et al., 2009). Silkworm fibroin silk has been functionalized by coupling enzymes such as horseradish peroxidase, cell-binding domains such as Arg–Gly–Asp (RGD) peptides and cell signaling factors such as parathyroid hormone (PTH) and bone morphogenetic protein-2 (BMP-2) to improve cell interactions and function or to form gradients on silk material surfaces (Karageorgiou et al., 2004, Sofia et al., 2001, Vepari and Kaplan, 2006).

10.7.2 Collagens Collagens represent a family of related structures widely distributed in nature and commonly used in drug delivery and pharmaceutical applications as well as in many medical devices. In humans, collagens are present


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in most connective tissues, blood vessels and in basement membranes in all organs where they participate in the cellular matrix structure and function. Animals such as cattle, pigs and horses as well as marine animals such as fish, sponges and the mussel byssus can being considered as sources of collagen (Olsen et al., 2003). Collagens are composed of three polypeptide chains (α-chains) with a characteristic sequence of (Gly–X–Y), where X and Y are usually proline and 4-hydroxyproline, respectively. The three α-chains twist around a common axis into a right-handed triple helix and the amino-acid sequences of three α chains can be the same or different. To date, 27 types of human collagen have been identified, differing in primary sequence chemistry. Of them, depending on morphology and function, collagens are classified into eight groups, fibril-forming collagens (types I, II, III) being the most extensively documented (Huang et al., 2007). The key to producing thermally stable collagens, with a melting temperature 39–40 °C (the normal melting temperature of human collagen) and with a proper triple helical conformation, in recombinant expression systems relies on the ability to effect appropiate post-translational processing of the recombinant collagen proteins. Recombinant collagens have been produced by transfected mammalian cells, insect cells, yeast, E. coli, transgenic tobacco, mice and silkworms. Of them, only mammalian cells transfected with a collagen gene and not with the enzyme prolyl-4-hydroxylase (PH4) genes expressed hydroxylated full-length collagens. In the other expression systems, overexpression of PH4 was required for the production of fully hydroxylated collagen (Olsen et al., 2003). Byssus threads of marine mussels are interesting elastomeric fibers with a great capacity for absorbing and dissipating energy that have been reported to have three distinct collagenous proteins in the thread, preCol P, preCol D and preCol NG, the last being a central collagen domain (stiff segment), and the first two being flanking domains having distinctive structural properties: preCol P has elastin-like flanking domains (soft segment) and preCol D has stiffer silk fibroin-like domains (hard segment) (Vaccaro and Waite, 2001). In common with other protein elastomers such as elastin, resiline and abductine, byssus threads are quite tough. Their Young’s modulus is low but the extensibility can be as high as 200%. Byssus threads have been reported where up to 70% of the total absorbed energy can be dissipated. Byssal threads are at least five times more extensible and five times tougher than an Achilles tendon. Indeed, unlike the tendon, byssal threads have a non-periodic microstructure and shrinkage and melting temperatures in excess of 90 °C (Qin et al., 1997). Purified collagens are capable of undergoing spontaneous alignment to form fibrils that have defined features characteristic of collagen fibers



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formed in vivo (Williams et al., 1978). The recombinant collagen expressed in yeast formed these fibrillar structures at neutral pH in phosphate buffer (Huang et al., 2001a, 2001b) and were the first to be electrospun as collagen scaffolds for wound dressing. Shortly thereafter, spun non-crosslinked collagen fibers exhibited a concentration dependence on the final fiber diameters produced, and showed suitable biological properties in preliminary in vitro tests (Matthews et al., 2002). Recently, collagen fibers have been produced without either organic solvents or blend formation with any synthetic and natural polymers, to provide new electrospinning experimental conditions to obtain biomimetic collagen self-assembled nanofibers (Foltran et al., 2008).

10.7.3 Elastin Native elastin is one of the most abundant ECM proteins. Working in partnership with collagen, elastin allows the body organs to stretch and relax. Thus, while collagen provides rigidity, elastin allows the recoil of elastic tissues. In its natural state, elastin is an insoluble protein owing to the presence of crosslinks. However, its soluble forms, such as tropoelastin (Mithieux et al., 2004) and α-elastin (Annabi et al., 2009), are frequently used as biomaterials. Natural elastin undergoes a self-aggregation process in its natural environment, leading to the formation of nanofibrils from a water-soluble precursor called tropoelastin (Urry, 2005). This ability resides in certain relatively short amino acid sequences, which are known to coacervate and form as fibrillar aggregates with a high degree of β-structure (Yang et al., 2002). Fibrous matrices as scaffolds for tissue engineering have been formed by α-elastin and tropoelastin fibers. The electrospinning process was optimized to provide uniform fibers in the range of microns showing, especially with tropoelastin, ‘quasi-elastic’ wave-like patterns at increased solution delivery rates (Li et al., 2005). Elastin-like polymers (ELPs) are a promising model for biocompatible protein-based polymers. The basic structure of ELPs is a repeat sequence found in the mammalian elastic protein elastin, or a modification thereof (Miao et al., 2003). Some of their main characteristics are derived from those of the natural protein. For example, the crosslinked matrices of these polymers retain most of the striking mechanical properties of elastin (Di Zio and Tirrell, 2003), which becomes important when this behavior is accompanied by other interesting properties, such as biocompatibility (Urry et al., 1991), stimuli-responsive behavior, and the ability to self-assemble. These properties are based on a molecular transition of the polymer chains called the inverse temperature transition (ITT). This transition is the key


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to the development of new peptide-based polymers as molecular devices and materials. The expansion of molecular biology has allowed the design of complex bioengineered ELPs as well-defined polymers (Girotti et al., 2004, Martino et al., 2002, Meyer and Chilkoti, 2002, Welsh and Tirrell, 2000). The most well known members within the ELP family are based on the pentapeptide VPGVG (or its permutations), and a wide variety of polymers with the general formula (VPGXG), where X represents any natural amino acid except proline, have been (bio)synthesized (Gowda et al., 1994, Martino et al., 2002, Urry, 2005). All polymers with this general formula that have been reported display functional properties such as acute stimuli-responsive behavior. The first elastin-mimetic protein fibers were produced from a genetically engineered ELP (Huang et al., 2000). Various morphological patterns, such as beaded fibers, thin filaments, or broad fibres with a ribbon-like appearance, were obtained by varying the solution concentration. Chemically crosslinked fibers were also prepared from two different ELPs containing glutamic acid and lysine residues, respectively, confirming their reversible anisotropy in swelling behavior and showing a fast response time upon temperature changes (Lee et al., 2001). In another study, electrospinning of ELP copolymer solutions in water and 2,2,2-trifluoroethanol provides fibers ranging from 100 nm to 3 μm showing the importance of the solvent during the fiber spinning (Nagapudi et al., 2005). Currently, the most often used method for laboratory scale production of ELPs is E. coli shake cultures that give typical yields of approximately 300 mg l−1. Other hosts that have been used for expression of ELPs or natural elastin sequences in which the ELP is produced and recovered from inside the cells are tobacco cell cultures, cultures of Aspergillus nidulands, transgenic tobacco plants and Saccharomyces cerevisiae. Finally, the yeast Pichia pastoris has been succefully used for secreted expression of ELPs (Schipperus et al., 2009). In biosynthesis using recombinant DNA techniques to combine polypeptide sequences derived from silkworm silk and mammalian elastin, silk-elastin-like proteins (SELPs) allow diverse physical, mechanical, and biological features to be abtained (Ferrari and Capello, 1997). Recently, genetically engineered silk-elastin-like protein copolymer (SELP) containing tandemly repeated amino acid sequence blocks from silk, GAGAGS, and elastin, GVGVP, was spun into clean microdiameter fibers using a wetspinning technique. Hydrated SELP-47K fibers were mechanically weak but displayed large strains to failure. This was modified by chemical crosslinking with glutaraldehyde enhancing their strength until 20 MPa and deformability of 200–700% showing potential applications in tissue engineering (Qiu et al., 2009).



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10.7.4 Resilin Resilin is a structural protein with elastomeric properties, which is represented in specialized regions of the cuticle of most insects, where it is known to play important roles in insect flight, the jumping mechanisms of fleas, and vocalization in cicadas (Kim et al., 2007). Naturally crosslinked resilin exhibits two outstanding material properties: high resilience (recovery after deformation), of 92%, resilin can be stretched three or four times its resting length before breaking, and can return to its original resting shape with very little deformation, and a very high fatigue lifetime. This performance is far superior to that of a known low resilience rubber, chlorobutyl rubber (56%), and even to that of high-resilience polybutadiene rubber (80%). Stress– strain data show it to have a modulus at 100% of about 2.5 kPa, considerably below that of typical unfilled synthetic elastomers and native elastin (Elvin et al., 2005). To date, partly because of difficulties in obtaining large amounts of pure resilin from natural sources and a lack of sequence data, structural studies of resilin have been limited, although early studies of resilin including electron microscopy and x-ray diffraction suggest that resilin is an unstructured amorphous protein matrix, formed by crosslinking of tyrosine residues as di- and trityrosine complexes (Lyons et al., 2007). The physical properties of the resilin polymer probably result from the three-dimensional amorphous and unstructured nature of the crosslinked protein matrix and the role of water as plasticizer, because dehydrated crosslinked resilin is very glassy and brittle, and in a partially dry environment (70% relative humidity) it is leathery with poor resilience. It is possible to store the dehydrated material for extended periods and then to recover its normal resilience on rehydration (Elvin et al., 2005). As for many elastomeric proteins, resilin contains distinct repetitive domains that appear to confer elastic properties to the protein although limited sequence homology between species has been reported. The conserved tetrapeptide YGAP seems to be the critical motif in conferring heat resistance and hydrophobic properties to resilin and related properties. Attempts to reproduce the desirable mechanical properties of resilin in synthetic biomaterials have generated recombinant proteins encoding multiple copies of consensus polypeptides based on repetitive domains within resilin-like genes, E. coli being the main host strain employed for overexpression. It is likely that the resilin-like proteins have a self-associating propensity, a property that similarly has been identified in elastin. It was shown that concentrated solutions of all resilin-like recombinant proteins tested have a tendency to form a concentrated protein lower phase at 4 °C, whereas the upper phase appears as an opaque solution at this temperature, suggesting that it has a micellar structure; heating of the cloudy solution


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makes it clear (Lyons et al., 2007). Further investigations of structural and functional properties of resilin are necessary for the development of novel highly resilient biomaterials.

10.8

Advances and limitations in the biosynthetic production of recombinant proteins

The increasing power of biotechnology opens new perspectives either to access rare and limited biological materials in sufficient amounts or to produce new materials that are based on natural models. Applications of recombinant protein fibrous materials in commodity materials, textiles, high-performance composites, and durable and tough materials in general, require the success of robust and low cost expression systems (Kluge et al., 2008). The main advantage of protein biosynthesis lies in the ability to directly and consistently produce high-molecular-weight polypeptides of exact aminoacid sequence with high fidelity, and, consequently, to have control over the folded structure functions and assemblies of the resulting materials. This, in turn, offers an enhanced safety profile, greater reproducibility and quality, and the ability of these materials to be tailored to enhance product performance (Gührs et al., 2000, McMillan et al., 1999, Olsen et al., 2003). Genetically tailorable composition and sequence to moderate specific features such as molecular weight, crystalinity and solubility, and the possibility of subsequent chemical modification confers a significant advantage compared with natural common proteins as they can be modified to the design of polymeric materials with improved physicochemical properties such as better defined biocompatibility, bio-recognition, biodegradation and stimuli sensitivity (Rodríguez-Cabello et al., 2006, Wong Po Foo and Kaplan, 2002). Protein-based polymers are environmentally clean over their entire life cycle from production to disposal; they can be produced from renewable sources and are biodegradable. The cost associated with the protein polymer synthesis depends on the cost of the materials used in the synthesis and scale of the synthesis. Indeed, the synthesis cost of these biosynthetic methods for producing a well defined and compositionally complex protein polymer does not increase with the length and complexity of the polymer after the initial construction of the target gene. This is in contrast to chemically synthesized polymers, in which the cost and complications of largescale synthesis often scale with increased architectural control and polymer molecular weight (Kiick, 2007). Most of the limitations and disadvantages of the biosynthetic process have been mentioned previously in connection with the recombinant expression systems: low accumulation levels of recombinant proteins, lack



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of sufficient information on post-translational events (proper folding, glycosylation, phosphorylation, formation of disulfide bridges) and downstream processing being the most relevant and commonly found. Choosing the best expression system mainly requires evaluating the options in terms of yields, glycosylation, proper folding of the protein of interest and economics of scale-up, but also of its chemical properties such as size, isoelectric point, stability at different pHs and temperatures and proteolitic resistance (Rai and Padh, 2001). Heterologous expression of proteins has to take into account other aspects such as gene size, because the size of expressible genes is limited, and the host codon usage.

10.9

Future trends

The textile industry has broadly benefited from the rapid development of strategic coalitions between the arts of protein engineering and heterologous protein expression and those of fermentation and bioprocessing technologies that have led to the possibility of ‘tailoring’ organisms in order to optimize the production of established or novel materials and of transferring genetic material from one organism to other. The advent of recombinant DNA technology and the ability to predictably manipulate the properties of the self-assembly of autonomous units either scientifically based or derived from functional biological macromolecules and elements of the biosynthetic process, in order to produce refined molecules with a degree of precision that was not possible using conventional synthetic technology, have provided an extremely rich and useful resource for the design of highly functional recombinant protein-based fibers with multiple applications. Designing new fibers with particular properties by using biotechnological approaches or transferring certain advantageous textile properties into micro-organisms, where they can be more readily reproduced by bulk fermentation, enable smart materials to be developed with biomimetic or stimuli-responsive properties for ‘intelligent clothes’. Because the potential of this genetic engineering technique has not yet been fully exploited, more complex and sophisticated materials and devices based on the mechanical properties of elastomeric proteins are expected in the near future. Taking into account that genetically engineering protein-based polymers are produced without the use of any oil-derived raw material and in an environmentally clean fabrication procedure, the process can be regarded as a long-term solution to environmental problems usually associated with many enzymatic transformation processes of textile processing and after care.


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10.10 Acknowledgments We acknowledge financial support from the MICINN (projects MAT 2007– 66275-C02–01, MAT 2009-14195-C03-03 and PSE-300100-2006-1), the JCyL (projects VA034A09 and VA030A08), the CIBER-BBN (project CB06-010003), the JCyL and the Instituto de Salud Carlos III under the ‘Network Center of Regenerative Medicine and Cellular Therapy of Castilla and León’ and the COST Action 868.

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11 Enzymatic functionalization of cellulosic fibres for textile and other applications: xyloglucan as a molecular anchor H. BRU M E R, Royal Institute of Technology (KTH), Sweden

Abstract: The inherent recalcitrance of paracrystalline cellulose, which restricts the performance and applications of cellulosic biofibres, has been circumvented by a biomimetic approach that avoids the limitations inherent in direct chemical methods for cellulose modification. The unique ability of the plant enzyme xyloglucan endo-transglycosylase (XET) to produce chemo-enzymatically functionalized xyloglucan (XG) polysaccharides was harnessed together with the inherently strong interaction of XG with cellulose to surface-anchor the XG derivatives. An overview of the use of XG-XET technology in the functionalization of diverse cellulosic substrates is presented. Specific examples are given to highlight the potential of the system for future applications in textiles, pulp and paper, and biomedicine. Key words: xyloglucan, xyloglucan endo-transglycosylase, cellulose, surface modification, biocomposites, biomimetics, functionalization.

11.1

Introduction

Increasing demand on global resources for the production of materials has long sustained interest in diverse applications of biofibres, ranging from paper and packaging to construction materials and textiles. Although synthetic, petrochemical-derived fibres sometimes have distinct performance advantages over their natural counterparts, wood pulp, cotton, and bast fibres are often desired, or even preferred, in many industrial and consumer products. In some cases, this is trivially because of economic considerations; (ligno)cellulosic biofibres are often cost-competitive with respect to synthetic alternatives. On the other hand, biofibres possess unique material and tactile properties that are often favoured by consumers, in addition to the general perception that these plant-derived materials are both natural and sustainable. The physical/chemical surface modification of cellulosic fibres is very often performed to improve functional properties for specific applications 266 © Woodhead Publishing Limited, 2010

Enzymatic functionalization of cellulosic fibres for textiles

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following liberation from the plant tissue source. This may involve selective removal of components, such as associated lignin, hemicelluloses and extractives, by pulping and bleaching chemicals to improve the optical properties of the final paper or textile product. Realization of the full potential of cellulosic fibres, however, requires the addition of chemistry to the fibre surface in many applications. Some common examples include the application of strength-building compounds, retention aids, hydrophobic molecules, and inorganic fillers during papermaking, as well as the binding of dyes, optical brighteners, and sizing agents to textile fibres and fabrics. Clearly, the need for surface functionalization extends to high performance products based on cellulose fibres, such as ‘smart’ textiles, electronic paper, cellulosic biocomposite materials, and biomedical devices. Indeed, a wide range of classical chemical methods have been developed to modify cellulosic fibre surfaces, either through physical adsorption or direct covalent attachment of molecules. Although powerful, the covalent derivatization of cellulose has certain limitations. Cellulose polysaccharide chains exist as insoluble, paracrystalline aggregates, which are characterized by low reactivity of the tightly hydrogen-bonded hydroxyl groups. Further, extensive reaction of these same hydroxyl groups leads to disruption of cellulose crystallinity, chain degradation, and, ultimately, a loss of fibre strength (Sassi and Chanzy, 1995; Sassi et al., 2000; Klemm et al., 2005). In some cases, reactions may require non-aqueous media, thus necessitating a solvent exchange or drying step, which may result in altered cellulose morphology or be technically impractical on a large scale. These concerns are particularly relevant for cellulosic hydrogels, such as bacterial cellulose or microfibrillated cellulose/nanocellulose suspensions, which are emerging as promising new biomaterial templates (Bodin et al., 2006, 2007a, 2007b; Henriksson et al., 2008; Iwamoto et al., 2005; Nakagaito and Yano, 2005; Yano et al., 2005). With the potential of cellulosic fibre functionalization in focus, a number of years ago our laboratory began work on a biomimetic approach to circumvent the inherent challenges of direct chemical approaches. In this approach, the strong interaction of cellulose with the cell-wall-matrix polysaccharide xyloglucan (XG) was harnessed, together with the unique catalytic properties of an endogenous plant transglycosylating enzyme, xyloglucan endo-transglycosylase (XET). The subsequent sections will provide an overview of some of the basic biochemistry of XG and XET in the context of the plant cell wall, and highlight how this system has been appropriated, in conjunction with organic chemistry, to install a range of functional groups on cellulose fibres. The practical application of this system has a strong foundation in the historical use of native XG as a sizing agent in textile and paper industries.


