Protein-Based Films And Coatings

PROTEIN-BASED FILMS and COATINGS Edited by Aristippos Gennadios, Ph.D. CRC PR E S S Boca Raton London New York Washin...

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PROTEIN-BASED FILMS and COATINGS Edited by

Aristippos Gennadios, Ph.D.

CRC PR E S S Boca Raton London New York Washington, D.C.

TX079 disclaimer Page 2 Thursday, December 20, 2001 2:01 PM

Library of Congress Cataloging-in-Publication Data Catalog record is available from the Library of Congress

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. 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 or for the consequences of their use. 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 prior permission in writing from the publisher. All rights reserved. Authorization to photocopy items for internal or personal use, or the personal or internal use of specific clients, may be granted by CRC Press LLC, provided that $1.50 per page photocopied is paid directly to Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923 USA. The fee code for users of the Transactional Reporting Service is ISBN 1-58716-1079/02/$0.00+$1.50. The fee is subject to change without notice. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. The consent of CRC Press LLC 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 CRC Press LLC for such copying. Direct all inquiries to CRC Press LLC, 2000 N.W. Corporate Blvd., Boca Raton, Florida 33431. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe.

Visit the CRC Press Web site at www.crcpress.com © 2002 by CRC Press LLC No claim to original U.S. Government works International Standard Book Number 1-58716-107-9 Printed in the United States of America 1 2 3 4 5 6 7 8 9 0 Printed on acid-free paper

Table of Contents

Introduction 1 Definitions, Functions, and Rationale 2 Film and Coating Composition 6 Film and Coating Formation 9 Properties of Protein Films and Coatings 10 Protein Film and Coating Application Opportunities Summary and Conclusions 31 References 32

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Introduction 43 Zein Structure 44 Physicochemical Properties of Zein 46 Glass Transition Temperatures 46 Zein Coatings 47 Zein Films 48 Resin Films 53 Conclusions 62 References 63

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Introduction 69 Composition, Structure, and Thermal Properties of Wheat Gluten Shaping Process of Wheat Gluten Materials 79 Postproduction Treatments of Wheat Gluten Films 95 Functional Properties of Wheat Gluten-Based Materials 99 Applications of Wheat Gluten-Based Films and Coatings 114 Conclusions 115 References 115

Introduction 123 Formation of Soy Protein Films 124 Properties of Soy Protein Films 126 Biodegradation of Soy Protein Films 133 Potential Applications and Future Research Needs References 134

Introduction 139 Protein Nature and Recovery 139 Film Formation Techniques and Mechanisms Film Properties 151 Biodegradability 155 Potential Uses 155 Conclusions 156 References 156

Introduction 159 Nature of Major Whey Proteins 160 Whey Protein Recovery 163 Whey Protein Film Formation 164 Whey Protein Film Properties 165 Applications 176

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Table of Contents Research Needs 177 References 177

Introduction 181 Nutritional Value of Casein 181 Chemical and Physical Characteristics of Casein and Caseinates Manufacture of Casein and Caseinates 184 Film Formation Mechanisms 186 Film Functional Properties 189 Uses of Casein Films and Coatings 207 The Future of Casein and Caseinate Films 208 References 209

Introduction 213 Myofibrillar Proteins 214 Physicochemical Properties of Myofibrillar Proteins 217 Formation of Myofibrillar Protein-Based Films 219 Properties of Myofibrillar Protein-Based Films 222 Conclusion 228 References 229

Introduction 233 Film Formation 234 Factors Affecting Mechanical and Barrier Properties Conclusions 249 References 250

Introduction 253 Molecular Characteristics of Keratins 254 Preparation of Reduced Keratins 256 Keratin Films 259 Keratin Microcapsules 267 Conclusions 271 References 271

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Introduction 275 Collagen and Gelatin Structure 276 Physical Properties 278 Rheology of Gelatin Films 290 Gas and Water Vapor Permeability of Gelatin Films 292 Optical Rotation and Circular Dichroism of Gelatin Solutions/Films Gelatin-Based Films 298 References 300

Introduction 305 Peanut Protein 306 Rice Protein 309 Pea Protein 312 Lupin Protein 314 Grain Sorghum Protein 316 Winged Bean Protein 318 Pistachio Protein 319 Cucumber Pickle Brine Protein Feather Keratin 322 Implication 323 References 323

Introduction 329 Water Sorption Isotherm 330 Mass Transfer through Edible Films Mathematical Models 333 Conclusion 336 References 337

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Pharmaceutical Applications of Protein Microparticles 343 Core and Wall Material Properties 345 Microencapsulation Methods 346 Microparticle Release Mechanisms 349 Limitations of Microparticles in Delivering Therapeutic Drugs and Proteins 350 Potential Causes of Protein Coating or Core Denaturation during Microparticle Processing 351 Stabilization of Proteins during Encapsulation 356 Stabilization of Proteins with Cosolvents 357 Summary 359 References 360

Introduction 367 Raw Materials 368 Manufacture of HGC 372 Capsule Filling 376 Drug Release from HGC 381 Conclusion 387 References 387

Introduction 393 Nature and Uses 394 Manufacturing 404 Softgels with Modified Release Characteristics Softgels with Gelatin Extenders or Substitutes Caplets/Tablets Enrobed/Coated with Gelatin References 433

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Introduction 445 History of Collagen Casing Manufacture 446 Collagen Casing Properties 453 Techniques and Procedures for Stuffing Collagen Casings 454 Techniques and Procedures for Smoking/Cooking Collagen Casings Summary 461 Acknowledgments 461 Glossary of Terms (Derro-Teepak, 1996) 461 References 464

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Introduction 467 Application of Edible Films 468 Properties and Types of Edible Films Antimicrobial Films 478 References 480

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Introduction 485 Antimicrobial Edible Films and Coatings 486 Antimicrobial Substances 488 Factors Considered in Designing Antimicrobial Edible Film and Coating Systems 490 Research and Development Trends for Antimicrobial Edible Protein Films and Coatings 496 Conclusion 498 References 498

Introduction 501 Postharvest Fruit and Vegetable Physiology 502 Types of Coatings Used on Fruits and Vegetables 503 Protein Materials Used in Fruit and Vegetable Coatings 504 Factors that Affect Coating Permeability 505 Protein-Based Coatings for Fresh Fruits and Vegetables 507 Protein as an Ingredient of Fruit and Vegetable Coatings 510 Labeling Issues 513 References 513

Introduction 517 Protein Films and Coatings 518 Film Properties 519 Heat and Mass Transfer Application of Coatings on Fried Foods Conclusion 523 References 523

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Importance of Nuts 527 Spoilage of Nuts 529 Traditional Strategies for Maintaining the Quality and Extending the Shelf Life of Nuts 532 Edible Protein Coatings for Maintaining the Quality of Nuts 535 Application of Edible Coatings on Nuts 539 Research Needs 543 Conclusions 544 References 544

Introduction 551 General Information 552 Unique Military Needs 553 Past Studies 555 Recent Studies 561 Future Studies 575 References 575

Introduction 579 Influence of Aroma-Packaging Interactions on Food Quality 580 Control of Flavor Interactions and Transfers by Edible Barriers 581 Aroma Barrier Properties of Edible Films and Coatings 583 Factors Affecting Aroma Transfers through Edible Barriers 588 Conclusion 597 References 598

Introduction 601 The Kosher and Halal Market 603 Kosher Dietary Laws 604 Halal Dietary Laws 607 Dealing with Kosher and Halal Supervision Agencies Gelatin 616 Biotechnology 617

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Table of Contents Kosher and Allergies References 620 Additional Reading

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Introduction 621 Experimental Materials 624 Results and Discussion 626 Extruded Soy Protein Foam Sheets Conclusions 635 References 637

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Preface

NTEREST in the development of films and coatings from renewable biopolymers has been intense during the last decade. Opportunities for adding value to agricultural biopolymers and concerns over the environmental impact of synthetic film materials are perhaps the two most important factors behind this interest. Several applications of biopolymer films and coatings have already been realized, mainly in the area of packaging, while a great number of additional applications are envisioned. Proteins of plant and animal origin, which often are abundantly available as by-products of the food processing industry, are among the biopolymers being used or investigated as feedstocks for the production of films and coatings. In recent years, the scientific literature worldwide has seen an explosion of published articles, often the product of interdisciplinary research, related to protein-based films. Food scientists and technologists, packaging scientists and technologists, protein chemists, and polymer chemists and technologists are among the disciplines involved with research endeavors in this area. It has become apparent that a book that comprehensively reviews the current body of knowledge on the formation, characterization, functional properties, and applications, both existing and potential, of protein-based films and coatings is needed. Protein-Based Films and Coatings aspires to meet this need by bringing together many researchers of varied backgrounds who are recognized leaders in this field. Each chapter was prepared as a comprehensive, stand-alone treatise on the covered topic. Readers will appreciate some inevitable overlap among chapters as an opportunity for exposure to the same issue from different perspectives. To enhance the thoroughness of coverage of the

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subject matter, a host of prominent researchers and professionals in this field provided peer-review for the chapters. It is hoped that newcomers to the challenging and promising field of protein-based films and coatings, as well as experienced researchers, will find valuable information in this book.

Acknowledgments

HE editor is truly indebted to the chapter authors for their fine efforts devoted to this project. He also expresses his gratitude to the following experts who kindly served as peer reviewers, thus further ensuring the accuracy and thoroughness of the covered topics: Erol Ayranci, Akdeniz University, Antalya, Turkey (Chapter 13); Christopher S. Brazel, University of Alabama, Tuscaloosa, AL (Chapter 14); Elena Castell-Perez, Texas A&M University, College Station, TX (Chapter 26); Sam K. C. Chang, North Dakota State University, Fargo, ND (Chapter 22); Luis Cisneros-Zevallos, Texas A&M University, College Station, TX (Chapter 21); David R. Coffin, U.S. Department of Agriculture, Wyndmoor, PA (Chapter 1); Kay D. Cooksey, Clemson University, Clemson, SC (Chapter 19); John Coupland, The Pennsylvania State University, University Park, PA(Chapter 4); Susan L. Cuppett, University of Nebraska, Lincoln, NE (Chapter 20); Jean-Luis Cuq, Université de Montpellier II, Montpellier, France (Chapter 10); Paul L. Dawson, Clemson University, Clemson, SC (Chapter 9); Jonn A. Foulk, U.S. Department of Agriculture, Clemson, SC (Chapter 5); Thomas B. Gold, DSM Catalytica Pharmaceuticals, Inc., Greenville, NC (Chapter 15); Ingolf U. Gruen, University of Missouri, Columbia, MO (Chapter 24); Jacques Guéguen, Institut National de la Recherche Agronomique, Nantes, France (Chapter 5); Harry Helén, University of Helsinki, Helsinki, Finland (Chapter 3); Thomas J. Herald, Kansas State University, Manhattan, KS (Chapter 18); Lyndon B. Kurth, Food Science Australia, Brisbane, Australia (Chapter 17); Tyre C. Lanier, North Carolina State University, Raleigh, NC (Chapter 8); Bertus Markies, Delft Gelatin B.V., Delft, The Netherlands (Chapter 11); Tara H. McHugh, U.S. Department of Agriculture, Albany, CA (Chapter 23); Dolores O’Riordan, University College Dublin, Belfield, Ire-

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Acknowledgements

land (Chapter 6); Nicholas Parris, U.S. Department of Agriculture, Wyndmoor, PA (Chapter 2); Geraldine Quinn, University College Dublin, Belfield, Ireland (Chapter 6); A. Estes Reynolds, University of Georgia, Athens, GA (Chapter 17); Elliot T. Ryser, Michigan State University, East Lansing, MI (Chapter 18); Diana C.W. Siew, Materials Performance Technologies, Auckland, New Zealand (Chapter 7); Mats Stading, The Swedish Institute for Food and Biotechnology, Goteborg, Sweden (Chapter 10); John G. Surak, Clemson University, Clemson, SC (Chapter 25); Zey Ustunol, Michigan State University, East Lansing, MI (Chapter 6); Jeroen J. G. van Soest, Agrotechnological Research Institute, Wageningen, The Netherlands (Chapter 3); Peter J. Vergano, Clemson University, Clemson, SC (Chapter 2). Many thanks are also due to the following individual chapter authors who provided peer review for their colleagues’ chapters: Elizabeth A. Baldwin (Chapter 22); Bill J. Bowman (Chapter 16); Manjeet S. Chinnan (Chapter 23); Karen R. Conca (Chapter 11); Nathalie Gontard (Chapter 26); Stéphane Guilbert (Chapter 1); Jung H. Han (Chapter 21); John M. Krochta (Chapter 24); Clyde M. Ofner III (Chapter 16); Hyun J. Park (Chapter 8); Graciela W. Padua (Chapter 12); Joe M. Regenstein (Chapter 16); Jong W. Rhim (Chapter 13); Thomas A. Trezza (Chapter 12). Finally, the editor thanks Technomic Publishing Co., Inc., and Dr. Eleanor S. Riemer in particular, for their support of this project.

List of Contributors

IRINA ANTONOVA Department of Biological Systems Engineering Virginia Polytechnic Institute & State University 312 Seitz Hall Blacksburg, VA 24061-0303 U.S.A. E-mail: [email protected] IOANNIS S. ARVANITOYANNIS Department of Agriculture Crop and Animal Production School of Technological Sciences University of Thessalia Fytoko Str., 38 446, Nea Ionia Greece E-mail: [email protected] ROBERT A. BAKER Citrus & Subtropical Products Laboratory Agricultural Research Service U.S. Department of Agriculture 600 Avenue S., N.W. Winter Haven, FL 33881 U.S.A. E-mail: [email protected]

ELIZABETH A. BALDWIN Citrus & Subtropical Products Laboratory Agricultural Research Service U.S. Department of Agriculture 600 Avenue S., N.W. Winter Haven, FL 33881 U.S.A. E-mail: [email protected] BILL J. BOWMAN Department of Pharmaceutical Sciences Philadelphia College of Pharmacy University of the Sciences in Philadelphia 600 South 43rd Street Philadelphia, PA 19104-4495 U.S.A. E-mail: [email protected] IAN J. BRITT Department of Food Science University of Guelph Guelph, Ontario N1G 2W1 Canada E-mail: [email protected] xv

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DIANE J. BURGESS Department of Pharmaceutical Sciences University of Connecticut 372 Fairfield Road Storrs, CT 06269-2092 U.S.A. E-mail: [email protected] PASCALE CHALIER Macromolecular Systems Unit Université de Montpellier II GBSA CC023 Place E. Bataillon 34095 Montpellier Cedex 5 France E-mail: [email protected] MUHAMMAD M. CHAUDRY Islamic Food and Nutrition Council 5901 N. Cicero Ave., Suite 309 Chicago, IL 60646 U.S.A. E-mail: [email protected] HONGDA CHEN Plant and Animal Systems Cooperative State Research, Education, and Extension Service U.S. Department of Agriculture 800 9th Street, S.W., Suite 3131 Washington, DC 20024-2220 U.S.A. E-mail: [email protected] MANJEET S. CHINNAN Center for Food Safety & Quality Enhancement The University of Georgia Griffin, GA 30223-1797 U.S.A. E-mail: [email protected]. peachnet.edu

KAREN R. CONCA Combat Feeding Program—Bldg. #36 U.S. Army Soldier and Biological Chemical Command Natick Soldier Center Natick, MA 01746 E-mail: Karen.Conca@natick. army.mil BERNARD CUQ Cereals and Agropolymers Technology Laboratory ENSAM—INRA 2 Place P. Viala 34060 Montpellier Cedex 1 France E-mail: [email protected] CATHERINE N. CUTTER Department of Food Science The Pennsylvania State University 111 Borland Laboratory University Park, PA 16802-2504 U.S.A. E-mail: [email protected] FRÉDÉRIC DEBEAUFORT Institut Universitaire de Technologie Départment de Génie Biologique Bd du Dr. Petitjean BP 17867 21078 Dijon France E-mail: [email protected] ARISTIPPOS GENNADIOS Materials Science & Clinical Supplies Product Development Banner Pharmacaps Inc. 4125 Premier Drive High Point, NC 27265-8144 U.S.A. E-mail: [email protected]


NATHALIE GONTARD Macromolecular Systems Unit Université de Montpellier II GBSA CC023 Place E. Bataillon 34095 Montpellier Cedex 5 France E-mail: gontard@arpb. univ-montp2.fr STÉPHANE GUILBERT Cereals and Agropolymers Technology Laboratory ENSAM—INRA 2 Place P. Viala 34060 Montpellier Cedex 1 France E-mail: [email protected] JUNG H. HAN Department of Food Science The University of Manitoba Winnipeg, Manitoba R3T 2N2 Canada E-mail: [email protected] MILFORD A. HANNA Industrial Agricultural Products Center University of Nebraska 211 L.W. Chase Hall, East Campus Lincoln, NE 68583-0730 U.S.A. E-mail: [email protected] NAVAM S. HETTIARACHCHY Department of Food Science University of Arkansas 272 Young Avenue Fayetteville, AR 72704 U.S.A. E-mail: [email protected]

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JAY-LIN JANE Department of Food Sciences & Human Nutrition and Center for Crops Utilization Research Iowa State University 2312 Food Science Ames, IA 50011-1061 U.S.A. E-mail: [email protected] ZHI Y. JU Department of Food Science University of Arkansas 272 Young Avenue Fayetteville, AR 72704 U.S.A. E-mail: [email protected] JOHN M. KROCHTA Departments of Food Science & Technology and Biological & Agricultural Engineering University of California—Davis One Shields Ave. Davis, CA 95616-8598 U.S.A. E-mail: [email protected] LOONG-TAK LIM Advanced Manufacturing Centre Husky Injection Molding Systems Ltd. 500 Queen Street South Bolton, Ontario L7E 5S5 Canada E-mail: [email protected] P. (KUMAR) MALLIKARJUNAN Department of Biological Systems Engineering Virginia Polytechnic Institute & State University 312 Seitz Hall Blacksburg, VA 24061-0303 U.S.A. E-mail: [email protected]

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CATHERINE MARQUIÉ Laboratoire de Technologie Cotonnière CIRAD-CA TA 70/16 Avenue Agropolis 34398 Montpellier Cedex 5 France E-mail: [email protected] VALERIE MICARD Cereals and Agropolymers Laboratory ENSAM—INRA 2 Place P. Viala 34060 Montpellier Cedex 01 France E-mail: [email protected] YOSHINORI MINE Department of Food Science University of Guelph Guelph, Ontario N1G 2W1 Canada E-mail: [email protected] MARIE HÉLÈNE MOREL Cereals and Agropolymers Laboratory ENSAM—INRA 2 Place P. Viala 34060 Montpellier Cedex 1 France E-mail: [email protected] PERMINUS MUNGARA Department of Food Sciences & Human Nutrition and Center for Crops Utilization Research Iowa State University 2312 Food Science Ames, IA 50011-1061 U.S.A. E-mail: [email protected]

CLYDE M. OFNER III Department of Pharmaceutical Sciences Philadelphia College of Pharmacy University of the Sciences in Philadelphia 600 South 43rd Street Philadelphia, PA 19104-4495 U.S.A. E-mail: [email protected] WESLEY N. OSBURN Department of Animal Sciences Michigan State University 3385 Anthony Hall East Lansing, MI 48824-1225 U.S.A. E-mail: [email protected] GRACIELA W. PADUA Department of Food Science & Human Nutrition University of Illinois at Urbana—Champaign 382-D Agricultural Engineering Sciences Bldg. 1304 West Pennsylvania Avenue Urbana, IL 61801 U.S.A. E-mail: [email protected] HYUN J. PARK Life Sciences Division/Graduate School of Biotechnology Korea University 1, 5Ka, Anam-Dong, Sungbuk-Ku Seoul 136-701 Republic of Korea E-mail: [email protected]


SANG K. PARK Center for Biomaterials and Biotechnology Department of Materials Science and Engineering Kwangju Institute of Science and Technology Kwangju 500-712 Republic of Korea MARÍA B. PÉREZ-GAGO Postharvest Department Valencian Institute of Agricultural Research Ctra. Moncada-Naquera Km. 5 46113 Moncada (Valencia), Spain E-mail: [email protected] JESÙS-ALBERTO QUEZADA-GALLO Departmento de Ingenieria Bioquimica Universidad Autonoma de Aguascalientes Edificio 36, Campus Universitario 20000 Aguascalientes Ags Mexico E-mail: [email protected] ALLISON E. RADWICK Verion, Inc. 254 Welsh Pool Road Exton, PA 19341 U.S.A. E-mail: [email protected] ANDRÉAS REDL Cereals and Agropolymers Technology Laboratory ENSAM—INRA 2 Place P. Viala 34060 Montpellier Cedex 1 France E-mail: [email protected]

JOE M. REGENSTEIN Cornell Kosher Food Initiative Department of Food Science Cornell University 8 Stocking Hall Ithaca, NY 14853-7201 U.S.A. E-mail: [email protected]

JONG W. RHIM Department of Food Engineering Mokpo National University 61 Dorim-ri, Chungmye-myon Muan-gun, Chonnan, 534-729 Republic of Korea E-mail: jwrhim@chungkye. mokpo.ac.kr

SUSAN S. SUMNER Department of Food Science & Technology Virginia Polytechnic Institute & State University Blacksburg, VA 24061-0418 U.S.A. E-mail: [email protected]

THOMAS A. TREZZA Frito-Lay, Inc. 7701 Legacy Dr. Plano, TX 75024-4099 U.S.A. E-mail: [email protected]

MARVIN A. TUNG (Deceased) Department of Food Science University of Guelph Guelph, Ontario N1G 2W1 Canada

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ANDRÉE VOILLEY Laboratoire de Génie des Procédés Alimentaires et Biotechnologiques ENS.BANA Université de Bourgogne 1 Esplanade Erasme 21000 Dijon France E-mail: [email protected] QIN WANG Department of Food Science & Human Nutrition University of Illinois at Urbana—Champaign 382-D Agricultural Engineering Sciences Bldg. 1304 West Pennsylvania Avenue Urbana, IL 61801 U.S.A. E-mail: [email protected] CURTIS L. WELLER Department of Biological Systems Engineering University of Nebraska 210 L.W. Chase Hall, East Campus Lincoln, NE 68583-0726 U.S.A. E-mail: [email protected]

ASAO YAMAUCHI Department of Biotechnology Faculty of Engineering Osaka University Suita, Osaka 565-0871 Japan E-mail: [email protected]. osaka-u.ac.jp KIYOSHI YAMAUCHI Department of Bioapplied Chemistry Faculty of Engineering Osaka City University Sumiyoshi-ku, Osaka 558-8585 Japan E-mail: [email protected]. osaka-cu.ac.jp JINWEN ZHANG Department of Plastics Engineering University of Massachusetts— Lowell Lowell, MA 01854 U.S.A. E-mail: [email protected] SUSHE ZHANG Ruco Polymers 125 New South Road Hicksville, NY 11802 U.S.A. E-mail: [email protected]

Proteins as Raw Materials for Films and Coatings: Definitions, Current Status, and Opportunities

N the past approximately 50 years, impressive advances have been made in the production of synthetic polymer films designed to protect foods, pharmaceuticals, and other products and to perform other functions such as mulching. With the increasing population and stress on limited resources and the environment, uses of renewable resources to produce edible and biodegradable films that can improve product quality and/or reduce waste disposal problems are being explored. A number of reviews and books have considered the materials, properties, and potential applications for edible films and coatings (Daniels, 1973; Guilbert, 1986; Kester and Fennema, 1986; Krochta, 1992, 1997a, b, c; Lindstrom et al., 1992; Conca and Yang, 1993; Cuq et al., 1994; Koelsch, 1994; Krochta et al., 1994; Baldwin et al., 1995a, b; Callegarin et al., 1997; Gennadios et al., 1997; Myllarinen et al., 1997; Debeaufort et al., 1998; Baldwin, 1999), biodegradable films and coatings (Huang, 1985; Kuman, 1987; Huang et al., 1990; Ching et al., 1993; Satyanarayana and Chaterji, 1993; Chapman, 1994; Fishman et al., 1994; Griffin, 1994; Jane, 1994; Narayan, 1994; Gebelein and Carraher, 1995; Fuller et al., 1996; Krochta and De Mulder-Johnston, 1996; Cuq et al., 1997; Guilbert et al., 1997; Petersen et al., 1999), or both edible and biodegradable films and coatings (Gontard and Guilbert, 1994; Guilbert and Gontard, 1995; Martin-Polo, 1995; Anker, 1996;

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2 Ahvenainen et al., 1997; Guilbert et al., 1997; Krochta and De Mulder-Johnston, 1997; Sessa and Willett, 1998; Arvanitoyannis and Gorris, 1999). Several reviews focused exclusively on either proteins (Gennadios and Weller, 1990; Gennadios et al., 1994a; McHugh and Krochta, 1994a; Torres, 1994; Krochta, 1997b; Cuq et al., 1998), polysaccharides (Nisperos-Carriedo, 1994; Nussinovitch, 1998), or lipids and resins (Hernandez, 1994; Baldwin et al., 1997; Callegarin et al., 1997; Shellhammer and Krochta, 1997a). The main focus of this book is on protein-based films and coatings. The objective of this introductory chapter is to (1) define edible and biodegradable films and coatings and the nature of proteins used to form such structures; (2) define methods available for protein film and coating formation and the properties measured to evaluate protein films and coatings; (3) summarize the barrier, mechanical, and other properties possessed by protein-based films and coatings; and (4) discuss existing and potential applications for protein films and coatings.

Films are normally regarded as stand-alone, being formed separate of any eventual intended use. These stand-alone films also are used as testing structures for determination of barrier, mechanical, solubility, and other properties provided by a certain film material. Such films can be used as covers, wraps, or separation layers; and they can be potentially formed into casings, capsules, pouches, and bags. Related products include molded items of greater thickness. Coatings involve formation of films directly on the surface of the object they are intended to protect or enhance in some manner. In this sense, coatings become part of the product and remain on the product through use and consumption. Table 1.1 summarizes different uses for protein films and coatings, distinguishing between edible and biodegradable products.

Films and coatings based on proteins are edible and/or biodegradable, depending on formulation, formation method, and modification treatments. As long as food-grade proteins and other food-grade additives (e.g., plasticizers, acid or base, salts, and enzymes) are used and only protein changes due to heating, pH modification, salt addition, enzymatic modification, and water removal occur, the resulting film or coating is edible (Krochta and De Mulder-Johnston, 1997).

Definitions, Functions, and Rationale

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A small percentage of the population is allergic to one or more proteins, and the formation of films or coatings from proteins does not generally reduce allergenicity. Thus, proper labeling is essential so that the affected population can avoid the relevant protein. Beyond this issue, protein films and coatings can enhance the nutritional quality of foods, based on the protein content and the potential for incorporation of nutritional supplements. Because edible films and coatings can normally support microbial growth, proper attention must be paid to water activity, pH, temperature, atmosphere, and time. Addition of antimicrobials to edible films can protect the films and coatings, as well as the related foods, from microbial growth. Edible films and coatings also are biodegradable. However, edibility is lost when the protein is reacted with other chemicals before or during film or coating formation (e.g., chemical grafting or chemical cross-linking), or when non-edible components are added to the film or coating. Biodegradable films and coatings for food packaging applications must be shown safe for such use (Krochta and De Mulder-Johnston, 1997). The challenge to biodegradable films and coatings for food packaging and other uses is that the film or coating must serve its function safely and effectively for the time needed. Only after the intended functional use has ended should biodegradation proceed. For the purposes of this chapter, biodegradable is taken to mean that the film or coating can be completely degraded by microorganisms in a composting process ultimately to only carbon dioxide, water, methane, and some biomass residue (Anonymous, 1993). A warm, moist environment with appropriate pH, nutrients, oxygen, and time for the appropriate microorganisms is necessary to allow the biodegradation process to proceed.