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11.2

Xyloglucans: a family of functional plant polysaccharides

11.2.1 Xyloglucan in the vegetative cell wall The XGs comprise a plant-specific family of polysaccharides based on a highly xylose-substituted β(1 → 4) glucan (cellulose) backbone (Carpita and McCann, 2000; Hoffman et al., 2005). Fuco-galacto-xyloglucans (Fig. 11.1) are widely distributed among land plants (excepting many grass species), in which they act as the primary crosslinking glycans of cellulose microfibrils in the primary cell wall (Carpita and Gibeaut, 1993; Carpita and McCann, 2000; Popper, 2008). As such, XGs constitute up to one-quarter of the dry weight of dicot cell walls (Busato et al., 2001 and references therein), and are intimately associated with cellulose by adsorption onto and entrapment within the paracrystalline structure (Pauly et al., 1999a). Indeed, XGs have a demonstrably tight and specific binding to cellulosic substrates, which is unique among polysaccharides (Zykwinska et al., 2005, 2008). Moreover, this binding is effectively irreversible over a broad pH range (de Lima and Buckeridge, 2001; Lima et al., 2004); strongly basic solutions (e.g. 2 M NaOH) are required to release cellulose-bound XGs, presumably through partial ionization of the polysaccharide chains (Edwards et al., 1985, 1986). This remarkable, inherent affinity of XGs for cellulose forms the basis for their use in biofibre modification. Although an exact structural explanation is still lacking, elucidation of the molecular details of the strong cellulose–XG interaction has been an area of continued interest since the mid-1970s (Hanus and Mazeau, 2006, and references therein; Valent and Albersheim, 1974). The seminal study of Vincken et al. (1995) is particularly illuminating in the context of the practical utilization of XG as a cellulose modification reagent, and the subsequent development of the XET/XGbased technology. This study was the first to demonstrate that quantitative binding of XG to microcrystalline cellulose occurs when the polysaccharide is comprised of four or more xyloglucan oligosaccharide (XGO) repeats, i.e. when the XG chain has a backbone of 16 or more Glc residues (n > 3, Fig. 11.1). Subsequent studies have confirmed and extended these results to indicate that the binding of XG to microcrystalline cellulose is largely independent of pH over the range 2–8 and temperature of 5–60 °C (de Lima and Buckeridge, 2001; Lima et al., 2004).

11.2.2 Xyloglucan in seeds In addition to their structural role in the vegetative cell wall, XGs have been recruited as the primary storage carbohydrates in the seeds of certain



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L-Fucpa1

(Galpb1)0–1

Xylpa1 6 4Glcpb1

2 (Galpb1)0–1

2 Xylpa1 6 4Glcpb1

2 Xylpa1 6 4Glcpb1

4Glcpb1 n

11.1 A common structure of XGs based on a Glc4Xyl3 core structure. Variable regioselective addition of galactosyl residues on one or two xylosyl branches further extends the core. Extension of Gal with an α-L-fucosyl residue (boxed) distinguishes XGs found in dicot primary cell walls from those found in seeds, cf. the molecular structure shown in Fig. 11.4.

species, including the tamarind (Tamarindus indica) and jatoba (Hymenaea courbaril) trees, as well as the ornamental nasturtium plant (Tropaeolum majus) (Buckeridge et al., 1997; Edwards et al., 1985; York et al., 1993). In general, the seed XGs have the same basic oligosaccharide repeat structures as those in the primary plant cell wall, but are distinguished by a lack of terminal α(1 → 2)-linked l-fucosyl residues on the β-d-Galp-(1 → 2)-α-dXylp(1 → 6)-d-Glcp sidechains (Buckeridge et al., 1997; Hoffman et al., 2005; York et al., 1993) (Fig. 11.1). The importance of fucosyl residues in the cellulose–XG interaction in vitro binding is unclear (Chambat et al., 2005; Whitney et al., 2006; Zykwinska et al., 2005), although they appear to be of little biological importance in vivo: mutant Arabidopsis plants that lack XG fucosylation have normal growth and cell wall strength (Reiter, 2002). However, selective enzymatic hydrolysis has indicated that the pendant galactose residues are important in maintaining XG solubility (Shirakawa et al., 1998; Whitney et al., 2006). The interested reader is referred to the review by Zhou et al. (2007) for a more detailed overview of XG structure– function relationships, including the effects on rheology and interactions with cellulose. Owing to their availability in large quantities, tree seeds, and especially tamarind seed kernels, comprise the most relevant sources of XG for fundamental studies and industrial applications. ‘Tamarind kernel powder (TKP)’ is a large-scale (ca. 100 000 tonne y−1) agricultural co-product from the production of tamarind fruit pulp for the food industry (Rao and Srivastava, 1973; Shankaracharya, 1998). De-oiled TKP contains ca. 60% XG by mass, which can be readily extracted by aqueous solutions to produce various crude preparations known as ‘tamarind gum’ (Gerard, 1980; Kumar


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and Bhattacharya, 2008; Rao and Srivastava, 1973). Repeated alcohol precipitation and dissolution, sometimes employing chemical or enzymatic treatment steps (Rao and Srivastava, 1973), can be used to prepare highpurity fractions of XG polysaccharide for diverse applications, including thickeners, stabilizers, or gelling agents in food and pharmaceutical products (Coviello et al., 2007; Gerard, 1980; Kumar and Bhattacharya, 2008; Yamatoya and Shirakawa, 2003).

11.3

Technical uses of native xyloglucans in cellulose modification

11.3.1 Applications in textiles and papermaking XG, extracted from tamarind kernel powder (TKP) in various purities, has long been used to modify cellulosic fibres from cotton, jute, and wood, especially in India, where the tree is indigenous (Gerard, 1980; Kumar and Bhattacharya, 2008; Rao and Srivastava, 1973; Shankaracharya, 1998). Sizing cotton, jute, and spun viscose with TKP improves yarn strength during weaving and imparts smoothness and stiffness to fabrics (Gerard, 1980; Kumar and Bhattacharya, 2008; Rao and Srivastava, 1973). Here, XGs are competitive with, or superior to, starches, and the strengthening effects are likely, at least in part, to be the result of fibre–fibre crosslinking mediated by the specific interaction of XG with cellulose. XG can also act as a functional barrier coating for cellulose fibres: adsorption of XG on to yarns or fabrics before dyeing, followed by treatment with a xyloglucanase (XGhydrolyzing enzyme) enhances partial dye removal, creating a desirable abraded look (Kalum, 1998). Regarding wood fibres, tamarind XG has been used in a crude form as a binder for sawdust briquette and plywood manufacture, whereas, in combination with cross-linking borate salts, the polysaccharide gum has been used as a paper adhesive (Gerard, 1980; Kumar and Bhattacharya, 2008; Rao and Srivastava, 1973; Shankaracharya, 1998). The use of TKP to replace starches and galactomannans for the improvement of paper strength properties has been long known (Gerard, 1980; Rao and Srivastava, 1973; Shankaracharya, 1998). More recently, the ability of XGs to improve paper properties has been re-examined using contemporary approaches. Isolated tamarind XG binds to chemical wood pulp fibres in aqueous suspension with high affinity, improving both strength properties as well as fibre distribution (formation) in paper sheets (Christiernin et al., 2003; Lima et al., 2003). In addition to wet-end application, XG has also been applied as a spray coating, where the polysaccharide gives considerably higher improvements in sheet strength properties for similar loading amounts (Ahrenstedt et al., 2008).



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Using a flow loop system, Yan et al. (2006) demonstrated that the effect of improved formation was the result of reduced fibre flocculation in the wet-end of the papermaking process. Rutland and co-workers, working in collaboration with our group, have used cellulose colloidal probe atomic force microscopy (AFM) to perform fundamental surface force measurements of the cellulose–XG interaction in the context of papermaking (Nordgren et al., 2008; Stiernstedt et al., 2006a, 2006b). These results suggest that XG forms a boundary lubricating layer, which reduces friction and allows fibres to more easily slip past one another in the wet state, thus rationalizing the observed improvements in sheet formation. Paradoxically, cellulose adhesion is also enhanced by XG (decreased friction is almost always associated with decreased adhesion behaviour). Wet cellulose surfaces in contact display almost no adhesion, but when coated with an adsorbed XG layer, a significant time-dependent adhesion is manifested. This implies that the XG, although essentially irreversibly adsorbed, is nonetheless able to reorient and bind to an opposing surface on time scales of the order of a few seconds to minutes. This probably contributes to the increased paper strength observed with XG-treated pulps, because fibre bonds achieve significant adhesion levels before drying owing to bridging effects (Stiernstedt et al., 2006a). Interestingly, the crosslinking of macroscopic cellulosic fibres by XG in paper and textiles may be analogous to the crosslinking of cellulose microfibrils by XG within the cell wall (Carpita and McCann, 2000). The capacity of wood pulp fibres to bind XG is closely correlated with the chemical composition of the fibre surface. Mechanical pulps, which have a high amount of residual surface lignin and extractives, bind less XG than chemical pulps with low lignin and hemicellulose content (Zhou et al., 2006a).

11.3.2 Composites from xyloglucan and cellulose fibrils In addition to whole plant fibres, complexes of pure cellulose micro/nanofibrils with XG have received continued attention owing to their relevance in understanding the molecular basis of plant cell wall morphology, see Whitney et al. (2006) and references therein. For bacterial cellulose composites, further processing with XG-active enzymes has been used to alter mechanical properties (Chanliaud et al., 2004). Several studies have underscored the remarkable physical properties of cellulose networks based on bacterial (Yano et al., 2005) and plant nanofibres (Iwamoto et al., 2005), including microfibrillated cellulose from wood pulp (Henriksson et al., 2008; Nakagaito and Yano, 2005). The admixture of cellulose nanofibres with XG thus represents an interesting new area of biofibre composite design that has, as yet, been little explored.


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11.4

Chemo-enzymatic modification of xyloglucans to functionalize cellulose surfaces

The rich history of studies on the in vivo and in vitro association of XG with cellulose, including industrially relevant applications, forms a solid basis for the use of XGs as molecular anchors to attach chemical functionality to cellulosic surfaces. Indeed, the realization that the selective modification of XG and subsequent aqueous adsorption on cellulosic fibres could circumvent some of the aforementioned difficulties associated with direct chemical derivatization has opened new possibilities to expand fibre properties. As a polymer, XG possesses a number of potentially reactive sites for functionalization (Fig. 11.1, cf. Fig. 11.4). The most obvious are the numerous primary and secondary hydroxyl groups of the polysaccharide, which can be derivatized using well-known organic chemical methods with varying selectivity. Indeed, alkylated, carboxymethylated, sulfated, and oxidized derivatives of tamarind seed XG have been synthesized (Gerard, 1980; Lang et al., 1992; Rao and Srivastava, 1973; Takeda et al., 2008), and some of these have been applied in papermaking (Gerard, 1980; Rao and Srivastava, 1973). In contrast to traditional, ‘chemical’ methods of polysaccharide modification, enzymes are well-known to offer certain advantages in terms of specificity and gentle reaction conditions. Although a multitude of enzymes capable of degrading polysaccharides is known, the number of enzymes suitable for adding chemical groups onto polysaccharides is currently limited; however, two alternative, chemo-enzymatic approaches have been devised to functionalize XG for cellulosic fibre modification.

11.4.1 Activation of xyloglucans with galactose oxidase Galactose oxidase catalyzes the regiospecific oxidation of the C-6 hydroxymethyl group (a primary alcohol) of galactose and galactosides to the corresponding aldehyde (Whittaker, 2003), which can be used as a reactive chemical handle for further derivatization. XGs are of course replete with subtending galactosyl residues along their backbones (Fig. 11.1). The seminal work by Gidley and colleagues on the oxidation of XG in solution using galactose oxidase formed the basis for the use of this enzyme to make polysaccharide conjugates (Lang et al., 1992). Subsequently, the catalytic ability of galactose oxidase was harnessed to activate the galactose branches of both locust bean galactomannan and tamarind XG for conjugation with proteins (Berry et al., 2001). In one example, the enzyme glucose oxidase and a monoclonal antibody were each covalently attached to the polysaccharides by direct reductive amination. In a complementary approach, a thiolated recombinant single-chain antibody fragment was coupled with the



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polysaccharides, which had been first activated by galactose oxidase and reacted with the thiophilic 4-(4-N-maleimidophenyl)butyric acid hydrazide. These protein– and enzyme–polysaccharide conjugates were readily adsorbed onto cotton cellulose powder in aqueous buffer. Most importantly, these gentle conditions retained the biochemical activities of the antibodies and enzyme, which were functional on the cellulosic substrate.

11.4.2 Xyloglucan endo-transglycosylase and xyloglucan as a toolkit for cellulose modification Xyloglucan endo-transglycosylases (XETs, EC 2.4.1.207) are plant enzymes that catalyze the endolytic cleavage and re-ligation of XG chains in the cell wall (Gilbert et al., 2008; Rose et al., 2002). This reaction, which proceeds without hydrolytic degradation of the XG polysaccharide, see Gilbert et al. (2008) for a full discussion, is implicated in transient wall loosening during cell growth, followed by reinforcement of the cellulose–XG network (Fig. 11.2) (Rose et al., 2002). In contrast, cell wall polysaccharide hydrolases and lyases irreversibly cleave cellulose cross-linking glycans leading to irreversible wall softening (e.g. fruit ripening, rotting). Notably, early work on the biochemistry of XETs indicated that the catalytic reaction is somewhat promiscuous; these enzymes can utilize XG oligosaccharide derivatives as alternative glycosyl acceptor substrates, thereby incorporating them into

Cellulose fibril XG

Tension

Cleavage

Expansion

XET

Re-ligation

Reinforcement

11.2 A biological role of xyloglucan endo-transglycosylases (XETs) in plant cell wall growth and restructuring. XET cleaves and re-ligates high molar mass XG chains, which bind and crosslink cellulose fibrils, without hydrolytic degradation. Transient wall loosening is believed to facilitate wall expansion, driven by turgor pressure.


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the XG chain by transglycosylation (Fig. 11.3) (Fry, 1997; Fry et al., 1992; Nishitani, 1992). Inspired by these observations, our laboratory has further developed this system as a biomimetic method to alter cellulosic fibre surface chemistry (Teeri et al., 2007; Zhou et al., 2007).

XG

R R

R

XGO-R

R

a XET

R R XG-R

b

R

R R

Cellulose surface

RR R

c

R′

R′

R

R′ R′

RR R

11.3 The use of XET and XG for cellulose modification: (a) XET produces modified XG (XG-R) by transglycosylation of XG onto XGO-R; (b) subsequent adsorption of XG-R from aqueous solution, driven by the strong cellulose–XG interaction, brings the functional group onto the cellulose surface; and (c) further reactions of surfacebound R groups are possible with appropriate reagents, including photoactivation, electrophilic/nucleophilic reactions, grafting from polymerization, and biomolecule capture.



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This technology requires three essential components (Fig. 11.3): (1) high molar mass XG polysaccharide, (2) chemically modified xyloglucan oligosaccharides (XGO-R), and (3) a xyloglucan endo-transglycosylase (XET). 1. Xyloglucan polysaccharide (XG). For reasons of convenience and largescale availability (vide supra), tamarind seed XG is the base material of choice. The oligosaccharide composition of this polysaccharide is wellestablished, and the native high molar mass is sufficiently high (Mw > 500 000). 2. Chemically modified xylogluco-oligosaccharides (XGO-R). These enzyme substrates are readily produced by the digestion of tamarind XG with microbial xyloglucanases (Gilbert et al., 2008; Pauly et al., 1999b; York et al., 1993), followed by reductive amination to install an amino group on the first Glc residue of the XGO (Fig. 11.4). This amino group provides a uniquely reactive chemical handle, which may be further functionalized, depending on the application. Reductive amination is practically convenient because the reaction occurs in aqueous solution without the need for hydroxyl protecting groups. Chemical handles may also be installed by anomeric glycosylation (Ibatullin et al., 2008), lipase-catalyzed esterification (Gustavsson et al., 2005), or via oxidation with galactose oxidase (Priem et al., 1997). Xyloglucanase digestion of tamarind XG produces a mixture of the four constituent oligosaccharides (Figs 11.1 and 11.4) and, although this may be simplified by β-galactosidase digestion and/or chromatographic separation (Greffe et al., 2005), it is practically more convenient to use the mixture directly on a large scale. 3. Xyloglucan endo-transglycosylase (XET). This enzyme is responsible for the recombination of the XGO-R substrates into the XG polysaccharide. Work in our laboratory has routinely employed a XET, PttXET16-34 from the hybrid aspen (Populus tremula × tremuloides), produced recombinantly in the yeast Pichia pastoris (Bollok et al., 2005; Kallas et al., 2005). Recombinant expression has the distinct advantage over extraction from native sources that it is highly scalable and industrially widespread (Bollok et al., 2005). PttXET16-34 is biochemically well-characterized (Kallas et al., 2005), including three-dimensional enzyme structural analysis (Johansson et al., 2004). Notably, these structural studies, which include XET-XGO ligand complexes, indicate that this enzyme has an active site cleft with the potential to tolerate a diversity of sterically bulky XGO-R functional groups. Admixture of these three components in buffered aqueous solution results in the XET-catalyzed incorporation of XGO-R into high mass XG, to yield functionalized XG, XG-R, with a concomitant reduction in polysaccharide chain length (Fig. 11.3, step a). Subsequent adsorption of XG-R


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Advances in textile biotechnology Tamarind xyloglucan

1. endo-(Xylo)glucanase/buffer/H2O 2. NH4HCO3/NaCNBH3/H2O

HO HO OH

OH O

HO x HO HO

O O

OH OH O

O HO

OH

O O

OH

HO O

O

HO

OH

O HO

OH

NH2

O

O OH

O O

O

O OH

HO

OH

OH

y

OH OH

OH OH

XGO–NH2

Further chemistry

OH (XGO)O HO

OH OH

H N

R

XGO–R

11.4 Chemo-enzymatic synthesis of amino-modified xyloglucooligosaccharides (XGO-NH2) and further derivatives (XGO-R) from tamarind XG. Digestion of the polysaccharide produces a mixture of oligosaccharides in the ratio 2 : 4 : 3 (Glc4Xyl3/Glc4Xyl3Gal/Glc4Xyl3Gal2) (York et al., 1993). XGO-NH2 and diverse XGO-R compounds are substrates for XET, which are incorporated into XG polysaccharide (see Fig. 11.3).



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onto cellulosic fibres in water results in gentle, surface-specific chemical functionalization (Fig. 11.3, Step b). The incorporation of XGO-R into XG by the enzyme is essential since, as discussed above, short XG oligosaccharides do not have a significant affinity for cellulose. Thus, the extended polysaccharide chain of the XG-R product functions as a molecular anchor, the length of which can be adjusted by the molar ratio of XG to XGO-R. The functional group density on the cellulosic surface can therefore be controlled either by adjusting the length of the XG-R anchoring tail, or varying the loading amount of XG-R with respect to the cellulose. Depending upon the nature of the R group, further elaboration of the surface chemistry is possible (Fig. 11.3, step c). Our initial proof-of-concept work (Brumer et al., 2004) focused on the well-known fluorophore fluorescein, which was conjugated to aminated XGOs (XGO-NH2) by reaction with fluorescein isothiocyanate (FITC) to yield XGO–FITC. XET-catalyzed incorporation of XGO–FITC into XG, following the scheme shown in Fig. 11.3, produced XG–FITC. Both the enzyme-catalyzed production and binding of XG–FITC to the model cotton cellulose substrate Whatman No. 1 filter paper were extensively characterized. To demonstrate the substrate versatility of the method, XG–FITC was adsorbed onto regenerated cellulose films and a XG–sulforhodamine conjugate (XG–SR) was bound to regenerated cellulose fibres (Brumer et al., 2004). Functionalized xyloglucan (XG-R) for the activation of cellulosic surfaces Although the production of yellow paper and pink fibres is somewhat trivial, the established methodology has a certain industrial relevance. The attachment of dyes and fluorophoric optical brightening agents (OBA) is widespread in both the paper and textile industries, whereas retention of these small organic molecules on cellulosic fibres is sometimes poor. Indeed, our laboratory collaborated with a large industrial wood pulp producer to test the potential of the XET/XG system to increase the efficiency of OBA retention. To this end, we generated a sulfated aminostilbene derivative of XG (XG–OBA) using XET, and demonstrated that it was quantitatively bound to pulp fibres, producing a dosage-dependent UV brightening effect (Fig. 11.5). Notably, the XG–OBA was retained on the pulp even after harsh mechanical refining (Zhou et al., 2006b and Brumer et al., unpublished results). Although inclusion of the final functionality on XG before cellulose binding may be preferred in some applications, in many cases it is desirable to produce activated (or activatible) cellulose surfaces that are capable of further reactions. As mentioned above, the amino group provides such a chemical handle for further elaboration, and cellulosic fibres activated with


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Advances in textile biotechnology (a) White light

(b)

UV light

11.5 An example of the use of the XET/XG method to retain an optical brightening agent on chemical wood pulp handsheets. Pulps were treated with six different amounts of XG-OBA and formed into sheets. Irradiation with ultraviolet light (b) shows a clear dose-dependent UV brightening effect, which is not observed under normal laboratory fluorescent tube lighting (a).

XG–NH2 have been reacted with a number of electrophiles, including isocyanates, isothiocyanates, thioesters, and activated carboxylic acids (Brumer et al., 2004; Teeri and Brumer, 2003). For example, XG–NH2-modified Whatman No. 1 filter paper can be reacted with FITC to yield paper effectively identical to that from direct XG–FITC modification (Brumer et al., 2004). Thiol groups on cellulosic fibres, from the adsorption of a thiolated XG derivative (XG–SH), can be specifically reacted with methanethiosulfonate reagents or disulfides in water to form derivatives of the general structure XG–S—S–R. Subsequently, reduction of the disulfide bond (represented by the em dash) under gentle, aqueous conditions, can be used to remove the functional groups with concommitant regeneration of the XG– SH thiol, which can be re-derivatized (Brumer et al., 2004; Gustavsson



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et al., 2005). As such, this chemical cycling represents a proof-of-concept for dynamically altering cellulose surface chemistry. Biomolecule capture The capture of biological molecules represents a special kind of surface activation, with numerous applications in separations, diagnostics, and biomedicine. Cellulose surfaces, including bacterial cellulose hydrogels, display low non-specific protein adsorption and high biocompatibility (Brumer et al., 2004; Helenius et al., 2006; Klemm et al., 2001; Miyamoto et al., 1989; Wan et al., 2006). Although desirable in certain applications, this biological ‘non-stickiness’ is a limitation in the use of cellulose to capture biological molecules or as a scaffold for tissue engineering. Here also the XET/XG system provides a solution. As a first example, adsorption of a XG bearing the small organic ligand biotin (XG–biotin) to Whatman No. 1 has yielded filters that are able to bind the protein streptavidin conjugated to the reporter enzyme alkaline phosphatase. Sensitive detection via conversion of a phosphatase substrate indicated that the XG–biotin-activated filters bound significant amounts of the protein conjugate, whereas control filters showed no detectable background binding (Brumer et al., 2004). We have recently extended this methodology to improve the adhesion of endothelial cells to artificial blood vessels composed of bacterial cellulose (Bodin et al., 2007a). Here, the adsorption of a XG–pentapeptide conjugate, XG–GRGDS, improves the ability of bacterial cellulose to function as a tissue scaffold and facilitates cell proliferation by interaction with integrin receptor proteins on endothelial cell surfaces. Importantly, adsorption of XG–FITC to the bacterial cellulose hydrogel, which is 95–99% water, indicates that the gentle aqueous binding conditions employed do not alter the morphology and, thus, the material properties of the substrate (Bodin et al., 2007a). Although the use of XG conjugates to anchor or capture biological molecules on cellulosic fibres is only in its infancy, one can readily envision a range of biomolecular probe–target systems of practical importance (Table 11.1). Extension to polymers: multivalent effects A potential limitation of the XET/XG-based method of cellulose modification is that in all of the examples described thus far, a single functional group is appended per XG chain. Given that a minimum XG-R chain length is required for binding, this ultimately limits the functional group density on the cellulosic material. Consequently, we have developed XG–initiator conjugates for the grafting of polymers from cellulose surfaces. This


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Table 11.1 Examples of molecular probes and their targets of importance in biological applications Probe

Target

Small molecule (e.g. biotin*) Carbohydrate Protein (e.g. RGD peptides*) Enzyme (e.g. alkaline phosphatase*) Substrate Oligonucleotide (DNA or RNA) Chelator

Protein (e.g. streptavidin*) Protein Protein (e.g. cell surface integrins*) Substrate (e.g. indolyl phosphates) Enzyme Oligonucleotide (DNA or RNA) Inorganics

* See text.

approach is conceptually identical to the surface activation described above, although here activation allows for the elaboration of complex polyvalent molecular architectures. In a first demonstration of the potential of XG-immobilized polymerization initiators, we successfully attached an atom-transfer radical polymerization (ATRP) initiator, the 2-bromopropionyl group, onto cellulose via the XG conjugate XG–INIATRP. Subsequently, methylmethacrylate and styrene were independently polymerized from filter paper sheets to create highly hydrophobic surfaces (Zhou et al., 2005). A particular advantage of controlled polymerization techniques, such as ATRP, is that polymer chains grow at the same rate, which results in low polydispersity and well-defined molecular properties (Matyjaszewski and Xia, 2001). Furthermore, the ‘living’ nature of ATRP facilitates the production of complex polymer block structures (Carlmark and Malmström, 2003), whereas the combination of graft polymerization with subsequent chemical modification opens a multitude of possibilities to further tailor cellulosic material properties (Golas and Matyjaszewski, 2007; Nyström et al., 2006). Attachment of the ring-opening polymerization (ROP) initiator 2,2-bis(methylol)propionic acid (bis-MPA) onto cellulose via the XG– INIROP conjugate has been used to extend the grafting-from-XG concept to so-called biodegradable polymers. Both poly(ε-caprolactone) (PCL) and poly(l-lactic acid) (PLLA) polyesters have been successfully grafted from XG–INIROP-modified filter paper to yield hydrophobic surfaces (Lönnberg et al., 2006). Notably, initiation from XG–INIROP produced papers that contained less polymer and were slightly less hydrophobic, for all polymers and graft lengths studied, than those resulting from direct esterification of the initiator onto cellulose. However, whereas the esterified and grafted cellulose paper surfaces were completely resistant to cellulase enzyme attack, papers onto which PCL and PLLA were grafted from XG–INIROP could be



281

degraded (Lönnberg et al., 2006). This suggests that the XG, which can be readily digested by endo-glucanases (endo-cellulases) (Gilbert et al., 2008; Pauly et al., 1999b), acts as a selectively cleavable linker that can facilitate recycling of grafted fibres in biocomposite materials. Indeed, existing technology involving the use of cellulases in paper and textile de-inking could potentially be adapted for such an application (Soni et al., 2008; Zeyer et al., 1994a; 1994b).

11.5


The use of XG as a molecular anchor to functionalize cellulose surfaces holds considerable promise for the development of new materials. Building upon a solid platform of knowledge of XG chemistry and enzymology, biomimetic adaptation of the unique XG–cellulose interaction is opening new possibilities for cellulosic substrates inherently derived from natural sources. The ability of XG conjugates to bind to all types of cellulose, from high-purity sources such as Whatman No. 1 filters (98% cotton cellulose, mainly Iα allomorph), cotton powder, bacterial and regenerated cellulose to complex industrial wood pulp fibres, highlights the broad potential of the system (Berry et al., 2001; Bodin et al., 2007a; Brumer et al., 2004; Zhou et al., 2006a). Moreover, there is currently an unprecedented focus on the use of cellulosic biofibres in diverse applications, from the nanoscale to the macroscale, as a consequence of their unique strength-to-density ratio, biodegradability, and renewability (Dufresne, 2008; John and Thomas, 2008; Mohanty et al., 2000; Pandey et al., 2005; Samir et al., 2005). Continued development of biological and chemical methods to control cellulose surface chemistry will be implicit in achieving the maximum performance of cellulosic materials in individual applications. Simultaneous with stepwise improvements in traditional commodities such as paper, packaging, and textiles, revolutionary smaller-volume, higher value-added products will be demonstrators of emerging technologies. In particular, the use of cellulosic fibres in nanocomposites, biomedical applications, and functional textiles has tremendous scope. Chemo-enzymatically modified XGs are thus a useful addition to the available toolkit necessary to realize this potential. The application of these molecules to bast fibres, or in combination with other types of enzymatic treatments (Mikolasch and Schauer, 2009), represent currently untapped areas that may bear fruit.

11.6


In the first instance, the reader is referred to the numerous articles and reviews cited in the text, which have been selected as a starting point for further exploration of the literature (the author regrets any omissions


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owing to space limitations or simple oversight). The following sources are particularly recommended: •

Zhou et al., 2007: a detailed review of the use of XG in cellulose modification, including an extensive bibliography. • Carpita and McCann, 2000: a highly recommended textbook introduction to the fundamentals of polysaccharide composition and plant cell wall structure. • Hoffman et al., 2005: an unparalleled overview of the structural diversity of XGs across plant species. • Gerard, 1980; Kumar and Bhattacharya, 2008; Rao and Srivastava, 1973; Shankaracharya, 1998: although each has its particular limitations, this quartet of reviews provides a holistic summary of traditional uses of XG. • Gilbert et al., 2008: a useful introductory primer on glycoside hydrolases and transglycosidases, including the XET enzyme. • http://www.biotech.kth.se/glycoscience/: homepage of the Division of Glycoscience (previously Wood Biotechnology) at the Royal Institute of Technology (KTH) in Stockholm, where the XET/XG cellulose modification biotechnology was elaborated. • http://www.swetree.com/: homepage of SweTree Technologies, a Swedish biotechnology company working toward biofibre improvement, including industrialization of the technology developed at KTH.

11.7

Acknowledgements

The author wishes to thank Professor Tuula Teeri and all of our co-authors who have contributed to the development of the XET/XG cellulose modification technology in Stockholm. Dr Peter Piispanen and Mr Kaj Kauko are especially thanked for acquiring the photographic images used to produce Fig. 11.5. Our work has been funded by the Knut & Alice Wallenberg Foundation, Swedish Foundation for Strategic Research, Formas, and the Swedish Research Council. The Swedish Research Council and the Swedish Foundation for Strategic Research (via Biomime, the Swedish Center for Biomimetic Fibre Engineering) are thanked for current funding during the writing of this chapter.

11.8

References

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12 Developments in the processing of chitin, chitosan and bacterial cellulose for textile and other applications A. F R A NC E SKO, M. DÍ A Z G ON Z Á L E Z, G. R. L OZ A NO and T. T Z A NOV, Universitat Politécnica de Catalunya, Spain

Abstract: A critical review is presented of the advances in the production and processing technologies such as fermentation and spinning of natural fibres comprising chitin, chitosan and bacterial cellulose. The main properties and applications of these natural polymers are described and their potential use in future applications, such as antimicrobial textiles for wound dressings, is outlined. Key words: natural fibres, chitin, chitosan, bacterial cellulose, fermentation, spinning, antimicrobial textiles, wound dressing.

12.1

Introduction

The textile industry of the future inevitably needs to utilize renewable resources, new fibres and environmentally friendly technologies to maintain competitive market positions. One approach toward this goal is to manipulate living organisms and renewable systems to generate materials in an environmentally friendly way. In this respect, biotechnology is gaining importance and several possibilities exist for producing entirely new, biopolymer-based fibre materials, using biotechnical tools. Natural fibres have been used by humans as construction materials and for making various fabrics for thousand years. Natural fibres include those produced by plants, animals, and geological processes. All plant fibres are composed of cellulose, the most abundant biopolymer on earth. The molecular and supramolecular structure of this macromolecule is suitable for many important applications. For many centuries, plant-derived cellulose has been utilized extensively by textile industries leading to a significant demand for wood biomass. According to the Food and Agriculture Organization of the United Nations (FAO), estimates of world production of natural fibres (cotton, jute, flax, hemp, wool, and natural skin) averaged 31 million tonne per year over the period 2003–2005 (Moir and Plastina, 2008). No significant increase in production is forseen for the future because cultivated areas, water resources, and the 288 © Woodhead Publishing Limited, 2010

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possibility of improving agricultural practices are significantly limited. Thus, by the middle of the 21st century, natural fibre production will be close to exhaustion. Besides the problem of limited natural sources, fibre production must be environmentally clean. It is obvious that cellulose and other polysaccharides, being from natural origin, are sustainable and healthy products, but some of the processes required for their isolation and modification present serious pollution problems. Over the past few years, a number of researchers have been involved in investigating the exploitation of non-traditional sources of cellulose and in designing biotechnological processes to produce this biopolymer. The production of cellulose by micro-organisms is considered to be one of the most promising ways to meet the environmental requirements. Apart from plants, certain bacteria, algae (Koyama et al., 1997), and fungi (Morita et al., 2003) also produce cellulose with an identical chemical composition to the cellulose of plant origin. In addition to cellulose, other polysaccharides such as chitin and chitosan have also become widely used over the past few years, especially in the chemical–pharmaceutical industry and in medicine. Chitin is the second most abundant polysaccharide found on earth after cellulose and is the main component in the shells of crustaceans, such as shrimp, crab, and lobster and it is also found in exoskeletons of molluscs and insects and in the cell walls of some fungi (Muzzarelli, 1994). The chitosan, usually produced commercially by alkaline deacetylation of chitin, is also found in some fungi. Recent high interest in large-scale chitin and chitosan production results from the need to process vast quantities of crab and shrimp shell wastes from seafood companies. The recycling of these wastes is the main commercial and sustainable way for large-scale chitin production. The main objective of this chapter is to overview recent progress in production and application of chitin, chitosan and bacterial cellulose fibres.

12.2

Types of new fibres: chitin, chitosan and bacterial cellulose

The natural polysaccharides chitin, chitosan and bacterial cellulose, although sharing a similar structure, exhibit different properties that make them useful for a large variety of applications.