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Most commonly, edible films and coatings are intended to function as a barrier to moisture, oxygen, flavor, aroma, and/or oil, thus improving food quality and shelf life. An edible film or coating may also provide some mechanical protection for a food, reducing bruising and breakage and thus improving food integrity. When an edible film or coating provides a moisture, flavor, aroma, or oil barrier between food components of different water activity, flavor, aroma, and/or oil content in a heterogeneous food, the quality and shelf life of the food are increased. When an edible film or coating prevents exchange of moisture, oxygen, aroma, or oil between the food and the environment, the quality and shelf life of the food also are increased. However, when functioning in this manner, edible films and coatings are not normally intended to eliminate the need for non-edible protective packaging. Rather, they are intended to work with conventional packaging to improve product quality and shelf life. However, the amount of conventional protective packaging may be reduced (source reduction); and the remaining, simpler package may be more recyclable. In addition, after the package is opened, an edible film or coating can continue to protect the product. The protective function of edible films and coatings may be enhanced with addition of antioxidants or antimicrobials to the film or coating. Depending on the nature of the food, an edible coating may also carry flavors, nutrients, etc., to enhance the quality of the food. Finally, an edible coating can provide additional important sensory attributes to foods, including gloss, color, and non-greasy, non-sticky, or non-color-bleeding surface. The various functions of edible films and coatings are summarized in Table 1.2.

Every food suffers from at least one mass transfer problem, whether it is moisture migration, oxygen intrusion, aroma loss or gain, or oil migration. Given the range of barrier attributes that edible films and coatings can provide, foods can benefit with improved stability, texture, taste, and aroma. Incorpora-

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tion of antioxidants and/or antimicrobials that are control-released can further enhance food quality. Many foods also suffer from breakage and disintegration in manufacture, packaging, transportation, and use. The higher yield, easier handling, and improved appearance from integrity-enhancing coatings are additional potential advantages. Finally, consumers often associate food quality with appropriate gloss, color, and tactile feel, properties that edible coatings can affect. All of these features are of even greater importance because of increased consumer interest in quality, variety, and convenience. At the same time, food processors are interested in worldwide markets that demand longer shelf life and in providing products with a minimum of packaging, which is ultimately recyclable. Thus, these factors have combined to spark interest in exploring the film-formation, barrier, mechanical, and sensory properties of proteins and other materials.

As with edible films and coatings, biodegradable films and coatings can function as barriers to moisture, oxygen, flavor, aroma, and/or oil to protect the quality of food and other products from the environment. In contrast with edible films and coatings, the intent with a biodegradable film or coating is generally to totally replace the conventional synthetic packaging or other conventional synthetic product. A biodegradable film (e.g., pouch) or coating (e.g., on paper) may also need to provide some mechanical protection for a food, drug, or other product. If a biodegradable material is formed into related structures, such as food service items, the items must also have appropriate mechanical properties. The function of biodegradable films and coatings may also be enhanced with the addition of antioxidants or antimicrobials. Finally, the visual sensory attributes of biodegradable films and coatings, such as transparency, gloss, and color, are also important. The various potential functions of biodegradable protein films and coatings are summarized in Table 1.2.

Any successful replacement of conventional synthetic films and coatings with biodegradable alternatives reduces use of non-renewable resources and reduces waste through biological recycling (e.g., composting). Successful development of biodegradable films for packaging and other applications is most likely to occur when recovery of conventional synthetic polymer products for recycling or energy recovery is difficult. An example of where biodegradable alternatives, including food packaging, would be especially useful is for ships, which are now legally prohibited from dumping persistent waste at sea. Also, as municipalities increase availability of curbside pickup for large-scale municipal composting, interest in biodegradable packaging is increasing. Many mu-

6 nicipalities also encourage small-scale domestic composting, even providing necessary equipment. Biodegradable polymer films can also be run through domestic garbage disposals to the municipal waste treatment plant.

Materials available for forming films and film coatings fall generally into the categories of proteins, polysaccharides, lipids, and resins. A plasticizer must often be added to reduce film or coating brittleness. A surface-active agent is also often necessary to aid film or coating formation. Other constituents can include antioxidants and antimicrobials to enhance the film or coating effectiveness. The U.S. Code of Federal Regulations provides the status of protein, polysaccharide, lipid, resin, plasticizer, emulsifier, preservative, and antioxidant materials related to acceptable use (Baldwin, 1999).

Proteins cover a broad range of polymeric compounds that provide structure or biological activity in plants or animals. Proteins are distinguished from polysaccharides because they are based on approximately 20 amino acid monomers, rather than just a few or even one monomer, such as glucose in the case of cellulose and starch. The amino acids are similar in containing an amino group ( NH2) and a carboxyl group ( COOH) attached to a central carbon atom. However, each amino acid has a different side group attached to the central carbon that lends unique character to that amino acid. The side group can be non-polar (hydrophobic), polar uncharged (hydrophilic), positively charged at pH 7, or negatively charged at pH 7 (Cheftel et al., 1985). Most proteins contain 100–500 amino acid residues. Depending on the sequential order of the amino acids (primary structure of the protein), the protein will assume different structures along the polymer chain (secondary structure of the protein), based on van der Waals, hydrogen bonding, electrostatic, hydrophobic, and disulfide cross-link interactions among the amino acid units (Cheftel et al., 1985). The tertiary protein structure reflects how the secondary structures organize relative to each other, based on the same types of interactions, to form overall globular, fibrous, or random protein structure. Finally, quaternary structure occurs when whole proteins interact with each other into associations to provide unique structure or biological activity. The secondary, tertiary, and quaternary structures of proteins can be modified by various physical and chemical agents, including heat, mechanical treatment, pressure, irradiation, lipid interfaces, acids and alkalis, and metal ions (Cheftel et al., 1985). Such agents are often used in the formation of protein films and coatings to optimize protein configuration, protein interactions, and resulting film properties.

Film and Coating Composition

7

Protein film-forming materials derived from animal sources include collagen, gelatin, fish myofibrillar protein, keratin, egg white protein, casein, and whey protein. Protein film-forming materials derived from plant sources include corn zein, wheat gluten, soy protein, peanut protein, and cottonseed protein. Proteins that have successfully been formed into films and/or coatings are listed in Table 1.3 along with the solubility properties of the native proteins before formation into films or coatings.

Polysaccharide film-forming materials include starch and starch derivatives, cellulose derivatives, alginate, carrageenan, chitosan, pectinate, and various gums. Proteins can be combined with polysaccharides to modify film mechanical properties (Shih, 1994; Arvanitoyannis et al., 1996, 1997, 1998a, 1998b; Arvanitoyannis and Biliaderis, 1998).

Edible lipids include beeswax, candelilla wax, carnauba wax, triglycerides (e.g., milkfat fractions), acetylated monoglycerides, fatty acids, fatty alcohols, and sucrose fatty acid esters. Edible resins include shellac and terpene resin. Because lipid and resin materials are not polymers, they do not generally form cohesive stand-alone films. However, along with often providing desirable gloss, they can be used to coat a food or drug surface to provide a moisture bar-

8 rier or to provide the moisture-barrier component of a composite film. Composite films can consist of a lipid layer supported by a protein or polysaccharide layer, or lipid material dispersed in a protein or polysaccharide matrix (Krochta, 1997a).

Protein films and coatings are often quite stiff and brittle due to extensive interactions between protein chains through hydrogen bonding, electrostatic forces, hydrophobic bonding, and/or disulfide cross-linking. Relatively small molecular weight hydrophilic plasticizers are often added, which mainly compete for hydrogen bonding and electrostatic interactions with the protein chains. The result of plasticizer addition is a reduction in protein chain-to-chain interactions, a lowering of the protein glass transition temperature(s), and an improvement in film flexibility (lowering of film elastic modulus). Also, film elongation (stretchiness or ductility) increases, and film strength decreases. Unfortunately, plasticizers generally also decrease the film’s ability to act as a barrier to moisture, oxygen, aroma, and oils. Plasticizers acceptable and generally used for protein edible films include glycerol, propylene glycol, sorbitol, sucrose, polyethylene glycol, fatty acids, and monoglycerides. Water is also an important plasticizer for protein films. Thus, film moisture content, as affected by the surrounding environment’s relative humidity (RH), has a large effect on film properties. The presence of hydrophilic plasticizers such as glycerol attracts additional moisture and additionally impacts film properties. Plasticizers that are commonly used with protein films and coatings are summarized in Table 1.4.

Emulsifiers are surface-active compounds with both polar and non-polar character, which absorb at the water-lipid interface and reduce surface tension. To produce protein-lipid or polysaccharide-lipid composite films from aqueous solution, it is often necessary to add an emulsifier to allow dispersion of the lipid material in the solution. Also, for some food-coating applications, addition

Film and Coating Formation

9

of a surface-active agent to a coating formulation may be necessary to achieve satisfactory surface wetting and spreading with the coating formulation and then adhesion of the dry coating. Some proteins are sufficiently surface-active that no emulsifier is necessary to form well-dispersed composite films or provide good surface wetting and adhesion.

As mentioned, edible films and coatings have the potential to be effective carriers and providers of antioxidants, antimicrobials, nutrients, flavors, and colors to enhance food safety, nutrition, and quality.

When edible films and coatings become part of a processed food or pharmaceutical product in the U.S., their constituents must be clearly indicated on the product label as ingredients. This allows individuals to avoid products that contain ingredients about which they have concerns. When edible coatings used with fresh fruits and vegetables are sold without labeled packaging in the U.S., the coating composition must be clearly displayed in the vicinity of the coated produce. Regulations vary greatly among countries as to coating of fresh fruits and vegetables (Baldwin, 1999).

Protein films and coatings are mainly formed by evaporation of solvent, usually water or aqueous ethanol, from a solution of the protein. With the exception of corn zein, wheat gluten, sorghum kafirin, and keratin, most film-forming proteins are soluble in water. Corn zein and wheat gluten films and coatings must be formed from aqueous ethanol solution or from an aqueous dispersion. Edible film and coating production that requires ethanol necessitates appropriate safety measures and attention to environmental release of solvent to the atmosphere. In this case, solvent recovery will become an increasingly important aspect of commercial operations. Table 1.3 lists the solvents required to solubilize various protein film-formers. For formation of a film or coating, the protein is first dissolved in the solvent. If heating or pH adjustment enhances film formation and/or properties, this is done next. If a composite film or coating based on an emulsion is desired, a lipid material, and possibly a surfactant, is added. Next, the mixture is heated to above the lipid melting point and then homogenized. Degassing is an important

10 step to eliminate bubble formation in the final film or coating. Finally, the protein film or coating is formed by applying the prepared formulation to the desired casting or product surface and allowing the solvent to evaporate. Providing heated air at low humidity and high velocity increases drying rates. For film production from such solvent casting, the formulation must be spread evenly on a surface that will release the film after drying. Formation of protein coatings on food products involves either dipping, spraying, enrobing, or panning the food with the coating formulation. These coating methods all require drying of solvent from a protein coating formulation after it is applied to the food surface. Gelatin capsules cast from aqueous solutions are used widely in the pharmaceutical and nutritional supplement industries. Hard capsule halves are formed by dipping steel pins into gelatin solutions. After drying, the capsules are removed from the pins. A drug or supplement is then filled into a hard capsule half and interlocking halves are joined to form the full capsule. Soft gelatin capsules containing drugs or nutritional supplements are formed from two previously formed sheets of plasticized gelatin by injection of the drug or supplement at the moment when the two sheets are brought together between the rotating halves of roller dies.

Collagen casings and film-wraps for meat products are produced by extruding a viscous (4–10% solids) aqueous suspension of purified acidified collagen into a neutralizing coagulation bath, followed by washing, plasticizing, and drying. However, thermoplastic extrusion would be an attractive way to form protein casings and films, avoiding the need to add and then remove solvent by drying. Some research suggests that some proteins display thermoplastic behavior. However, inducing protein thermoplastic behavior generally has not been much explored or exploited for edible film production. Successful, efficient production of protein edible films using conventional extrusion equipment would certainly improve commercialization potential. However, little has been published on this approach.

Table 1.5 lists the properties generally of interest with protein films and coatings, along with the pertinent testing method references.

The main interest in edible films and coatings is generally based on their

Properties of Protein Films and Coatings

11

potential to provide some combination of moisture, oxygen, flavor, aroma, color, or oil barrier for a food or drug, with a resulting increase in quality and shelf life. Thus, the permeability of edible films to these substances is of interest. Permeability is a steady-state property that describes the extent to which a permeating substance dissolves and then the rate at which the permeant diffuses through a film, with a driving force related to the difference in concentration of the permeant between the two sides of the films. Permeability is thus defined as

Permeability

steady-state rate of permeation (film thickness) through film in amount per time permeant concentration or partial (film area) pressure difference across film

where the concentration or partial pressure difference is between the phases adjacent to the two sides of the film. The polar character of proteins determines the barrier properties of protein films. Protein films have high permeability to polar substances, such as water vapor, and low permeability to non-polar substances, such as oxygen, aromas, and oils. Because plasticizers, including water, generally increase film permeability, permeability of protein films to both polar and non-polar substances increases with plasticizer content and RH. The challenge in use of the barrier properties of protein films is to select the

12 protein, plasticizer, and film-formation conditions that optimize the desired barrier properties, while achieving other desirable properties such as film flexibility, strength, and solubility. Lack of knowledge and data on edible films, in general, still prevents design of edible films to desired specifications. Tables 1.6 and 1.7 list selected protein film water vapor permeability (WVP) and oxygen permeability (OP) data, along with synthetic film data for comparison. It is important to note that plasticizer content (type and amount) and test conditions (temperature and RH) have important effects on film properties. Increasing plasticizer amount, temperature, and RH generally increases permeability. Thus, film properties should be compared at as near identical testing conditions as possible.

Protein films have quite high WVP compared to edible waxes, which are often used as moisture barrier coatings on fruits, vegetables, confections, and drugs, and low-density polyethylene (LDPE) packaging film, which commonly is used to protect food and drugs from moisture (Table 1.6). Protein film WVP values are two to four orders of magnitude greater than that of LDPE. The aqueous ethanol-soluble proteins, corn zein and wheat gluten, and also fish myofibrillar protein appear to form films that have the lowest WVP among the proteins and are comparable to the WVP of cellulose derivatives. Table 1.6 also shows the effect of higher plasticizer concentration and higher RH test conditions on increasing WVP.

At low to intermediate RH, protein films have OP values that are lower than those of the polyethylenes (which are not good oxygen barriers), comparable to those of modest oxygen barriers such as polyesters, and approaching those of the best oxygen barriers, ethylene-vinyl alcohol copolymer (EVOH) and polyvinylidene chloride (PVDC) (Table 1.7). Protein films also appear to have somewhat lower OP than cellulose-based films. Table 1.7 also shows the effect of higher RH on increasing OP of protein films, which is similar to the effect of RH on the excellent synthetic oxygen-barrier EVOH. The low OP of protein films would appear to make them useful for coatings and pouches for oxygen-sensitive products.

Limited research has been done on quantifying the aroma permeabilities of protein films. However, results showed that wheat gluten film was a better barrier to 1-octen-3-ol (mushroom aroma) than low-density polyethylene (LDPE)

13

14

15

16 film or methylcellulose (MC) film (Debeaufort and Voilley, 1994). Other research results showed that whey protein film was better than vinylidene chloride copolymer (co-VDC) film and was comparable to ethylene-vinyl alcohol copolymer (EVOH) film as a barrier to limonene (citrus aroma) at similar conditions (Miller and Krochta, 1998b). Similarly, limited data exist on oil permeability of protein films. Corn zein was shown to have excellent grease resistance, both as a film and as a coating on paper (Trezza and Vergano, 1994). These results showed the potential for fully compostable paper-based wraps and boxes for the food service industry. Research results also showed that whey protein film (De Mulder-Johnston, 1999) and whey protein coating on paper (Chan, 2000) provided excellent oil-barrier properties.

The properties usually measured to mechanically characterize films are tensile strength (TS) (pulling force per film cross-sectional area required to break the film), elongation (E) (degree to which film can stretch before breaking), and elastic modulus (EM) (film stiffness as determined by ratio of pulling force/area to degree-of-film-stretch). Film toughness is approximated by the product of TS and E. Protein films appear to have lower TS than most polysaccharide films and synthetic polymer films, and lower E than synthetic films (Table 1.8). Plasticizer level has a dramatic effect on film properties, with TS and EM decreasing and E increasing with increased plasticizer content. When plasticizer level is reduced to obtain protein film TS values similar to those of polyethylene (PE) or polypropylene (PP) films, the E values are one to two orders of magnitude lower than those of PE or PP. Thus, protein films have nowhere near the toughness of conventional synthetic polymer films. However, mechanical properties are generally sufficient to allow use of protein films for many products, such as casings, wraps, pouches, and coatings.

Film and coating solubility is an important property that relates to intended use. In some cases, a film or coating readily soluble in water is desirable, such as a readily soluble pouch containing food ingredients. In other cases, a water-insoluble film or coating is preferred to provide some water resistance and improve food integrity. Film formers such as corn zein and wheat gluten that are not soluble in water produce films and coatings that are not water soluble. Protein film-formers that are soluble in water produce films of varying solubility, depending on the protein and the conditions of film formation and treatment. For example, native whey protein isolate produces totally water-soluble

17

18 films; but heat-denatured solutions of whey protein isolate produce films in which the protein is insoluble (Perez-Gago et al., 1999).

An important trend in recent years has been the investigation of many approaches for enhancing the barrier, mechanical, and solubility properties of protein films. Generally, these approaches involve modification of protein structure and/or interactions among protein molecules. Another approach has involved the creation of blended or composite films by introduction of polysaccharides or hydrophobic (lipid or wax) materials, respectively. Tables 1.9, 1.10, and 1.11 summarize the effects of selected treatments on protein film properties.

As mentioned earlier, type and amount of plasticizer affects interactions between protein molecules, with resulting effect on film properties. Plasticizers are added to decrease film stiffness (EM) and increase film elongation (E). Normally, as the plasticizer amount increases, permeability values and E increase, while TS and EM decrease. At least with certain proteins, some plasticizers allow achievement of desired mechanical properties with less effect on barrier properties than alternatives (Table 1.9). This is an area worthy of additional research to allow more effective selection of plasticizers. Unfortunately, plasticizer selection is not capable of reducing protein film WVP by more than a factor of two to three. However, optimum selection of plasticizer can improve the already excellent oxygen barrier ability of protein films.

Depending on the method of protein recovery, protein fraction, film-solution treatment, and film-forming conditions, protein structure and interactions will be influenced. For example, different means of recovering casein can influence casein structure and thus film properties (Chick and Ustunol, 1998; Tomasula et al., 1998). On the other hand, the -lactoglobulin fraction of whey protein gives the same film properties as the whole whey protein (Maté and Krochta, 1996a). Ultrasound treatment of film-forming solutions likely disrupts existing protein interactions, thus resulting in increased molecular interaction (Banerjee et al., 1996). Modified protein interaction can also occur due to film-forming solution pH adjustment (Gontard et al., 1992; Brandenburg et al., 1993) or film adjustment to the protein isoelectric point (pI) (Avena-Bustillos and Krochta, 1993; Gennadios et al., 1993c). Heat treatment of film-forming solution causes denaturation due to intramolecular disulfide

19

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22

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24 bond breakage. The result is opportunity for protein cross-linking due to intermolecular disulfide bond formation and changed film properties. In fact, most protein film formation involves such heat treatment of film-forming solutions. The effect on some protein film properties is dramatic (Perez-Gago et al., 1999). Protein cross-linking can also occur due to an enzyme (Stuchell and Krochta, 1994; Lim et al., 1998), addition of a cross-linking agent (Avena-Bustillos and Krochta, 1993; Gennadios et al., 1993a; Marquie et al., 1995; Parris et al., 1998; Rhim et al., 1999; Were et al., 1999), or ultraviolet radiation (Gennadios et al., 1998b). Film drying or curing conditions can also modify protein structure and interactions (Gennadios et al., 1996b; Ali et al., 1997; Miller et al., 1997; Alcantara et al., 1998; Jangchud and Chinnan, 1999). Modifying protein structure and interaction by the methods discussed above can reduce protein film WVP by a factor of two to four (Table 1.10). Such change is impressive, but not sufficient to convert protein films to good moisture barriers. However, reduction of protein film OP by enhancing protein structure and interaction improves the already excellent oxygen barrier properties of protein films (Table 1.10). The largest effect of changes in protein structure and interaction can be on film mechanical properties and film solubility. Protein film mechanical properties can change by several factors, and the film protein content can go from totally soluble to totally insoluble (Table 1.10).

A polysaccharide material, such as starch, can be combined with protein to produce a blended film system. Limited data exist on such blended films (Arvanitoyannis et al., 1996, 1997, 1998a, b; Arvanitoyannis and Biliaderis, 1998). It appears there is little effect on film WVP, but that addition of polysaccharide may increase film OP and TS and decrease film E (Table 1.11). The greatest reduction in protein film WVP is achieved by addition of a lipid or wax to the film structure. The protein provides structural integrity for the film while the hydrophobic lipid or wax provides the moisture barrier. Dispersing a lipid or wax in the protein film structure to achieve an “emulsion film” can reduce the film WVP by an order of magnitude. However, this still places the film’s WVP one to three orders of magnitude greater than the WVP of the pure lipid or wax, or of a PE film. Thus far, the only way to achieve a good moisture barrier is to laminate a preformed protein or polysaccharide film with a layer of lipid or wax to achieve a bilayer film (Table 1.11). While addition of a lipid or wax to a protein film can reduce film WVP, such addition usually increases film OP, reduces film TS and EM, and increases film E (Table 1.11). Also, the possibility of resulting cracks and pinholes in the film increases.

Protein Film and Coating Application Opportunities

25

The goal of biodegradable films and other biodegradable products is simple: replace existing synthetic, non-biodegradable products for uses such as those listed in Table 1.1 at the lowest cost possible. However, edible films and coatings are normally not meant to be replacements for existing non-edible films and coatings. The main focus is on improving food quality and shelf life by reducing the effect of moisture, oxygen, migration, etc., protecting food from microbes, maintaining food product integrity, and enhancing product appearance. However, this must be related to the cost of coating materials and to the cost of the coating process. Evaluating the financial benefit of improved food quality and shelf life is difficult, but must be done. In addition, potential savings related to reduction of packaging may factor into the analysis. Finally, if the resulting packaging is simpler and, thus, more recyclable, this advantage should also be included in the analysis. Edible film and coating application goals are listed in Table 1.12.

Table 1.13 lists commercial edible film and coating materials along with intended applications. These include forming an edible film as a coating directly on a food or drug product, as well as filling a food or drug product into a preformed edible film casing, pouch, or capsule. Table 1.14 lists the functions edible coatings serve in different food products. Edible wax and shellac coatings are used on fruits and vegetables to prevent moisture loss and improve appearance. Wax, shellac, and zein coatings are used on confections to provide moisture resistance and improve appearance. Sucrose esters of fatty acids are used in formulations to reduce respiration, moisture loss, and skin marking and bruising in fruits and vegetables. Hydroxypropyl methylcellulose (HPMC), hydroxypropyl cellulose (HPC), methylcellulose (MC), shellac, and zein coatings and gelatin capsules are used for pharmaceuticals to improve product appearance, structural integrity, and ingestibility, as well as to protect against moisture and oxygen. HPMC and MC

26

are also used for coatings on fried foods to reduce moisture loss and fat gain. MC is also used in commercial formulations to extend the life of fresh whole or cut fruits and vegetables. HPMC pouches are marketed for delivery of pre-measured food ingredients to food formulations with subsequent dissolution of the pouch. Starch-based materials are marketed for coating drug tablets. Starch and dextrin formulations are also marketed as protective coatings, integrity maintainers, appearance enhancers, and seasoning adhesives for nuts, snacks, cereals, and meats. Collagen casings and wraps inhibit moisture loss and oxygen transport and provide structural integrity for meat products.

While the potential applications in Tables 1.13 and 1.14 are noteworthy, actual applications represent a small fraction of the foods that could benefit from an edible film or coating. Every food suffers from at least one mass transfer problem, whether it be moisture migration, oxygen intrusion, aroma loss, or oil migration. Many foods could also benefit from improvement in integrity or appearance. However, applications have not become more widespread because commercial film and coating materials are either too expensive or too difficult to form, and/or commercial materials do not function efficiently. Thus, extensive research continues in this area. Certain new edible film and coating materials, including proteins, are targeted for replacing materials currently used in ex-

27

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Summary and Conclusions

31

isting applications. Beyond existing applications, edible films and coatings are generally seen as having considerable potential for applications far beyond current usage for improving product quality and, in some instances, reducing use of synthetic packaging films. Several reviews list edible protein film and coating applications that have been explored over the years (Baker et al., 1994; Baldwin, 1994; Gennadios et al., 1994a, 1997; Krochta, 1997c; Krochta and De Mulder-Johnston, 1997). Table 1.15 lists protein film and coating research that has been conducted to provide improved materials and coating formation for existing applications, as well as to explore new food applications. Compared to the large number of studies performed on film formation and properties (see Tables 1.6–1.11), a relatively small number of application studies have been performed. Thus, information is generally lacking on approaches to coating foods, as well as the resulting effectiveness of edible films and coatings in food systems. This makes it very difficult for food processors to decide on the “value-added” merit of an edible film or coating relative to the additional cost involved.

Based on their chemical composition and structure, proteins make excellent oxygen, aroma, and oil barrier films and coatings at low to intermediate RH, similar to the best synthetic barrier films. Research on the effect of plasticizer type and amount may provide opportunities for achieving desired mechanical properties while further improving these barrier properties. The hydrophilic nature of proteins results in high film and coating WVP, which is several orders of magnitude greater than synthetic moisture barrier films. Increasing protein interaction (e.g., chain cross-linking) can reduce the WVP by several factors, but nowhere near that required to achieve a good moisture barrier. The main effects of such protein modifications are usually to increase film strength and stiffness, and decrease film elongation and solubility. However, addition of lipid or wax materials improves the moisture barrier ability. Additional research on protein-lipid/wax composite film formation and properties is needed to improve the performance and reduce the cost of protein-based films and coatings as moisture barriers. Protein films have mechanical properties that, overall, are inferior to those of synthetic and polysaccharide films but are sufficient for most applications. Addition of polysaccharide material to protein film can improve film strength, elongation, and toughness. Additional research in this area will likely help optimize properties of protein-polysaccharide films and coatings. Much good research has been devoted to forming various protein films and quantifying the protein film properties. However, it is a large step from that point to determination of the film’s effectiveness in improving the quality of foods or other products. Considerable research is needed to improve the effi-

32 ciencies of forming protein films as coatings on foods and to quantify the effects of the coatings on food quality using both instrumental and sensory evaluation. Investigation of protein coating adhesion, gloss, and color as affected by coating formulation and coating process is needed. Demonstration of protein coating effectiveness at reducing moisture change, oxidation, aroma, and oil migration, and improving food integrity is necessary to help protein coatings reach their full potential. Recent research on coating fruits and vegetables, meats, eggs, starch foods, nuts, other dry foods, and paper has provided needed data and helped demonstrate the potential of protein coatings. Research on the effectiveness of adding antioxidants, antimicrobials, and other additives to protein films and coatings is also needed. Advantages would result for certain food applications from the ability to extrude protein films as an alternative to solvent casting. Achieving such capability will require research ranging from establishing the thermal properties of proteins to extrusion conditions suitable for optimum film formation and properties. Such capability is also essential for realizing the potential of protein films and related structures for biodegradable products. Continuing interest in increasing food quality, reducing use of limited resources, and reducing the environmental impact of synthetic polymers will likely result in increased use of edible and biodegradable films and coatings in the future. To achieve this potential, continuing advances in formation, properties, and economics of edible and biodegradable protein films and coatings are necessary.