12.2.1 Chitin structure and properties Chitin, a polysaccharide (aminopolyglucan) composed of β(1–4)-linked N-acetyl-d-glucosamine residues (Fig. 12.1a), occurs in exoskeletons, in the


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(a) NH

HO

O O

HO

O

NH

HO

O O

O

HO

O

NH

HO

OH

H3COC

OH

OH

H3COC

OH

H3COC

n

OH NH H3COC

(b) OH NH2

HO O

O O

O

HO

O O

O

HO

OH

H3COC N O

NH2

nY

nX (c)

OH

OH OH

O O

O OH

NH2

OH

OH

HO HO

OH

HO

OH

HO

HO

O O

O

O

OH

OH

HO OH

OH n

12.1 Molecular structure of: (a) chitin, (b) chitosan (nX > nY) and (c) cellulose.

cuticles of many invertebrates, and in the cell walls of green algae, some fungi, and yeasts (Muzzarelli, 1994). Chitin exhibits structural similarity to cellulose and differs from it with the replacement of C-2 hydroxyl residues by acetamide groups. Depending on the polysaccharide source and isolation conditions, chitin has a different degree of acetylation. The length of the chitin molecule in nature varies widely. Although 5000–8000 N-acetyl-dglucosamine residues are found in crab chitin, the one from yeast contains only up to 100 residues (Synowiecki and Al-Khateeb, 2003). Chitin molecules, through hydrogen bonds, are assembled in fibrils occuring in antiparallel and parallel arrangement, named α- and β-chitin, respectively. The chitinous microfibrils are arranged in sheets, which in the case of α-chitin adhere strongly by hydrogen bonding. α-Chitin has very low hydrophilicity and water permeability. β-Chitin, with a lower content of intersheet hydrogen bonds, swells readily and is more permeable. Chitin is insoluble in water, diluted acidic or basic solutions and in most of the organic solvents. It is usually dissolved in concentrated acids as well as in hexafluoroisopropanole or hexafluoroacetone.



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12.2.2 Chitosan structure and properties Chitosan (Fig. 12.1b) is a chitin derivative, produced by chitin deacetylation with concentrated alkali solutions at elevated temperatures (Agboh and Qin, 1997). By controlling temperature in chitin processing, it is possible to obtain the desired degree of deacetylation and molecular weight of the chitosan derivative, varying from 30 kDa to well above 1000 kDa. The rigid d-glucosamine structures, high crystallinity and capacity for hydrogen bond formation between the chitosan molecules lead to the poor solubility of chitosan in common organic solvents and complete insolubility in aqueous solutions above pH 7. Reducing the molecular weight and lowering the crystallinity by random deacetylation improves its solubility in dilute acids where the free amino groups are protonated and the molecule becomes fully soluble below pH 5. Chitosan is an interesting polymer because of its physicochemical properties, including its solid-state structure and the dissolved state conformation (Pa and Yu, 2001). In the solid state, relatively rigid crystallites form because of the regularly arranged hydroxyl and amino groups, whereas in solution, hydrogen bonding drives the formation of microfibrils, depending on chitosan concentration (Ohkawa et al., 2004). A number of unique properties were identified for both chitin and chitosan, such as their biocompatibility, biodegradability, non-toxicity and the ability to chelate heavy metals. Such characteristics make chitin and chitosan among some of the most interesting biopolymers for exploitation in the field of fibre preparation.

12.2.3 Bacterial cellulose structure and properties Amongst the cellulose-forming bacteria, Gluconacetobacter (formerly Acetobacter), Agrobacterium, Aerobacter, Achromobacter, Azotobacter, Rhizobium, Sarcina, Salmonella and Escherichia (Deinema and Zevehvizen, 1971; Römling, 2002; Ross et al., 1991), Acetobacter strains have been reported as the most efficient producers of cellulose. These Gram-negative and strictly aerobic bacteria are not pathogenic and are commonly found in naturally grown fruits and in fruit products. The production of cellulose by Acetobacter xylinum was reported for the first time by A. J. Brown (1886). Strains of the Acetobacter xylinum species produce extracellular cellulose that is easily isolated as a fibre material. The primary structure of bacterial cellulose is similar to that of plant cellulose: an unbranched polymer of β-1,4-linked glucopyranose residues (Fig. 12.1c). However, its degree of polymerization (DP) ranges from 2000 to 8000, and its crystallinity is between 60 and 90%. The major advantage of bacterial cellulose is that it is obtained free of lignin, hemicellulose or pectin in a 3-D network


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composed of a bundle of nanometric size microfibrils (Fig. 12.2). Because of its characteristic microstructure, bacterial cellulose has a high elastic modulus (78 GPa), a high tensile strength (at least 2 GPa), and a large surface area (El-Saied et al., 2004; Guhados et al., 2005). Bacterial cellulose also has good biocompatibility and an extremely high water content of 90% or more. Upon removal of water by air drying, the bacterial cellulose will only rehydrate to the same low extent as that of plant celluloses (about 6%) after re-exposure to water. After freeze-drying, however, it can absorb up

(a)

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12.2 SEM images (×5000) of cellulose production from: (a) Acetobacter and (b) wood pulp. Microbial cellulose has a very smooth network of microfibrils. Wood pulp has similar microfibrils, but they are part of a larger aggregation of the cell wall remains.



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to 70% of the original water content by re-swelling. Through a stepwise exchange of water for other solvents, it is possible to introduce into bacterial cellulose methanol, acetone, or n-hexane in the same volume as water, while maintaining the hollow space and network structure (Klemm et al., 2005; Schrecker and Gostomski, 2005).

12.3

Basic principles, methods and technologies

The optimal biopolymer production process should be environmentally clean and feasible for large-scale production at an acceptable cost. In the following, progress in chitin, chitosan and bacterial cellulose fibres production is summarized.

12.3.1 Chitin and chitosan fibres production Several methods for chitin and chitosan production from microbial sources such as Allomyces, Aspergillus, Penicillium, Fusarium, Mucor and Rhisopus are known (Allan et al., 1978; Knorr and Klein, 1986; Muzzarelli et al., 1980; White et al., 1979). However, these techniques are not currently used on an industrial scale. The conventional way to produce chitin from common sources, e.g. crab and shrimp shells, includes an extraction process to remove the minerals, typically calcium carbonate, followed by repeated treatments with a dilute alkali solution to remove the proteins from the shell wastes. A bleaching process is usually involved to remove the pigments. Commercially, chitosan is produced by chitin deacetylation with concentrated alkaline solutions at elevated temperature. During this process, the acetamide groups of chitin undergo hydrolysis and chitosan is formed. Several companies including France Chitine (www.france-chitine.com), Primex (www. primex.is) or Heppe Biomaterial (www.biolog-heppe.de) are using these processes for chitin and chitosan production, and the Belgian company KitoZyme (www.kitozyme.com) uses renewable fungal resources to produce chitin and chitosan. This process permits excellent control of the molecular characteristics, in particular the degree of deacetylation and the molecular mass (length of polymer chains) of the biopolymer. Research on fibre fabrication methods has attracted interest from both academia and industry. Several fabrication techniques such as spinning techniques, melt-blown, phase separation and self-assembly have been employed to produce fibres suitable for various purposes (Zhang et al., 2005). Fibres based on chitin and chitosan have been known for a long time and, in the early stages of man-made fibre development, much of the attention was focused on a chitin as a potential raw material for making artificial silk. This resulted in many attempts during 1920s and 1930s to produce chitin fibres from a number of solvent systems. Only after recognition of


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the natural renewable resources as materials with potential for fibres production, were major breakthroughs in preparation of chitin and chitosan fibres achieved, i.e. since the 1970s. In most studies, chitin and chitosan fibres are obtained by wet-spinning, comprising dissolving the polymer in an appropriate solvent followed by extrusion of the polymer solution through the spinneret into a non-solvent to solidify the fibres. The polymer precipitates in the form of filament, which can be washed, drawn and dried to form the fibres (Agboh and Qin, 1997). Traditional chitin solvents are organic and can contain di- or trichloroacetic acid alone or in conjunction with other organic solvents such as formic acid, acetic acid, chloral hydrate, and methylene chloride. Moreover, amide compounds such as dimethylacetamide (DMAc) and N-methyl-2pyrrolidone (NMP) or mixtures of these amides in conjunction with LiCl form a stable spinning solution. The most commonly used chitosan solvent is aqueous acetic acid (normally 2%). Additionally, 2% acetic acid–methanol (1 : 1, v/v) aqueous solution is considered to be a particularly suitable spinning solvent for preparation of chitosan filaments (Hirano et al., 1999). The mechanical properties of chitin/chitosan fibres produced by wet-spinning depend on both the chemical nature (e.g. degree of deacetylation) of the fibre and production (spinning) conditions. Fibres obtained from polymers with a lower degree of deacetylation showed dry and wet strength higher than those fibres obtained using more deacetylated polymers. Higher fibre strengths can be achieved also by using special spinning conditions such as the dry-jet wet-spinning technique where the biopolymer solutions are extruded, loaded to an air gap of different lengths still as a solution and precipitated in coagulation bath (Kim and Pak, 2005) and the use of special solvent system to obtain a liquid crystal phase chitin and chitosan solutions. Different drying conditions can affect fibre arrangement in e.g. 3D meshes where the fibres are stuck to each other resulting in better mechanical properties (Tuzlakoglu et al., 2004). However, chitin and chitosan fibres formed by the wet-spinning method have relatively low tensile strength and are partially soluble at pH below 5.5, thus need further improvement for specific applications. A recent breakthrough in nanofibre technology is the use of electrospinning as a convenient method for preparation of polymer fibrous materials with very fine diameters, enormous surface-to-weight area and superior mechanical properties. In this process, nanofibres are produced from the polymer solution by electrostatic forces. In a simplified form, the electrospinning process consists of a syringe that holds polymer solution and two electrodes where the positive one is connected to the syringe and the negative one is the collector under a direct current (dc) voltage supply in the kV range. The electrified jet of polymer solution drops from the tip of the syringe and onto the collector, where it sets into a fibre owing to



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evaporation of the solvent (Fig. 12.3). The solvent plays two crucial roles in the electrospinning process: first, to solvate the polymer molecules to form the electrified jet and carry the solvated polymer molecules towards the collector, then to leave the polymer fibres by rapid vaporization of the solvent molecules (Park et al., 2007). Variations in the morphology of the prepared nanofibres are influenced by both system and process parameters. System parameters include polymer solubility and concentration, molecular weight, molecular weight distribution, pH, architecture (branched or linear) of the polymer and solution properties (viscosity, conductivity and surface tension). For example, higher polymer concentration leads to increased uniformity and fibres with cylindrical shape (Deitzel et al., 2001). Thinner fibres are obtained from solutions with lower polymer concentrations (Shin et al., 2001a). Although solvent viscosity typically ranges from several dozens to few hundreds of millipascals per second and conductivity from nano- to microsiemens per centimetre, general recommendations for system parameters are usually avoided because the ideal values vary considerably with the polymer-solvent system (Greiner and Wendorff, 2007). The formation and structure of the electrospuns are also greatly affected by process parameters such as: electric potential, polymer flow rate, capillary–collector distance, ambient parameters (temperature, humidity and air velocity in the chamber) and motion of

Syringe

Polymer or blend solution Metallic needle

Electrified jet High-voltage power supply Fibre mat

Collector

12.3 Schematic representation of electrospinning fibre production.


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the collector. Deitzel et al. (2001) reported that there is an optimal range of electric field strengths for a certain polymer/solvent system, as either too weak or too strong will lead to the formation of beaded fibres. Increase in the polymer flow rate results in an increase in fibre diameter and porosity and, in addition, to bead formation owing to the inability of fibres to dry completely before reaching the collector, as recently described by Sill and von Recum (2008). The same report shows that capillary–collector distance is less significant for the produced fibre size, but rather determines the final result as electrospinning or electrospraying. Upon an inadequate drying jet the final result is small droplets instead of fine fibres. The fibre diameters of the electrospuns are strongly affected by ambient parameters, such as temperature and relative humidity. A change in temperature affects two parameters with two opposing effects on the average diameter: solvent evaporation rate which increases and viscosity of the polymer solution which decreases with increase of the temperature. The final result will depend on which parameter prevails in the system (De Vrieze et al., 2009). The relative humidity leads to thinner or thicker fibres, only depending on the chemical nature of the polymer (Tripatanasuwan et al., 2007). Electrospinning has the potential to be further developed for large-scale production of nanofibres. Modern electrospinning technology is able to generate continuous fibres with diameters in the range of nanometers to a few micrometers (Doshi and Reneker, 1995). Natural chitosan has poor fibre-forming properties, which makes difficult the preparation of pure chitosan nanofibres. Trifluoroacetic acid (TFA) has excellent volatility, which allows rapid solidification of the TFA–chitosan solution and preparation of chitosan nanofibres (Ohkawa et al., 2004). Geng et al. (2005) showed that pure chitosan nanofibres could also be prepared directly from the solution in concentrated acetic acid. The increase of acetic acid concentration allows the strength of the electric field applied to be reduced resulting in low-diameter, uniform and bead-free nanofibres. However, to reduce toxicity concerns, chitosan electrospuns are usually prepared from traditional dilute aqueous acetic acid solvent using fibre-forming facilitating additives like poly(ethylene oxide) (PEO) (Bhattarai et al., 2005; Zhang et al., 2008) or poly(vinyl alcohol) (PVA) (Li and Hsieh, 2006), resulting in very fine nanofibrous structure (Fig. 12.4). Electrospuns of chitosan derivatives such as hexanoyl (Neamnark et al., 2006) and carboxymethyl chitosan (Du and Hsieh, 2008a) were also reported. Chitin electrospuns can be prepared using hexafluoroisopropyl alcohol as a solvent (Noh et al., 2006). Electrospinning from such a solution led to 400 nm diameter nanofibres (Fig. 12.5). The versatility of the electrospinning technique is in the formation of fibres with different morphologies and prepared from different materials. Therefore, different polymers, blends, mixtures or precursors can be made



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12.4 SEM images (×5000) of chitosan electrospuns. Electrospinning solution, chitosan/PEO; 80/20 doping ratio. Chitosan nanofibres obtained with fine nanofibre structure.

into fibres to suit specific applications (Liao et al., 2006). Chitin and chitosan electrospuns were prepared using a variety of additives and/or in blends with other biopolymers for targeted applications. Silk fibroin was used to prepare blend nanofibres with both chitin (Park et al., 2006) and chitosan (Park et al., 2004). Du and Hsieh (2008b) developed a two-step approach to prepare cellulose/chitosan hybrid nanofibres starting from their derivatives: cellulose acetate (CA) and dibutyryl chitin (DBC). The method included electrospinning of the biopolymers mixture followed by simultaneous alkaline hydrolysis.

12.3.2 Bacterial cellulose production By selecting the substrates, cultivation conditions, various additives, and, finally, the bacterial strain, it is possible to control the molecular mass, the molecular mass distribution, and the supramolecular structure of bacterial cellulose. Thus, it is possible to control the important properties of cellulose, and also the course of biosynthesis. A number of articles have been published to provide detailed information on cellulose biosynthesis (Cannon and Anderson, 1991; Kimura and Kondo, 2002; Ross et al., 1987, 1991) and the factors affecting the production of bacterial cellulose (El-Saied et al., 2004). Owing to the very interesting properties of bacterial cellulose for


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Advances in textile biotechnology (a)

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12.5 SEM images (×500) of: (a) chitin electrospuns and (b) commercial chitin microfibres. Chitin nanofibres have diameters of 50–460 nm, whereas commercial chitin fibres have an average diameter of 8.77 μm.

different industries, in recent years, the design of economical large-scale production processes has attracted considerable attention (Shoda and Sugano, 2005). It is clear that the conventional method of microbial cellulose production by static cultivation using sugars such as glucose, fructose, and sucrose is not applicable to large-scale industrial production because



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of the long cultivation time and expensive carbon source. To date, fermentation processes using stirred-tanks or airlift reactors have been reported as the most effective way to produce microbial cellulose on a large scale. Although agitated culture, where cellulose is synthesized in the form of fibrous suspensions, pellets or irregular masses (Czaja et al., 2004), causes some problems such as strain instability, non-Newtonian behavior or proper oxygen supply (Kouda et al., 1996, 1997a,b), this process has been proposed as the most suitable technique for economical scale production. One of the most important problems of agitated culture of bacteria is the generation of non-producing mutants. The addition of ethanol to the medium has been found to be effective to repress this spontaneous mutation (Park et al., 2003). However, to raise a large-scale production, the selection of stable strains is indispensable (Toyosaki et al., 1995). There have been many studies on bacterial cellulose production in stirred-tank reactors. Various parameters including reactor design (Kouda et al., 1997b), carbon and nitrogen sources (Bae and Shoda, 2005a; Naritomi et al., 1998), and pH and dissolved oxygen (Hwang et al., 1999) have been studied in order to optimize bacterial cellulose production. The addition of water-soluble polysaccharides such as agar has also been found to enhance the productivity (Bae et al., 2004). For the optimization of all the experimental parameters that affect bacterial cellulose production statistical methods have also been described (Bae and Shoda, 2005b). In addition to bacterial cellulose production in stirred-tanks, an intensive investigation on production using airlift reactors has been made in last years. This type of reactor has a low power consumption but requires the supply of oxygen-fortified air to improve the oxygen transfer in the culture (Chao et al., 2001; Zuo et al., 2006). As for stirredtanks, culture conditions, including pH, carbon and nitrogen sources (Noro et al., 2004), and the presence of water-soluble polysaccharides (Ishida et al., 2003) have been studied for airlift reactors. The main advantage of the stirred-tank reactor is its ability to prevent the inhomogeneity of the culture broth by strong mechanical agitation, whereas the disadvantage is its high energy cost (0.66 kW h−1 for 1 g L−1) for generating the mechanical power. Conversely, an airlift reactor has a low energy cost (0.11 kW h−1 for 1 g L−1) but its agitation power is limited, resulting in low fluidity of the culture broth, especially at high bacterial cellulose concentrations. One example of a large-scale production of bacterial cellulose is Cellulon® fibre. This cellulosis fibre, acquired by Monsato (US) in the mid-1990s, was first developed by two US-based companies, Weyerhaeuser Co. and Cetus Co. using a deep-tank fermentation technique and a patented, genetically improved Acetobacter strain. The Cellulon® fibres produced by agitated aerobic bacterial fermentation, are approximately 0.1 μm wide and of indeterminate length, providing a large surface area.