Ahvenainen, R., Myllarinen, P. and Poutanen, K. 1997. “Prospects of Using Edible and Biodegradable Protective Films for Foods,” Europ. Food Drink Rev., Summer:73, 75, 77–80. Alcantara, C.R. 1996. Barrier and Mechanical Properties of Whey-Protein-Isolate-Based Films: Film Drying Rate and Coated-Food Integrity Studies. Ph.D. Thesis, University of California, Davis, USA. Alcantara, C.R., Rumsey, T.R. and Krochta, J.M. 1998. “Drying Rate Effect on the Properties of Whey Protein Films,” J. Food Proc. Preserv., 21:387–405. Ali, Y., Ghorpade, V.M. and Hanna, M.A. 1997. “Properties of Thermally-Treated Wheat Gluten Films,” Industr. Crops Prod., 6:177–184. Alikonis, J.J. 1979. Candy Technology. Westport, CT: AVI Publishing Company, Inc. Alikonis, J.J. and Cosler, H.B. 1961. “Extension of Shelf Life of Roasted and Salted Nuts and Peanuts,” Peanut J. Nut World, 40(5):16–17. Andres, C. 1984. “Natural Edible Coating Has Excellent Moisture and Grease Barrier Properties,” Food Proc., 45(13):48–49. Anker, M. 1996. Edible and Biodegradable Films and Coatings for Food Packaging—A Literature Review. Goteborg, Sweden: SIK. Anker, M., Stading, M. and Hermansson, A.-M. 1998. “Mechanical Properties, Water Vapor Permeability, and Moisture Contents of -Lactoglobulin and Whey Protein Films Using Multivariate Analysis,” J. Agric. Food Chem., 46:1820–1829. Anker, M., Stading, M. and Hermansson, A.-M. 1999. “Effect of pH and the Gel State on the Me-

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Based on Whey Proteins or Mixtures of Whey Proteins and Lactose,” Food Structure, 12:457–468. Mullen, J.D. October 26, 1971. U.S. patent 3,615,715. Myllarinen, P., Rantamaki, P., Latva, K. and Ahvenainen, R. 1997. Minimization of Food Packaging Using Active Edible Coatings: Possibilities and Challenges. Finland: VTT Information Service. Narayan, R. 1994. “Polymeric Materials from Agricultural Feedstocks” in Polymers from Agricultural Coproducts, eds., M.L. Fishman, R.B. Friedman and S.J. Huang, Washington, DC: American Chemical Society, pp. 2–28. Nisperos-Carriedo, M.O. 1994. “Edible Coatings and Films Based on Polysaccharides” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Co., Inc., pp. 305–335. Noznick, P.P. and Bundus, R.H. April 18, 1967. U.S. patent 3,314,800. Nussinovitch, A. 1998. “Hydrocolloid Coating of Foods: A Review,” Leatherhead Food RA Food Industry Journal, 1:174–188. Olson, S. and Zoss, R. April 16, 1985. U.S. patent 4,511,583. Park, H.J. and Chinnan, M.S. 1995. “Gas and Water Vapor Barrier Properties of Edible Films from Protein and Cellulosic Materials,” J. Food Engr., 25:497–507. Park, H.J., Chinnan, M.S. and Shewfelt, R.L. 1994a. “Edible Coating Effects on Storage Life and Quality of Tomatoes,” J. Food Sci., 59:568–570. Park, H.J., Chinnan, M.S and Shewfelt, R.L. 1994b. “Edible Corn-Zein Film Coatings to Extend Storage Life of Tomatoes,” J. Food Process. Preserv., 18:317–331. Park, J.W., Testin, R.F., Park, H.J., Vergano, P.J. and Weller, C.L. 1994c. “Fatty Acid Concentration Effect on Tensile Strength, Elongation, and Water Vapor Permeability of Laminated Edible Films,” J. Food Sci., 59:916–919. Park, H.J., Rhim, J.W. and Lee, H.Y. 1996. “Edible Coating Effects on Respiration Rate and Storage Life of Fuji Apples and Shingo Pears,” Food Biotechnol., 5:59–63. Parris, N., Coffin, D.R., Dickey, L.C. and Craig, J.C. 1998. “Composition Factors Affecting the Physical Properties of Hydrophilic Zein Films” in Paradigm for Successful Utilization of Renewable Resources, eds., D.J. Sessa and J.L. Willett, Champaign, IL: AOCS Press, pp. 255–265. Pérez-Gago, M.B., Nadaud, P. and Krochta, J.M. 1999. “Water Vapor Permeability, Solubility, and Tensile Properties of Heat-Denatured versus Native Whey Protein Films,” J. Food Sci., 64:1034–1037. Petersen, K., Nielsen, P.V., Bertelsen, G., Lawther, M., Olsen, M.B., Nilsson, N.H. and Mortensen, G. 1999. “Potential of Biobased Materials for Food Packaging,” Trends Food Sci. Technol., 10:52–68. Rankin, J.C., Wolff, I.A., Davis, H.A. and Rist, C.E. 1958. “Permeability of Amylose Film to Moisture Vapor, Selected Organic Vapors, and the Common Gases,” Ind. Engr. Chem., 3:120–123. Rhim, J.W., Wu, Y., Weller, C.L. and Schnepf, M. 1999. “Physical Characteristics of a Composite Film of Soy Protein Isolate and Propyleneglycol Alginate,” J. Food Sci., 64:149–152. Rice, J. 1994. “What’s New in Edible Films?” Food Process., 55(7):61–62. Rosenberg, M. and Young, S.L. 1993. “Whey Proteins as Microencapsulating Agents. Microencapsulation of Anhydrous Milkfat—Structure Evaluation,” Food Structure, 12:31–41. Roy, S., Weller, C.L., Gennadios, A., Zeece, M.G and Testin, R.F. 1999. “Physical and Molecular Properties of Wheat Gluten Films Cast from Heated Film-Forming Solutions,” J. Food Sci., 64:57–60. Salame, M. 1986. “Barrier Polymers” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker, New York: John Wiley & Sons, pp. 48–54.

40 Satyanarayana, D. and Chaterji, P.R. 1993. “Biodegradable Polymers: Challenges and Strategies,” J. Macromolecular Sci.-Rev. Macromolecular Chem. Phys., 33:349–368. Schilling, E.D. and Burchill, P.I. July 4, 1972. U.S. patent 3,674, 506. Sessa, D.J. and Willett, J.L. 1998. Paradigm for Successful Utilization of Renewable Resources. Champaign, IL: AOCS Press. Shellhammer, T.H. and Krochta, J.M. 1997a. “Edible Coating and Film Barriers” in Lipids—Industrial Applications and Technology, eds., F.D. Gunstone and F.B. Padley, New York: Marcel Dekker, pp. 453–479. Shellhammer, T.H. and Krochta, J.M. 1997b. “Water Vapor Barrier and Rheological Properties of Simulated and Industrial Milkfat Fractions,” Trans. ASAE, 40:1119–1127. Shellhammer, T.H. and Krochta, J.M. 1997c. “Whey Protein Emulsion Film Performance as Affected by Lipid Type and Amount,” J. Food Sci., 62:390–394. Shellhammer, T.H. and Krochta, J.M. 1998. “Microstructural Evaluation of Heat Annealed Edible Whey Protein and Beeswax Emulsion Films,” ASAE Paper No. 98-6073, St. Joseph, MI: American Society of Agricultural Engineers. Shifrin, G.A. November 19, 1968. U.S. patent 3,411,921. Shih, F.F. 1994. “Interaction of Soy Isolate with Polysaccharide and Its Effect on Film Properties,” JAOCS, 71:1281–1285. Siew, D.C.W., Heilmann, C., Easteal, A.J. and Cooney, R.P. 1999. “Solution and Film Properties of Sodium Caseinate/Glycerol and Sodium Caseinate/Polyethylene Glycol Edible Coating Systems,” J. Agric. Food Chem., 47:3432–3440. Stuchell, Y.M. and Krochta, J.M. 1994. “Enzymatic Treatments and Thermal Effects on Edible Soy Protein Films,” J. Food Sci., 59:1332–1337. Stuchell, Y.M. and Krochta, J.M. 1995. “Edible Coatings on Frozen King Salmon: Effect of Whey Protein Isolate and Acetylated Monoglycerides on Moisture Loss and Lipid Oxidation,” J. Food Sci., 60:28–31. Suderman, D.R., Wiker, J. and Cunningham, F.E. 1981. “Factors Affecting Adhesion of Coating to Poultry Skin: Effects of Various Protein and Gum Sources in the Coating Composition,” J. Food Sci., 46:1010–1011. Taylor, C.C. 1986. “Cellophane” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker, New York: John Wiley & Sons, pp. 159–163. Tomasula, P.M., Parris, N., Yee, W. and Coffin, D.R. 1998. “Properties of Films Made from CO2-Precipitated Casein,” J. Agric. Food Chem., 46:4470–4474. Torres, J.A. 1994. “Edible Films and Coatings from Proteins” in Protein Functionality in Food Systems, eds., N.S. Hettiarachchy and G.R. Zeigler, New York: Marcel Dekker, pp. 467–507. Torres, J.A. and Karel, M. 1985. “Microbial Stabilization of Intermediate-Moisture Food Surfaces. III. Effects of Surface Preservative Concentration and Surface pH Control on Microbial Stability of an Intermediate Moisture Cheese Analog,” J. Food Process. Preserv., 9:107–119. Torres, J.A., Motoki, M. and Karel, M. 1985. “Microbial Stabilization of Intermediate Moisture Food Surfaces. I. Control of Surface Preservative Concentration,” J. Food Process. Preserv., 9:75–92. Trezza, T.A. 1999. Surface Properties of Edible, Biopolymer Coatings for Foods: Color, Gloss, Surface Energy and Adhesion. Ph.D. Dissertation, University of California, Davis, USA. Trezza, T.A. and Vergano, P.J. 1994. “Grease Resistance of Corn Zein Coated Paper,” J. Food Sci., 59:912–915. Turbak, A.F. August 8, 1972. U.S. patent 3,682,661. Watters, G.G. and Brekke, J.E. 1961. “Stabilized Raisins for Dry Cereal Products,” Food Technol., 15:236–238. Were, L., Hettiarachchy, N.S and Coleman, M. 1999. “Properties of Cysteine-Added Soy Protein-Wheat Gluten Films,” J. Food Sci., 64:514–518. Whitman, G.R. and Rosenthal, H. January 19, 1971. U.S. patent 3,556,814.

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Wolff, I.A., Davis, H.A., Cluskey, J.E., Gundrum, L.J. and Rist, C.E. 1951. “Preparation of Films from Amylose,” Ind. Engr. Chem., 43:915–919. Wong, Y.C., Herald, T.J. and Hachmeister, K.A. 1996. “Evaluation of Mechanical and Barrier Properties of Protein Coatings on Shell Eggs,” Poultry Sci., 75:417–422. Wu, P.J. and Schwartzberg, H.G. 1992. “Popping Behavior and Zein Coating of Popcorn,” Cereal Chem., 69:567–573.

Formation and Properties of Corn Zein Films and Coatings

Z

EIN comprises a group of alcohol-soluble proteins (prolamins) found in corn

endosperm. It accounts for 50% or more of total endosperm protein, and its only known role is the storage of nitrogen for the germinating embryo (Larkins and Hurkman, 1978). Zein occurs in small, compact protein bodies embedded in the glutelin protein matrix and distributed chiefly in the outer layers of the corn endosperm (Wolf et al., 1967; Dombrink-Kurtzman and Bietz, 1993). It can be extracted with aqueous alcohol and dried to a granular powder. Commercial zein is essentially a by-product of the corn wet-milling industry. After cleaning, corn is steeped in water containing sulfur dioxide to soften the grains. Coarse milling separates the hulls and germ from the endosperm, which is further milled into a fine slurry. Centrifugal separation of starch from the endosperm slurry leaves a protein-rich mass, or corn gluten meal, from which zein is extracted. Annual zein production in the U.S. is more than 1 million pounds, mostly used in formulations of specialty food and pharmaceutical coatings. The potential supply of zein, estimated at 750 million pounds per year in the U.S., calls for expanded markets and drives research and development of novel value-added applications (Shukla, 1992). New methods for isolating zein from dry milled corn that may reduce the cost of zein extraction have been proposed (Parris et al., 1997). 43

44 The film-forming properties of zein have been recognized for decades, and they are the basis for its commercial utilization (Winters and Deardorff, 1958; Kanig and Goodman, 1962; Mendoza, 1975; Andres, 1984). Coating films are formed on hard surfaces by covering them with zein solutions and allowing the solvent to evaporate off. The dried zein residue forms hard and glossy, scuff-proof, protective coatings that also are resistant to microbial attack (Reiners et al., 1973). Free-standing films or thin films, prepared in the laboratory by peeling off dried zein coatings cast on flat surfaces (Aydt et al., 1991), are potentially useful for a range of applications from edible films to biodegradable packaging. Zein films are brittle and must be plasticized to make them flexible. Films were also prepared in the laboratory by drawing them from a moldable, putty-like, resin formed from plasticized zein (Lai and Padua, 1997). Zein resins were also extruded into films (Ha, 1999), seeking to facilitate manufacture process scale-up. Performance of zein-based films and coatings in edible/biodegradable applications depends on their composition and morphology. Addition of plasticizers and cross-linking agents affects mechanical and barrier properties. Processing methods are reported to affect film morphology. This chapter reviews basic studies on zein structure and behavior, current methods for preparation of zein-based films and coatings, mechanical and barrier properties of films, and morphology characterization studies.

Zein occurs as aggregates linked by disulfide bonds in whole corn; however, some of those bonds may be cleaved by reducing agents during extraction or commercial wet-milling operations (Reiners et al., 1973). Based on solubility differences, zein consists of three protein fractions, i.e., -zein, -zein, and -zein (Esen, 1987). -Zein accounts for 75 to 85% of the total protein and is dominated by two groups of proteins, Z19 and Z22, according to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Argos et al., 1982). Z19 refers to the faster band with apparent molecular weight of 19 kDa, and Z22 is the slower band with apparent molecular weight of 22 kDa. However, according to the complete amino acid sequences of -zeins, the true molecular weights of Z19 and Z22 are 23 to 24 kDa and 26 to 27 kDa, respectively (Pedersen et al., 1982; Heidecker and Chaudhuri, 1991). Zein amino acid sequence was derived from chromatography on zein itself and from determination of the genes that encode the protein. Z19 and Z22 consist of 210 and 245 amino acids, respectively (Pedersen et al., 1982; Heidecker and Chaudhuri, 1991; Thompson et al., 1992). Both Z19 and Z22 have sequence homology: N-terminals contain 35 to 37 amino acids, C-terminals have eight amino acids, and the central domain consists of nine (for Z19) or 10 (for Z22) repetitive domains. These repetitive domains contain blocks of 14 to 25

Zein Structure

45

amino acid residues with an average length of 19 and 20 amino acids for Z19 and Z22, respectively (Heidecker and Chaudhuri, 1991; Thompson et al., 1992). Determination of secondary structure utilizing optical rotation, optical rotary dispersion, and circular dichroism (Watson et al., 1936; Elliot and Williams, 1939; Foster and Edsall, 1945; Danzer et al., 1975; Argos et al., 1982; Pedersen et al., 1982; Heidecker and Chaudhuri, 1991; Thompson et al., 1992; Tatham et al., 1993) suggested that the average -helical proportion of zein is 50 to 60%. Also, by using the sequence prediction method of Chou and Fasman (1978), it was determined that the -helices predominate in the central domains of zein (Tatham et al., 1993). Early studies on zein conformation in alcoholic solutions using ultracentrifugation, birefringence, and dielectric experiments (Watson et al., 1936; Elliot and Williams, 1939; Foster and Edsall, 1945; Tatham et al., 1993) indicated that zein particles have an asymmetric ellipsoid or rod shape with 7:1 to 28:1 axial ratios. Based on hydration potential, polarity, and secondary structure properties of the residue, a possible nine- or 10-helix structure was proposed by Argos et al. (1982). In their model, the nine successive helical segments are folded upon each other in an anti-parallel fashion. Helical segments are arranged in a ring of “pencils” held together, side-by-side, by hydrogen bonds and linked at each end by glutamine-rich turns or loops. Garratt et al. (1993) proposed that -helices lie in a hexagonal arrangement instead of an open ring. Hexagonal packing would increase packing efficiency and help explain the self-association ability of zein units. However, both models considered the -zein molecule to approximate a globular conformation of a length-to-width ratio of only about 2:1. Small-angle X-ray scattering (SAXS) was later used to study the shape and dimensions of zein in alcoholic solutions (Danzer et al., 1975; Chou and Fasman, 1978; Tatham et al., 1993; Matsushima et al., 1997). Tatham et al. (1993), using SAXS and viscometry, determined a more asymmetric structure for the solution conformation of -zeins. They obtained values of Rg (radius of gyration) of 44.1 Å and Rc (radius of gyration of the cross section) of 2.45 Å in 70% aqueous methanol at concentrations of 2.5 to 8 mg/mL. Data were interpreted in terms of zein particles being either a prolate ellipsoid of length 196 Å and cross-section diameter of 7 Å or a rod of 153 Å in length and 13.8 Å in diameter. Viscometric measurements yielded values of 238 and 18 Å for length and diameter, respectively, regardless of shape. Those measurements seemed too small for a zein molecule because the -helical portion of the molecule alone (50%) would have a length > 200 Å. Tatham et al. (1993) proposed a model consisting of long rigid rods formed of an -helix folded back on itself forming a two-helix bundle held together by rigid bonds. They demonstrated that an asymmetric model could explain the film-forming ability of zein. Long, rodshaped molecules could undergo orientation into two- and three-dimensional arrays stabilized by hydrogen bonds between side chains of polar residues.

46 Matsushima et al. (1997) reported SAXS measurements of zein in 70% aqueous ethanol over a concentration range of 2 to 40 mg/mL. They determined Rg and Rc values of 40 and 13.9 Å, respectively, for reduced -zein in solutions containing 0.1 or 2% (v/v) -mercaptoethanol. For the non-reduced -zein, they obtained Rg and Rc values of 49.8 and 19 Å, respectively. From those measurements, they proposed that the reduced -zein structural unit has a rectangular prism shape measuring 130 Å for the longest dimension and 34 Å for each of the other two dimensions. For the non-reduced zein, the longest dimension was determined at 160 Å and 46 Å for the other two lateral dimensions. Matsushima et al. (1997) assumed a model related to that of Argos et al. (1982) consisting of nine (Z19) or 10 (Z22) helical segments folded in anti-parallel fashion linked by glutamine-rich turns and held in place by hydrogen bonds. They proposed that the helical segments were aligned to form a compact prism rather than an open ring. A portion of the N-terminus formed an additional helical segment at the end of the ribbon. This model would explain film formation by the alignment of rods and their self-association into two-dimensional films.

Water sorption isotherms were determined for zein powder and cast films (Baianu and Wu, 1998). Due to the hydrophobic nature of zein, water sorption was extremely low in the low water activity (aw) range. Water sorption, though, increased exponentially with aw. For zein powder and unplasticized films, water absorption was minimal at aw < 0.8, while for glycerol-plasticized films water absorption was minimal at aw < 0.7. No significant difference in water absorption was observed between zein powder and unplasticized films. Both powder and films exhibited relatively low water content at high aw levels (< 50 g/100 g dry matter at aw ~ 1). In contrast, glycerol-plasticized films had five times higher water sorption capacity than unplasticized films at high aw. The higher water-holding capacity of plasticized films was attributed to the hydrophilicity of glycerol. Kanig and Goodman (1962) measured water absorption of zein films cast from ethanol-water solutions and plasticized with oleic acid. They reported low water absorption (1.5%) for films at 10 to 52% relative humidity (RH), while films softened considerably at 93% RH. Softening changed the structure and appearance of films and greatly affected their mechanical and barrier properties. Films plasticized with polyethylene glycol (PEG) or lactic acid had lower water absorption compared to glycerol-plasticized films (Kanig and Goodman, 1962).

Zein films are brittle at ambient temperature and need plasticization to be-

Zein Coatings

47

come flexible. Plasticization increases polymer mobility, decreases glass transition temperature (Tg), and markedly changes rheological properties (Kokini et al., 1995). The Tg depends largely on molecular conformation and the presence of plasticizers. Common zein plasticizers include glycerol, glyceryl monoesters, PEG, and fatty acids. Zein, although not soluble in water, is readily plasticized by it. Madeka and Kokini (1996) reported that the Tg of zein decreased rapidly from 139 C for bone-dry zein to 47 C for zein with 6.6% moisture content. The plasticizing effect of water on zein decreased above 10% moisture content. At sufficiently high moisture levels, Tg fell below the freezing point of water, making it impossible to obtain true Tg values for water-plasticized zein. A state diagram for zein above 20% moisture content predicted a rubbery state at ambient temperature and an entangled polymer flow above 60 C (Madeka and Kokini, 1996). Lai and Padua (1997) determined a Tg value of zein films plasticized with oleic acid (33% oleic acid) at 94 C. Their result was consistent with predictions of the state diagram presented by Kokini et al. (1995). In another study, Magoshi et al. (1992) reported a Tg of 165 C for zein cast from aqueous ethanol. Differential scanning calorimetry (DSC) scans showed endothermic peaks at 210 and 320 C, which were attributed to crystallization of amorphous zein to -form crystals and to zein degradation, respectively.

Corn zein is one of a few proteins, such as collagen and gelatin, used commercially as an edible coating. Zein coatings are used as oxygen, lipid, and moisture barriers for nuts, candies, confectionery products, and other foods (Cosler, 1958; Andres, 1984). Rice fortified with vitamins and minerals has been coated with zein/stearic acid/wood resin mixtures to prevent vitamin and mineral losses during washing in cold water (Mickus, 1955). Pharmaceutical tablets are zein-coated for controlled ingredient release and protection (Gennadios and Weller, 1990). Zein-based coatings effectively maintained a high concentration of sorbic acid at the surface of an intermediate moisture cheese analog (Torres and Karel, 1985). Zein coatings reduced moisture and firmness loss and delayed color change (reduced oxygen and carbon dioxide transmission) in fresh tomatoes (Park et al., 1994a, b). Use of zein-based coatings has been suggested for reducing oil uptake by deep-fat fried foods (Feeney et al., 1993); for protecting active ingredients in chewing gum (Broderick, 1998); for achieving controlled release of active ingredients in pharmaceutical tablets (Oshlack et al., 1994); and for masking the taste of orally administered drugs (Meyer and Mazer, 1997).

Zein coatings are usually prepared in three steps. Zein powder is dissolved in

48 warm, aqueous ethyl alcohol or isopropanol. FDA-approved plasticizers such as propylene glycol, glycerin, and acetylated glycerides are added to increase coating flexibility. Products to be coated are dipped into (Wu and Schwartzberg, 1992; Wong et al., 1996), sprayed with (Mallikarjunan et al., 1997), or brushed with (Park et al., 1994a, 1994b; Trezza and Vergano, 1994) the zein/plasticizer solution. Antioxidants, such as BHA and BHT, are often added to prevent lipid oxidation (Gennadios and Weller, 1990). Upon solvent evaporation, a shiny, protective zein film is formed on the product surface. Vegetable oils (e.g., cottonseed oil or soybean oil) are often added to coating mixtures to enhance film shine (Andres, 1984).

Thickness is critical to the properties of zein coatings. To measure thickness, a thin section of the coated product may be observed under a light microscope (Park et al., 1994b). Another method involves spraying the film-forming solution on a flat surface simulating the coating process on the actual product; data on surface area and density of film-forming solution may be used to calculate film thickness (Mallikarjunan et al., 1997). Water vapor permeability (WVP) and oxygen permeability (OP) are barrier properties commonly investigated to evaluate the ability of edible coatings to protect foods from the environment and from adjacent food components with different aw. Zein-coated, deep-fat fried mashed potato balls (Mallikarjunan et al., 1997) and shell eggs (Wong et al., 1996) retained higher interior moisture levels than uncoated samples. However, water absorption of zein coatings increased at high RH (Kanig and Goodman, 1962). Zein coatings had oxygen and carbon dioxide transmission rates lower (by one or two orders of magnitude) than those of low-density polyethylene (LDPE), methylcellulose (MC), and hydroxypropyl cellulose (HPC) films and similar to those of polyester films at the same testing temperature and RH (Guilbert, 1986; Kester and Fennema, 1986; Hanlon, 1992; Krochta, 1992; Park et al., 1994b). OP of zein coatings increased with increasing plasticizer levels (Butler and Vergano, 1994). Films of moderate oxygen and carbon dioxide barrier ability reduced respiration rates of vegetables by limiting exposure to ambient oxygen and by increasing internal carbon dioxide concentration (Park et al., 1994a, 1994b).

Formation of free-standing zein films for food and packaging applications has become the focus of intensive research and development (Guilbert, 1986; Aydt and Weller, 1988; Aydt et al., 1989; Gennadios and Weller, 1990, 1994;

Zein Films

49

Gennadios et al., 1993a; Wu, 1995; Herald et al., 1996; Park et al., 1996b; Krochta and De Mulder-Johnston, 1997; Lai and Padua, 1998). Zein films are generally cast from alcohol solutions. Zein is dissolved into warm (65 to 85 C) aqueous ethanol or isopropanol with added plasticizers. The solution is cooled (40 to 50 C), allowing bubbling to cease prior to casting (Gennadios et al., 1993b). The zein solution is then poured into a well-shaped plate, usually a glass plate, where films are formed as the alcohol evaporates from the film surface. Dried films are peeled off the plate surface. Plasticizers are added to polymers to reduce brittleness. They decrease intermolecular forces between polymer chains, thus increasing flexibility and extendibility. Commonly used plasticizers are liquid organic compounds such as polyols, mono/di/oligosaccharides, lipids, and lipid derivatives (Reiners et al., 1973). Glycerol is often used to plasticize zein films. However, it tends to migrate from the bulk of the film matrix to the surface. According to Park et al. (1994c), the interaction between protein molecules and glycerol is weak, and excess glycerol easily migrates through the film matrix. Cast zein films are initially transparent, but after only a few hours they appear greasy and “cloudy” because of glycerol “sweating out.” Plasticizer migration results in loss of film flexibility (Park et al., 1992). Park et al. (1994c) used a mixture of glycerol and PEG (average molecular weight 400). They observed that migration rates of glycerol-PEG mixtures in zein-based films were slower than that of glycerol alone and such mixtures could slow down the deterioration of mechanical properties during film storage. Hydrophilic plasticizers (e.g., glycerol) act as humectants, retaining the moisture that plasticizes zein. However, they are not effective at temperatures below freezing or in low RH environments. As water freezes or evaporates, zein reverts to a stiff and brittle material. Fatty acids have been used to plasticize zein films (Kanig and Goodman, 1962; Reiners et al., 1973; Park et al., 1994d; Masco-Arriola, 1996; Lai et al., 1997; Lai and Padua, 1998). Unsaturated fatty acids remain effective plasticizers at temperatures below freezing and in low RH environments. However, Kanig and Goodman (1962) reported that oleic acid-plasticized zein films became brittle and almost colorless over a storage period of six weeks in a light stability chamber. Property changes were attributed to oleic acid oxidation.

Several studies reported quantitative information on tensile properties, such as tensile strength (TS) and elongation at break (E), of zein films (Table 2.1). Multiple factors, such as film composition, temperature, RH, and storage time, affect tensile properties. TS of zein films was similar to that of wheat gluten films and two- to threefold lower than that of MC and HPC films (Gennadios et al., 1993a). Generally, adding plasticizers decreased TS and increased E (Park

50

Zein Films

51

et al., 1992, 1994c; Santosa and Padua, 1999). Zein films containing only glycerol as a plasticizer showed low E (3%) (Park et al., 1994c), but E increased (76%) as the PEG/glycerol ratio increased from 0:100 to 50:50. Results were attributed to differences in structure and molecular size between glycerol and PEG. Also, TS was highest at the 50:50 PEG/glycerol ratio for several plasticizer concentrations tested. However, after 20 days of storage (at 25 C and 50% RH), samples became brittle showing higher TS and low E (Park et al., 1994c). Gennadios et al. (1993a) reported that TS of zein films varied substantially with conditioning at different combinations of RH (23 to 75%) and temperature (5 to 45 C). TS decreased with RH and increased with temperature. A regression model linear in RH and quadratic in temperature fitted well (R2 0.96) the TS data of zein films (Gennadios et al., 1993a). Kanig and Goodman (1962) reported that zein films plasticized with oleic acid softened after one week of storage at 93% RH. TS of zein films increased, about twofold, by adding cross-linking agents to the alcoholic film-forming solutions (Yamada et al., 1995; Yang et al., 1996; Parris and Coffin, 1997). Cross-linking agents included formaldehyde, glutaraldehyde, epichlorohydrin, citric acid, 1,2,3,4-butanetetracarboxylic acid, polymeric dialdehyde starch, and 1,2-epoxy-3-chloropropane. Park et al. (1994d) increased TS of cast zein films by laminating them with MC films. Also, E increased by laminating zein films with soy protein isolate or wheat gluten films (Foulk and Bunn, 1994).

Zein films have WVP values lower than or similar to those of other protein films (Guilbert, 1986), cellulose ethers, and cellophane (Kanig and Goodman, 1962; Taylor, 1986; Krochta, 1992). However, their WVP is notably higher than that of LDPE (Smith, 1986) or ethylene-vinyl alcohol copolymer (EVOH) (Foster, 1986). Plasticization affects WVP of zein films (Table 2.1). Films containing no plasticizer had lower WVP values than those plasticized with glycerol or mixtures of glycerol with PEG or polypropylene glycol (PPG) (Mendoza, 1975; Park et al., 1994c; Park and Chinnan, 1995). Plasticizer incorporation into a polymer, either during fabrication or subsequently by permeation, increases the mobility of chain segments and consequently increases the effective diffusion coefficient of permeates (Seymour and Carraher, 1984). Gontard et al. (1993) and Cherian et al. (1995) explained that higher gas permeability was due to an increase in free volume between biopolymer molecular chains. WVP also increased with RH, likely due to moisture plasticization and to increasing vapor pressure differences between the two sides of the film (Park and Chinnan, 1990; Butler and Vergano, 1994; Lai and Padua, 1998). In moist environments, protein films tend to swell and change their macromolecular

52 structure due to high water sorption (Watt, 1983); as a result, diffusion of water molecules dramatically increases leading to high WVP. However, the low water vapor barrier ability of zein films could be an advantage for zein-based food packaging films. It would allow water vapor movement across the film, thus preventing water condensation inside the package, which could lead to microbial spoilage (Ben-Yehoshua, 1985). Parris et al. (1997) proposed a new method for isolating zein from rinsed, dry milled corn by extraction with 70% (v/v) ethanol. Films prepared from isolated zein, containing 1% starch, had lower WVP values and were more water resistant than films prepared from commercial zein. However, water resistance decreased with increasing starch content. Preparation of protein-based films with low WVP normally requires addition of lipid materials or treatment with cross-linking agents (Avena-Bustillos and Krochta, 1993; Gontard et al., 1994; McHugh and Krochta, 1994a, b). Application of a carnauba wax layer notably decreased WVP of zein films from 9.07 to 0.115 g mm/m2 h kPa (Weller et al., 1998). Substantial reductions in WVP of zein films by the application of wax layers were expected because waxes are resistant to moisture transport (Kester and Fennema, 1986). Bilayer films were produced by combining zein films with MC films (Park et al., 1994d, 1996a, b) or with soy protein/wheat gluten films (Foulk and Bunn, 1994). Generally, bilayer films are prepared by pouring, spraying, or rolling a second film directly on top of a previously prepared one. Foulk and Bunn (1994) sprayed the second layer with a worm screw, which allowed an electric paint sprayer to move at a constant speed while dispensing a uniform amount of solution. Spraying gave a very thin second layer that dried fast, thus shortening exposure of the dried first layer to the solvent of the second layer. Bilayer films were also formed by laminating together two independently prepared single-layer films using a solvent between them while applying uniform pressure. WVP of laminated MC/zein-fatty acid films decreased as chain length and concentration of lauric, stearic, and palmitic acid increased (Park et al., 1994d). Zein/soy protein and zein/wheat gluten films had lower WVP than single zein films (Foulk and Bunn, 1994). Another method to laminate two individually formed films used heat and pressure from a set of rolls (Foulk and Bunn, 1994). Another approach to reduce WVP is the use of cross-linking agents. Polymeric dialdehyde starch (PDS) and 1,2-epoxy-3-chloropropane have been used to improve water resistance of zein films (Yamada et al., 1995; Parris and Coffin, 1997). Parris and Coffin (1997) reported a reduction in WVP when PDS was added (at 20% w/w of zein) to zein films without using any plasticizers. However, PDS did not affect WVP when glycerol or PPG were added to the films. Oxygen and carbon dioxide permeability values of zein films were low at 0 to 50% RH (Gennadios and Weller, 1990; Park and Chinnan, 1990; Aydt et al., 1991; Gennadios et al., 1993b). They were lower than those of other

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polysaccharide and composite polysaccharide/lipid films (Greener and Fennema, 1989). However, they were higher than those of wheat gluten films (Park and Chinnan, 1990; Aydt et al., 1991; Gennadios et al., 1993b). Kretschmer (1957) attributed this to the high -helical content of zein. Oxygen molecules apparently can more readily permeate through the zein helical conformation than through the highly cross-linked gluten structure. OP of zein films also was lower than that of common plastic films, such as LDPE, high-density polyethylene, polypropylene, polystyrene, and unplasticized polyvinyl chloride (Billing, 1989). Polyamide-6, a packaging material regarded as a good oxygen barrier (Billing, 1989), was slightly more permeable to oxygen than zein, wheat gluten, or wheat gluten/soy protein isolate films (Gennadios et al., 1993b). Plasticizers affect gas permeability of films. Increasing plasticizer content in zein films increased OP (Park and Chinnan, 1995; Yamada et al., 1995). Linear relationships were observed between gas permeability and film thickness (Yamada et al., 1995). RH markedly affects oxygen barrier characteristics of protein films. Moisture has a plasticizing or swelling effect on hydrophilic polymers resulting in increased gas permeability (Ashley, 1985). Moisture-sensitive, oxygen-barrier synthetic polymers must be either co-polymerized with a hydrophobic polymer, “sandwiched” between hydrophobic polymer layers, or treated with cross-linking agents to limit their exposure to moisture. Temperature also affects film OP. Gennadios et al. (1993b) determined OP of zein films at 7, 15, 25, and 35 C under 0% RH. The data followed the Arrhenius activation energy model (lnP lnP0 (Ea/RT)). Ea (activation energy) and P0 (Arrhenius constant) values were estimated at 11.1 kcal/mol and 7.9 mol/m s Pa, respectively.

Zein may be plasticized in solution with long chain fatty acids and collected as a soft solid mass after precipitation with cold water. Zein films may be drawn from wet, freshly precipitated resins or formed by extrusion of dry resin pellets (Figure 2.1). Laboratory preparation of drawn films involves (1) a plasticization step in which a zein solution in warm aqueous ethanol (70%) is stirred with fatty acids (0.5 to 1 g fatty acid/g zein); (2) a resin formation step, in which cold water is added to the solution to precipitate the plasticized zein; and (3) a kneading step where the precipitate is collected and kneaded into a cohesive and elastic moldable mass. The hydrated resin is very ductile and extendible. Films are drawn from the soft resin, extended over rigid frames, and allowed to dry at room conditions (Lai et al., 1997; Lai and Padua, 1997). Dry films (~0.030 mm thick) are translucent, flexible, ductile, and heat sealable (Figure 2.2). Under the microscope, they show a fiber network structure with pinholes and structural gaps (Figure 2.3).

Figure 2.1 Extruded zein resin pellets.

Figure 2.2 Zein resin rolled films and drawn film.

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Figure 2.3 Light microscope image of drawn zein films showing fiber network.

Fusion lamination, performed by heating under pressure (11,000 psi and 100 C) several sheets together, produced transparent zein films that were more ductile and pliable than the original ones. Rakotonirainy and Padua (1999) suggested that films melted and flowed under the applied pressure, filling pinholes and gaps in the structure, thus reducing macro-defects and increasing film uniformity. To facilitate storage and handling, wet resins were rolled into sheets and dried in a cabinet drier (at 90 C for four hours). Dried resins could be melted and formed into films by extrusion or softened with aqueous ethanol for cold rolling (Ha, 1999). Films produced by either method were translucent and flexible. They could be thinned to gauge by hot rolling.

Typical TS, E, and Young’s modulus values of drawn zein films at 25 C and 50% RH were 9 MPa, 12%, and 267 MPa, respectively (Lai and Padua, 1997). For comparison, TS and E values of 3 to 11 MPa and 25 to 50%, respectively, were reported for collagen films (Hood, 1987). Typical TS and E values for LDPE films are 9 to 17 MPa and 500%, respectively (Briston, 1986). Ductility

56 of drawn zein films increased at high RH, as expected for protein films. For example, E increased from 12 to 30%, and Young’s modulus decreased from 267 to 150 MPa when RH increased from 50 to 98%, while TS was not significantly (P > 0.05) affected (Lai and Padua, 1998). This was attributed to zein plasticization by moisture. Films softened when stored at 98% RH, in agreement with previous studies (Kanig and Goodman, 1962; Gennadios et al., 1993a). Tensile properties of zein/fatty acid films were affected by fatty acid content. Increasing oleic acid from 0.5 to 1.0 g/g zein decreased TS and increased E. Films containing 0.5 g oleic acid/g zein had 9 MPa TS and 12% E, while films prepared with 0.6 g oleic acid/g zein had 6 MPa TS and 44% E (Santosa and Padua, 1999). Cold-rolled films could be re-plasticized, to increase ductility, by heating them in fatty acid baths. Re-plasticization increased fatty acid content from 40 to 60%. Re-plasticized films showed increased E (154%) and decreased TS (3.3 MPa). Tensile properties of extruded films were similar to those of rolled films (Ha, 1999).

Sorption isotherms of drawn zein films showed slow water absorption at aw values up to 0.85 (Figure 2.4). Water was more rapidly absorbed at aw > 0.85. Drawn films had lower equilibrium moisture content (7 g water/100 g solids) than cast zein films of identical composition (12 g water/100 g solids) at aw of

Figure 2.4 Moisture sorption isotherms of drawn (䊏) and cast (䉬) zein films plasticized with oleic acid (adapted from Lai and Padua, 1998).

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0.95. Differences were attributed to extensive zein-fatty acid interactions occurring in drawn films, which reduced zein exposure to water (Lai and Padua, 1998). Water absorption of rolled films was affected by plasticization. Due to their hydrophobic nature, zein films resisted wetting. However, prolonged immersion in water resulted in sheet swelling and loss of dimensional stability (Santosa and Padua, 1999). Fatty acid content of films affected their water absorption (Santosa and Padua, 1999). Films containing 0.6 g oleic acid/g zein absorbed 20% water after 24 hours of immersion at 25 C, while films containing 0.8 g oleic acid/g zein absorbed 7% water under the same conditions (Santosa and Padua, 1999). Films prepared with linoleic acid showed less water absorption than those plasticized with oleic acid. Re-plasticization further decreased water uptake to ~4% at 48 hours of immersion (Figure 2.5). Water absorption of extruded films was similar to that of rolled films.

WVP of drawn zein films was relatively low (0.06 g mm/m2 h kPa at 25 C and 0/98% RH gradient) (Lai and Padua, 1998) compared to hydrophilic biopolymers such as wheat gluten (2.2 g mm/m2 h kPa at 21 C and 0/85% RH gradient) (Park and Chinnan, 1990) and cellophane (0.3 g mm/m2 h kPa at 38 C and 0/90% RH gradient) (Taylor, 1986), but an order of magnitude higher than that of LDPE films (0.003 g mm/m2 h kPa at 38 C and 0/90% RH gradient) (Smith, 1986). WVP varied with RH, changing from 0.04 to 0.06 g mm/m2 h kPa when RH increased from 50 to 98% (Lai and Padua, 1998). Temperature also affected permeability with WVP being lower at 25 C than at 5, 15, or 35 C (Lai and Padua, 1998). Polymer films often show an increase in WVP with increased temperature resulting from a higher diffusion coefficient due to increased molecular mobility of plasticizers and polymer chains. However, WVP of zein films also increased at lower temperatures (5 and 15 C). This was possibly due to water condensation on the film surface, which plasticized the protein and increased WVP (Lai and Padua, 1998). Films drawn from zein resins were coated with drying oils (linseed or tung oils) to decrease WVP. Upon exposure to ambient air, drying oils polymerize to form tough, water-repellent coatings. Rakotonirainy and Padua (1999) measured WVP of drawn zein films dipped in laboratory-prepared and commercial formulations of tung and linseed oils. Coating films with a commercial formulation of tung oil reduced WVP by tenfold to 0.005 g mm/m2 h kPa at 25 C and 0/98% RH gradient (Rakotonirainy and Padua, 1999).

Morphological differences were observed between cast and drawn films plasticized with oleic acid at 0.5 g/g zein (Lai and Padua, 1997). Cast films

58

Figure 2.5 Water absorption rate of zein films plasticized with linoleic acid (adapted from Santosa and Padua, 1999). Filled symbols represent data points for films with 0.8 g linoleic acid/g zein; open symbols represent re-plasticized films containing 60% linoleic acid.

showed a different finish between their two sides. The side in contact with the casting surface was smooth and glossy, whereas that in contact with air during drying was irregular and opaque. Drawn film surfaces were both smooth and of low gloss. Scanning electron microscopy (SEM) images showed differences in surface morphology between the two types of films. The glossy side of cast films appeared featureless, whereas the opaque side showed globular deposits, possibly due to fatty acids separated from the zein matrix by the evaporating solvent. DSC scans showed differences in thermal behavior between cast and drawn films of identical composition (Figure 2.6). Melting peaks in cast films indicated fatty acid phase separation, while no phase separation was detected in drawn films. Thermograms suggested a more extensive interaction between zein and fatty acids in drawn films than in cast films. SEM images of drawn films in Figure 2.7 show the effect of resin drawing and stretching on film morphology. Figure 2.7(a) shows the central portion of the film, where stretching was more effective, while Figure 2.7(b) shows the outer edge of films where stretching was not apparent. DSC scans of cold-rolled films (0.7 g oleic acid/g zein) showed endothermic peaks corresponding to the melting of oleic acid (Figure 2.8), which indicated fatty acid phase separation in those films. Melting peaks disappeared after films were heated above 100 C, suggesting that heat treatment promoted fatty acid

Figure 2.6 DSC scans of drawn (solid lines) and cast (dotted lines) zein films plasticized with oleic acid. Reprinted with permission from Lai and Padua, 1997, Cereal Chemistry, 74: 771–775.

Figure 2.7 SEM images of drawn zein films showing the effect of stretching. (a) Central portion of film showing the effect of stretching. (b) Outer edge of films where stretching was not apparent (adapted from Lai and Padua, 1997).

59

60

Figure 2.8 DSC scans of rolled zein films plasticized with oleic acid (adapted from Santosa, 1999).

binding to zein and prevented phase separation. Extruded films of similar fatty acid content showed no evidence of phase separation (Ha, 1999). Zein binding of fatty acids was proposed by Izzo and Ho (1989) to explain an observed reduction in extractable fatty acids after extrusion of zein and corn oil mixtures. Chiue et al. (1994, 1997) suggested that the observed antioxidant activity of zein may be attributable to its capacity to bury unsaturated fatty acids in its inter- or intramolecular spaces, a phenomenon that likely involves the amide groups of zein.

Among cereal proteins, those of wheat gluten are the most extensively studied due to their bread-making properties. Electron micrographs and X-ray measurements on wheat gluten have yielded a structural model (Grosskreutz, 1960, 1961) that could provide insight on the structure of zein resins. Wheat proteins and phospholipids (7% of gluten) are critical to the structure of wheat gluten. Grosskreutz (1961) proposed that stretched gluten consisted of films predominantly parallel to the surface. Each sheet is formed of protein platelets hydrogen-bonded together through an interstitial aqueous phase under the action of hydration and mechanical working. Phospholipids, organized in bilayers, were thought to be located between the platelet films acting as slip planes between them. The structure of zein films was investigated by wide-angle (WAXS) and small-angle (SAXS) X-ray scattering (Lai et al., 1999). WAXS results indicated that both cast and drawn films had similar d-spacings of 4.6 and 10.5 Å

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corresponding to the -helix backbone distance along the chain and the inter-chain spacing between helices, respectively. Granular zein gave slightly smaller values. Apparently, film formation by resin deformation or solution casting did not change the basic internal structure of the -helix. SAXS results, on the contrary, showed that drawn films had a strong periodicity across the film plane that was not observed for cast films. Results were interpreted in terms of a platelet structure in drawn films developed during the doughing process and aligned during film formation. Oleic acid appeared to play an important role in the formation of platelet structures. Contrary to phospholipids in wheat gluten films examined by Grosskreutz (1960), oleic acid was present in sufficient concentration (~35%) to develop a periodic protein-fatty acid layered complex throughout the zein film. Granular zein did not show any SAXS periodicity. Lai et al. (1999) proposed a structural model for drawn films based on X-ray measurements (Figure 2.9) and on the zein molecular dimensions determined by Matsushima et al. (1997) for non-reduced zein. Zein structural prism-like units of 160 46 46 Å, consisting of ribbons of folded (antiparallel) -helical segments, formed protein tetramers. Layers of double-stacked zein units, measuring 160 x 92 x 46 Å, alternated with bilayers of fatty acid (42 Å), which conferred flexibility to the films. The structural model suggested that zein plasticization with oleic acid involved dispersion of zein molecules in aqueous alcohol followed by electrostatic adsorption of fatty acids to the charged residues on the zein surface. Addition of cold water during resin precipitation resulted in hydrophobic aggregation of zein-oleic acid units to form films. Sheet formation was facilitated by kneading and rolling actions applied to the resin mass. The absence of melting peaks in DSC scans of zein-oleic acid resins indicated that oleic acid was well dispersed and adsorbed to the protein. Heat treatment applied during hot rolling of resin films increased their clarity and flexibility. The effect was attributed to increased interaction between zein and oleic acid. Heating possibly induced unfolding of the prism-like zein molecule and exposed new sites for binding of oleic acid. Excessive heating or mechanical action of the initial zein and oleic acid dispersion caused protein gelation, which interfered with resin formation upon addition of cold water. It was hypothesized that zein unfolded under heat and mechanical action, promoting zein interchain bonds and preventing oleic acid binding to the protein. Zein gels readily expelled oleic acid upon mechanical treatment, preventing resin formation.

The potential increase in zein availability has spurred considerable research on its properties and utilization. Pharmaceutical coatings and confectioner’s glaze remain the bulk of zein commercial applications. However, intensive re-

Figure 2.9 Structural model of drawn zein films. From Lai et al., 1999, Journal of Applied Polymer Science, 71: 1267–1281. Copyright 1999. Reprinted by permission of John Wiley & Sons, Inc.

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search on zein utilization for edible and biodegradable film formulations has resulted in improved performance. Plasticization with polyols and fatty acids reduces stiffness and increases flexibility of films. Heat treatment under high pressure improves clarity and texture of films. Improvement of water barrier properties is accomplished by coating or laminating zein films with water-resistant lipid materials. Scalable preparation methods based on extrusion technology have recently been proposed. However, film mechanical properties remain a challenge to researchers. The high tensile strength of polymeric films used in packaging allows use of smaller thickness gauge, thus reducing packaging weight and cost. Present methods to increase tensile strength of zein films/sheets often result in increased stiffness and reduced flexibility. However, competitive performance levels of zein-based packaging films seem attainable. Zein-based films are resistant to insect and rodent attack, freezing and thawing cycles, and long-time exposure to sunlight and rainfall. Also, lipid-coated films resist exposure to high RH environments. Continuing research addresses the use of cross-linking agents, film morphology characterization, time-dependent mechanical properties, plasticizer oxidation rates, biodegradability rates, and processing technology. Results are expected to contribute to increased utilization of agricultural products.

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66 McHugh, T.H. and Krochta, J.M. 1994b. “Water Vapor Permeability Properties of Edible Whey Protein-Lipid Emulsion Films,” JAOCS, 71:307–312. Mendoza, M. 1975. Preparation and Physical Properties of Zein Based Films, M.S. Thesis, University of Massachusetts, Amherst, USA. Meyer, G.A. and Mazer, T.B. February 4, 1997. U.S. patent 5,599,556. Mickus, R.R. 1955. “Seals Enriching Additives on White Rice,” Food Engr., 27:91–93, 160. Oshlack, B., McGinity, J., Chasin, M. and Bodmeier, R. October 18, 1994. U.S. patent 5,356,467. Park, H.J. and Chinnan, M.S. 1990. “Properties of Edible Coatings for Fruits and Vegetables,” ASAE Paper No. 90-6510, St. Joseph, MI: American Society of Agricultural Engineers. Park, H.J. and Chinnan, M.S. 1995. “Gas and Water Vapor Barrier Properties of Edible Films from Protein and Cellulosic Materials,” J. Food Engr., 25:497–507. Park, H.J., Chinnan, M.S. and Shewfelt, R.L. 1994a. “Edible Coating Effects on Storage Life and Quality of Tomatoes,” J. Food Sci., 59:568–570. Park, H.J., Chinnan, M.S. and Shewfelt, R.L. 1994b. “Edible Corn Zein Film Coatings to Extend Storage Life of Tomatoes,” J. Food Process. Preserv., 18:317–331. Park, H.J., Bunn, J.M., Weller, C.L., Vergano, P.J. and Testin, R.F. 1994c. “Water Vapor Permeability and Mechanical Properties of Grain Protein-based Films as Affected by Mixtures of Polyethylene Glycol and Glycerin Plasticizers,” Trans. ASAE, 37:1281–1285. Park, J.W., Testin, R.F., Park, H.J., Vergano, P.J. and Weller, C.L. 1994d. “Fatty Acid Concentration Effect on Tensile Strength, Elongation, and Water Vapor Permeability of Laminated Edible Films,” J. Food Sci., 59:916–919. Park, J.W., Testin, R.F., Vergano, P.J., Park, H.J. and Weller, C.L. 1996a. “Fatty Acid Distribution and Its Effect on Oxygen Permeability in Laminated Edible Films,” J. Food Sci., 61:401–406. Park, J.W., Testin, R.F., Vergano, P.J., Park, H.J. and Weller, C.L. 1996b. “Application of Laminated Edible Films to Potato Chip Packaging,” J. Food Sci., 61:766–768, 777. Park, H.J., Weller, C.L., Vergano, P.J. and Testin, R.F. 1992. “Factors Affecting Barrier and Mechanical Properties of Protein-based Edible, Degradable Films,” Paper No. 428, presented at the Annual Meeting of the Institute of Food Technologists, June 20–24, New Orleans, LA. Parris, N. and Coffin, D.R. 1997. “Composition Factors Affecting the Water Vapor Permeability and Tensile Properties of Hydrophilic Zein Films,” J. Agric. Food Chem., 45:1596–1599. Parris, N., Dickey, L.C., Kurantz, M.J., Moten, R.O. and Craig, J.C. 1997. “Water Vapor Permeability and Solubility of Zein/Starch Hydrophilic Films Prepared from Dry Milled Corn Extract,” J. Food Engr., 32:199–207. Pedersen, K., Devereux, J., Wilson, D.R., Sheldon, E. and Larkins, B.A. 1982. “Cloning and Sequence Analysis Reveal Structural Variation among Related Zein Genes in Maize,” Cell, 29:1015–1026. Rakotonirainy, A.M. and Padua, G.W. 1999. “Effect of Coating and Lamination Processes on Tensile Properties and Water Vapor Permeability of Zein Films,” Paper No.11B-25 presented at the Annual Meeting of the Institute of Food Technologists, July 24–28, Chicago, IL. Reiners, R.A., Wall, J.S. and Inglett, G.E. 1973. “Corn Proteins: Potential for Their Industrial Use” in Industrial Uses of Cereals, ed., Y. Pomeranz, St. Paul, MN: American Association of Cereal Chemists, pp. 285–298. Santosa, F.X.B. 1999. Preparation and Properties of Zein-based Flexible Films, Ph.D. Dissertation, University of Illinois at Urbana-Champaign, USA. Santosa, F.X.B. and Padua, G.W. 1999. “Tensile Properties and Water Absorption of Zein Films Plasticized with Oleic and Linoleic Acids,” J. Agric. Food Chem., 47:2070–2074. Seymour, R.B., and Carraher, C.E. 1984. Structure-Property Relationships in Polymers. New York: Plenum Press, pp. 107–111. Shukla, P.T. 1992. “Trends in Zein Research and Utilization,” Cereal Foods World, 37:225. Smith, S.A. 1986. “Polyethylene, Low Density” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker, New York: John Wiley & Sons, pp. 514–523.

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Formation and Properties of Wheat Gluten Films and Coatings

HE very high molecular weight, markedly apolar character, complexity, and diversity of their fractions are features of wheat gluten (WG) proteins that can be utilized to make films with novel functional properties, such as selective gas barrier properties and rubber-like mechanical properties. WG-based materials are homogeneous, transparent, mechanically strong, and relatively water resistant. They are biodegradable and a priori biocompatible, apart from some WG-specific characteristics such as allergenicity. They also are edible when food-grade additives are used, and the presence of impurities is avoided. The moisture barrier properties of WG-based films are relatively poor (water vapor permeability around 5 10 12 mol/m s Pa) as compared to synthetic films, such as low-density polyethylene (LDPE) (0.05 10 12 mol/m s Pa). However, this feature could be interesting for packaging cheese, fruits, and vegetables, or for films for agricultural uses, or cosmetic applications. The barrier properties of WG-based films can be substantially improved to approximate those of polyethylene (PE) films by incorporating fatty compounds (e.g., beeswax and paraffin) in the film structure. Many mechanical properties of WG-based films have been determined and modeled. For the most resistant WG-based materials, critical fracture deformation (Dc 0.7 mm) values are slightly lower than those of synthetic reference materials, such as LDPE (Dc 2.3 mm) and cellulose films (Dc 3.3 mm). Addition of fibers to form compos-

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70 ite materials can considerably enhance the mechanical properties of WG films. The thermoplastic properties of WG proteins and their moisture resistance are especially interesting for natural “resin” applications. The gas (O2, CO2, and ethylene) barrier properties of WG-based films are highly interesting, as they are exceptionally good at low relative humidity (RH) conditions. O2 permeability values (around 1 amol/m s Pa) of WG-based films at dry conditions are close to those of ethylene-vinyl alcohol (EVOH) (0.2 amol/m s Pa) and are much lower than those of LDPE (1000 amol/m s Pa). Gas barrier properties closely depend on RH and temperature. This effect is more pronounced for hydrophilic gases (CO2) than for hydrophobic gases (O2). Thus, the CO2/O2 selectivity coefficient can rise from 3 to more than 50 when RH increases from 0 to 100% and temperature increases from 5 to 45°C. Monitoring changes in the gas composition around fresh mushrooms packaged with WG films confirmed the high selectivity of the films, as CO2 and O2 concentrations remained around 1–2% despite product respiration levels (Barron et al., 2001). Solute (especially antimicrobial and antioxidant agents) retention properties of WG films have been studied and modeled, thus paving the way for potential applications involving controlled release of beneficial agents for foods (e.g., active coatings), agriculture (e.g., coated seeds), pharmacy (“drug delivery”), or cosmetics. Common cross-linking agents (such as formaldehyde) or specially designed cross-linking agents have been shown to significantly improve the mechanical properties of protein films. Unfortunately, chemically lipophilized WG films did not have enhanced water barrier or water resistance properties (Micard and Guilbert, 2000). Multilayer agromaterials [WG/paper or WG/other biopolymers such as poly(lactic acid)] as well as composite agromaterials (with fibers of cotton, sisal, coconut, or straw) gave good test results in terms of compatibility and properties. Films made from grafted macroperforated PE and WG were formed by hot-press molding of WG-thermosetting resin, and their properties were characterized (Perez-Perez, 1997). Developing such materials should give rise to protein-based films with novel gas, vapor, and solute permeability properties and with mechanical properties similar to those of conventional synthetic films. The thermoplastic characteristics of these protein-based raw materials have also been studied. Water and other plasticizers can lower the glass transition temperature (Tg) of the WG and enable processing at temperatures below those that lead to protein decomposition, which means that protein-based films can be formed by using techniques that are conventionally used with synthetic polymers (e.g., extrusion, injection, and molding). The depressive effect of plasticizers on Tg is relatively well described by the Couchman-Karasz relation, as long as the system remains homogeneous (at temperatures above 0–10 C) (Gontard and Ring, 1996).