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12.4

Applications

In this section the most relevant applications of chitin, chitosan and bacterial cellulose are briefly described with special focus on textile and medical (wound dressing) fields.

12.4.1 Chitin and chitosan The growing interest in chitin and chitosan exploitation is tightly connected to their unique properties, such as biocompatibility, biodegradability, nontoxicity and ability to chelate heavy metals. Chitin and its derivatives are widely used as a coating material for nylons, cotton, and wool fibres. The use of such modified fibres includes the production of wound dressings, medical textiles, sanitary absorbents and non-allergenic, deodorizing, and antimicrobial underwear, sportwear and socks. The addition of chitin into the coating of waterproof textiles causes a large increase in its water vapour permeability. Moreover, the finishing of wool, cotton and polyester fibres with chitin derivatives such as chitosan improves their dyeability (Julià et al., 2000) and colorfastness (Najafi et al., 2008). Numerous publications report on chitin and chitosan based materials with high-value applications especially in the chemical–pharmaceutical industry (Felse and Panda et al., 1999) and medicine (Khor and Lim et al., 2003). A wide variety of chitosanbased materials are commercially available for this kind of application. Some commercially available chitosan-based products for medical applications are: CeloxTM (www.celoxmedical.com); HemCon® 100% chitosan sponge bandages, haemostatic and surgical dressings, dental dressings, and plasters (www.hemcon.com); Chitoskin, another sponge-like product, comprising 50% chitosan and 50% bovine gelatine (www.sangui.de); and KiOnutrime-CsTM and KiOmedineTM (www.kytozyme.com). There are no textile bandages coated with chitosan currently on the market. Chitin and chitosan in combination with other biopolymers and their derivatives have been used in their fibre forms to design various types of wound dressing. Loke et al. (2000) and Wang et al. (2002) demonstrated that intrinsically antimicrobial chitosan can serve as a good platform to be further upgraded for application as a wound-healing promoter. The biocompatibility of some derivatives such as dibutyryl chitin (DBC) in the context of wound dressings has been studied by Muzzarelli et al. (2005). DBC fibres were obtained by the wet-spinning method followed by preparation of a non-woven material further reinforced with 5-methylpyrrolidinone chitosan. It was found that DBC fibres are suitable for contacting both intact and wounded human tissues. Pielka et al. (2003) coated polypropylene nonwoven materials with regenerated chitin and its derivative DBC. The materials obtained caused no cytotoxic effects in in vitro or in vivo studies and



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had a positive influence on acceleration of the wound-healing process. Cotton-like fibres were prepared from chitosan-combined glycosamynoglycans, namely hyaluronic acid, chondroitin sulfate and heparin using the wet-spun technique by Hirano et al. (2001). Between 5 and 33% of glycosaminoglycans were present in the fibre materials making them mechanically weaker, but also allowing the development of targeted release strategies as an alternative wound healing approach. The effect of the molecular weight and treatment concentration on the antimicrobial activity of chitosantreated cotton fabrics has been investigated by Shin et al. (2001b). The antimicrobial activity of chitosan-treated fabrics seems to be more affected by the treatment concentration rather than by the molecular weight of the biopolymer. Chitosan has also been immobilized onto cotton fabrics using polycarboxylic acid (BTCA) and this treatment significantly affected the antimicrobial activity of the fabrics against Gram-positive and Gramnegative bacteria and fungi tested (El-tahlawy et al., 2005). Lim and Hudson (2004) studied the antimicrobial activity of O-acrylamidomethyl-N-[(2hydroxy-3-trimethylammonium)propyl] chitosan chloride (NMA-HTTC), a water-soluble derivative of chitosan. This derivative contains acrylamidomethyl functional groups, which can react with cellulose under alkaline conditions. The fibre-reactive chitosan derivative showed complete bacterial reduction within 20 min. Chitosan was blended with polyurethane (Shih and Huang, 2003) and the mixture was used to treat woollen fabrics. The experimental results indicated an improvement in both the shrinkproof and the antimicrobial properties of the fabric, but yellowing of the fabric and softness decrease were also observed. Chitin and chitosan and their derivatives have been shown to be promising materials in tissue engineering and regeneration. Their nanofibres are continuous and potentially allow for integrated manufacturing of 3D nanofibrous scaffold with high porosity and high spatial interconnectivity (Dzenis, 2004; Ma et al., 2005). Another application of chitin and chitosan in waste water treatment (chelation) has been described by Gerente et al. (2007). One of the most effective methods for purification and isolation of the enzyme chitinase is chitin affinity chromatography, which employs chitin as the adsorbent to selectively adsorb chitinase from the fermentation broth. To increase its adsorption capacity, chitin is usually processed in a chemical or controlled enzymatic way. Recently, Kao et al. (2009) demonstrated a fermentation approach to increase the chitinase adsorption capacity of chitin in the enzyme affinity purification process. The method involves the controlled bacterial (Paenibacillus sp.) treatment of chitin where enzymes from bacteria serve to partially hydrolyse chitin and obtain fibre-like structures with deep pores to provide more sites for subsequent adsorption of chitinases. It was also suggested that the fermentation process provided not only the chitinase hydrolytic effect, but also an unknown modifying effect


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from Paenibacillus sp. So-called fermentation-processed chitin (FPC) was compared for chitinase adsorption capacity to other chitin adsorbents obtained using chemical and enzymatic processes and results showed superior behaviour of FPC for enzyme adsorption and subsequent enzyme activity of immobilized chitinase. This new method for chitin processing has the advantages of low energy consumption and waste emission over traditional chemical/physical methods and shows potential for scale-up from a 5 L stirred tank-bioreactor to industrial-scale production.

12.4.2 Bacterial cellulose Owing to its purity, remarkable mechanical properties in both the dry and the wet state, porosity, water absorbency, mouldability, biodegradability and excellent biological affinity, a wide range of applications of bacterial cellulose can be envisaged in the food, medical, paper and textile fields. A new type of artificial leather with a soft touch was produced from bacterial cellulose. Bacterial cellulose has also been investigated for paper production and companies like Ajinomoto Co. along with Mitsubishi Paper Mills in Japan are currently active in this area. Shah and Brown (2005) described some interesting uses of this type of ‘electronic paper’ such as e-book tablets, e-newspapers, dynamic wallpapers, rewritable maps and learning tools. Because of its outstanding sound reproducibility, Sony Corporation has now commercialized high-quality headphones using bacterial cellulose sheets. Other interesting uses of bacterial cellulose, most of them described as possible applications in the patent WO 8912107 (Brown, 1989) include: water treatment, carrier for battery fluids and fuel cells, mixing agent, viscosity modifier, light-transmitting optical fibres, biological substrate medium, food or food substitute and lint-free specialty clothing. Despite of all these interesting uses, to date, the biomedical field has been the main area of application of bacterial cellulose (Czaja et al., 2007). This includes healthcare textiles (Wan and Millon, 2005), artificial skin (Czaja et al., 2007), catheter covering dressing (Wan and Millon, 2005), dialysis membrane (Wan and Millon, 2005), membranes for tissue-guided regeneration (Czaja et al., 2007; Wan and Millon, 2005), controlled-drug release carriers (Wan and Millon, 2005), scaffolds for tissue engineering (Czaja et al., 2007), and artificial blood vessels (Backdahl et al., 2006). Various commercial preparations of bacterial cellulose such as Biofill®, Bioprocess®, Gengiflex® and BASYC® have been successfully applied for some of the medical applications described above. A family of wound care products based on bacterial cellulose named XCell® (www.xyloscorp.com) has been marketed in the US since 2003. Owing to its unique nano- and microfibrils 3D network bacterial cellulose served as a template or matrix for the synthesis of nanoparticles and nanowires (Li et al., 2009). For bacterial cellulose to be



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suitable for these diverse applications some modifications of the fibres such as acetylation process (Barud et al., 2008) or forming composite with other materials (Pommet et al., 2008; Wan et al., 2006) have been reported.

12.4.3 Bacterial cellulose–chitosan blends Since the chemical structure of the chitosan backbone is very similar to that of cellulose, it was expected that chitosan could be miscible with cellulose and the blending might improve the mechanical and biological properties of the developed material. Ciechanska (2004) reported on the modification of bacterial cellulose during microbiological synthesis by introducing selected bioactive polysaccharides, such as various chitosan forms and their derivatives, into the culture medium. This led to the incorporation of glucosamine and N-acetylglucosamine units in the cellulose chains. The modified bacterial cellulose consists of microfibres having diameters in the order of tenths of micrometer that form a 3D network and are characterized by excellent moisture-retaining properties. Such features make bacterial cellulose/chitosan a dressing material suitable for treatment of various kinds of wounds, burns and ulcers. The same method has been exploited recently to produce bacterial cellulose–chitosan films (Phisalaphong and Jatupaiboon, 2008). Modifying bacterial cellulose by adding 0.25–0.75 (% w/v) of chitosan (85% DAC) of MW 30 000 and 80 000 in the culture medium during biosynthesis by A. xylinum provided improved mechanical properties in the wet and the dry state, and large water absorption capacity and average surface area.

12.5


Chitin, chitosan and bacterial cellulose are natural biopolymers with numerous intrinsic properties for various applications. The natural abundance of chitin and its derivative chitosan in combination with their biocompatibility, biodegradability, antimicrobial activity and non-toxicity is a reason for increased interest in their exploitation, especially in biomedical and pharmaceutical applications. It is expected that electrospinning will become the method of choice for the production of chitin and chitosan fibres. The major advantage of this method is the ability to produce long and continuous nanofibres in a cost-effective way, which makes it attractive for industrialization. However, as in all nanotechniques, its low productivity is an important disadvantage. In order for electrospinning to be commercially viable, it is essential to increase the production rate of the nanofibres. The other major constraint for large-scale production of pure chitin and chitosan nanofibres still remains the selection of appropriate non-aggressive and non-toxic solvents.


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Compared with cellulose from plants, bacterial cellulose possesses higher water holding capacity, higher crystallinity, higher tensile strength, and a finer web-like network. However, the production cost of bacterial cellulose is still high compared with that of other natural fibres. Nevertheless, largescale microbial cellulose production, through environmentally friendly fermentation processes, appears to be feasible in the future based on the increased understanding of the biochemical and genetic background of cellulose biosynthesis.

12.6

Future trends

Basic electrospinning can only yield random nanofibres rather than aligned nanofibres. This can be overcome by using a rotating drum collector (Subramanian et al., 2005) and the simple or parallel knife-edge disc collector (Xu et al., 2004), which improves the degree of alignment of the fibres. The low productivity of the electrospuns can be solved using multi-nozzle electrospinning devices where several syringes in parallel alignment are used to introduce the same polymer solution. This is also a method of choice to obtain blended and even multicomponent nanofibres (e.g. for complex tissue architectures) when the syringes are loaded with different polymer solutions. Optimization of the system (by choosing an appropriate solvent and fibre-forming facilitating product) and process parameters (e.g. electric potential) in combination with use of a rotating collector and multiple nozzles is necessary to increase the productivity of aligned continuous fibres. Multicomponent nanofibres can be also produced by coaxial electrospinning. In coaxial electrospinning, two concentrically aligned nozzles are used for spinning and the same voltage is applied to both nozzles. In this instance, a core–shell nanofibre is created for application in hollow nanofibres and nanochannels (Sun et al., 2003). This method is also suitable for fabricating solid fibres using electrically dissimilar materials. For example, one polymer may be selected to enhance the mechanical strength, whereas the other could be chosen to increase the wettability. The development of an efficient large-scale fermentation technology for microbial cellulose production at a competitive cost is also required in the future. For this purpose, a combined use of airlift and stirred-tank reactors, the use of rotating disk reactors, or continuous cultivation, might be possible ways to obtain high-productivity bacterial cellulose synthesis at lower energy cost. Another important approach to raise a good bacterial cellulose production is the optimization of the culture and the bacterial strain selection. Improvement in media compositions by selecting appropriate carbon and nitrogen sources, and incorporating substances that stimulate cellulose production could also contribute to cost reduction. Escherichia coli and Salmonella are able to improve the cellulose production with Acetobacter



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xylinum. Although the amounts of cellulose produced by these bacteria were significantly smaller than that from Acetobacter xylinum, they exhibit a faster growth rate. As it is believed that the cellulose synthetic genes of all these bacteria are similar, genetic modification of Acetobacter xylinum, in order to increase its growth rate is expected to have a direct effect on the bacterial cellulose production time.

12.7


Numerous articles and books dealing with chitin, chitosan and bacterial cellulose have been published over the past few years. Biopolymers, Journal of Applied Polymer Science, Journal of Biomedical Materials Research, Polymers for Advanced Technologies, Biotechnology & Bioengineering, Cellulose, Polymer Bulletin, Applied Microbiology and Biotechnology, and Bioresource Technology are some of the journals devoted to the dissemination of research and scientific and technological progress in the field of biopolymers. Advances in Chitin Science is a series of chitin proceedings books initiated in 1996 following the 1st International Conference of the European Chitin Society, and since then 10 volumes have been published. Extensive research in chitin and chitosan chemistry and application has been carried out by Professor Muzzarelli from University of Ancona, Italy (Muzzarelli and Muzzarelli, 2002; Muzzarelli et al., 1998, 2005; Muzzarelli, 1994). The main focus on the science and technology of marine polysaccharides, in particular chitosan chemistry and applications, has been defined by the research group of Professor Vårum from the Norwegian University of Science and Technology (www.biotech.ntnu.no/nobipol). There have been many studies in the area of bacterial cellulose research, since the discovery in 1886 of the cellulose-producing activity of Bacterium xylinum by Adrian J. Brown. The production of cellulose by this Gram-negative bacterium has long been regarded as an archetype for the study of cellulose biogenesis. A number of excellent review articles have been published to provide detailed information of cellulose biosynthesis (Brown and Saxena, 2000; Kimura and Kondo, 2002; Ross et al., 1991). The first breakthrough in this area came from the purification of Acetobacter cellulose synthase from the Professor Malcolm Brown’s research group in 1989. Since this important finding, successful research in bacterial cellulose has been carried out in Brown’s laboratory at The University of Texas, USA. Further information about the work of this group on bacterial cellulose biosynthesis, production and applications can be found at www.botany.utexas.edu/mbrown. In recent years, the University of Jena, Germany (www.polymet-jena.de) and the Virginia Tech Wake Forest University School of Biomedical Engineering (www.sbes.vt.edu), USA have been active in bacterial cellulose development, e.g. of nanocellulose hollow filaments for medical applications such


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as blood vessels (www.jenpolymers.de, www.arterion.se). The European Chitin Society (www.euchis.org) organizes the annual International Conference of the European Chitin Society with the aim of encouraging basic and applied scientific studies of all aspects of chitin, chitosan and their derivatives, and serving as a major tool for dissemination of the results primarily in Europe. In the USA, the Cellulose and Renewable Materials Division from the American Chemical Society serves its members by providing an integrated platform to identify, process, and disseminate knowledge that advances the science and technology of renewable materials and biopolymers.

12.8

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13 Hydrolysis of regenerated cellulose fibres for textile and other applications T. BE C H T OL D and C. B. S C H I M P E R, Leopold-Franzens University Innsbruck, Austria

Abstract: The use of steaming, drying, and alkalisation as pretreatment processes for regenerated cellulose – such as lyocell type fibres, viscose and modal fibres – is described. These processes overcome the problems caused by the size of the enzymes and the average pore diameters of regenerated cellulose fibres and allow cellulases to access the pores in the fibres so that hydrolysis is not restricted to the surface of the fibre. An efficient combination of pre-treatment steps and cellulase hydrolysis exhibits significant potential to produce new materials. Differences in crystal structure between native cellulose fibres and mercerised cotton and regenerated cellulose fibres are described and the factors that determine the effects of cellulase on substrates are outlined. Key words: cellulose fibres, textiles, cellulase, hydrolysis, alkalisation, viscose, lyocell, modal.

13.1

Introduction

Cellulose is the most abundant polymer provided by nature. Fibrous structures can be easily separated from plant sources, e.g. seed (cotton) and stem (flax, hemp, jute), or are available via dissolution/regeneration steps as regenerated cellulose fibres, e.g. lyocell, viscose or modal fibres. The ability of cellulose polymers to interact with water by sorption in the polymer structure, by swelling and uptake of liquid water into fibre pores explains the favourable wear and comfort properties of textiles containing cellulose fibre. In 2007, the share of man-made cellulose fibres of the total textile fibre production of 76.192 million tonnes per year can be estimated at 3.592 million tonnes, which corresponds to 4.7% (Anon., 2008). In the same year an amount of 26.704 million tonnes of cotton was produced, which indicates the relative size of the cellulose fibre market and demonstrates the substantial potential of man-made cellulosics. Owing to the limited farmland available for cultivation of cotton and increasing concerns about environmental aspects of cotton farming such as water consumption for irrigation and the use of pesticides and herbicides, 312 © Woodhead Publishing Limited, 2010

Hydrolysis of regenerated cellulose fibres for textiles

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growth in production of cellulose fibres is expected to occur mainly in the share of regenerated cellulose fibres. Thus, growing interest is directed towards modification of regenerated cellulose fibres e.g. by cellulase treatment. The processing of cellulose fabrics and garments with cellulase enzymes has allowed a wide range of applications, including novel finishes, soft hand, fuzz and pilling prevention, increase of smoothness and softness, increase of lustre and colour brightness, improvement of handle and drape, and fashionable wash down effects (Andreaus et al., 2000; Cavaco-Paulo, 1998; Cavaco-Paulo and Gübitz, 2003; Cavaco-Paulo et al., 1997; Kumar et al., 1995). Although cellulose fibres such as flax, hemp and cotton have been used for more than 2000 years, the structure of cellulose fibres has not yet been described in sufficient detail. Differences in the crystal structure between native cellulose fibres (cellulose I) and mercerised cotton and regenerated cellulose fibres (cellulose II) have already been detected, but structural models describing the behaviour of cellulose fibres in aqueous conditions still need to be developed. In particular, the differences in fibre structure of various native and regenerated cellulose fibres in the swollen state will influence the effect and intensity of a cellulase treatment. Thus, the final effect of a certain treatment will be influenced by three groups of factors (Fig. 13.1): •

type of enzyme and treatment conditions (type of cellulase, temperature, pH, concentration); • material structure, including fibre swelling and accessibility (cotton, viscose, lyocell); • preceding treatments modifying fibre properties and thus observed enzyme reactivity (retting of flax, bleaching, mercerisation of cotton, alkalisation of cellulose fibres, drying steps, dyeing). Cellulase treatment (endo-, exo-, total crude cellulase, pH, T)

Cellulase reactivity Material structure (surface, accessibility, swelling, porosity)

Preceding treatments (retting, bleach, alkalisation, drying)

13.1 Schematic presentation of the main parameters influencing the observed cellulase reactivity.


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Therefore, the effects of cellulases observed on a certain substrate are dependent on several factors which will be discussed in more detail below. This interlinkage between treatment conditions, fibre properties and history of preceding treatments explains the sometimes unexpectedly high weight loss and strength loss obtained in practice under similar treatment conditions and using similar cellulosic substrates, when the effect of treatments preceding enzymatic hydrolysis had not been considered.