Composition, Structure, and Thermal Properties of Wheat Gluten

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Formation of a homogeneous malleable phase (plasticization), using different thermal (at temperatures above the Tg), mechanical (shear), or chemical (additives and degradation) treatments, was investigated on the basis of the viscoelastic and flow properties of malleable materials as a function of temperature, moisture content, and time. “Plasticized gluten” is similar to a structured viscoelastic solid with pseudoplastic behavior (Redl et al., 1999a). Analysis of rheometric functions of plasticized WG revealed that the time/temperature combination principle could be applied. For given mixing conditions, a power law as a function of temperature and plasticizer (e.g., glycerol) content can characterize the complex viscosity of plasticized WG. A study also was conducted using a corotating twin-screw extruder for simulating the flow properties and extrusion conditions (Redl et al., 1999b).

To determine the properties and potential applications of material proteins, it is essential to compare their structural features with those of chemically synthesized organic polymers used to produce plastic materials. Proteins, contrary to homopolymers or copolymers in which one or two monomers are repeated, are heteropolymers comprised of more than 20 different amino acids, each with specific sequences and structures. Due to this molecular diversity, proteins have considerable potential for the formation of linkages that differ with respect to their position, nature, and/or energy (Guilbert and Graille, 1994; Cuq et al., 1998). Novel research on “material proteins” has led to the application of polymer science concepts and tools to investigate macromolecular structure and function relationships. This involves investigating the structure of proteins at different levels (atomic, molecular, and supramolecular arrangements); studying structural variations according to temperature and the presence of functional additives; and simulating the macroscopic properties of macromolecular arrangements (mechanical, optical, thermal, and electrical properties). Protein molecular weights have a substantial effect on protein network structure. They also determine the presence of molecular overlapping, leading to formation of physical nodes. As is the case for synthetic macromolecules, overlapping could occur beyond a critical molecular weight (Mc). But, a high mean molecular weight restricts polymer flow during material formation, which can lead to defects in the end product. The -helix and -sheet secondary structures of proteins are highly stabilized by cooperative hydrogen bonds and can resemble crystalline zones. These zones are important in WG proteins where many repetitive sequences are present. They could be responsible for the formation of regular “crystal lat-

72 tice”-type arrangements, which are known to have a marked effect on the final material properties of conventional synthetic polymers. WG protein-based materials could, thus, be defined as mainly amorphous three-dimensional arrangements stabilized by low-energy interactions that are partially reinforced by regular “crystal lattice”-type arrangements and strengthened by some covalent bonds. The molecular weight is fixed by genetic code, which means that for a given subunit, there is no variability, and, if considering the different classes of grain protein, the polydispersity is low contrary to synthetic polymers. But, during grain maturation and thermomechanical processing, aggregation of subunits occurs, hence, creating polydispersity. Similar to synthetic polymers, protein materials with high polydispersity are generally easier to process but have poorer mechanical properties. The formation of protein-based materials and their final characteristics and properties depends, therefore, on protein structure and structural heterogeneity, heat sensitivity, and hydrophilic/hydrophobic characteristics. It is still ambitious to elaborate structure/function relationships in this field of protein-based materials, but it appears to be important to have some knowledge of the structure and thermal properties of WG proteins to understand the specificity of WG-based materials.

Vital WG is the cohesive and elastic mass that is leftover after starch is washed away from wheat flour dough. Commercially, it is an industrial by-product (worldwide annual production of 400,000 metric tons) of wheat starch production via wet milling. Whereas dry wheat flour comprises 9–13% protein and 75–80% starch, WG consists mainly of wheat storage protein (70–80%, dry matter basis) with traces of starch and non-starch polysaccharides (10–14%), lipids (6–8%), and minerals (0.8–1.4%). WG is suitable for numerous food and nonfood uses. Its main application is in the bakery industry, where it is used to strengthen weak flours rendering them suitable for bread baking. It is widely accepted that the unique rheological properties of WG are derived from its protein composition, with lipids and carbohydrates being contaminants entangled in the protein matrix. Wheat proteins have been studied for over 200 years; however, it was not until 1907 when a first comprehensive study was undertaken by Osborne. He distinguished four wheat protein classes based on their solubility in different solvents, namely, albumins, globulins, gliadins, and glutenins. The albumins and globulins (15–22% of total protein), which are, respectively, water- and salt-soluble, are removed with starch granules during gluten processing. In contrast, the gliadins, which are alcohol-soluble, and the glutenins, which are soluble (or at least dispersible) in di-


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lute acid or alkali solutions, are being collected into gluten. Gliadins and glutenins amount to up to 85% (w/w) of total wheat flour protein and are evenly distributed in gluten [33–45% (w/w) gliadin, 40–45% (w/w) glutenin] (Wrigley and Bietz, 1988).

Gliadin was also named prolamin owing to its large proline (14–17 g/100 g) and glutamine (38–40 g/100 g) contents. Its total content of charged amino acids (basic or acidic) is below 6 g/100 g, making gliadin a poorly ionized species across the entire pH range. The solubility of gliadin in 60–70% alcohol results from its peculiar amino acid composition. Gliadin includes several protein bands when analyzed by acid or sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) under reducing or nonreducing conditions (Lafiandra and Kasarda, 1985). Genes encoding for gliadin are located on the chromosomes of groups 1 and 6. More than 75 allelic blocks, each coding for several proteins, have been characterized (Metakovsky, 1991). Polymorphism of gliadin proteins has been widely studied using various electrophoretic or chromatographic techniques, thus providing a useful basis for identification of wheat genotypes (Bietz and Cobb, 1985). As a rule, wheat gliadin patterns comprise 20 to 30 different polypeptides ranging in molecular mass from 30 to 74 kDa (with most above 50 kDa). According to their increasing mobility at acidic pH, polypeptides are classified into -gliadin, -gliadin, -gliadin, and -gliadin. Analysis of genes encoding for gliadin revealed homology of strong sequences from - and -gliadin, whereas complete sequence for -gliadin is still required. Gliadin contains repetitive sequences rich in proline and glutamine that form loose helical structures consisting of repetitive -reverse turns, usually associated with more extensive nonrepetitive regions, which are compact and globular (Tatham et al., 1990). The lack of cysteine residues distinguishes -gliadin polypeptides. Other gliadin types include 6 ( - and -gliadin) or 8 ( -gliadin) cysteine residues, all involved in intramolecular disulfide (SS) bonds. Owing to its peculiar composition, -gliadin is among the more heat-resistant WG proteins, remaining extractable after severe heating (Wrigley et al., 1980). Gliadin molecules may interact together or with glutenin molecules via hydrophobic interactions and hydrogen bonds. Evidence of increasing intermolecular -sheet structures has been obtained by infrared (IR) spectroscopy for -gliadin when the water content increased from the dry state up to 40% (w/w) (Wellner et al., 1996). In the fully hydrated state, gliadin exhibits viscous flow properties without significant elasticity. For cereal technologists, gliadin accounts for the extensibility of wheat flour dough and acts as a filler diluting glutenin interactions.

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Glutenin has similar amino acid composition (rich in proline and glutamine) to gliadin, with a slightly lower content of hydrophobic amino acids (Wrigley and Bietz, 1988). Contrary to gliadin, which is comprised of distinct polypeptide chains, glutenin consists of polymers made from polypeptide chains (also named subunits) linked end-to-tail by SS bonds (Kasarda, 1999). These polymers cover a broad range of molecular masses up to several million Da. Two groups of subunits are distinguished according to their molecular size and their gene location, namely, high and low molecular weight glutenin subunits. High molecular weight glutenin subunits (HMW-GS), also referred to as A subunits, comprise polypeptide chains ranging from 70–105 kDa. Genes (Glu-1) located on the long arm of chromosomes of group 1 encode the polypeptides. From three to five distinct polypeptides are encoded depending on genotype. The allelic variations have been thoroughly studied by Payne and Lawrence (1983). The HMW-GS have been numbered: first, according to their increasing mobility in SDS-PAGE, and thereafter, in the order of their discovery. They are also classified into x or y types. The x type subunits are larger than the y type, and typically have four cysteine residues instead of seven for y subunits. The HMW-GS composition of wheat cultivars has been related to their bread-making potential, and Payne et al. (1983) proposed a quality score for each subunit, which is relevant in breeding strategy. Most of the genes encoding for HMW-GS have been sequenced, and the primary structures of the subunits are accurately known. The amino acid sequences show strong homology, with N and C extremities consisting of preserved nonrepetitive sequences and with a large central domain including hexa (PGQGQQ) and nona (GYYPTSPQQ) repetitive peptide sequences (Halford et al., 1987). Structure prediction indicated that the N- and C-terminal domains are predominantly -helical, while the repetitive domains are rich in -turns. Cysteine positions are well preserved, with one residue located at the C-termini of the molecules, another present in the central domain (for y subunits only), and all others placed at the N-termini. The presence of cysteine residues at both ends of the molecules would insure the formation of linear polymers through inter-polypeptide SS bonds. In addition, it is believed that all cysteine residues are involved in either intramolecular or intermolecular bonds with no free sulfhydryl (SH) groups available. SS mapping is still under study, and the exact pairing of cysteine residues will have to be defined before an accurate description of the polymeric structure of glutenin could be achieved. Low molecular weight glutenin subunits (LMW-GS) have been less studied in terms of genetic polymorphism and primary and secondary structures, although they represent 70–80% of the glutenin mass. Compared with HMW-GS, LMW-GS are numerous, comprising up to 40 different polypeptides ranging from 30–65 kDa. The LMW-GS are encoded by genes


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(Glu-3) located on the short arm of chromosomes of group 1, close to those encoding for gliadin polypeptides (Gli-1). Variations at the Glu-3 loci are still under study, and already, 20 different allele blocks have been identified (Gupta and Shepherd, 1990). Only a few gene sequences, encoding for LMW-GS B and C are available (Cassidy and Dvorák, 1991). After a short preserved N-terminal domain, there is a repetitive domain rich in proline and glutamine (QQPPFS peptide) followed by a large nonrepetitive domain, in which seven cysteine residues with well-preserved location are found. An eighth cysteine residue with another one from the C-termini would be involved in intermolecular SS bonds, the others form intramolecular linkages (Masci et al., 1995).

Several studies aimed at characterizing the molecular composition of glutenin polymers in terms of specificity of SS bond formation or subunit associations are presented here. SS mapping of peptide digests were obtained after enzymatic hydrolysis of glutenin polymer was investigated by Köhler et al. (1993). They demonstrated specific intermolecular bonding between y-HMW-GS and LMW-GS, and between the different types of HMW-GS. Analysis of subunit oligomers produced from stepwise reduction of glutenin polymers (e.g., by reducing agents), or study of polymer formation under oxidation of partly purified glutenin subunits, led to the conclusion that association of glutenin subunits is a nonrandom process (Werner et al., 1992). Based on experimental investigation of the viscosity drop under SS reduction, the glutenin polymer is thought to be a linear SS-bonded polymer (Ewart, 1988). Despite the lack of accurate data on association of subunits, several models of glutenin polymer structure have been proposed (Khan and Bushuk, 1979; Graveland et al., 1985; Kasarda, 1989; Gao et al., 1992). All of these models proposed that intermolecular SS bonds join the glutenin subunits.

Glutenin is believed to be one of the largest natural polymeric molecules with estimated molecular mass of over 107 (Kasarda, 1999). The large molecular size of glutenin impairs its total extractability in SDS buffer, a solvent able to unfold polypeptide structures and known to disrupt protein complexes stabilized by weak forces. Consequently, the total glutenin polymer size distribution cannot be accurately measured by classical methods such as ultracentrifugation and gel permeation chromatography (GPC). Sonication was shown to bring total glutenin polymers into solution, probably by reducing their size distribution range through very limited scissions along the polymer chains (Singh et al., 1990). Nevertheless, even after sonication, the polymers remained too large to

76 be resolved on GPC column. To overcome this problem, recent investigations proposed to combine field flow fractionation (FFF) and multi-angle laser light scattering (MALLS) detection to estimate the size of sonicated glutenin (Stevenson et al., 1999). Sonicated glutenin polymers were shown to have hydrodynamic diameters up to 35 nm, corresponding to a molecular weight of 11 million Da (assuming spherical shape). An increase in SDS-unextractable glutenin is supposed to result from an increasing weight-average molecular weight (Mw) of glutenin polymer. Southan and MacRitchie (1999) proposed that assumptions relevant to synthetic polymers might apply to glutenin polymers. In that respect, the mixing properties of wheat flour dough were related to the amount of SDS-unextractable polymers in flour (Gupta et al., 1993). Also, Tsiami et al. (1997) showed that the rheological properties of gluten subfractions were related to their size distribution ranges.

Deamidated WG, obtained with acidic or basic treatments at high temperature, is commercially available. The treatment provokes some peptide bond scission but, especially, converts glutamine into glutamic acid. This change insures increased charge density on the protein and enhances electrostatic repulsion, thereby promoting water solubility of the deamidated WG. Deamidation levels as low as 2–6% were adequate to obtain water-soluble WG (Matsudomi et al., 1982). Protein conformation changes upon deamidation and hydrophobic residues are exposed at the surface of the molecules. The combination of charged amino acids and hydrophobic surfaces provides amphiphilic properties to deaminated WG. Consequently, it gains emulsifying and foam-stabilizing capabilities. Limited hydrolysis of WG by proteases also increases water solubility within the entire pH range. Emulsifying properties are especially good at low pH (around 4) and for a limited degree of hydrolysis (1–2%) (Mimouni et al., 1994). Using a water-soluble WG derivative allows the formation of film-forming solutions without ethanol as a co-solvent.

Wheat-based food products can be toxic for people suffering from celiac disease. The condition of celiac disease results in lesions of the small intestinal mucosa (villi atrophy), which impair absorption of nutrients. It has been recognized as being caused by wheat proteins in 1952. The onset of illness most commonly occurs around the age of two, after wheat has been introduced into the diet, and in early adult life (third and fourth decades). However, celiac disease


77

can begin at any age. This gluten-sensitive enteropathy is not a food allergy but a genetically inheritable disease linked to specific histocompatibility cell antigen (HCA system). Adherence to a gluten-free diet abolishes symptoms, and intestinal mucosa reverses to completely normal histology and function. Specific peptide sequences especially rich in proline and glutamine (Pro-Ser-Gln-Gln and Gln-Gln-Gln-Pro) activate the cellular immune system (T cells), which probably produces the actual enterocyte injury (Schuppan, 2000). Those peptides are found in wheat gliadins, glutenin, rye secalins, barley hordeins, and, in a much lower amount, oat avenins.

The structure/properties relationships of hydrated biopolymers could be better understood through the theories of glass transition used in polymer science, in terms of critical variables of time, temperature, and plasticizer content (Slade, 1984; Slade et al., 1989). Glass transition of an amorphous or partially amorphous polymer is typically described as a transition from a brittle glass to a highly viscous or liquid-like rubber. The glass transition separates the two domains according to clear structural differences, and the properties of polymers are seriously modified when the temperature rises above the Tg. In isothermic conditions, the addition of plasticizers has theoretically the same effect on molecular mobility as increased temperature. Plasticizers, which are generally low molecular weight components, increase free volume of the polymer system, allowing increased backbone chain segmental mobility. Despite their wide variety of structure, reflecting the diversity of the amino acids from which they are built, it is well established that most proteins undergo a glass transition. A variety of techniques, including differential scanning calorimetry (DSC), dynamic mechanical thermal analysis (DMTA), and pulsed nuclear magnetic resonance, demonstrated that WG is a highly amorphous multipolymer system, which is water plasticizable but not water soluble (Slade, 1984; Hoseney et al., 1986; Slade et al., 1989; Kalichevsky et al., 1992). The Tg of dry WG (without any plasticizer) can be estimated by using the Matveev equation (Matveev, 1995). The calculation is based on the additive contributions of structural parameters (Matveev, 1995; Di Gioia et al., 1999):

Tg 1

20

Ni Vi Tgi 1 20

Ni Vi

i 1

(1)

i 1

where V is the van der Waals volume, Tgi the partial contribution to Tg of the ith amino acid, and Ni the number of amino acids of the ith type. Di Gioia et al. (1999) showed that the prediction was very close to the measured Tg for corn

78 gluten proteins. The same authors also found a good predictive power of the Matveev equation for WG proteins (unpublished results). The Tg of plasticized WG has been studied as a function of water, glycerol, and sorbitol contents using DMTA (Pouplin et al., 1999). For these plasticizers, the general behavior of Tg broadly followed the Couchmann-Karasz relation using a heat capacity ( Cp) value of 0.4 J/g K for WG. Compared on such a fractional weight basis, it was concluded that the plasticizing effect of glycerol and sorbitol on WG was less important than the plasticizing effect of water. However, a unique linear curve was obtained with the three plasticizers when the evolution of Tg was presented on the fractional molecular basis. This was related to the similar chemical structure of these three components that contain hydroxyl groups. Generally, by comparing plasticizer efficiency on a weight basis, similar plasticization was obtained with plasticizers of different molecular structure (hydroxyl or amino groups). Plasticizer efficiency (i.e., decrease of Tg) at equal molar content was generally proportional to the molecular weight of the plasticizer and inversely proportional to the percentage of hydrophilic groups on the plasticizer. The migration rate of the plasticizers in the polymer was related to their physicochemical characteristics. It was assumed that polar substances interacted with readily accessible polar amino acids, whereas amphiphilic substances interacted with nonpolar zones, which are, in most proteins, buried and accessible with difficulty (Di Gioia et al., 1998; Di Gioia and Guilbert, 1999). Micard and Guilbert (2000) studied the Tg of hydrophobized and native WG and its protein fractions using modulated DSC at moisture contents ranging from 0–20% (w/w on wet basis). The Tg values of unplasticized products were ~175 C regardless of the treatment (hydrophobization) or the fraction tested, except for the gliadin-rich fraction (162 C) (Figure 3.1). The experimental change in heat capacity ( Cp) at the glass transition ranged from 0.32–0.50 J/g K depending on the WG fractions. The Gordon-Taylor (0.5 J/g K) fit of Tg evolution as a function of water content showed that glutenin-rich fractions were more sensitive to water plasticization than gliadin-rich fractions (Figure 3.1). Prediction of the Tg from the water content of each protein fraction using the Gordon-Taylor model gave intermediary values of adjustable parameter k for gluten compared to those of the gliadin- and glutenin-rich fractions. This demonstrated that native WG is a molecular blend, instead of a physical blend, of gliadin and glutenin. The Kwei equation (Kwei, 1984) gave better fit to experimental data and demonstrated that the plasticization of WG and its fractions by water is influenced by secondary interactions. Because water is the most ubiquitous and uncontrollable plasticizer, the properties of WG-based materials are highly dependent on the usage conditions, such as RH and temperature. This temperature and water content dependence has been long interpreted in terms of water activity, disruptive wa-

Shaping Process of Wheat Gluten Materials

79

Figure 3.1 Glass transition temperature (Tg) of hydrophobized wheat gluten and of native wheat gluten and its fractions as a function of water mass fraction. (䊉): native gluten; (䉱): glutenin-rich 1; (䉭): glutenin-rich 2; (䊊): gliadin-rich; (*): hydrophobized gluten. (Adapted from Micard and Guilbert, 2000.)

ter-polymer hydrogen bonding in a polymer hydrogen bonded network. Gontard and Ring (1996) reported that WG-based films behaved as typical amorphous materials, and that the temperature and water mass fraction at which large changes in film mechanical and barrier properties were observed were well correlated with Tg.

Processing films, coatings, or other materials based on polymers of agricultural origin requires three main steps (Figure 3.2): (1) Break intermolecular bonds (non-covalent and covalent, if necessary) that stabilize polymers in their native forms by using chemical or physical rupturing agents. Polymer chains become mobile. (2) Arrange and orient mobile polymer chains in the desired shape. (3) Allow formation of new intermolecular bonds and interactions to stabilize the three-dimensional network. The shape obtained in step (2) is maintained by eliminating agents used in step (1) to rupture intermolecular bonds.

80 Based on these three steps, two technological processes are used to make protein-based materials (Figure 3.2): (1) The “solvent process” is based on dispersing and solubilizing proteins in various solvents and then casting, spraying, or dipping followed by drying. (2) The “thermoplastic process” is based on thermoplasticization of proteins. Using plasticizers and temperature allows for passing through the glass transition of proteins with formation of a rubbery mass that can be shaped and stabilized by cooling and/or by eliminating volatile plasticizers.

Figure 3.2 Two main approaches for shaping polymers of agricultural origin, i.e., casting or thermoplasticizing. (Adapted from Gontard, 1998.)


81

Film formation from WG solutions or dispersions has been extensively studied (Wall and Beckwith, 1969; Gennadios and Weller, 1990; Gontard et al., 1992; Gennadios et al., 1993a,b,c,d; Park and Chinnan, 1995). This process requires formation and drying of a thin layer of protein matrix. WG films have also been produced by collecting the surface “skin” formed during heating of WG solutions to temperatures near boiling (Watanabe and Okamoto, 1976). Several studies about using this process to prepare WG films were reviewed by Gennadios et al. (1994). WG proteins are insoluble in water and require a complex solvent system with basic or acidic conditions in the presence of alcohol and SS bond-reducing agents. Generally, changing the pH of the medium disrupts hydrogen and ionic interactions, while ethanol disrupts hydrophobic interactions (mainly occurring among gliadin molecules). Intermolecular (for glutenins) and intramolecular (for gliadins and glutenins) covalent SS bonds are cleaved and reduced to SH groups when dispersing WG in alkaline environments (Okamoto, 1978). Reducing agents, such as sodium sulfite, cysteine, or mercaptoethanol, should be used in acidic environments. A wide variety of acids can be used, including citric acid, acetic acid, phosphoric acid, propionic acid, and lactic acid. On the other hand, bases such as sodium hydroxide, potassium hydroxide, and, mainly for its volatility, ammonium hydroxide are used. During drying, all of these volatile disruptive agents are progressively eliminated. Solvent removal increases the concentration of WG proteins. Consequently, active sites for bond formation become free and close enough to each other to create new interactions. New hydrogen bonds, hydrophobic interactions, and SS bonds contribute to formation of a three-dimensional network. Solvent removal can be fractionated to obtain film-forming solutions with very low alcohol concentration. Colloidal dispersions of WG were obtained by dissolving WG in an aqueous alcohol medium in which acid was added (Cook and Shulman, 1998). Then, the alcohol concentration was lowered by adding water or by diafiltration to a level at which the proteins were no longer soluble. WG proteins precipitated in the form of microparticles, and the resulting colloidal dispersions were homogeneous and stable (physically and microbiologically). When cast onto a substrate and dried at ambient temperature, the particles fused to form continuous glossy coatings. Opta Food Ingredients, Inc. (Bedford, MA, USA) commercialized such film-forming colloidal dispersions. Recently, an alternative method for preparing WG films at neutral pH and without using ethanol as a co-solvent was developed by Lens et al. (2001). The addition of plasticizers is essential to avoid film brittleness. This brittleness results from extensive intermolecular associations (Wall and Beckwith, 1969). Plasticizers are small molecules of low volatility which, when added to polymeric materials, modify the three-dimensional structure, decrease attractive intermolecular forces, and increase free volume and chain mobility. As a

82 result of these changes in molecular organization, added plasticizers modify the functional properties of films by increasing film extensibility and flexibility and by decreasing cohesion and mechanical resistance. Sorbitol, mannitol, diglycerol, propylene glycol, triethylene glycol, maltitol, polyvinyl alcohol, and polyethylene glycol are used as plasticizers for WG films, but glycerol and ethanolamine are preferred. These molecules are small enough to insert between protein chains and to form hydrogen bonds through their hydroxyl groups. In terms of level of use, plasticizers are employed in amounts sufficient to lower the Tg to ambient temperature. Typical plasticizer levels are 15–40% of the weight of protein (preferably 20–30%). Polar plasticizers increase the hygroscopicity of WG-based materials. In addition, they diffuse easily if the material is in contact with a moisture-rich product. For these reasons, the research for nonpolar, low diffusivity plasticizers is essential for the development of WG-based plastics (and more generally for all hydrocolloid-based bioplastics). The use of amphipolar plasticizers, such as fatty acids and derivatives, should be recommended. The arrangement of protein chains; type, localization, and density of intermolecular and intramolecular bonds; and accessibility of residues play a fundamental role in the functional properties of the resulting three-dimensional networks. They are highly dependent on film formation conditions, including the nature, concentration, and order of the addition of solvents; drying conditions; and the nature and amount of plasticizers (Gontard, 1998). For example, Gontard et al. (1992) demonstrated that WG film properties were highly dependent on film-forming conditions. Strong interactions were observed between pH and ethanol concentration affecting film opacity, solubility, and water vapor permeability. WG concentration and pH of the film-forming solution were the most important factors determining film mechanical properties (Figure 3.3). Response surface analysis showed that high WG concentration induced formation of a resistant film with high puncture strength. The most resistant film (puncture strength of 4.5 N) was obtained at high WG concentration (12.5 g/100 mL) and pH above 5. When pH decreased, the effect of high concentration of WG on film puncture strength was less pronounced. The saddle surface shape indicated an optimum pH at about 4. Film puncture strength increased as WG concentration increased from 7.5–12.5 g/100 mL. Greater film strength could involve a higher number and/or a better localization of bonds among protein chains. During drying of the cast film-forming solutions, ethanol and acetic acid, which were responsible for protein dispersion, evaporated first, thus allowing bond formation among protein chains. During this stage, the proximity of protein chains induced by high WG concentrations could facilitate the formation of such cross-bonds. The effect of WG concentration on WG film structure was confirmed by studying electronic scanning microstructures of film sections (Figure 3.3). A very dense microstructure [Figure 3.3(b)] was observed for high


83

Figure 3.3 Effect of pH and wheat gluten concentration of film-forming solution on puncture strength and electronic scanning microstructure of wheat gluten films: (a) wheat gluten concentration 7.5 g/100 mL ; pH 4; ethanol concentration 45 mL/100 mL (left micrograph); (b) wheat gluten concentration 12.5 g/100 mL; pH 4; ethanol concentration 45 mL/100 mL (right micrograph).

WG concentration (12.5 g/100 mL) at pH 4 and 45% ethanol, compared to the microstructure obtained with the same pH and ethanol concentration, but at 7.5 g/100 mL WG concentration [Figure 3(a)]. Decreasing the pH from 6 to 4 increased film puncture strength, which could be due to the effect of acid on protein conformation. Sufficient protein unfolding, induced by pH levels below the isoelectric point, was required to enhance the probability of cross-bonding between protein chains during drying, thus improving film hardness. But at pH below 4, the film lost its cohesive structure, and a rapid, substantial decrease of hardness was observed (Gontard et al., 1992).