13.2

Regenerated cellulose fibres

Regenerated cellulose fibres offer a high variability in fibre structure and numerous options to modify the material, thus differences in observed cellulase reactivity can be more distinct than in cotton fibres. Regenerated cellulose fibres commercially available in the textile fibre market at present, can be roughly divided into two major groups with regard to their production process. The first group of fibres is produced by the viscose process and the second group are lyocell-type fibres. Viscose fibres and modal fibres are produced by dissolution of cellulose xanthogenate in alkaline solution and precipitation/coagulation, and cellulose regeneration in acid baths, respectively. The combination of precipitation, coagulation and cellulose regeneration leads to the core skin structure known for viscose fibres (Abu-Rous et al., 2007). During the coagulation/ regeneration of the cellulose considerable shrinkage of the coagulated structure occurs and the fibre obtains its characteristic multi-lobal form. Modal fibres are produced following the xanthogenate process, but special conditions are applied, such as addition of modifiers to the spin-dope and Zn salts in the spin bath. Cellulose polymers with a higher degree of polymerisation then can be regenerated to high wet modulus fibres which exhibit an all-skin structure. Lyocell fibres are formed from NMMO solution (N-methylmorpholineN-oxide monohydrate) by a wet-spinning process. The fibre shows high crystallinity, high longitudinal orientation of crystallites, high amorphous orientation, low lateral cohesion and relatively large void volumes (Öztürk et al., 2006, 2009; Schurz, 1994; Schurz and Lenz, 1994). The fibrillar structure of the lyocell fibres explains their high longitudinal stability in the swollen state, but it also causes significant weakness of the fibre against fibrillation. Penetration of swelling agents into amorphous regions of the fibre weakens the interfibrillar bonding and, as a result, the long macrofibrils can be split off when shear stress is applied on swollen fibres. Typical fibre properties of viscose, modal, lyocell fibres and cotton fibres, which are related to swelling in water and sorption of water molecules are summarised in Table 13.1. In addition to the water retention value determined by the centrifugation method, the porosity Vp characterised by



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Table 13.1 Representative fibre data of man-made cellulosic fibres and cotton: degree of polymerisation, porosity and BET moisture regain Parameter Water retention value WRV Pore volume Vp Average pore diameter Pore surface Moisture content (20 °C, 60% RH) BET moisture regain Vm

Unit

Viscose −1

c

Modal d

c

Lyocell d

b

Cotton c

0.56d

ml g

0.88 , 0.89

0.62 , 0.64

ml g−1 Å

0.64a 29a

0.49a 24a

0.53a, 0.58b 29a, 32b

0.31a 35a

m2 g−1 %

438a 9.98d

408a 9.64d

365a, 369b 9.23e

177a 5.54d

mmol g−1

3.05c

2.99c

2.73c, 2.79e

1.59e

0.71 , 0.71

a Determined by inverse size exclusion chromatography (ISEC) (Bredereck and Gruber, 1995). b Öztürk et al. (2009). c Siroka et al. (2008). d Okubayashi et al. (2005). e Okubayashi et al. (2004).

inverse size exclusion chromatography (ISEC) and data describing moisture sorption are given. The Brunauer-Emmet-Teller (BET) value Vm characterises the moisture regain corresponding to a monomolecular layer on the substrate. A more extensive comparison of fibre properties of lyocell, viscose and modal has been presented by Kreze and Malej (2003). The values in Table 13.1 show distinct differences between the man-made cellulosic fibres and cotton fibres, which exhibit approximately half the sorption capacity and internal surface area. The much lower pore volume Vp and also the lower water retention value (WRV) indicate lower expansion and swelling of the cellulose structure for cotton.

13.3

Cellulases

For cellulose hydrolysis, the cellulases have to access the surface of the cellulose fibre, then the enzyme binds with the cellulose binding domain to the polymer structure. Thus, porosity of the cellulose fibre and reactivity of surface will influence the degradation rate of the enzymes. For the swollen cellulose fibre, the cellulose surface is best illustrated by a water-containing boundary layer formed between highly ordered parts of the cellulose fibre and fully hydrated, less ordered parts of the three-dimensional polymer structure. This layer can be assumed to show a gradual change in ‘cellulose concentration’, from high polymer density inside the fibre to pure solvent outside the fibre. The molecular size of enzyme proteins e.g. Trichoderma


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reesei cellulase is assumed to be 35–75 Å for a spherical shape, and is estimated between 20 × 110 and 40 × 250 Å for an ellipsoid form (BuschleDiller et al., 1994). Comparing the size of enzyme proteins with the pore size distribution of cellulose fibres, Bredereck and Gruber (1995) concluded that the hydrolysis is expected to proceed mainly at the surface of the swollen fibres. Degradation inside the fibre structure can only proceed in a few pores, which have a diameter large enough to permit access to the cellulase proteins. Thus, enzymatic hydrolysis of viscose, modal and lyocell starts from the surface of the fibres. Analysis of pore size distribution by ISEC indicates a reduction of pore volume after enzymatic hydrolysis for lyocell fibres, whereas an increase was measured for viscose fibres and to a lower extent for modal fibres. These differences were explained by Bredereck and Gruber (1995) in terms of the different pore size distribution in the cross-section of lyocell type fibres and viscose/modal fibres. The hydrolysis of viscose fibres was also studied by scanning electron microscopy (SEM) and atomic force microscopy (AFM). Differences between a total crude cellulolytic complex and endoglucanase-enriched neutral cellulolytic enzymes were observed for viscose fibres. A total crude enzyme removes fuzz, pills and impurities from the fabrics surface and leaves distinct cavities and indentations along the length of macro-fibrils. When viscose fibres are treated with endoglucanase-enriched enzymes, the major process is peeling and smoothing of fibre surface (Kokol, 2003). Similar differences between the action of exoglucanases (cellobiohydrolase, CBH) and endoglucanases (EG) were reported for cotton fibres, where CBHs were shown to produce indentations in the fibre structure, whereas EGs smoothen and peel the surface of the fibres (Lee et al., 2000). In a systematic study of the adsorption of a total crude cellulose, Shen et al. (2002, 2004) compared the adsorption on cotton, viscose and flax fibres. As a result it could be demonstrated, that viscose fibres offered the greatest number of sites for adsorption of enzymes, the largest surface area available for enzyme coverage, and the maximum volume that can be occupied by cellulases. The relative magnitude of these parameters being 4 : 2 : 1 for viscose, cotton and flax, respectively. Compared with cotton fibres, man-made cellulose fibres show a greater variability in sorption and swelling behaviour depending on the processing steps in textile production, thus observed cellulose reactivity is influenced by the conditions applied during the preceding treatment steps.

13.4

Cellulase hydrolysis of regenerated cellulose fibres

In a total crude cellulose, three main types of enzymes are active. Endoglucanases or endocellulases hydrolyse cellulose polymers randomly along the © Woodhead Publishing Limited, 2010


317

chains. As the activity of the enzymes is directly related to the accessibility of a certain part of the polymer chain, the action of EGs is preferably observed in the non-crystalline regions. Exocellulases or CBH attack the polymer chain ends, producing cellobiose as the primary product. Depending on the cellulase formulation, cellobiose and small chain oligomers are released into solution and are hydrolysed by β-glucosidases to glucose, which is released into the solution as a major product of the cellulase activity. To identify the site of action of the different enzymes in a total crude cellulose, the relative sizes of enzymes, glucose units, fibrils in the fibre and pores have to be considered (Bredereck and Gruber, 1995; Lee et al., 2000). In Table 13.2, average dimensions of elements of fibre architecture and of proteins have been summarised. As shown Table 13.2, the enzymes are too large to enter into the micropore system of the fibres and, thus, primary action of cellulases occurs at the surface of the fibres. Mechanical agitation can support the action of the enzymes by formation of macropores e.g. owing to fibrillation of the fibre. Progressive degradation removes the more reactive amorphous parts of the fibres resulting in formation of indentations, which enable enzymes to access to inner parts of the fibre. Figure 13.2 gives a schematic presentation of the action of CBH, EG and β-glucosidase on an insoluble cellulose substrate. Both EG and CBH reactions with the insoluble cellulose substrate are heterogeneous chemical reactions whereas the β-glucosidase reaction with the dissolved short-chain oligomers and cellobiose is a homogenous process in the liquid phase. In total crude cellulase mixtures, this homogeneous reaction can be supposed to be more rapid and less rate determining than the heterogeneous reactions of EG and CBH. In addition to the specific activity of the enzymes applied, the heterogeneous reactions of EG and CBH will directly be influenced by the ‘surface’ properties of the insoluble cellulose substrate treated. Thus, besides the Table 13.2 Representative average dimensions of fibre elements and cellulases (Bredereck and Gruber, 1995; Lee et al., 2000; Öztürk et al., 2009) Structural element

Dimension (nm)

Length of unit cell in crystalline cellulose I Elementary fibril (36 cellulose chains) Average pore diameter of viscose, modal fibre (ISEC) Average pore diameter of cotton fibre (ISEC) Microfibrils Macrofibrils Fibre Cellulase

1.0 3.5 2.3–2.9 4.1 10–30 60–360 10 000–30 000 2 × 11 to 4 × 25


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Advances in textile biotechnology Glucose

Cellobiose and small chain oligomers b-Glucosidase CBH

EG

Crystalline regions

Amorphous regions

13.2 Reaction scheme of enzymes on insoluble cellulose substrates.

three-dimensional structure of a fabric and the accessibility of the yarn used, the fibre structure and porosity will determine the ability of the cellulose binding domain to couple to the surface of the material. Fibre structure on the molecular scale and polymer organisation in the material thus directly influences the observed activity of EG and CBH on a given substrate. For textile processing, the strength loss that correlates with weight loss occurring during the hydrolysis process is most critical (Kumar and Purtell, 1994). Although fabric construction, yarn twist, type of enzyme and process conditions can be controlled easily, every preceding step applied during textile processing has to be considered carefully with regard to possible modification in fibre properties. Any change in fibre reactivity will also modify hydrolysis rate and thus overall weight loss. Carrillo et al. studied the cellulose hydrolysis rate of various viscose, modal and lyocell fibres by analysis of the formation of reducing sugars in the treatment baths. The highest concentration of reducing sugars was found by hydrolysis of viscose fibres, followed by modal fibres, whereas lyocell fibres showed the lowest rate of hydrolysis. Fibrillated lyocell fibres showed a higher degradation rate, most probably because of the higher accessibility and surface area available in the fibrillated fibre (Carrillo et al., 2003a). Lyocell fibres which had been defibrillated by cellulase treatments also exhibited a higher rate of dye uptake, which was explained by the increased accessibility of the fibre structure owing to morphological and structural changes (Carrillo et al., 2003b). Study of the protein sorption on the fibres showed the highest sorption for viscose, followed by modal and lyocell. Although the protein sorption on modal fibres is only slightly lower than on viscose fibres, a much lower hydrolysis rate was observed for modal fibres. Thus, in addition to the avail-



319

able surface and protein sorption, the surface reactivity of the swollen cellulose structure also has to be considered. Restructuring of the cellulose fibre surface will change the reactivity for cellulose hydrolysis with cellulases. During textile processing e.g. pretreatment, dyeing and finishing, goods are treated in aqueous solutions of auxiliaries at elevated temperatures. Many steps involve an intermediate drying of the wet material, which can be thought of as a steam and dry heat treatment. Even during consumer use, consecutive wet/dry cycles are applied in from of wear and household wash cycles, which are known to change fibre sorption properties such as water retention value, moisture sorption or iodine sorption (Siroka et al., 2008). As the heterogeneous hydrolysis reaction with cellulases involves the sorption and binding of the cellulose binding domain, any change of cellulose surface can be expected to modulate the cellulose hydrolysis rate for a given substrate.

13.5

Restructuring by heat and steam treatment

Reducing sugars, cG (g l–1)

The influence of fibre morphology and cellulose polymer chain order in the fibre on the hydrolysis rate of cellulose fibres is shown in Fig. 13.3. In the experiments shown in Fig. 13.3, the production of reducing sugars as function of hydrolysis time has been studied for cotton (cellulose I crystal structure), lyocell (cellulose II structure, NMMO process), viscose (cellulose II structure, xanthogenate process) and modal (cellulose II structure, modified xanthogenate process).

20

15

10

5

0 0

2

4 Time (h)

6

8

13.3 Production of reducing sugars as a function of hydrolysis time. Liquor ratio 1 : 25; 0.96 g l−1 protein (30 ml l−1 of enzyme solution); T = 55 °C; 䊏, cotton; 䊉, lyocell; 䉱, modal; 䉲, viscose; cG, concentration of reducing sugars (glucose) in g l−1.


320


The highest degradation rate is observed for viscose. This can be attributed to several fibre properties which favour sorption and access of cellulases. As shown in Table 13.1, viscose fibres show the highest expansion in the wet state, resulting in water retention values of 0.88–0.89 ml g−1; thus in the wet state, the fibre structure is highly expanded and reactive. Furthermore, owing to their multi-lobal shape, viscose fibres offer the highest fibre surface for protein sorption. As the molecular size of proteins prevents their access into the pore system inside the cellulose fibres, the effective fibre surface influences directly the degradation rate. Owing to their oval cross section, lyocell and modal fibres offer substantially lower surface for cellulase adsorption and, as a result, lower degradation rates are observed. The pore surface determined by ISEC (Table 13.1) is of minor influence on the degradation rate as cellulases are too big to access the small pores inside the fibres. Schimper et al. (2004) studied the influence of dry heat treatments and steam treatment on the hydrolysis rate of cotton, lyocell viscose and modal fibres. In this study, the highest production of reducing sugars was observed with viscose fibres, which exceeded 20 g l−1 glucose after 8 h hydrolysis at 55 °C, with a liquor ratio of 1 : 25 and a concentration of 30 ml l−1 of a commercial total crude cellulose (Fig. 13.4). For the same hydrolysis time approximately half of the concentration of glucose was observed for hydrolysis of cotton. An even lower hydrolysis rate was observed for modal fibres and lyocell had lowest degradation rate.

Reducing sugars, cG (g l–1h–1)

3.0 2.5 2.0 1.5 1.0 0.5 0.0 Control

100

120 140 160 Temperature (°C)

180

200

13.4 Concentration of reducing sugars cG formed per hour as a function of hydrolysis time for different fibres after dry heat treatment at different temperatures. Liquor ratio 1 : 25; 0.96 g l−1 protein (30 ml l−1 of enzyme solution); T = 55 °C; 䊏, cotton; 䊉, lyocell; 䉱, modal; 䉲, viscose; cG, concentration of reducing sugars (as glucose) in g l−1.



321

Determination of the protein concentration in solution did not show a correlation between the adsorbed protein and total hydrolysis rate. When the hydrolysis experiment was performed after a dry heat treatment, an increase in cellulase hydrolysis rate was observed for all fibres at 100–130 °C. For a 45 s steam treatment at 100–130 °C, a remarkable increase in hydrolysis rate appears for all fibres tested. The thermal treatment, particularly steaming, thus causes an activation of the cellulose surface. At higher temperatures, e.g. 160–190 °C, dry heat treatment leads to a deactivation of the fibre surface, a phenomenon often called ‘hornification’. Similar results were obtained with steaming processes between 160 and 190 °C, as superheated steam also acts mainly as a drying medium and thus induces hornification of the fibre similar to that with hot air treatment.

13.6

Treatment in alkaline swelling solutions

Treatment in swelling solutions, e.g. alkalisation processes, leads to an intensive reorganisation and activation of the cellulose (Klemm et al., 1998). In mercerisation of cotton fibres, in addition to reorganisation of the fibre, there is also a change in the cellulose crystalline structure from cellulose I to cellulose II structure. In addition to the effects of alkali concentration, the amount of applied tension affected the overall activity during cellulase hydrolysis. Buschle-Diller and Zeronian (1994) compared the influence of slack mercerisation and mercerisation with tension on weight loss and tensile strength loss during enzymatic hydrolysis of cotton. Cellulase hydrolysis occurred at the highest rate in slack mercerised fabric, followed by in tension mercerised fabric. Alkalisation as a whole activates the cellulose fibres for enzymatic hydrolysis, which is attributed to higher accessibility and decreased crystallinity of the fibres after slack mercerisation. Regenerated cellulose fibres show even higher sensitivity towards reorganisation in concentrated alkali solutions, and at certain concentrations even dissolution of the fibre can occur. Viscose fibres exhibit a solubility maximum at approximately 10% (w/w) NaOH solution, and high swelling in alkaline solution. Modal and lyocell-type fibres exhibit higher stability against alkali treatment and thus modal fibres can be mercerised in blends with cotton. Alkalisation of regenerated cellulose fibres can be used to improve dimensional stability, dyeability, fibrillation and pilling behaviour, lustre and smoothness (Bui et al., 2008a; Siroky et al., 2009). The alkalisation process changes accessibility, porosity and sorptive properties for moisture, elemental iodine and dyestuff (Bui et al., 2008b). These modifications are dependent both on the type of alkali used and the concentration applied. Alkalisation of lyocell fibres influences the rate of access and distribution of a fluorescent whitening agent (FWA) through the cross-section of lyocell and modal fibres. After alkalization, higher penetration depth and higher


322


FWA uptake were observed, with more distinct effects being observed for NaOH-treated samples (Öztürk et al., 2009). In particular, regenerated cellulose fibres show remarkable potential for reorganisation of the fibre structure which also alters the cellulose hydrolysis rate. The effect of treating lyocell fabric with 8% NaOH solution on the cellulase hydrolysis rate and the tensile strength loss has been studied by Shin et al. (2000). Fabric samples which had been alkalised under tension were found to exhibit a smaller loss of tensile strength for the same weight loss compared with control samples, this was attributed to the more uniform enzyme reaction. Shin et al. (2000) also studied the progress of enzymatic hydrolysis by scanning electron microscopy. As the enzymatic hydrolysis progressed, fibrils were produced and damage occurred deep inside the fibre. The higher crystallinity, moisture regain and dye adsorption of NaOH pretreated fabrics undergoing the same cellulose treatment conditions can be explained by the extraction of short molecular chains from the amorphous regions, and by change of the amorphous regions into more open structures. Similar changes in fibre morphology owing to swelling processes were also shown for cotton fibres, where a distinct increase in the cellulose hydrolysis rate could be observed after various treatments in alkaline swelling agents (Nicolai and Nechwatal, 2002). Treatment of cotton fabric in 25% (w/w) aqueous solution of NH3, in liquid NH3 at −40 °C, in 2% (w/w) NaOH solution at 100 °C and 300 g l−1 NaOH at 20 °C, all led to an increase in the enzyme hydrolysis rate. Remarkably, almost up to twice the hydrolysis rate was observed for samples that were not dried after the swelling treatment and were used for the cellulose hydrolysis in the wet state. When the samples were dried after the treatments in swelling solution and used as dry samples for the hydrolysis, the activity of the cellulases was still 10–40% higher than for untreated materials. The treatment of cellulose fibres in swelling solutions leads to a substantial change in fibre reactivity and accessibility, similar to the ‘never dried state’ of freshly spun regenerated cellulose fibres before their first drying. During drying, the swollen expanded structure of the fibres collapses and, as a result, the hydrolysis rate decreases. Compared with cotton fibres, the regenerated cellulose fibres show more swelling in water and alkaline swelling agents. Thus, the effects of alkali pretreatment on the hydrolysis rate of cellulases are even more pronounced than for cotton. In an extensive experimental study, Schimper et al. (2009) investigated the influence of alkali pretreatment on the hydrolysis rate of viscose fabric. In addition to the influence of alkali concentration, the effects of intermediate drying were also studied. Figure 13.5 shows the influence of alkali concentration on the hydrolysis rate observed during the first



323

14 60 min 180–240 min

Hydrolysis rate (g l–1h–1)

12 10 8 6 4 2 0 0

1

2

3

4

5

Liq. NH3

[NaOH] (mol l–1)

13.5 Cellulose effect of alkali pretreatment on hydrolysis rate of viscose fabric as function of alkali concentration, hydrolysis activity given as concentration of glucose formed per hour and per litre (Liq. NH3 indicates pretreatment in liquid ammonia) (Schimper et al., 2009).

60 min of cellulase treatment and during the 4th hour of hydrolysis, of samples, which were treated in the ‘never dried state’ without intermediate drying. During the first 60 min of treatment, the distinct increase in hydrolysis rate can be explained by the activation of the viscose fibre, obtained during the preceding alkalisation. Thus, the maximum activation is observed above a NaOH concentration of 2.3 mol l−1, near the swelling maximum reported for viscose fibres. For interpretation of the activation effects in 3.5 and 4.9 mol l−1 NaOH, it has to be considered that a fibre treated in these solutions passes the swelling maximum twice, during impregnation and during the washing procedure. Samples that were dried before enzymatic hydrolysis showed a considerably lower hydrolysis rate. Both line-dried and freeze-dried samples were hydrolysed much more slowly than samples hydrolysed directly in the wet state without drying (Table 13.3). The distinct effect of activation is explained by the creation of more easily accessible cellulose regions owing to enlarged pores and a greater inner surface area. Intensive structural reorganisation of the fibres makes the surface of the fibres more susceptible to cellulase sorption and hydrolysis reactions. During drying, weakly adsorbed water is removed and a part of the pores formed or expanded during the alkalisation step collapses. As a result, overall cellulose activity decreases.


324


Table 13.3 Enzymatic hydrolysis of pre-alkalised viscose fibres. Weight loss (%) after 4 h treatment for different concentrations of NaOH solution (duration of treatment 1 min) and drying methods (wet = hydrolysis without drying) (Schimper et al., 2009) Control

Wet Line dried Freeze dried

13.7

24.9

NaOH concentration 1.0 mol l−1

2.2 mol l−1

3.5 mol l−1

32.1 26.1 23.6

55.2 43.3 43.2

81.5 52.9 46.9

Technical processing

The strong activation of viscose fibres at higher NaOH concentration is accompanied by a considerable change in fibre properties e.g. stiffness and tensile strength/elongation behaviour, which limits the maximum NaOH concentration for textile processing to values below the swelling maximum. In the production of special embroidery, polyester yarn is embroidered on felt or fabric woven from another material, e.g. polyvinylacohol (PVA), cellulose acetate or cellulose fibres, which can be removed in a subsequent treatment step. PVA can be removed by dissolution in water, cellulose acetate is dissolved in acetone, and cellulose fibres are removed by acid hydrolysis. Dissolution of cellulosic fibres e.g. viscose by cellulase hydrolysis has been proposed as an alternative green process (Döbel, 1999). However, for intensive activation of viscose fibres to achieve complete removal from viscose–polyester blends with enzymatic hydrolysis, concentrations above the swelling limit are favourable (Schimper et al., 2008; Vasconcelos and Cavaco-Paulo, 2006). In a special variation of the two-step process, the activation agent is applied only superficially, e.g. by spraying or printing, thus permitting a localised activation on the surface of a textile. Such effects are of interest to generate surface modifications during garment washing in ring-dyed denim (Schimper et al., 2007).

13.8

Conclusion

Man-made cellulose fibres offer great potential future for applications because they are based on natural resources, which can be processed to give a very wide range of different fibres. In viscose, modal and lyocell fibres, the cellulose polymer shows high chemical reactivity, water sorption and accessibility for lower molecular weight chemicals. During treatment with swelling agents, including water, fibre reorganisation in intercrystalline regions



325

occurs and with use of stronger swelling agents, e.g. alkali solutions, intracrystalline parts begin to reorganise, thus yielding modified fibre properties such as porosity, accessibility and reactivity (Bui et al., 2008b; Jaturapiree et al., 2008). Changes in fibre structure and morphology also influence the observed reactivity of enzymes for cellulose hydrolysis. As these changes are more distinct in regenerated cellulose fibres, than in cotton fibres, the effects of preceding treatments in swelling solutions on enzyme hydrolysis rate are more critical for man-made cellulose fibres. For technical processing, this interlinkage between effects of pretreatment (drying steps, alkali treatments, crosslinking operations, dyeing with bifunctional reactive dyes) and the final result of an enzyme treatment has to be considered to achieve stable and reproducible process conditions.

13.9

References

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Index

α-amylases, 11, 140 α-chitin, 290 α-elastin, 255 α-factor, 40 AATCC 79 water drop test method, 142 acetate fibres, 104 acetic acid system, 104 Acetobacter, 291, 299 Acetobacter xylinum, 291, 304–5 acoustic cavitation, 53 acrylics, 100–1 actin, 156–7 Activa, 197, 198 adipic acid bishexylamide, 86 adsorption–desorption process, 50 affinity tags, 244 Agaricus bisporus, 205, 206, 213, 214, 215, 225 Agrobacterium tumefaciens, 112, 155 alkaline scouring, 139, 140, 144 alkalisation, 321 amidases, 87 amylases, 10–12, 140, 141 AmyUS100ΔIG mutant, 11–12 apyrases, 157 Arabidopsis thaliana, 152, 253 arabinogalactan proteins, 155 aromatic polyesters, 78 Aspergilli, 241 Aspergillus nidulands, 256 Aspergillus oryzae, 11 atom-transfer radical polymerisation, 280 Autographa californica, 242 avidin, 62 Avogadro number, 48 2,2′-azinobis-(3-ethylbenzthiazoline-6sulphonic acid), 82

Bacillus amyloliquefaciens subtilisin, 16 β-chitin, 290 BAA see Bacillus amyloliquefaciens α-amylase Bacillus amyloliquefaciens α-amylase, 10 Bacillus lentus, 16 Bacillus licheniformis α-amylases, 10 Bacillus sp., 11, 15 Bacillus subtilus, 141 Bacillus thuringiensis cotton, 153 bacterial amylases, 141 bacterial cellulose advantages and limitations, 303–4 applications, 302–3 –chitosan blends, 303 future trends, 304–5 processing developments for textile, 288–305 production, 297–9 structure and properties, 291–3 Acetobacter and wood pulp cellulose production, 292 bast fibres, 151 BASYC, 302 biocatalysis, 98 biofabrication, 200–1 Biofill, 302 biomolecule capture, 279 biopolymer conjugation, 211–13 BioPrep 3000L, 141 bioprocessing, 15, 302 bioremediation, 210 bioscouring, 141, 143 biosensing, 209–10 biotechnology, 194 biotin, 62 bisulphite, 180

328 © Woodhead Publishing Limited, 2010

Index BLA see Bacillus licheniformis α-amylases bleaching, 33, 145 Boehmeria nivea, 151, 156 boiling off see scouring Bombyx mori, 182, 183, 184, 193, 242, 248 Brunauer-Emmet-Teller value, 315 byssus threads, 254 caffeoyl coenzyme A O-methyl transferase, 158 calcium hydroxide, 138 carbohydrate-binding modules, 63, 121–2 carboxyl ester hydrolases, 18 catalase, 59 CBM see cellulose-binding molecule cell membrane complex, 193 cell wall, 151 cellobiohydrolase, 316, 317 cellobiohydrolase I, 122 cellulases, 13–15, 143, 315–16 Cellulon fibre, 299 cellulose, 312 cellulose acetate, 59 cellulose acetate fibres basic concept and principles, 100–6 anhydroglucose repeating unit, 104 cellulose acetate biodegradation main reactions, 106 side chains as substrates for enzymes, 104–6 biomodification, 120–4 advantages and limitations, 123–4 cellulose diacetate and triacetate K/S values, 123 esterases, 120–3 recombinant wild-type cutinase, 122 enzymatic modification for textile and other applications, 98–125 fibre biomodification assessment current methodologies, 106–9 future trends, 124–5 textile chemical finishers distribution, 100 cellulose acetates, 104 cellulose-binding domain– β-glucuronidase protein, 15 cellulose-binding molecule, 14 cellulose diacetate, 104, 120

329

cellulose fibres cellulase hydrolysis, 316–19 fibre elements and cellulases average dimensions, 317 reaction scheme of enzymes on insoluble cellulose substrates, 318 enzymatic functionalisation for textiles, 266–81 future trends, 281 native xyloglucans in cellulose modification, 270–1 xyloglucans, 268–70 xyloglucans chemo-enzymatic modification, 272–81 hydrolysis for textile and other applications, 312–25 cellulase reactivity, 313 cellulases, 315–16 technical processing, 324 regenerated, 314–15 representative fibre data, 315 restructuring by heat and steam treatment, 319–21 reducing sugars as a function of hydrolysis, 319 reducing sugars concentration, 320 treatment in alkaline swelling solutions, 321–3 alkali concentration effect on hydrolysis rate, 323 pre-alkalised viscose fibres enzymatic hydrolysis, 324 cellulose triacetate, 104, 120 CeloxTM, 300 cetyltrimethylammonium bromide, 179 chemotherapy, 66 chimeric enzyme, 15 chitin, 289 advantages and limitations, 303–4 applications, 300–2 fibres production, 293–7 electrospuns and commercial microfibres, 298 future trends, 304–5 processing developments for textiles, 288–305 structure and properties, 289–90 molecular structure, 290 chitin affinity chromatography, 301 chitosan, 91, 200, 211, 289 advantages and limitations, 303–4 applications, 300–2


330

Index

–bacterial cellulose blends, 303 fibres production, 293–7 electrospuns, 297 future trends, 304–5 processing developments for textiles, 288–305 structure and properties, 291 molecular structure, 290 chitosan blends, 303 ChitoSkin, 300 chymotrypsin, 185 cinnamyl alcohol dehydrogenase, 157 circular dichroism, 246 clone, 239 cloning sites, 239 collagen scaffolds, 198 collagens, 198, 253–5 comonomers, 101 composites, 271 concatemerisation, 238 conifers, 151 conventional chemical processing cotton properties effect, 139 environmental problems, 139–40 vs enzymatic treatment of cotton, 132–45 cotton fibre chemistry and structure, 133–5 cotton seed coat fragments, 135–7 enzymatic treatments, 140–4 future trends, 144–5 copolymerisation, 81 cortex cells, 193 cotton, 322 see also Gossypium hirsutum bioscouring, 141–4 effect on cotton properties, 143–4 conventional chemical treatments and conditions, 138–40 cotton properties effect, 139 environmental problems, 139–40 enzymatic treatment vs conventional processing, 132–45 enzymatic treatments, 140–4 costs comparison, 144 cotton seed coat fragment removal, 142–3 desizing, 140–1 mechanical and sonic power application to increase mass transfer, 143 fibre chemistry and structure, 133–5 cellulose and non-cellulosic constituents connections, 134

cellulose structure, 133 cotton structure model, 134 cotton wax composition, 136 future trends, 144–5 seed coat fragments, 135–7 fragments in fabric and under magnification, 135 structure, 138 xylan structure, 137 cotton pre-treatment, 33–5 discontinuous pre-treatment, 34–5 mechanical energy representation, 35 full-continuous pre-treatment, 33–4 pad batch–hot bleach process, 34 cresolase, 205 cuticle cells, 193 cutinase, 120 cutinases, 90, 173 Dacron, 58 degumming, 172, 182–3, 186 Denilite II Assist, 82 desizing, 33, 144 dextran, 65 dibutyryl chitin, 300 differential frictional effect, 174 diffusion, 64 diffusion controlled systems, 64 direct ELP tagging, 245 direct nesslerisation method, 107 DNA cloning, 4 DNA ligases, 4, 5 DNA oligomerisation, 238 DREBIA, 152 Duramyl, 11 Durazyme, 16 dynamic light scattering, 246 EDTA see ethylenediamine tetra-acetic acid elastic protein fibres developments for textile and other applications, 235–59 elasticity, 246 elastin, 255–6 elastin-like polymers, 17, 245, 255 elastomeric proteins, 247 electron paramagnetic resonance, 221 electroporation, 6 electrospinning, 247–8, 255, 294, 296, 304 schematic representation, 295


Index elementary fibre, 151 ELP see elastin-like polymers ELP-mediated affinity capture, 245 endoglucanase C, 122 endoglucanases, 316, 317 energy dispersive x-ray spectroscopy, 109 5-enolpyruvylshikimate-3-phosphate synthase, 153 ENZUP, 181 enzymatic finishing, 177–82 enzymatic functionalisation cellulose fibres for textile and other applications, 266–81 chemo-enzymatic modification of xyloglucans, 272–81 future trends, 281 native xyloglucans in cellulose modification, 270–1 xyloglucans, 268–70 enzymatic hydrolysis core polymer fibres for textile and other applications, 77–91 advantages and limitations, 90–1 applications, 90 basic principles, methods and technologies, 80–90 future trends, 91 polyesters, 78–9 enzymatic modification core polymer fibres for textile and other applications, 77–91 advantages and limitations, 90–1 applications, 90 basic principles, methods and technologies, 80–90 future trends, 91 polyamide, 79–90 polyesters, 78–9 polyacrylonitrile and cellulose acetate fibres for textile and other applications, 98–125 basic concept and principles, 100–6 cellulose acetate biomodification, 120–4 fibre biomodification assessment current methodologies, 106–9 future trends, 124–5 polyacrylonitrile biomodification, 109–20 textile chemical finishers distribution, 100 enzymatic scouring, 144, 175–7 enzymatic textile treatments

331

adsorption limitation in textile pores, 47–52 enzymes adsorption at a substrate surface, 49 enzymes adsorption dynamics, 52 modelling, 49–52 cotton pre-treatment current processes, 33–5 discontinuous pre-treatment, 34–5 full-continuous pre-treatment, 33–4 mechanical energy representation, 35 pad batch–hot bleach process, 34 future trends, 53–4 mass transfer limitations in textiles, 36–8 diffusion equation graphical solution, 38 fabrics inter- and intra-yarn pores, 36 yarn model, 37 mass transfer model, 43–7 calculations results, 47 processes and equipment developments, 32–54 squeezing factor α relationship with pressure force, 40 short-cut method for determination, 42 squeezing model, 39–43 enzyme transport into a fabric, 42, 43 fabric enzyme concentration, 41 pore velocities and deformation rate relationship, 41 textile slab model, 39 ultrasound application, 52–3 effect on enzymes performance, 53 enzymatic treatment application to silk fibres, 182–6 Bombyx mori fibroin heavy chain gene, 184 sericin properties and composition, 184 application to wool fibres, 174–5, 174–82 enzymatic finishing, 177–82 enzymatic scouring, 175–7 wool cuticle, 175 wool fibre micrograph, 174 bioscouring, 141–4 effect on cotton properties, 143–4 costs comparison, 144