84 Finally, it appeared that film formation conditions should be selected based on the intended specific uses of the films. For example, if the film is intended for use as a superficial coating for handled products, the main properties to optimize would be mechanical properties and appearance. High WG concentration (12.5 g/100 mL), pH about 4, and ethanol concentration about 32.5% would be recommended to obtain a resistant, homogeneous, and transparent film without insoluble particles or excessively increased water vapor permeability. If the film is intended for use as a moisture barrier between two components of heterogeneous food products, a film solution pH between 5 and 6 and a low ethanol concentration (20%) would be required to obtain the lowest water vapor permeability (Gontard et al., 1992).

The knowledge of rheological and flow properties of “malleable” plasticized WG is of particular importance for the control of thermoplastic processes such as thermoforming or extrusion.

The general aspect of a frequency sweep curve of WG plasticized with glycerol is presented in Figure 3.4. Storage and loss moduli increased linearly with frequency in a log/log scale. The slope of the loss modulus curve (0.40) was superior to the one of the storage modulus (0.33). This behavior compared well with those observed for WG/water dough (Le Grys et al., 1981; Kokelaar et al., 1996) and for xanthan gum gels (Richardson and Ross-Murphy, 1987). Values of storage and loss moduli at 1 rad/s were 4.7 104 Pa and 2.2 104 Pa, respectively. The order of magnitude of the storage modulus was comparable to that of low moisture flour dough (2 104 to 1 105 Pa for 35–44% moisture content) reported by Navickis et al. (1982). The dependence of the storage modulus on frequency for flour/water dough has been reported as being somewhat lower, with slopes close to 0.29 (Bohlin and Carlson, 1980) and 0.27 (Le Grys et al., 1981). The complex viscosity * followed a power law behavior over the whole frequency window investigated, without a Newtonian plateau at low shear rate: *

K

n 1

(2)

with consistency K 49,800 Pa sn and n 0.31. The frequency dependence (n) of the dynamic moduli exhibited intermediate behavior between a permanent network and a temporary, entangled network. For such an intermediate behavior, Ross-Murphy (1995) proposed the term of partial network systems, in which polymeric chains are “physically”

85


Figure 3.4 Evolution of storage G (䊉) and loss G (䊊) modulus, and complex viscosity * (䊐) with the strain amplitude (pulsation at 1% strain amplitude) for glycerol-plasticized wheat gluten (80 C, 45% glycerol, t 1.5 tpeak). (Adapted from Redl et al., 1999a.)

cross-linked into networks, where the cross-links themselves are of small but finite energy and/or lifetime. Such rheological behavior would be consistent with the molecular structure of WG, where the amino acids of the polypeptide chains offer a wide range of physical cross-links, essentially hydrogen bonds and/or hydrophobic interactions. The influence of temperature and plasticizer content was evaluated with the principle of time-temperature and time-plasticizer superposition. Superposition was fairly good for the loss modulus G , but did not really apply for the storage modulus G (Redl et al., 1999a). This indicated that mostly local elastic interactions influenced the temperature-dependent dynamics. The WG-glycerol mixture did not behave as a thermorheologically simple material. Nevertheless, the effect of temperature and glycerol content can be estimated using time-temperature and time-plasticizer superposition, and a general expression for describing the viscous behavior of the product can be derived. The proposed law of viscosity is expressed as follows: *

K 0 exp n

Ea 1 R T

GC

SME

n 1

(3)

where T, GC, and SME are the temperature, glycerol content (expressed as g

86 glycerol/g WG), and specific mechanical energy, respectively. The used material constants are: K0 2.73Pa sn, n 0.31, Ea/R 13,300 K, 17.7, and 43 g/kJ. Similar rheological behavior was previously proposed for molten or plasticized starches (Vergnes and Villemaire, 1987; Willett et al., 1995; Della Valle et al., 1996, 1998), and the factors for temperature and glycerol dependency were comparable. An important difference for plasticized WG was that the energy term ( ) was positive, whereas, for thermoplastic starch, the energy term was negative. Thermoplastic starch is depolymerized by mechanical energy, whereas plasticized WG is rather structured, but the mechanisms involved are not yet completely understood. The general expression proposed for describing the viscous behavior of a WG/glycerol mixture might seem simplistic considering the observed complex rheological behavior, but it was sufficient for modelling the flow behavior in a twin-screw extruder (Redl et al., 1999b).

Extrusion of plasticized WG was studied in a corotating, intermeshing twin-screw extruder (Redl et al., 1999b). The used extruder configuration and an example of extracted screws after dead stop operation are shown in Figures 3.5 and 3.6, respectively. The extracted screws after dead stop operation show three different zones, as typically encountered in twin-screw extrusion: (1) powder solid conveying, (2) partially filled “melt” flow, and (3) completely filled “melt” flow. In this case, the solid conveying section was very short, especially for extrusion conditions with a barrel temperature of 80 C. Homogeneous and glossy dough was formed before the mixing elements. “Plastification” seemed to occur with low mechanical energy input, contrary to starch extrusion where “plasticization” or melting generally occurs before a restrictive element (Colonna et al., 1982; Barrès et al., 1990; Della Valle et al., 1997). Until the last screw elements before the die, the screws (including the mixing block) were only partially filled with little bullets of plasticized WG that were not adhering to the screws. The mixing block did not appear to be very

Figure 3.5 Schematic representation of the Brabender DS25 twin-screw extruder. Reprinted with permission from Redl et al., 1999b, Cereal Chemistry, 76:361–370.


87

Figure 3.6 Extracted screw after dead stop operation during extrusion of glycerol-plasticized wheat gluten (glycerol content 34.6% w/w; flow rate 1.9 kg/h; screw speed 100 rpm; barrel regulation temperature 80 C).

efficient in mixing due to its right-handed configuration. The last screw elements before the die were completely filled to create the pressure necessary for the flow through the die. For extrusion conditions with a barrel temperature of 60 C (not shown), the formed dough remained mat and sticky until the last elements of the screw. However, the physical or biochemical meaning of such a transition from a mat and sticky to a glossy and non-sticky dough is not very clear. At a given plasticizer content (34.6% w/w), extrusion was possible in a quite narrow window of operating conditions. Generally, the obtained extrudates were solid-like and exhibited a marked elastic behavior. At low temperature and low shear conditions, the extrudates had smooth surfaces and substantial die swell (up to 200%). By increasing thermomechanical severity (i.e., increasing barrel temperature, increasing screw speed, and decreasing flow rate), the observed die swell decreased, and important surface distortions appeared to complete rupture of the extrudates. In general, extrudate die swelling is primarily due to the elastic recovery of the deformation that an elastic polymer was exposed to in the die. The notable extrudate swell that we observed indicated the highly elastic nature of the WG polymer. For continuous extrudates, there is still sufficient molecular mobility to recover the experienced strain. In the case of extrudate rupture, we believe that the mobility of the polymeric chains is reduced due to cross-linking reactions, and elastic recovery without rupture is no longer possible. Consequently, the “protein melt” might no longer be able to support the strain experienced during its extrusion through the die. The dynamic rheological properties and molecular size distribution indicated increased density of cross-links. An example of storage moduli of three different extrudates is given in Figure 3.7. For the examples shown, samples were extruded at constant feed rate (Q 1.9 kg/h) and increasing screw speed (N 50, 100, or 200 rpm). In these conditions, the specific mechanical energy input and the maximum temperature varied from 640–3700 kJ/kg and from 97–134 C, respectively. We observed an important increase of the storage moduli over the whole frequency range investigated. The plateau modulus GN0 that can be associated with the network density was evaluated with a Cole Cole model as explained by Tschoegl (1989). It increased from 23–86 kPa, reflecting a nearly threefold increase in network density. Increasing cross-link density was also evidenced by an increase of the high molecular weight fraction from

88 5.4–34.8% of total proteins (fraction Fi, insoluble in a 2% SDS phosphate buffer, corresponding to proteins with molecular mass above 7 MDa). Extrudate breakup appears to be caused by increased cross-linking within the material. The driving mechanisms responsible for such cross-linking reactions remain unknown. The increase of insoluble fraction appeared to be closely related to the specific mechanical energy input and the maximum temperature reached. But, the relative importance of each parameter was not clear, as these two variables are related. From sampling along the screw, we observed that the most important changes in the molecular size distribution occurred in the converging section of the die. Assuming that the reaction is temperature controlled, the rate of protein insolubilization was estimated by dividing the amount of SDS insoluble fraction (Fi) with the residence time in the die. Figure 3.8 shows the corresponding Arrhenius plot. The calculated activation energy from this Arrhenius plot was 71 kJ/mol. This activation energy was lower than that observed for heat denaturation of proteins in static conditions (100–125 kJ/mol). Mechanical shear probably enhances the cross-linking effect of temperature. Plasticized WG is not a simple thermoplastic material. Extrusion of WG is possible in a quite narrow window of operating conditions. Extrudates are

Figure 3.7 Evolution of storage G modulus with pulsation for wheat gluten extrudates. Lines represent the fit of Cole Cole functions. Screw speed N 50 rpm (䊊), 100 rpm (ⵧ), 200 rpm (䉭); feed rate Q 1.9 kg/h; regulation temperature Tr 80 C. Reprinted with permission from Redl et al., 1999b, Cereal Chemistry, 76:361–370.


89

Figure 3.8 Arrhenius plot of the SDS-insoluble wheat gluten protein fraction (Fi) versus maximum temperature in the extruder die. The residence time (RT) in the die was 57 s (䊉) or 22 s (䊊). Activation energy was 71 kJ/mol. (Adapted from Redl et al., 1999b.)

solid-like and elastic. No viscous flow can be observed. Under thermomechanical severe conditions, cross-linking reactions occur. We hypothesize that these cross-linking reactions are temperature controlled, the activation energy as compared with static conditions is probably lowered by mechanical shear.

The Tg and change in heat capacity ( Cp at Tg) of WG samples from solvent (casting) and thermoplastic (thermal treatment, mixing, thermomolding, and mixing followed by thermomolding) processes were determined by modulated DSC as a function of water content (Micard et al., 2000b). Increase of Tg and drastic decrease of Cp were recorded when WG was dry processed (Table 3.1) as compared to being cast. The casting process gave WG film with calorimetric properties ( Cp) close to those of native WG. Thermomolding and use of chemical cross-linker during casting resulted in a drastic drop of SDS-soluble proteins. Contrarily, the mixing of WG, even by using high specific mechanical

90

energy, resulted in only slight polymerization of the proteins. Therefore, except for treatments where high temperature is applied to WG, changes in calorimetric parameters do not correlate well with the SDS-insoluble protein content. Tg, Cp, and protein extractability by SDS could account for different kinds of protein networks. To investigate the development of a network structure, rheological and biochemical changes were followed in a measuring mixer recording torque and temperature. A typical evolution of torque and temperature during mixing of WG and glycerol is shown in Figure 3.9. The torque evolution was characterized by a lag phase, followed by an exponential increase to a maximum and then a continuous decrease. The temperature evolution was characterized by a sigmoid-shaped curve, with the turning point at the maximum torque. Regardless of the mixing conditions (i.e., rotation speed, filling ratio, and glycerol content), maximum torque appeared to always occur in the temperature range of 55–60 C (Redl et al., 1997, 1999a). Samples collected during the lag phase, at time intervals below 0.5 tpeak, showed the consistency of a sand/water blend. During the torque increase phase, a cohesive, plastic, and very sticky dough was formed. Just beyond tpeak, the WG/glycerol dough changed to a fairly glossy, non-sticky, and very elastic material. The evolution of dough can very well be compared to the observation made with the extracted screws of the extrusion trials (Figure 3.6, evolution from powder > sand/water-like > plastic, sticky > elastic, non-sticky). Figure 3.9 summarizes the main rheological and biochemical changes observed during the development of structure in the WG/glycerol dough. Before reaching maximum torque (t < tpeak), the storage and loss moduli, G and G , remained quite stable. Also, the time dependency of the dynamic moduli (slopes in a double logarithmic plot n and n , Figure 3.7) did not change significantly. At maximum torque, the dynamic moduli remained more or less unaltered, whereas their time dependency decreased. After tpeak, moduli increased (especially storage modulus G ), and their time dependency decreased further. This


91

Figure 3.9 Evolution of the storage modulus, G (䊊) and loss modulus, G (䊉) at 1 rad/s; the slope of storage modulus, n (䊐), and loss modulus, n (䊏); the SDS-insoluble protein fraction, Fi (䉭), with reduced mixing time during mixing of wheat gluten and glycerol. (Adapted from Redl et al., 1999a.)

observed increase of the elastic modulus and decrease of time dependency indicated enhanced gel-like behavior. However, the established feature of a sol-gel transition (equal slopes of dynamic moduli; Winter and Mours, 1994) could not be observed. We may conclude that the development of gel-like behavior and elastic structure is not due to a sol-gel transition (i.e., the growth of clusters due to polymerization reactions), but rather to a rearrangement of polymeric proteins and the creation of new interactions. The high molecular weight fraction (Fi) of WG decreased up to 1.5 tpeak, and the intermediate molecular weight fractions (F2 and F3, not shown) increased. A progressive de-polymerization from Fi and F1 to F2 and F3 might be the cause. Then, at 2 tpeak, the evolution was inverted, and the insoluble fraction (Fi) nearly doubled at the expense of fractions F2 and F3. The evolution of the molecular size of proteins during processing has been reported to be an equilibrium reaction between depolymerization due to mechanical stress and polymerization depending on the time-temperature history of the sample (Morgan et al., 1983; Strecker et al., 1995; Weegels and Hamer, 1998). In our case, depolymerization was dominant in the first stage of mixing, as commonly observed in bread dough mixing (Weegels et al., 1996). Protein polymerization occurred only after 1.5 tpeak, once the dough temperature reached 70 C. From these observations, we propose a hypothetical mechanism for devel-

92 opment of the dough structure. In the first stage, plasticization occurs. We believe that mainly hydrophilic interactions promote the gel-like behavior and that the molecular size is reduced due to increasing mechanical shear. The reason for the drastic change at maximum torque and at 50–60°C may be an exposure of hydrophobic sites, reinforcing the gel-like behavior. Thereafter, the network structure might be stabilized with covalent cross-links, probably via SS bonds through free radical reaction mechanisms, leading to the observed increase of molecular size. This structure development mechanism is most probably a temperature-controlled phenomenon and only indirectly dependent on mechanical energy input, as mechanical energy is partly responsible for the temperature evolution through viscous heat dissipation.

Multilayer agromaterials [WG/paper, WG/other biopolymers such as poly(lactic acid)], as well as composite agromaterials combining WG with fibers of cotton, sisal, coconut, or straw, have given promising test results and could possibly be used for different applications. One of the main developments in multicomponent WG-based materials involves incorporation of lipid compounds into the film structure.

Films made from WG and lipids can combine the water vapor resistance of the lipids with the relatively good mechanical properties of the WG. Lipids can either be incorporated in WG film-forming solutions to obtain composite films (emulsion technique) or deposited as layers onto the surfaces of preformed WG films to obtain bilayer films (coating technique) (Figure 3.10). The effects of lipids incorporated into WG film-forming solutions can vary from destruction of the protein network to improvement of the mechanical and barrier properties of the cast films. These effects are complex and depend on the nature and structure of the lipids. Also, they depend on the chemical interactions between the lipids and the protein. Gontard (1991) incorporated 22 different lipids (at 20% w/w dry matter) into WG film-forming solutions. Among these lipids, 11 were selected because of their ability to form composite films with satisfactory appearance and mechanical properties. Water vapor permeability, opacity, solubility in water, and mechanical properties were determined for each type of WG-lipid film as a function of lipid concentration and compared with a control WG film without any lipid component (Gontard et al., 1994). For all tested lipids, film dispersion in water increased sharply above a lipid content of about 20% (w/w dry matter). Above this threshold, the presence of lipids probably


93

Figure 3.10 Water vapor permeability ( 10 12 mol/m s Pa) of a wheat gluten film, a wheat gluten/beeswax emulsion film, and a wheat gluten/beeswax bilayer film. Beeswax was added at 5.3 mg/cm2 of film surface. x is film thickness.

partially destabilized the protein structural matrix (reduction of intermolecular interactions among protein chains) or decreased water resistance when the lipid component carried polar, hydrophilic groups. Film opacity highly intensified as lipid concentration increased. This opacity increase was quite low with diacetyl tartaric ester of monoglyceride. Beeswax, a solid and highly hydrophobic lipid, was the most effective lipid in improving the moisture barrier ability of films. Water vapor permeability decreased by sixfold (Figure 3.10) from 5.08 to 0.83 10 12 mol/m s Pa as beeswax concentration increased from 0–36.8% (w/w dry matter). But, films with beeswax were opaque, brittle, and disintegrated easily in water. These drawbacks were reduced when sucroglycerides or diacetyl tartaric ester of monoglycerides were used as the lipid component in the composite films. Combining WG with a diacetyl tartaric ester of monoglycerides (at 20% w/w dry matter) produced films that, when compared to the control, glycerol-plasticized WG films, were as transparent, had higher mechanical resistance, and had about 50% lower water vapor permeability. However, improving moisture barrier properties of such composite films by increasing lipid concentration proved to be of limited value, because the hydrophilic protein matrix altered the resistance of the lipid components to water vapor transmission.

94 The coating technique for preparing multicomponent films from hydrocolloids and lipids was less effective than the emulsion technique in certain studies (i.e., Koelsch, 1994, with methylcellulose-based films) but was more effective in other studies (i.e., Schultz et al., 1949, with pectinate-based films). For WG-based films, Gontard et al. (1995) investigated the effect of a lipid layer on the water vapor permeability of films prepared from WG, diacetyl tartaric ester, and glycerol. The molten lipid application was superior to the solvent (ethanol) method for obtaining bilayer films with low water vapor permeability. The best results were obtained with a 0.09 mm-thick beeswax/WG bilayer film that had water vapor permeability of 0.023 10 12 mol/m s Pa, which was more than 200-fold lower than the value obtained with an uncoated control WG film. With the same amount of beeswax per unit surface area of WG film (5.3 mg/cm2), the coating technique was more effective than the emulsion technique (water vapor permeability of 0.023 10 12 mol/m s Pa for beeswax/WG bilayer film versus 0.83 10 12 mol/m s Pa for beeswax/WG emulsion film) (Figure 3.10).

Bilayer films composed of a WG film and a modified PE film were prepared by Perez-Perez (1997). The objective was to combine the gas selectivity of the WG films (Mujica-Paz and Gontard, 1997) with the excellent mechanical properties and moisture resistance of the PE films. WG films were prepared first using a casting procedure and then hot-pressed with the PE films at different temperatures. Because of chemical incompatibility, no adhesion between the WG layer and the synthetic layer could be obtained using simple PE. Three types of modified PE films with different reactive groups were tested for their ability to offer enhanced adhesion with the WG films: ethylene/acrylic ester/maleic anhydride terpolymer (EAMAT); ethylene/glycidyl methacrylate copolymer (EGMC); and PE grafted with maleic anhydride (PGMA). Various time and temperature combinations for hot-pressing were tested, and the adhesion of the WG films on the synthetic films was evaluated. Optimum conditions for hot-pressing were 110 C and 6 min. At these conditions, good adhesion was obtained only with PGMA. The adhesion strength (0.04 N/mm width) was equivalent to that of WG/WG or PGMA/PGMA bilayer films hot-pressed at the same conditions. Adhesion of WG films to EAMAT or EGMC films was not sufficient, probably due to the lower chemical compatibility of these synthetic materials with WG than with PGMA. Tensile strength and percent elongation at break values of PGMA/WG bilayer films were not affected by RH, unlike those of WG films. Also, the gas and water vapor permeability values of the PGMA/WG films were considerably reduced as compared to those of control WG films. However, the high selectivity of the WG films to oxygen and carbon dioxide was highly reduced when the WG films were combined with PGMA films.

Postproduction Treatments of Wheat Gluten Films

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WG films are relatively water resistant and water insoluble, but their high water vapor permeability, due to their inherent hydrophilic nature, remains the greatest limitation for their use (Gennadios et al., 1993b). Furthermore, their mechanical and water vapor barrier properties are strongly affected by water or other plasticizers (Gontard et al., 1993). The functional properties of WG-based films can be improved by appropriately selecting the casting conditions (as discussed earlier) or by subjecting the film-forming solutions to various treatments prior to casting. Such treatments can be enzymatic (most notably transglutaminase), chemical (e.g., cations and aldehydes), thermal, or radiative (e.g., ultraviolet- or -radiation) treatments (Gennadios and Weller, 1990; Gontard et al., 1992; Herald et al., 1995). Another approach is to apply various treatments on preformed films.

Heat treatments promote protein aggregation through SS bonding and hydrophobic interactions (Weegels et al., 1994; Gennadios et al., 1996; Ali et al., 1997; Cuq et al., 2000a; Micard et al., 2000a). This may be responsible for the observed changes in mechanical and barrier properties of heat-treated protein films. For example, aggregation resulted in almost total insolubilization (98%) of WG proteins in SDS when a short time-high temperature treatment was applied to WG film (Micard et al., 2000a).

Heat-curing of WG films prepared by casting or by using a heating press increased film tensile strength and decreased film elongation (Ali et al., 1997; Cuq et al., 2000a; Micard et al., 2000a). Tensile strength and elongation of cast films heated at 140 C reached up to 300% and 43%, respectively, of the value obtained with the films heated at 80 C (Table 3.2). WG films obtained by a thermal process (using a heating press; 10 min at 200 bar) showed even more pronounced tensile strength improvement when heated at 135 C versus 80 C. Specifically, the tensile strength of the films heated at 135 C was 784% of the value obtained for films heated at 80 C (Cuq et al., 2000a). The inverse relationship observed between tensile strength and elongation for cast (Table 3.2) or thermomolded heat-cured WG films could be attributed to differences in plasticizer content as suggested by Sánchez et al. (1998) for wheat gliadin films. However, the glycerol loss of heat-cured WG films (measured loss 11%) cannot explain such a change in their mechanical properties, confirming the predominant effect of thermal treatment (Micard et al., 2000a). Similar to heat-cured films from soy protein (Gennadios et al., 1996) or

96

whey protein (Miller et al., 1997), a significant interaction between temperature and exposure time was registered for heat-cured WG films (Table 3.2) (Ali et al., 1997; Cuq et al., 2000a; Micard et al., 2000a). When short time (15 min) treatments were applied, at temperatures of over 95 C or 110 C, mechanical properties of the films improved greatly (Table 3.2) (Cuq et al., 2000a; Micard et al., 2000a). Reducing RH, similar to increasing curing temperature, may accelerate an increase in film strength and loss of extensibility as demonstrated by Miller et al. (1997) for whey protein films.

Water vapor permeability of WG films at 25 C and 60% RH decreased as the exposure time increased at a given temperature (Ali et al., 1997). A decrease of 16 or 24% was reported for heat-cured WG films treated for 2 or 24 h at 95 C, respectively. Soy protein films showed even greater decreases in water vapor permeability (31% or 54% after heat curing for 2 or 24 h at 95 C, respectively) (Gennadios et al., 1996).

WG films appeared to be highly sensitive to color changes following heat-curing treatments as compared to soy protein films (Gennadios et al., 1996; Ali et al., 1997). The main effect of heating on the color of WG films was increased yellowness, as evidenced by increased +b color values (Ali et al., 1997). Similar to mechanical properties, color changes mainly occurred at temperatures above 95 C for short treatments (15 min) (Micard et al., 2000a). Also,

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short (10 min) heating up to 95°C of WG film-forming solutions prior to casting did not affect color coordinates of cast WG films (Roy et al., 1999).

Formaldehyde is the main chemical that has been used to treat preformed protein films. It leads to a drastic SDS insolubilization of proteins by reinforcing the protein network through covalent linkages (i.e., methylene bridges). Both soaking (Guéguen et al., 1998) and vapor exposure (Micard et al., 2000a) have been used to apply formaldehyde to protein films, the latter approach offering more reproducible results. Exposure of WG films to formaldehyde vapor increased film tensile strength and stiffness by 400% and decreased film elongation by 62% (Micard et al., 2000a). Pea protein (Guéguen et al., 1998), cottonseed flour (Marquié et al., 1995), and corn zein (Parris and Coffin, 1997) films (or film-forming solutions) treated with formaldehyde before or after casting had similar increases (165–470%) in tensile strength and decreases in elongation (30–60%, except for cottonseed flour films). Formaldehyde treatment of preformed films (by soaking or vapor exposure) from WG (Micard et al., 2000b) or pea protein (Guéguen et al., 1998) was more effective than treatment of film-forming solutions prior to casting. Immersion of cast WG films in calcium chloride solution or buffer solution at the isoelectric point of WG (pH 7.5), contrary to immersion in lactic acid solution, increased film tensile strength (by 46% and 8%, respectively) (Gennadios et al., 1993c).

No significant change in water vapor permeability of cast WG films treated with formaldehyde was observed (Micard et al., 2000a). Soaking in calcium chloride solution or in buffer solution at the isoelectric point of WG lowered the water vapor permeability of WG films by about 15% (Gennadios et al., 1993c). However, these two soaking treatments did not change the oxygen permeability of the WG films (Gennadios et al., 1993c).

Ultraviolet (UV) radiation (0.25 or 1 J/cm2) had no effect or only minor effect on mechanical properties of cast WG films (Micard et al., 2000a). Mechanical properties of UV-treated pea protein films were also unchanged (Guéguen

98 et al., 1998). In contrast, soy protein films subjected to a lower UV radiation dosage (0.0104 J/cm2) had higher tensile strength (by 65%) and lower elongation (by 31%) than control, untreated soy protein films (Gennadios et al., 1998). The observed changes in mechanical properties of UV-treated soy protein films were attributed to the notable amounts of tyrosine and phenylalanine in soy protein, because these two amino acid residues can participate in cross-linking reactions induced by UV radiation (Gennadios et al., 1998).

Gamma radiation (10 kGy) increased tensile strength (by 43%) and stiffness (by 79%) and decreased elongation (by 32%) of WG films (Micard et al., 2000a). This effect could be explained by the formation of dityrosine cross-linkages within polypeptide macromolecules as was shown by Brault et al. (1997) for irradiated caseinate films. Above 10 kGy, the improvement of mechanical properties of WG films was less pronounced (Micard et al., 2000a), which could be attributed to a decrease in the amount of the largest polymers of glutenin (Köksel et al., 1998). The water vapor permeability of -irradiated (10 or 20 kGy) WG films increased by 19%, probably because of glutenin degradation.

The storage duration and conditions (temperature and RH) of WG films greatly influenced thiol oxidation (Morel et al., 2000). This increase in thiol oxidation led to formation of protein polymers of large molecular size, which might have been responsible for the observed changes in mechanical properties of the films. Indeed, increasing aging of WG films at 20°C and 60% RH from 48–360 h increased film tensile strength by 75%, increased film stiffness (Young’s modulus) by 314%, and decreased film elongation by 36% (Micard et al., 2000a). In contrast, the water vapor permeability of films was unchanged.

Comparing the effects of different posttreatments of WG films is difficult, as relevant studies were mainly focused on the effects of a single posttreatment on the properties of a single type of film obtained with a certain casting method. However, an attempt to make such a comparison is presented in Table 3.3. A recent study comparing the effects of formaldehyde, temperature, UV radiation, and -radiation on mechanical properties of WG films cast from aqueous ethanol solvent at low pH clearly showed that the most effective treatments were the exposure to formaldehyde vapors and the high temperature-short time

Functional Properties of Wheat Gluten-Based Materials

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treatment (Micard et al., 2000a). UV radiation and, to a lesser extent, a milder heat treatment (80 C for 15 min) were less effective in modifying the mechanical properties of the films.