332

Index

cotton seed fragment removal, 142–3 desizing, 140–1 enzymes for processing protein fibres, 172–4 peptidases classification, 173 mechanical and sonic power application to increase mass transfer, 143 vs conventional chemical processing of cotton, 132–45 conventional chemical treatments and conditions, 138–40 cotton fibre chemistry and structure, 133–5 cotton seed coat fragments, 135–7 future trends, 144–5 wool and silk fibres, 171–87 future trends, 186–7 enzyme immobilisation, 58, 59–63 on fabrics, 59–61 strategies, 60, 61–3 biotinylated enzymes immobilisation, 64 immobilisation on polyamide, 63 enzyme ligase, 3 enzymes, 10, 98–9 design and engineering for textile applications, 3–22 protein fibres processing, 172–4 enzymes engineering amylases, 10–12 α-amylases–Termamyl LC, 12 cellulases, 13–15 design for textile applications advantages and limitations, 19–21 future trends, 21–2 rDNA molecular cloning basic principles, 3–6 enzyme production, 7–8 lipases/esterases, 18–19 Fusarium solani pisi cutinase, 20 pectinases, 13 proteases, 15–18 Eri silkworm, 182 Escherichia coli, 7, 59, 240, 256, 304 Esperase, 176 ethylene, 157 ethylenediamine tetra-acetic acid, 138 Eudragit polymer, 181 exoglucanases, 316, 317

fermentation-processed chitin, 302 ferments, 98 fibre biomodification cellulose acetate, 120–4 advantages and limitations, 123–4 cellulose diacetate and triacetate K/S values, 123 esterases, 120–3 recombinant wild-type cutinase, 122 current methodologies for assessment, 106–9 polyacrylonitrile, 109–20 acrylic fibres cross-section, 118 acrylic K/S values, 111 advantages and limitations, 115–20 conditions and major results from published treatments, 116–17 nitrile-metabolising enzymes, 109–15 physical characteristics after treatment with nitrile hydratase, 111 surface erosion model, 114 total nitrogen and PAA concentration, 113 working solution ingredients, 111 fibres see also specific fibre quality improvement by plant engineering, 150–63 fibrohexamerin, 194 fibroin, 182, 183, 184, 185, 194, 215, 253 Fick’s diffusion equation, 37 filling, 201 FITC see fluorescein isothiocyanate Flavobacterium sp, 89 flax, 157–9 flourescein microscopy, 121 fluorescein isothiocyanate, 115 fluorescence spectroscopy, 246 fluorescent molecules, 90 food processing, 210–11 France Chitine, 293 fuco-galacto-xyloglucans, 268 fungal cellulases, 13 Fusarium solani, 19 Fusarium solani pisi, 83, 86, 87 Fusarium spp., 157 fusion tags, 245

fall armyworm see Autographa californica

G. barbadense, 133 G. hirsutum, 133


Index galactose oxidase, 272–3 gas chromatography, 107 gelatine, 198 genetically modified organisms, 155 Gengiflex, 302 gentamicin, 67 Geobacillus stearothermophilus US100 strain, 11 GhADF1 gene, 156 GIRD-CT-2002-00695, 181 glow discharge techniques, 86 glucanases, 157 glucose oxidase, 61 glutamine, 202, 204 glutathione S-transferase tag, 241 Gossypium, 133 Gossypium hirsutum, 156–7 Hansenula, 241 HemCon, 300 Heppe Biomaterial, 293 homologous recombination, 9 hornification, 321 horseradish peroxidase, 59, 219 redox cycle, 220 Humicola insolens Ce17B endoglucanase, 14 Humicola lanuginosa, 19 hydrogels, 65 hydrolases, 89 1-hydroxybenzotriazole, 82, 88 hydroxyethylcellulose, 64 Hymenaea courbaril, 269 hypermannosylation, 241–2 immobilised metal affinity chromatography, 245 insulin, 65 inter-yarn pores, 36 inverse temperature transition, 255–6 inverse transition cycling, 245 isothermal titration calorimetry, 246 iterative technique, 237 jatoba see Hymenaea courbaril jet, 34 jigger, 34 Kevlar, 249 KiOmedine, 300 KiOnutrime-Cs, 300 KitoZyme, 293

333

l-dopa, 209 Laccase Acid Red 1, 220 laccases, 61, 82, 90, 218–19 redox cycle, 218 LAMY α-amylase, 11 leather processing, 201 Leu181Ala mutant, 19 lignin, 137, 159 linear polyester, 78 lipases, 173 lipases/esterases, 18–19 lipid transfer proteins, 155 Lock and Key Model, 98 low-temperature plasma treatment, 179 lyocell fibres, 314, 320 lysozyme, 58 Marseilles soap, 183 mass transfer, 119 limitations in textiles, 36–8 diffusion equation graphical solution, 38 fabrics inter- and intra-yarn pores, 36 yarn model, 37 model, 43–7 calculation results, 47 Maxapem, 16 mechanical agitation, 119 mega-primer method, 8 Merino fleece, 174 Met197Ala, 12 modal fibres, 314, 320 molecular genetic tools, 121 monooxygenase, 205 Muga silkworm, 182 mulberry silk, 182 mulberry silkworms see Bombyx mori Muta-Gene M13, 9 Mutagenesis Kit, 9 nanofibres, 247, 295 NaOH, 138 naringinase, 59 nasturtium see Tropaeolum majus non-Fickian behaviour, 65 non-proteolytic enzymes other enzymes for protein fibre functionalisation, 218–23 horseradish peroxidase redox cycle, 220 laccase redox cycle, 218 transglutaminases, 195–201


334

Index

applications, 197–201 biological properties and biotechnological utility, 195–7 catalysed reactions, 195 tyrosinases, 205–13 applications, 209–13 biological properties and biotechnological utility, 205–8 protein-bound tyrosine-catalysed oxidation, 206 wool and silk fibres functionalisation, 193–226 future trends, 223–6 using transglutaminases, 201–5 using tyrosinases, 213–18 nuclear polyhedrosis virus, 242 nylon-degrading peroxidases, 88 nylon fibres, 79 orb-web spinning spiders, 248 organophosphorus hydrolase, 58 origin of replication, 239 orthodox type II endonucleases, 4 oxidases, 88 oxidoreductases, 194 Paenibacillus sp., 301 PAN see polyacrylonitrile particle bombardment method, 156 Peclet number, 40 pectate lyases, 145 pectin, 142 pectinases, 13, 141, 143 pectinolytic enzymes, 13 peptidases, 173 peroxidases, 218, 219 PET see poly(ethylene terephthalate) Pichia, 241 Pichia pastoris, 7, 241, 253, 256, 275 plant fibres, 151–5 fibre improvement, 152–3 fibre quality and genes, 153–5 plants engineering defining plant fibres, 151–5 fibre improvement, 152–3 fibre quality and genes, 153–5 engineering examples in fibre species, 156–61 cotton, 156–7 fibre qualities improvement, 150–63 future trends, 161–2 methods, 155–6 some engineering examples in fibre species

flax, 157–9 novel properties introduction, 159–61 plasma treatment, 62, 81 poly-β-hydroxybutyrate, 159 polyacrylonitrile basic concept and principles, 100–6 acrylic polymer structure, 102 side chains as substrates for enzymes, 100–4 biomodification, 109–20 acrylic fibres cross-section, 118 acrylic K/S values, 111 advantages and limitations, 115–20 conditions and major results from published treatments, 116–17 nitrile-metabolising enzymes, 109–15 physical characteristics after treatment with nitrile hydratase, 111 surface erosion model, 114 total nitrogen and PAA concentration, 113 working solution ingredients, 111 chemical transformations, 103 enzymatic modification for textile and other applications, 98–125 fibre biomodification assessment current methodologies, 106–9 future trends, 124–5 textile chemical finishers distribution, 100 polyamide, 79–80 advantages and limitations, 90–1 applications, 80, 90 compositions, 80 enzymatic hydrolysis and modification, 77–91 fibres modification, 85–90 cutinase single exchanges in amino acids, 88 methods to quantify enzymatic hydrolysis, 89–90 nylon oligomers degradation, 89 polyamide 6.6 enzymatic hydrolysis, 87 future trends, 91 properties, 80 polyesters, 78–9, 139, 324 poly(ethylene terephthalate), 78–9 advantages and limitations, 90–1 applications, 79, 90


Index

335

QuickChange kit, 9 quinones, 205

examples and applications, 247–58 collagens, 253–5 elastin, 255–6 fibrous materials mechanical properties, 252 polymer nanofibres applications, 248 resilin, 257–8 silks, 248–53 spider silks, 250–1 experimental characterisation, 245–7 expression systems, 239–44 bacteria, 240–1 fungi, 241–2 in vitro systems, 243–4 insects, 242–3 mammalian cells, 243 plants, 242 future trends, 259 purification, 244–5 rDNA methods principles applied in production, 236–7 secondary structural motifs in proteins, 237 recursive directional ligation, 238 regenerated cellulose fibres, 314–15 cellulase hydrolysis, 316–19 resilin, 257–8 restriction endonucleases, 4 restriction site, 4 retting, 158 ribonucleic acid, 3 ring-opening polymerisation, 280

ramie see Boehmeria nivea raw cotton fibres, 133 reaction engineering, 87 recombinant DNA host cell transformation, 6 molecular cloning basic principles, 3–6 obtaining, cutting and joining the DNA, 4–6 restriction endonucleases cleavage patterns, 5 recombinant expression systems, 239–44 recombinant proteins advances and limitations in biosynthetic production, 258–9 biomimetic design, 237–8 DNA oligomerisation techniques, 238

S. faecalis, 59 Saccharomyces, 241 Saccharomyces cerevisiae, 256 Salmonella, 304 sand washing, 186 scales, 193 scouring, 33, 138, 141 seed coat fragments, 135, 143 selection markers, 239 sericin, 182, 185, 194, 215 properties and composition, 184 sericin gum, 182, 183 shrink-resist finishing, 174 silk, 182, 248–53 biomaterials, 249 developments for textile and other applications, 235–59 enzymatic treatment, 171–87 application, 182–6

enzymatic hydrolysis and modification, 77–91 fibres modification, 80–5 analytical techniques, 84–5 cutinase, 82–4 enzymatic hydrolysis, 83 laccase, 82 future trends, 91 properties, 79 polyhistidine tag, 241 polyhydroxyalkanoate, 63 polylinkers, 239 polymer nanofibres, 248 polyphenol oxidases, 205 polyvinyl alcohol, 140 polyvinylalcohol hydrogel, 67 Primex, 293 procambium, 157 proline, 19 promoter, 239 protease enzymes, 186 proteases, 15–18, 181 protein disulphide isomerases, 222 proteins adsorption, 90–1 site-specific modification, 199–200 tagging, 212–13 proteolytic enzymes, 172, 178 PROTEX project, 181 PttXET16–34, 275 Purastar OxAm, 11


336

Index

enzymes used for protein fibre processing, 172–4 future trends, 186–7 functionalisation using nonproteolytic enzymes, 193–226 future trends, 223–6 other enzymes, 218–23 protein fibres functionalisation using transglutaminases, 201–5 protein fibres functionalisation using tyrosinases, 213–18 transglutaminases, 195–201 tyrosinases, 205–13 silk-elastin like proteins, 256 silk fibroin filament, 182 silk-like proteins, 253 silk proteins, 253 silkworm see Bombyx mori smart materials, 56–67 containing enzymes or enzyme substrates functional packaging materials, 59 functional textiles, 58–9 wound dressings, 58 enzyme immobilisation, 59–63 biotinylated enzymes immobilisation, 64 immobilisation on polyamide, 63 on fabrics, 59–61 strategies, 60, 61–3 future trends, 67 responding to enzymes as triggers, 63–7 active agent release, 65 active agent release triggered by different external conditions, 66 controlled-release systems and mechanisms, 63–6 enzyme substrates covalent attachment, 66–7 soda ash see sodium carbonate sodium carbonate, 138 sodium carboxymethylcellulose, 64 sodium salt, 180 sol-gel transition, 246 spider silks, 249, 250–1 squeezing model, 39–43 Staphylococcus aureus, 59 starches, 140 Streptomyces mobaraensis, 196, 197, 204 structure-guided protein engineering, 14 subtilisin BPN see B. amyloliquefaciens subtilisin

subtilisin Carlsberg, 16 subtilisinE-VPAVG, 18 subtilisins, 15–18 sulfydryl oxidases, 222 surface plasmon resonance spectroscopy, 245 surfactant, 138 swelling-controlled mechanism, 65 synthetic hydrogels, 66 T4 DNA ligase, 6 tamarind see Tamarindus indica tamarind kernel powder, 269, 270 Tamarindus indica, 269 technical fibre, 151 Termamyl LC, 11 terminator, 239 Terylene, 79 textile pores adsorption limitation, 47–52 enzymes adsorption at a substrate surface, 49 enzymes adsorption dynamics, 52 modelling, 49–52 textiles advantages and limitations, 19–21 amylases, 10–12 α-amylases–Termamyl LC, 12 cellulases, 13–15 cellulosic fibres enzymatic functionalisation, 266–81 functional plant polysaccharides, 268–70 future trends, 281 native xyloglucans technical uses, 270–1 xyloglucans chemo-enzymatic modification, 272–81 chitosan, chitin and bacterial cellulose processing developments, 288–305 advantages and limitations, 303–4 basic principles, methods and technologies, 293–303 future trends, 304–5 types of new fibres, 289–93 core polymer fibres enzymatic hydrolysis and modification, 77–91 enzyme production, 7–8 enzymes design and engineering applications, 3–22 enzymes engineering, 8–10


Index directed evolution, 9–10 site-directed mutagenesis, 8–9 future trends, 21–2 lipases/esterases, 18–19 Fusarium solani pisi cutinase, 20 pectinases, 13 polyacrylonitrile and cellulose acetate fibre enzymatic modification, 98–125 proteases, 15–18 recombinant DNA molecular cloning basic principles, 3–6 host cell transformation, 6 obtaining, cutting and joining the DNA, 4–6 restriction endonucleases cleavage patterns, 5 recombinant silk and other elastic protein fibres developments, 235–59 advances and limitations of recombinant proteins, 258–9 biomimetic design of recombinant proteins, 237–8 examples and applications, 247–58 expression systems, 239–44 future trends, 259 recombinant DNA methods in production of recombinant proteins, 236–7 recombinant proteins experimental characterisation, 245–7 recombinant proteins purification, 244–5 TGase see transglutaminases thrombin functionalised fabrics, 58 thrombin-sensitive peptide, 67 time-of-flight secondary ion mass spectrometry, 178 tissue engineering, 198–9 Trametes hirsuta, 211 transcriptome approach, 154 transferases, 194 transformation, 3, 6 transgene, 155 transglutaminases, 3, 173, 181–2, 196, 224–5 applications, 197–201 biofabrication, 200–1 leather processing, 201 proteins site-specific modification, 199–200 tissue engineering, 198–9

337

biological properties and biotechnological utility, 195–7 catalysed reactions, 195 protein fibres functionalisation, 201–5 TRI auto porosimeter, 142 triacylglycerol hydrolases, 18 Trichoderma, 241 Trichoderma reesei, 207, 210–11, 211, 215, 225, 316 trifluoroacetic acid, 296 Triton X-100, 82 Tropaeolum majus, 269 tropoelastin, 255 Tussah silkworm, 182 tyrosinases, 61, 224–5 applications, 209–13 biopolymer conjugation, 211–13 biosensing and bioremediation, 209–10 food processing, 210–11 l-dopa production, 209 biological properties and biotechnological utility, 205–8 catalysed oxidation, 206 fungal, 208 protein fibres functionalisation, 213–18 ultrasound, 52 urease, 61 van der Waals forces, 105 vector, 239 vinylsulphone dyes, 108 viscose fibres, 314 wild silk, 182 winch, 34 wool, 201 enzymatic treatment, 171–87 application, 174–82 enzymes used for protein fibre processing, 172–4 future trends, 186–7 functionalisation using nonproteolytic enzymes, 193–226 future trends, 223–6 other enzymes, 218–23 protein fibres functionalisation using transglutaminases, 201–5 protein fibres functionalisation using tyrosinases, 213–18


338

Index

transglutaminases, 195–201 tyrosinases, 205–13 wool fibre, 174 wound dressing, 67 x-ray crystallography, 246 XET see xyloglucan endotransglycosylase XGO see xylogluco-oligosaccharide xylem, 151 xyloglucan and XET as toolkit for cellulose modification, 273–81 amino-modified XGO chemoenzymatic synthesis, 276 biomolecule capture, 279 functionalised xyloglucan, 277–9 molecular probes and targets, 280 multivalent effects, 279–81 use, 274 XET biological role, 273 XET/XG method to retain optical brightening agent, 278

cellulosic fibres enzymatic functionalisation, 266–81 chemo-enzymatic modification, 272–81 activation with galactose oxidase, 272–3 functional plant polysaccharides family, 268–70 seeds, 268–70 structure, 269 vegetative cell wall, 268 technical uses in cellulose modification, 270–1 composites, 271 textiles and papermaking, 270–1 xyloglucan endo-transglycosylase, 267, 273–81, 275 biological role, 273 use for cellulose modification, 274 xyloglucan polysaccharide, 275 xylogluco-oligosaccharide, 268, 275, 276 yield points, 247 Young’s modulus, 246


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