The mechanical properties of materials are largely associated with distribution and density of intermolecular and intramolecular interactions allowed by the primary and spatial structures. Cohesion of WG-based materials depends mainly on the type and density of intramolecular and intermolecular interactions, but also results from interactions with other constituents. Cooperative phenomena generally allow optimal thermodynamical stability of systems. The effects of interactions depend on interaction probabilities and interaction energies. The mechanical properties of polymer films depend on major interactions that may stabilize the networks. From a simplistic point of view, when covalent bonds stabilize the network or when density of bond energy is high, films are very resistant and relatively elastic. On the other hand, when most interprotein interactions are of low energy, films are easily distended. WG films show low

100 tensile strength but high elongation at break as compared to other films (Table 3.4). Mechanical properties of WG films depend greatly on processing conditions, plasticizers and other additives (e.g., lipids), chemical treatments (e.g., cross-linking), and external conditions (e.g., RH and temperature). Glycerol and water act as plasticizers of WG films, generally reducing mechanical resistance and increasing extensibility (Gontard et al., 1993). However, at very low water content and, thus, low RH (below 15 g/100 g dry matter and 60%, respectively), hydration had a positive effect on film mechanical resistance, probably due to formation of supplementary hydrogen bonds among protein chains (Figure 3.11). It should be noted that film mechanical properties can be highly modulated by adjusting the processing conditions based on the intended application. The effects of thermal treatments on mechanical properties of pressure-molded WG films were investigated by Cuq et al. (2000a). They prepared films from hand-mixed WG/glycerol doughs of sand/water-like consistency (corresponding to a reduced mixing time of 0.3 tpeak, Figure 3.9). The tensile properties of these materials are shown in Figure 3.12. Increasing the processing temperature from 80–135°C induced an increase in mechanical resistance of the WG network (tensile strength increased from 0.26–2.04 MPa) and a decrease in elongation (from 468–236%). This also was correlated with an important increase of the Young’s modulus (from 0.09–6.39 MPa). Variations in the mechanical properties of WG films as a function of temperature followed a sigmoidal shape that could be fitted with the model described by Peleg (1994). This model was used to determine a characteristic inflexion point at 116°C.

Figure 3.11 Influence of water activity and temperature on mechanical properties (puncture strength and deformation) of a wheat gluten film. (Adapted from Gontard et al., 1993.)

101

102

Figure 3.12 Tensile properties (determined at 60% RH and 20 C) of pressure-molded wheat gluten films. Experimental values (䊉) and corresponding Mooney-Rivlin modeling (—). [according to Equation (4) in text]. Films were molded for 10 min at various temperatures [numbers indicate the molding temperature ( C)]. Glycerol content was 40% (w/w).

Thus, increasing the processing temperature of the films from 95–125 C significantly increased film cohesion. This could be explained by an increased cross-linking of the polymeric network induced by the thermal treatment. To describe the behavior of cross-linked rubbers in unidirectional extension, an empirical formula, known as the Mooney-Rivlin equation, was used (Cuq et al., 2000b). The model is based on the assumptions of flexible long-chain molecules, weak intermolecular forces, and cross-linking of molecules at a few sites along their length to form a network. For the case of simple extension, the stress-strain relation can be written as follows: 2

1 2

C1

C2

(4)

where is tensile stress, is elongation ratio ( L/L0), and C1 and C2 are characteristic constants representing, respectively, the origin and the slope of the Mooney plot.


103

The Mooney model fitted quite well the experimental tensile data (Figure 3.12). Both Mooney parameters, C1 and C2, increased with film processing temperature (Table 3.5). The obtained Mooney constants compared quite well with those of natural or synthetic rubbers (C1 30–380 kPa, C2 30–480 kPa) (van Krevelen, 1990). Interpretation of the Mooney parameters in molecular terms is not straightforward. The parameter C1 is generally related to the network structure and the density of covalent cross-links in the material. The interpretation of parameter C2 is more ambiguous; an interpretation in terms of physical cross-links or the failure to achieve equilibrium are considered the most appropriate (Erman and Mark, 1997; Yeoh and Fleming, 1997; Furukawa et al., 1999). The nature of interactions in the WG-based films was studied by determining protein solubility in various solvents. Increasing the temperature from 80–135 C substantially reduced (to 0%) the SDS solubility of the proteins that could be recovered if -mercaptoethanol, an agent reducing SS bonds, was added to the solubilization buffer (Table 3.1) (Cuq et al., 2000a). This indicated that SS bonds were mainly involved in network formation. Furthermore, the protein solubility in SDS was strongly correlated to the Young’s modulus and, concurrently, to the molecular size of entanglements derived thereof (Figure 3.13). Thermoplastic processed WG films plasticized with glycerol behaved like rubber, and their tensile properties could be modeled well using the statistical theory of rubber-like elasticity. Increasing processing temperature induced cross-linking reactions that were reflected in an increase of elastic modulus and a decrease of solubility in a 2% SDS buffer. A very close relationship could be evidenced between the elastic modulus and the protein solubility in 2% SDS.

Mass transfer is important when selecting packaging or coating materials. In the absence of pores and physical faults, permeability (P) of the polymer film is

104 equal to the product of the diffusion coefficient (D), representing the mobility of permeant molecules in the polymer, and of the solubility coefficient (S), representing the permeant concentration in the film in equilibrium with the external pressure: P D S. Temperature and the size, shape, and polarity of the diffusing molecules affect the diffusion and solubility of permeants. Moreover, diffusion and solubility depend on film characteristics, including the type of forces influencing molecules of the film matrix, the extent of cross-linking, the degree of crystallinity, and the presence of plasticizers or other additives.

Substantial moisture barrier ability is required for protein-based films to reduce moisture exchange of the packaged or coated product with the external atmosphere. Water vapor permeability values of selected protein-based films are shown in Table 3.6. The water vapor permeability of WG films is equivalent to that of other protein- or polysaccharide-based films, but it is relatively high compared to that of synthetic polymer films. The water vapor transmission rate through WG films increased with increasing RH (Gontard et al., 1993) and tem-

Figure 3.13 Tensile modulus of thermomolded wheat gluten films versus protein extractability in a 2% SDS phosphate buffer, Fsol. Error bars represent the standard deviation of five replicates. (Adapted from Cuq et al., 2000b.)


105

perature (Figure 3.14). At 30 C, water vapor permeability was tenfold lower at 50% RH than at 93% RH. Water vapor permeability of WG films was also highly dependent on the nature and amount of the added plasticizer. A linear relationship was observed between glycerol concentration and water vapor transmission rate of WG films (Gontard et al., 1993). Due to their relatively low water vapor barrier ability, protein-based films can be used as protective barrier layers to limit moisture exchange for only short-term applications or in low-moisture foods. Coating WG films with lipids, such as beeswax, greatly reduced film water vapor permeability, which became twofold lower than that of LDPE (Gontard et al., 1994, 1995). Composite WG/lipid films can thus be used as protective barrier layers to limit moisture exchange between the external atmosphere and the packaged food or within heterogeneous foods (e.g., pizza, pies, and cakes with moist centers) to limit moisture transfer among food components.

Similar to water vapor permeability, oxygen, carbon dioxide, and ethylene

106

Figure 3.14 Effect of water activity and temperature on the water vapor transmission rate of a wheat gluten film. (Adapted from Gontard et al., 1993.)

permeabilities of WG films are highly dependent on RH and temperature. At low RH and temperature, the gas permeabilities of WG films are very low. At 0% RH, a WG film had very low O2 and CO2 permeability values (1.24 and 7.4 amol/m·s·Pa, respectively) compared to other biopolymer or synthetic polymer films (Table 3.7). For example, at 0% RH, the O2 permeability of the WG film was 800-fold lower than that of LDPE and 9.6-fold lower than that of polyamide-6, a well-known potent oxygen barrier polymer at 0% RH (Billing, 1989). Also, the WG film was 2.6- and 570-fold less permeable to CO2 than polyamide-6 and LDPE, respectively. Interesting applications of WG films as oxygen and carbon dioxide barriers at dry conditions could thus be considered (e.g., for low water activity products or sandwiched between two layers of a highly water-resistant material). However, the gas permeability values of WG films increased substantially with increasing RH and temperature (Figure 3.15) (Mujica-Paz and Gontard, 1997). For example, when RH increased from 0%–100% at 24 C, CO2 permeability increased from 88–55,580 amol/m s Pa (Figure 3.15), ethylene permeability from 1–3098 amol/m s Pa, and oxygen permeability from 77–1970 amol/m s Pa. For these three gases, permeability values remained low at RH below 50%, regardless of temperature, and increased steeply at RH over 50%. Such exponential increase of permeability as a function of RH is typical for hydrophilic polymers, such as cellophane, ethylene-vinyl alcohol, or methylcellulose/palmitic acid films. The increase of gas permeability as a function of RH may be related to the hydrophilicity of WG due to its high content of amide groups (Gontard et al.,


107

1992). Interactions between water molecules and amide groups contribute to a high water content in the film at high RH and to a modification of the protein network structure. The disruption of hydrogen bonds may create additional sites for dissolution of oxygen and increase mobility of the gas molecules within the polymer bulk phase (Gennadios and Weller, 1990; Gontard et al., 1993, 1996). But, the effect of RH on gas permeability varies with temperature. From theoretical considerations, gas permeability of films should vary with temperature following an Arrhenius-type relationship (Paine and Paine, 1983; Mannheim and Passy, 1985): ln P

ln Po (Ea / R)(1/ T )

(5)

where Ea is the activation energy of permeability (J/mol), R is the universal gas constant (8.314 J/mol K), T is the absolute temperature (K), and Po is the Arrhenius constant (amol/m s Pa). For oxygen permeability of WG films, Gennadios et al. (1993d) reported a good fit of the Arrhenius model at the temperature range of 5–35°C at 0% RH. Calculated Ea values for oxygen permeation through WG films were quite similar to those of synthetic films. For carbon dioxide and ethylene (Gontard, 1998), Arrhenius plots showed that a change occurred in the slopes, which were lower at low temperatures (Figure 3.16). Ea values below the inflexion point were very low. Ea values above the inflexion point were higher indicating that the permeability became more sensitive to temperature changes and comparable to that of synthetic films (Gontard, 1998).

108 Figure 3.15 Effect of temperature and relative humidity on oxygen permeability, carbon dioxide permeability, and selectivity (CO2/O2 permeability ratio) of a wheat gluten film. (Adapted from Mujica-Paz and Gontard, 1997.)


109

The temperature at which the inflexion point occurred decreased with increasing RH. This observation was in agreement with the evolution of glass transition with RH (Gontard et al., 1996). Tg is one of the most important properties of polymers directly related to structure modifications that affect the diffusivity and, subsequently, the permeability of gases. Gontard and Ring (1996) studied the depression of Tg of WG films by water using DMTA and DSC. The WG films behaved as typical amorphous polymers. The temperature and water mass fraction at which large changes of WG film mechanical and barrier properties were observed (Gontard et al., 1996) correlated well with Tg (Figure 3.17). The temperature at which the inflection point occurred at a defined RH could also be correlated with the evolution of glass transition with RH (Figure 3.17). Increased RH promotes both gas diffusivity (due to the increased mobility of hydrophilic macromolecular chains) and gas solubility (due to swelling of the matrix by absorbed water), leading to a sharp increase in gas permeability. With CO2, the sharp increase of permeability is more pronounced than with O2. The selectivity coefficient between CO2 and O2 (defined as the ratio of the respective permeabilities of the two gases) was very sensitive to moisture variations of WG films (Figure 3.15). The selectivity coefficient of WG films varied from 4.0 at water activity of 0.30 to 25 at water activity of 0.95, whereas the selectivity coefficient for synthetic films remains generally relatively constant from 4 to 6. This could be explained by the differences in water solubility of these gases (i.e., CO2 is highly water soluble) but is mainly explained by specific interactions between CO2 and the water-plasticized protein matrix, as shown in Figure 3.18. The exceptional selectivity of WG films can be qualified as sorption selec-

Figure 3.16 Carbon dioxide and ethylene permeability of a wheat gluten film as a function of relative humidity (RH) and temperature (Arrhenius plots and activation energy values (Ea) below and above the glass transition temperature (Tg). (Adapted from Gontard, 1998.)

110

Figure 3.17 Glass transition temperature of a wheat gluten film determined by dynamic mechanical thermal analysis (E inflexion point and tan ) and differential scanning calorimetry (DSC) as a function of water content. The crosses indicate water mass fraction and temperature at which changes in mechanical and barrier properties of the wheat gluten film occurred. (Adapted from Gontard et al., 1996.)

tivity (while the main conventional selective films have a selectivity of diffusion). From this basis, interesting applications of WG films could be developed for modified atmosphere packaging of fruits and vegetables.

Aroma components are present in foods at low levels, but their loss greatly affects sensory properties by reducing flavor intensity or by changing the aroma of the food. Packaging reduces the transfer of small molecules, such as aroma compounds, between foodstuffs and the environment. A range of synthetic polymers and laminates is available for food packaging, but one of the most important limitations of these packaging materials is that polymers allow the mass transport of low molecular weight substances. Polymers with lipophilic characteristics may be unsuitable for packaging products containing nonpolar substances, such as fats, oils, or aroma compounds, as these may be sorbed into the polymer or permeate through it (Munoz et al., 1999). Protein films, which are hydrophilic in nature, may be used as stand-alone film wraps or as supplements to synthetic packaging materials. The aroma permeability of polymer films is often related to their O2 permeability. However, aroma barrier properties of protein films and other types of biopolymer films have not been studied enough to elucidate their aroma transport behavior (Sadler and Braddock, 1991; Miller et al., 1998; Gallo et al., 1999).

Figure 3.18 Carbon dioxide solubility in a wheat gluten film and potential interactions between carbon dioxide (in a combined or dissolved form) and the wheat gluten network. (Adapted from Gontard, 1998.)

111

112 Debeaufort and Voilley (1994) reported that the permeability of WG films 10 12 g/m s Pa) was 20 times lower than that of to 1-octen-3-ol (4.6 methylcellulose films (122 10 12 g/m s Pa) and 10 times lower than that of LDPE films (51 10 12 g/ms Pa). It seems that in hydrophobic films, such as LDPE, the permeation of 1-octen-3-ol was influenced more by the amount of sorbed permeant than in hydrophilic films such as WG. Similarly, permeability of d-limonene in whey protein films was not influenced by permeant concentrations across a large range of concentration (Miller et al., 1998). Other studies using methyl ketones (blue cheese odor) or esters allow the conclusion that WG films were better barriers, to aroma compounds than methylcellulose-based films or several plastic films. WG films show great promise as aroma barriers, but a better knowledge of the aroma transport properties of WG films is necessary to explain the transfer process. The barrier efficiency could depend on the WG characteristics (e.g., type of polymerization, three-dimensional structure, and Tg) and on the properties of the volatile compounds (e.g., shape, size, nature, and polarity). As proposed by Debeaufort and Voilley (1994), the use of WG films for wrapping foodstuffs such as cheese or mushrooms to reduce off-flavors during refrigerated storage seems very interesting.

The ability of WG films to control solute transport may be very important, especially for applications involving food contact. For such applications, the migration of plasticizers or additives from the film or coating into the food structure is generally undesirable unless a controlled release of a chosen additive is wanted (“active packaging”). Improved microbial stability of foods is possible by using active layers as surface retention agents to limit the diffusion of food additives (particularly antioxidant and antimicrobial agents) into the food interior (Torres and Karel, 1985; Torres et al., 1985; Cuq et al., 1995a; Guilbert et al., 1996; Redl et al., 1996). Locally maintaining a high effective concentration of a preservative may allow considerable reduction of its total amount for achieving the same effect. It is then important to be able to predict and control surface preservative migration. The mathematical theory of diffusion in isotropic substances is based on the hypothesis that the rate of transfer of diffusing substances through the unit area of a section is proportional to the concentration gradient normal to the section, i.e., Fick’s first law. One method to quantify the diffusion is to study the desorption of a solute, such as sorbic acid, from a thin layer of solute into a stirred liquid medium. In the case of Fickian diffusion, the initial desorption is linear versus the square root of time, and a diffusion coefficient can be derived (Crank, 1975). As for the rheological properties, morphological or physicochemical changes of the matrix are reflected in the migration properties of the diffusing molecules. Studying the influence of variables such as temperature and composition of the


113

matrix can therefore be of great importance for a structural insight. The diffusion coefficient of sorbic acid in a WG-based film was 7.5 10 12 m2/s (Table 3.8). The addition of lipids reduced sorbic acid diffusivity by 20 to 50%. But the diffusivity was still higher than the diffusivity through the corresponding pure lipid films (2.4 10 16 m2/s and 2.7 10 13 m2/s for beeswax and acetylated monoglycerides, respectively) (Table 3.8). The effect of temperature on the diffusion could be described by an Arrhenius type relationship with Ea values ranging from 30.0 to 39.8 kJ/mol. An explicit numerical method for modeling the migration of sorbic acid through a WG-based film in contact with a food surface was proposed allowing researchers to trace the evolution of the concentration profiles of sorbic acid within the surface layer and the food (Figure 3.19). The model was validated with experimental data obtained with WG and beeswax films placed on agar gels as model foods. Modeling showed that in the case of a WG film, the surface concentration dropped below 10% of the initial value after 1 h, whereas, in the case of a beeswax film, the surface concentration remained above 75% of the initial value after one week. Simulation led to the conclusion that, to get a significant surface retention, sorbic acid diffusivity in the edible surface layer has to be lower than 10 15 m2/s. Microbiological analysis confirmed the efficiency of preservative retention within surface coatings. The antimicrobial effect of various WG/sorbic acid-based films was evaluated against Penicillium notatum. The study was conducted at two temperatures on an acid model food. Control WG-based films had no fungicidal effect by themselves. However, the addition of lipids (e.g., beeswax or diacetyl tartaric ester of monoglycerides) to the WG films delayed the growth of fungi. Sorbic acid retention by all films was highly dependent on temperature. At 4 C, WG-based films containing sorbic acid delayed growth of P. notatum for four days, whereas no effect was observed at 30 C. The WG/lipid-based films showed good sorbic acid retention and a strong fungicidal effect, at both 4 and 30 C, delaying growth of P. notatum by more than 21 days.

114

Figure 3.19 Theoretical evolution of concentration profile of sorbic acid in wheat gluten film placed on a model food as a function of time. Calculated values are obtained using: DFilm 7.5 10 12 m2/s and Dmodel food 9 10 10 m2/s (C sorbic acid concentration at time t, C0 initial concentration of sorbic acid in film).

Little information exists on the biodegradation of WG-based films and on the absence of environmental toxicity from degraded products of WG-based materials generated during natural degradation or composting. The structure and conformation of protein networks can change significantly so that the resistance to enzymatic hydrolyses and to chemical attacks can be affected (Swaisgood and Catignani, 1991). However, Garcia-Rodenas et al. (1994) showed that the sensitivity of films made from WG or casein to in vitro proteolyses was not significantly different from that of the native proteins.

Application of WG-based materials can be envisioned for the coating of seeds, pills, and foodstuffs, and for making cosmetic masks and polishes or drug capsules. However, the allergenic character of WG-based products for people suffering from celiac disease could limit such applications to cereal-based products in which the presence of WG or similar cereal proteins is

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known and expected. For example, recent commercialization of cheese that was coated with WG-based film as a wax replacement had to be ceased in the Netherlands because of that. Therefore, applications are generally geared toward making flexible films for agriculture (e.g., mulching and films for banana culture), packaging films, and coatings for paperboards or objects (e.g., biodegradable materials). Such WG-based biodegradable materials can be reinforced with fibers to prepare composite isolating materials for the construction or automobile industries. The very specific moisture, gas, and solute barrier properties of WG-based films could be interesting for applications involving controlled release of beneficial agents (e.g., active packaging and drug delivery concepts) or for modified atmosphere packaging techniques (Barron et al., 2001). Various current and envisioned applications of protein films in general are extensively discussed elsewhere in this book.

Homogeneous, transparent, mechanically strong, and water-resistant WG-based materials have been obtained either by casting or by thermoplastic processing. Casting is generally suitable for applying coatings on food and nonfood items. Formation of WG-based materials using thermal techniques that are generally implemented with synthetic thermoplastic polymers (e.g., extrusion, injection, and molding) is more cost-effective. There is considerable potential for modulating the properties of WG-based films through appropriate selection and formulation of raw materials, careful use of fractionation techniques and rheological modifying additives, and adjustment of film-forming processes (e.g., characteristics and concentration of solvents, dispersing and plasticizing agents, temperature, protein concentration, and drying conditions). The moisture, gas, and solute barrier properties of WG-based films could be specifically interesting for applications in the fields of active coatings, active packaging, drug delivery systems, or modified atmosphere packaging.

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Slade, L. 1984. “Starch Properties in Processed Foods: Staling of Starch-Based Products.” Presented at the 69th Annual Meeting of the American Association of Cereal Chemists, Washington, DC, paper no. 112. Slade, L., Levine, H. and Finlay, J.W. 1989. “Protein-Water Interactions: Water as a Plasticizer of Gluten and Other Protein Polymers” in Protein Quality and the Effects of Processing, eds., R.D. Phillips and J.W. Finlay, New York: Dekker, pp. 9–23. Southan, M. and MacRitchie, F. 1999. “Molecular Weight Distribution of Wheat Proteins,” Cereal Chem., 76:827–836. Stevenson, S.G., Ueno, T. and Preston, K.R. 1999. “Automated Frit Inlet/Frit Outlet Flow Field-Flow Fractionation for Protein Characterization with Emphasis on Polymeric Wheat Proteins,” Anal. Chem., 71:8–14. Strecker, T.D., Cavalieri, R.P., Zollars, R.L. and Pomeranz, Y. 1995. “Polymerization and Mechanical Degradation Kinetics of Gluten and Glutenin at Extruder Melt-Section Temperatures and Shear Rates,” J. Food Sci., 60:532–537, 557. Swaisgood, H.E. and Catignani, G.L. 1991. “Protein Digestibility: In Vitro Methods of Assessment,” Adv. Food Nutr. Res., 35:185–236. Tatham, A.S., Shewry, P.R. and Belton, P.S. 1990. “Structural Studies of Cereal Prolamines, Including Wheat Gluten” in Advances in Cereal Science and Technology, ed., Y. Pomeranz, St. Paul, MN: American Association of Cereal Chemists, pp. 1–52. Torres, J.A. and Karel, M. 1985. “Microbial Stabilization of Intermediate Moisture Food Surfaces. III. Effects of Surface Preservative Concentration and Surface pH Control on Microbial Stability of an Intermediate Moisture Cheese Analog,” J. Food Process. Preserv., 9:107–119. Torres, J.A., Motoki, M. and Karel, M. 1985. “Microbial Stabilization of Intermediate Moisture Food Surfaces. I. Control of Surface Preservative Concentration,” J. Food Process. Preserv., 9:75–92. Tschoegl, N.W. 1989. The Phenomenological Theory of Linear Viscoelastic Behavior: An Introduction. Berlin: Springer-Verlag. Tsiami, A.A., Bot, A., Agterof, W.G.M. and Groot, R.D. 1997. “Rheology of Mixtures of Glutenin Subfractions,” J. Cereal Sci., 26:279–287. Van Krevelen, D.W. 1990. Properties of Polymers, 3rd edition. Amsterdam: Elsevier Science. Vergnes, B. and Villemaire, J.P. 1987. “Rheological Behaviour of Low Moisture Molten Maize Starch,” Rheol. Acta, 26:570–576. Wall, J.S. and Beckwith, A.C. 1969. “Relationship between Structure & Rheological Properties of Gluten Proteins,” Cereal Sci. Today, 14:16–18, 20–21. Watanabe, K. and Okamoto, S. 1976. “Formation of Yuba-Like Films and Their Physical Properties,” New Food Industry, 18(4):65–77. Weegels, P.L. and Hamer, R.J. 1998. “Temperature-Induced Changes of Wheat Products” in Interactions: The Keys to Cereal Quality, eds., R.J. Hamer and R.C. Hoseney, St. Paul, MN: American Association of Cereal Chemists, pp. 95–130. Weegels, P.L., Verhoek, J.A., De Groot, A.M.G. and Hamer, R.J. 1994. “Effects on Gluten of Heating at Different Moisture Contents. I. Changes in Functional Properties,” J. Cereal Sci., 19:31–38. Weegels, P.L., van de Pijpekamp, A.M., Graveland, A., Hamer, R.J. and Schofield, J.D. 1996. “Depolymerization and Re-polymerization of Wheat Glutenin during Dough Processing. I. Relationship between Glutenin Macropolymer Content and Quality Parameters,” J. Cereal Sci., 23:103–111. Wellner, N., Belton, P.S. and Tatham, A.S. 1996. “Fourier Transform IR Spectroscopic Study of Hydration-Induced Structure Changes in the Solid State of -Gliadins,” Biochem. J., 319:741–747. Werner, W.E., Adalsteins, A.E. and Kasarda, D.D. 1992. “Composition of High-Molecular-Weight Glutenin Subunit Dimers Formed by Partial Reduction of Residue Glutenin,” Cereal Chem., 69:535–541.

122 Willett, J.L., Jasberg, B.K. and Swanson, C.L. 1995. “Rheology of Thermoplastic Starch: Effects of Temperature, Moisture Content, and Additives on Melt Viscosity,” Polym. Engr. Sci., 35:202–210. Winter, H.H. and Mours, M. 1994. “Rheology of Polymers near Liquid-Solid Transitions,” Adv. Polym. Sci., 134:165–234. Wrigley, C.W. and Bietz, J.A. 1988. “Protein and Amino Acids” in Wheat Chemistry and Technology, Vol. 1, 3rd edition, ed., Y. Pomeranz, St. Paul, MN: American Association of Cereal Chemists, pp. 159–275. Wrigley, C.W., du Cros, D.L., Archer, M.J., Downie, P.G. and Roxburgh, C.M. 1980. “The Sulphur Content of Wheat Endosperm Proteins and Its Relevance to Grain Quality,” Austr. J. Plant Physiol., 7:755–766. Yeoh, O.H. and Fleming, P.D. 1997. “A New Attempt to Reconcile the Statistical and Phenomenological Theories of Rubber Elasticity,” J. Polym. Sci., 35:1919–1931.

Formation and Properties of Soy Protein Films and Coatings

OYBEAN is one of the most important legume crops, representing 52% of the total oilseed production worldwide (USDA, 1999). The United States accounted for 46% of the world’s production of soybeans and for 60% of the world trade market, making it the world leader in soybean production (USDA, 1999). Soybean yields high protein soy meal, concentrate, and isolate, which are inexpensive protein sources. The major use of soybean in the food industry is as a source of oil, while soy protein concentrate and isolate are readily available as co-products of the oil processing industry (Pszczola, 1998). The use of soy protein as a food source is increasing due to its functional, nutritional, and nutraceutical properties (Liu, 2000). Soy protein used in the food industry is classified as soy flour, concentrate, or isolate based on the protein content. Soy flour contains 50–59% protein and is obtained by grinding defatted soy flakes. Soy protein concentrate contains 65–72% protein and is obtained by aqueous liquid extraction or acid leaching process. Soy protein isolate contains more than 90% protein and is obtained by aqueous or mild alkali extraction followed by isoelectric precipitation (Soy Protein Council, 1987). Soy protein is an abundant and relatively cheap ingredient source for various food applications, with soy protein meal costing about 15 cents per pound (Soya & Oilseed Bluebook, 1999). The functional properties that make soy protein useful in foods include cohesiveness, adhesiveness, emulsification, dough and fi-

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124 ber formation, whippability, solubility, and foaming (Wolf and Cowan, 1975). The introduction of “second generation soy foods” (e.g., soy ice cream, soy yogurt, vegetable burgers, soy sausages, and soy pancakes) has increased the uses of soy protein (Liu, 1997). Soy protein also is used in infant formulas (soy protein formulas account for a quarter of infant formula sales) and in baked, meat, and dairy products (Witherly, 1990). The consumption of industrial soy protein in the United States is broken down to 60% as food ingredient, 20% as adhesive, 10% as plastic, and 10% for other applications (USB, 2000). However, despite its high protein quality, low cost, and plentiful supply, currently only a small percentage of soy protein isolate (SPI) is used as food ingredient, and is thus an underutilized product. The use of soy protein as a film-forming agent can add value to soybeans by creating new channels for marketing soy proteins (Rayner et al., 2000). Soy protein is a viable and renewable resource for producing edible and environmentally friendly biodegradable films. This chapter discusses the formation and properties of soy protein-based films.

Soy protein films are typically prepared by drying thin layers of cast film-forming solutions. SPI is most often used to prepare such film-forming solutions, although 7S and 11S soy protein fractions have also been used (Kunte et al., 1997; Subirade et al., 1998). The drying temperature and relative humidity (RH), which determine the drying rate, of cast solutions can affect the film structure and properties. In general, rapid drying of cast polymer solutions limits the development of intermolecular associations within the film structure as solvent removal restricts the mobility of the molecular chains (Banker, 1966). Alcantara et al. (1998) studied the effect of drying rate on the properties of cast films from whey protein isolate. Rapid drying at 95 C and 30% RH yielded films that were thinner, stiffer, stronger, and less extendable than films dried more slowly at 21 C and 50% RH. Also, films dried at 95 C and 30% RH had lower water vapor permeability than films dried at 21 C and 50% RH. Similarly, cast peanut protein films dried at 90 C had lower water vapor permeability and greater tensile strength than films dried at 70 or 80 C (Jangchud and Chinnan, 1999). Films need to have good texture and flexibility to prevent cracking during handling and storage (Torres, 1994). Therefore, plasticizers are added to soy protein film-forming solutions to increase the flexibility and “soften” the structure of formed films. Generally, plasticizers act by entering between polymeric molecular chains, physicochemically associating with the polymer, reducing cohesion within the film network, and effectively extending and softening the film structure (Banker, 1966; Arnold, 1968; Guilbert, 1986). Examples of food-grade plasticizers incorporated into edible films are polyols (e.g., glyc-

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erol, sorbitol, and polyethylene glycol), sugars (e.g., glucose and honey), and lipids (e.g., monoglycerides, phospholipids, and surfactants). Glycerol and sorbitol are the most commonly used plasticizers for soy protein-based films (Gennadios et al., 1994). Extruding soy protein formulations into films is also possible. Such an extrusion process, using soy protein, plasticizer (e.g., polyhydric alcohols), and water as the main formulation ingredients, was described by Naga et al. (1996). They used pressure of 5–50 kg/cm2 and temperature of 110–180 C in the extruder to knead and melt the raw material. Films have also been extruded from blends of soy protein isolate, poly(ethylene oxide), and low-density polyethylene (Ghorpade and Hanna, 1996). Yuba, a Japanese word for soymilk film, is the dried film formed on the surface of heated soymilk (Wu and Bates, 1972a, b; Farnum et al., 1976; Chuah et al., 1983; Sian and Ishak, 1990a, b; Gennadios and Weller, 1991). Protein and lipids are the main components in yuba with carbohydrates and ash being present in smaller amounts. A typical yuba composition (w/w, dry basis) of 55% protein, 28% lipids, 12% carbohydrates, and 2% ash was reported by Wu and Bates (1972a). Temperatures close to the boiling point (e.g., 95 C) were considered optimum for yuba formation (Wu and Bates, 1972b). A soymilk pH value around nine maximized film yield and protein incorporation into films (Wu and Bates, 1972b; Sian and Ishak, 1990a, b). Soy protein polymerization was attributed to disulfide bonds and, to a lesser degree, hydrophobic interactions and hydrogen bonds (Farnum et al., 1976; Sian and Ishak, 1990a). Yuba-like films have also been formed on the surface of heated solutions of SPI (Wu and Bates, 1972a) and 11S or 7S soy protein fractions (Shirai et al., 1974; Okamoto, 1978). Dried yuba is quite brittle, has a relatively long shelf life, and is used as a wrapper for other foods, cooked with other foods, or used in soups (Liu, 1997). Recently, film preparation by spinning SPI in a coagulating buffer was described (Rampon et al., 1999). Specifically, they prepared a film-forming dope by mixing SPI, water, and 1.3 M sodium hydroxide solution in a cutter to a homogeneous, viscous, alkaline (pH 12.5) mass. This dope was poured into a cylinder and forced through a filter using compressed air. The dope was then extruded through a rectangular nozzle immersed into a coagulating bath of acetate buffer. The buffer had a pH of 4.7, which corresponded to the isoelectric region of SPI. Upon contact with the buffer, the proteins in the dope coagulated into a wet film that was collected on a polyethylene roller and dried overnight at 15 C (Rampon et al., 1999). Fourier transform IR spectroscopy showed that no secondary structure differences existed between the protein film and the isolate. Protein aggregation within the isolate increased by immersing and dispersing the SPI into the isoelectrical pH buffer. Fluorescence spectroscopy did not reveal any tertiary structural differences between the isolate and the film. Stretching the film in two orthogonal directions did not change signifi-

126 cantly (P < 0.05) the tensile properties, thus indicating that no network orientation existed (Rampon et al., 1999). Therefore, film structure could have resulted from a random aggregation of proteins. Finally, another technique for preparing soy protein films was described by Ogale et al. (2000) and Cunningham et al. (2000). They thermally compacted mixtures of SPI and glycerol at a temperature of 150–160 C and pressure of 10 MPa. Processing temperatures above 180 C could not be used due to substantial thermal degradation of the protein films as evidenced by thermogravimetric analysis (Ogale et al., 2000).

In general, soy protein films have poor moisture resistance and water vapor barrier ability due to the inherent hydrophilicity of the protein and the substantial amounts of hydrophilic plasticizers used to impart film flexibility (Rhim et al., 2000). Compared to low-density polyethylene films, cast SPI films had greater water vapor permeability (WVP) values by roughly four orders of magnitude (Gennadios et al., 1993b). In contrast, soy protein films are potent oxygen barriers, at least at low RH environments (Gennadios et al., 1993a; Ghorpade et al., 1995b). In general, good oxygen barrier ability is a characteristic of protein films (Krochta and De Mulder-Johnston, 1997; Miller and Krochta, 1997). Mechanical properties of protein films, such as tensile strength (TS) and percentage elongation at break (E), offer an indication of expected film integrity under conditions of stress that would occur during processing, handling, and storage. Mechanical property testing evaluates film performance and the fundamental structure of the material (Stuchell and Krochta, 1994). A three-dimensional matrix, constructed by interaction of protein molecules, is presumably the supporting structure that dictates the mechanical properties of the protein film. In the case of SPI films, as mentioned, development of the film structure involves disulfide bonds, hydrophobic interactions, and hydrogen bonds. Interactions between proteins and small molecules, including water, plasticizers, lipids, and other additives dispersed in the space of the matrix, also contribute to the mechanical behavior of protein films (Chen, 1995). Several factors, including surface charges, hydrophobicities, structures and structural stability, polymer chain length, geometry, molecular mass and distribution, and even genetic variants, may significantly affect the mechanical properties of protein films (Kester and Fennema, 1986; Chen, 1995). In general, soy protein films, similar to films from other proteins, have only moderate mechanical properties compared to commonly used plastic films (e.g., polyethylene, polypropylene, and polyvinylidene chloride) (Gennadios et al., 1994; Krochta and De Mulder-Johnston, 1997).

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The effects of various factors, including physical, chemical, and enzymatic treatments, on the properties of soy protein films are discussed below.

The protein concentration of film-forming solutions is an important factor in preparing soy protein-based films. The mechanical properties of films cast from aqueous film-forming solutions containing 5, 8, or 10% (w/w) defatted soy flour (DSF), soy protein concentrate (SPC), or SPI are shown in Table 4.1 (Park and Hettiarachchy, 2000). Film TS increased with increasing concentrations of DSF, SPC, or SPI in the film-forming solutions. For all types of films, the greatest values of TS were observed at 10% concentration of soy products. Most likely, insoluble components in DSF and SPC contributed to the lower TS values of DSF and SPC films compared to SPI films.

It is generally believed that heating film-forming solutions of sulfur-containing proteins (e.g., soy protein, wheat gluten, whey proteins, and egg albumen) prior to casting results in cast films with improved mechanical properties and greater water resistance (McHugh et al., 1994; Stuchell and Krochta, 1994; Handa et al., 1999; Pérez-Gago et al., 1999; Roy et al., 1999). Heating above 60 C and alkaline conditions (below pH 10.5) promote soy protein polymerization by altering the three-dimensional structure through unfolding of polypeptide chains, thus exposing sulfhydryl and hydrophobic groups (Kelley and Pressey, 1966). The alkaline conditions also favor thiol-disulfide inter-

128 change reactions by de-protonating sulfhydryl groups and allowing them to act as nucleophiles. Upon drying of the cast solutions, unfolded macromolecules approach each other and are linked by disulfide bonds and hydrophobic interactions. Table 4.1 shows the effect of heating alkaline (pH 9) film-forming solutions at 75 or 95 C for 45 minutes on mechanical properties of cast DSF, SPC, and SPI films plasticized with glycerol (50% w/w of DSF, SPC, or SPI) (Park and Hettiarachchy, 2000). DSF, SPC, and SPI films from solutions heated at 95 C had greater TS and E values than films from solutions heated at 75 C. It is noted that DSF, SPC, and SPI films cast from non-heated solutions cracked into small pieces upon drying due to limited intermolecular interaction.

Protein film formation is pH dependent. Soy protein films can be formed at both alkaline and acidic conditions. In a study on interfacial film formation, lipids (corn oil or glycerol trioleate) were added to SPI solutions, and satisfactory films were produced at pH values lower than 4.6 (as low as 1), which is the isoelectric region of soy protein, and up to pH 6.5 (Flint and Johnson, 1981). Above pH 7.4, the increasing solubility of SPI limited film formation. The observed film formation at very low pH values, where SPI solubility is high, suggested that lipophilic groups were probably available for binding at the oil-water interface (Flint and Johnson, 1981). Films prepared at acidic or alkaline conditions may have advantageous antimicrobial activity that merits investigation. Cast SPI films formed at alkaline conditions (pH 8 to 11) had greater TS, greater E, and lower WVP than films formed at acidic conditions (pH 1 to 3) (Gennadios et al., 1993b). Similar to heating, alkaline conditions facilitate soy protein denaturation, thus promoting the formation of disulfide bonds within the structure of the dried films (Gennadios et al., 1993b). The natural pH of DSF film-forming solutions is 6.7. DSF films were formed at pH between 6 and 10. Below pH 6, DSF precipitated, and films could not be formed. Table 4.2 shows that DSF, SPC, or SPI films prepared at pH 6 had substantially greater WVP than films prepared at pH 7, 8, 9, or 10. This likely resulted from partial inhibition of thiol-disulfide interchange reactions and thiol oxidation reactions at the acidic pH value of 6. Similarly, Brandenburg et al. (1993) reported that SPI films cast from solutions at pH 6 had inferior properties (greater WVP and oxygen permeability, and lower TS and E) compared with films cast from solutions at pH 8, 10, or 12.

Besides heating soy protein film-forming solutions prior to casting, cast SPI films have also been subjected to heat treatments (heat curing) to modify their properties (Gennadios et al., 1996; Rangavajhyala et al., 1997; Rhim et al.,


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2000). Heating SPI films at 80 or 95 C for 2, 6, 14, or 24 hours increased film TS and reduced film E, WVP, and water solubility (Gennadios et al., 1996). These effects were more pronounced as heating temperature and time increased (Gennadios et al., 1996). SPI films heat-cured at 90 C for 24 hours had substantially lower protein solubility in various solvents than control non-heated films (Rhim et al., 2000). For example, the protein solubility in water for heated films was 0.36 mg/mL versus 0.03 mg/mL for non-heated films (Rhim et al., 2000). Improvements in mechanical toughness and moisture resistance have also been documented for films from other proteins such as gelatin (Yannas and Tobolsky, 1967), collagen (Weadock et al., 1984), corn zein (Julius, 1967), wheat gluten (Ali et al., 1997), and whey protein (Miller et al., 1997). In general, thermal treatments of proteins promote formation of intra- and intermolecular cross-links, which mainly involve lysine and cystine amino acid residues (Cheftel et al., 1985). The increased cross-linking within the protein network directly affects film properties (e.g., increased TS and reduced E). Moreover, this heat-induced cross-linking probably involves polar protein groups, thus enhancing film hydrophobicity. For example, SPI films heat-cured in an air-circulating oven at 90 C for 24 hours had lower moisture contents than untreated SPI films after conditioning at 50% RH and 25 C for two days (Rhim et al., 2000). Because water has a plasticizing effect on pro-

130 tein-based films (Gontard et al., 1993), the decreased moisture uptake by heat-cured protein films likely contributes further to their increased mechanical toughness.

Use of enzymes that promote protein cross-linking to improve protein film properties appears feasible. Stuchell and Krochta (1994) incubated (37 C for 24 hours) SPI film-forming solutions with horseradish peroxidase prior to casting. This enzyme catalyzes oxidation of tyrosine amino acid residues, thus forming di-, tri-, and tert-tyrosine. Treatment with horseradish peroxidase did not affect film WVP, but it increased film TS and protein solubility and decreased film E. Because of the increase in protein solubility of enzyme-treated SPI films, the authors concluded that horseradish peroxidase caused protein degradation in addition to cross-linking (Stuchell and Krochta, 1994). Transglutaminase (TG), an enzyme that catalyzes formation of -( -glutamyl)-lysyl cross-links between the glutamine and lysyl groups of protein molecules, has been receiving attention in the food industry (Nielsen, 1995). Formation of heterologous biopolymers combining soy protein and casein through TG cross-linking has been reported (Motoki et al., 1987). Also, TG has been used to prepare cross-linked biopolymers from 11S soy protein fraction and whey protein (Yildirim et al., 1996; Yildirim and Hettiarachchy, 1997). Cast, heterologous films prepared from TG-cross-linked mixtures (1:1 w/w) of 11S soy globulin and whey protein isolate had lower solubility in water and two times greater TS than control homologous films (Yildirim and Hettiarachchy, 1998). However, the use of cross-linking enzymes, such as TG, is costly.

The ability of low molecular weight aldehydes (e.g., formaldehyde) to react with primary amino groups and sulfhydryl groups in proteins, thereby forming intra- and intermolecular cross-links, is well documented (Feeney et al., 1975). The effect of cross-linking by formaldehyde on SPI films has been studied (Ghorpade et al., 1995b; Rhim, 1998; Rhim et al., 2000). Formaldehyde can be directly added to SPI film-forming solutions or it can be applied by immersing dried SPI films into formaldehyde solutions (Rhim et al., 2000). Treatment with formaldehyde resulted in larger than twofold increases in TS and puncture strength, while also reducing the WVP (by about 6%) and the water solubility (by about 42%) of SPI films (Ghorpade et al., 1995b). The oxygen permeability of the formaldehyde-treated SPI films increased slightly (Ghorpade et al., 1995b). Similarly, increased oxygen permeability following treatment with formaldehyde has been reported for collagen films (Lieberman and Gilbert,


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1973). In another study, formaldehyde rendered SPI films practically insoluble in water and other solvents (i.e., 0.01 N hydrochloric acid, 0.01 N sodium hydroxide, 4 M urea, and 0.2 M 2-mercaptoethanol) (Rhim et al., 2000). Dialdehyde starch, a polymeric aldehyde obtained by reacting native starch with periodic acid (Pfeifer et al., 1960), also showed potential in cross-linking SPI films (Rhim et al., 1998, 2000). SPI films containing dialdehyde starch at 5 or 10% w/w of SPI had increased TS (by about 20%) compared to control SPI films (Rhim et al., 1998). Also, SPI films containing dialdehyde starch (10% w/w of SPI) were practically insoluble in water and other solvents (Rhim et al., 2000). Although aldehydes are effective protein cross-linkers, they are generally toxic, and this limits their possible use with protein films to non-edible applications.

Reportedly, aromatic amino acids, such as tyrosine and phenylalanine, can absorb UV radiation and recombine to form covalent cross-links in proteins (Fujimori, 1965; Forbes and Sullivan, 1966; Tomihata et al., 1992). Gennadios et al. (1998b) subjected cast SPI films to UV irradiation (253.7 nm wavelength). The irradiated films had increased TS and reduced E compared to control non-irradiated films. Film WVP was not affected by UV irradiation. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis patterns for UV-treated SPI films showed bands of aggregates, increasing in intensity with UV dosage, that were lacking in electrophoretic patterns of control films (Gennadios et al., 1998b). Also, UV-treated SPI films had lower protein solubility in water than control films (0.11 versus 0.36 mg/mL) (Rhim et al., 2000). In contrast to UV radiation, subjecting SPI films to -irradiation (5, 10, 20, or 30 kGy) did not affect film TS and E (Ghorpade et al., 1995a).

The water barrier ability of protein films can be improved through combination with hydrophobic lipid materials (Gennadios et al., 1994; Krochta and DeMulder-Johnston, 1997). One approach is to add the lipids to protein film-forming solutions, which are then cast to prepare emulsified, bi- or multi-component films. Alternatively, molten lipids can be deposited (laminated) onto pre-formed protein films to prepare bi- or multi-layer films. Fatty acids (i.e., lauric, myristic, palmitic, or oleic acid) were incorporated into SPI film-forming solutions at 10 to 30% w/w of SPI to prepare soy protein-fatty acid bi-component films (Gennadios et al., 1998a). Sucrose stearate, an emulsifier promoting oil-in-water emulsions, was also added (5% w/w of the fatty acid) to the film-forming solutions. Fatty acid-containing SPI films had notably lower WVP values than control SPI films. Also, oleic acid had a plasticization

132 effect on the films. For example, SPI films containing oleic acid at 10% w/w of SPI had an E of 228% versus an E of 70% for the control SPI films. However, because lipids lack the structural integrity of proteins or polysaccharides, the addition of fatty acids substantially reduced film TS (Gennadios et al., 1998a). Another concern regarding protein-lipid films would be their potential susceptibility to lipid oxidation.

The formation of covalent cross-links between proteins and propylene glycol alginate (PGA) has been documented (Ranganayaki and Stainsby, 1978; McKay et al., 1985). The -amino groups of lysine are involved in this interaction of proteins with PGA. For example, the available lysine content of SPI was reduced by about 18% after interaction with PGA (Mohamed and Stainsby, 1984). Alkaline conditions (pH 8) favor the protein-PGA interaction (McKay et al., 1985; Shih, 1994). Formation of SPI-PGA films has been reported (Shih, 1994; Rhim et al., 1999). SPI-PGA films prepared with low levels of PGA (10% w/w of protein) had increased TS and water resistance (Shih, 1994; Rhim et al., 1999). Although PGA is edible, its use as a cross-linker for protein films would reduce the nutritional value of the protein by impairing the availability of lysine. Nevertheless, in most cases, an edible film or coating is unlikely to contribute notably to the nutritional content of foods.

Preparation of SPI films following interaction with succinic anhydride was reported by Ghorpade et al. (1995b). Succinylation did not affect film TS, puncture strength, WVP, or oxygen permeability. However, the succinylated SPI films had notably greater water solubility than control SPI films (Ghorpade et al., 1995b). A concern with succinylation is that it lowers the nutritional value of proteins by forming succinyl-lysine isopeptides, which are resistant to cleavage by pancreatic enzymes (Cheftel et al., 1985).

As mentioned, soy protein films, similar to soy protein gels, are formed through polymerization of heat-denatured proteins, with disulfide bonds and hydrophobic interactions being the main forces maintaining the film network (Shimada and Cheftel, 1988). In sulfur-containing proteins, cysteine groups can undergo polymerization via sulfhydryl-disulfide interchange reactions during heating to form a continuous covalent network upon cooling (Arntfield et al., 1991). Addition of cysteine (1% w/w) to SPI film-forming solutions at pH 7 increased the number of disulfide bonds in the solutions by 56.6% and

Potential Applications and Future Research Needs

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also increased the TS of cast films from 5.7 to 8.1 MPa (Were et al., 1999). Therefore, the addition of cysteine to SPI films may be advantageous due to disulfide rearrangement.

Development of a standard method for evaluating biodegradability of biopolymer-based plastics is required to develop and evaluate new biopolymer films. Methods used to assess biodegradation of plastics include following changes of mechanical properties in soil; BOD changes or CO2 evolution in slugs; molecular changes by enzymatic or microbial treatment; mechanical property changes or CO2 evolution in soil; gas evolution by anaerobic digestion; and mechanical property changes in various compost soil conditions. Such methods help establish the degradation rate (change in chemical structure, decrease in mechanical strength, fragmentation, or weight loss) of the polymeric material or plastic product under laboratory scale composting conditions. SPI films were stored in containers with compost of 20 or 35% water content at 15 or 25 C for 30 days or until the test specimens disappeared completely, whichever occurred first (Park and Hettiarachchy, 2000). TS of films exposed to the aerobic composting environment was determined after washing the specimens free of any residue using a solvent-free, neutral pH cleaning agent. When the water content of the compost during storage was 20%, film color and shape remained relatively stable up to seven days. However, when the water content of the compost was 35%, film TS decreased rapidly. The degree of degradation of SPI films was greater at 25 C than at 15 C (Park and Hettiarachchy, 2000). The degradability of the SPI films during storage appeared to be more sensitive to moisture than to temperature.

The good oxygen barrier ability of SPI films could be utilized in the manufacture of multilayer packaging where the protein films would function as the oxygen barrier-providing layer. SPI coatings on pre-cooked meat products could control lipid oxidation and limit surface moisture loss. Incorporation of antioxidant and flavoring agents in SPI coatings could improve overall quality characteristics of food products (Kunte et al., 1997). In recent studies, SPI films carrying antimicrobial compounds (nisin or lysozyme) showed inhibitory potential against Lactobacillus plantarum (Padgett et al., 1998) and Listeria monocytogenes (Ko et al., 2000). Also, SPI films may find application as microencapsulating agents of flavors and medications or in coatings of fruits, vegetables, and cheese (Petersen et al., 1999). Protective SPI coatings could also be used on certain food products, such as meat pies and high moisture/low

134 sugar cakes, which require films that are highly permeable to water vapor (Gennadios et al., 1993a). However, considering films from SPI or other biopolymers for edible packaging applications will require the completion of relevant sensory studies, which are scarce in the literature. At present, production of edible protein-based films is considered to be expensive compared to plastic films, and research on cost reduction and production in large scale is necessary to promote the application of films from soy protein or other proteins.

Alcantara, C.R., Rumsey, T.R. and Krochta, J.M. 1998. “Drying Rate Effect on the Properties of Whey Protein Films,” J. Food Process Engr., 21:387–405. Ali, Y., Ghorpade, V.M. and Hanna, M.A. 1997. “Properties of Thermally-Treated Wheat Gluten Films,” Industr. Crops Prod., 6:177–184. Arnold, L.K. 1968. Introduction to Plastics. Ames, IA: The Iowa State University Press. Arntfield, S.D., Murray, E.D. and Ismond, M.A.H. 1991. “Role of Disulfide Bonds in Determining the Rheological and Microstructural Properties of Heat-Induced Protein Networks from Ovalbumin and Vicilin,” J. Agric. Food Chem., 39:1376–1385. Banker, G.S. 1966. “Film Coating Theory and Practice,” J. Pharm. Sci., 55:81–89. Brandenburg, A.H., Weller, C.L. and Testin, R.F. 1993. “Edible Films and Coatings from Soy Protein,” J. Food Sci., 58:1086–1089. Cheftel, J.C., Cuq, J.L. and Lorient, D. 1985. “Amino Acids, Peptides, and Proteins” in Food Chemistry, 2nd edition, ed., O.R. Fennema, New York: Marcel Dekker, pp. 245–369. Chen, H. 1995. “Functional Properties and Applications of Edible Films Made of Milk Proteins,” J. Dairy Sci., 78:2563–2583. Chuah, E.C., Idrus, A.Z., Lim, C.L. and Seow, C.C. 1983. “Development of an Improved Soya Protein-Lipid Film,” J. Food Technol., 18:619–627. Cunningham, P., Ogale, A.A., Dawson, P.L. and Acton, J.C. 2000. “Tensile Properties of Soy Protein Isolate Films Produced by a Thermal Compaction Technique,” J. Food Sci., 65:668–671. Farnum, C., Stanley, D.W. and Gray, J.I. 1976. “Protein-Lipid Interactions in Soy Films,” Can. Inst. Food Sci. Technol. J., 9:201–206. Feeney, R.E., Blankenhorn, G. and Dixon, H.B.F. 1975. “Carbonyl-Amine Reactions in Protein Chemistry,” Adv. Protein Chem., 29:135–203. Flint F.O. and Johnson R.F.P. 1981. “A Study of Film Formation by Soy Protein Isolate,” J. Food Sci., 46:1351–1353. Forbes, W.F. and Sullivan, P.D. 1966. “The Effect of Radiation on Collagen. I. Electron-spin Resonance Spectra of 2537-Å-Irradiated Collagen,” Biochim. Biophys. Acta, 120:222–228. Fujimori, E. 1965. “Ultraviolet Light-Induced Change in Collagen Macromolecules,” Biopolym., 3:115–119. Gennadios, A. and Weller, C.L. 1991. “Edible Films and Coatings from Soymilk and Soy Protein,” Cereal Foods World, 36:1004–1009. Gennadios, A., Weller, C.L. and Testin, R.F. 1993a. “Temperature Effect on Oxygen Permeability of Edible Protein-based Films,” J. Food Sci., 58:212–214, 219. Gennadios, A., Brandenburg A.H., Weller C.L. and Testin, R.F. 1993b. “Effect of pH on Properties of Wheat Gluten and Soy Protein Isolate Films,” J. Agric. Food Chem., 41:1835–1839. Gennadios, A., McHugh, T.H., Weller C.L. and Krochta, J.M. 1994. “Edible Coating and Films Based on Proteins” in Edible Coating and Films to Improve Food Quality, ed., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 201–277.

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Gennadios, A., Ghorpade, V.M., Weller, C.L. and Hanna, M.A. 1996. “Heat Curing of Soy Protein Films,” Trans. ASAE, 39:575–579. Gennadios, A., Cezeirat, C., Weller, C.L. and Hanna, M.A. 1998a. “Emulsified Soy Protein-Lipid Films” in Paradigm for Successful Utilization of Renewable Resources, eds., D.J. Sessa and J.L. Willett, Champaign, IL: AOCS Press, pp. 213–226. Gennadios, A., Rhim, J.W., Handa, A., Weller, C.L. and Hanna, M.A. 1998b. “Ultraviolet Radiation Affects Physical and Molecular Properties of Soy Protein Films,” J. Food Sci., 63:225–228. Ghorpade, V.M. and Hanna, M.A. 1996. “Mechanical Properties of Soy Protein-Polyethylene Ribbon and Film Extrudates,” Trans. ASAE, 39:611–615. Ghorpade, V.M., Gennadios, A., Hanna, M.A. and Weller, C.L. 1995a. “Soy Protein Isolate/Poly(ethylene oxide) Films,” Cereal Chem., 72:559-563. Ghorpade, V.M., Li, H., Gennadios, A. and Hanna, M.A. 1995b. “Chemically Modified Soy Protein Films,” Trans. ASAE, 38:1805–1808. Gontard, N., Guilbert, S. and Cuq, J.-L. 1993. “Water and Glycerol as Plasticizers Affect Mechanical and Water Vapor Barrier Properties of an Edible Wheat Gluten Film,” J. Food Sci., 58:206–211. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation: Theory and Practice, ed., M. Mathlouthi, Essex, England: Elsevier, pp. 371–394. Handa, A., Gennadios, A., Froning, G.W., Kuroda, N. and Hanna, M.A. 1999. “Tensile, Solubility, and Electrophoretic Properties of Egg White Films as Affected by Surface Sulfhydryl Groups,” J. Food Sci., 64:82–85. Jangchud, A. and Chinnan, M.S. 1999. “Peanut Protein Film as Affected by Drying Temperature and pH of Film Forming Solution,” J. Food Sci., 64:153–157. Julius, A. July 4, 1967. U.S. patent 3,329,509. Kelley, J.J. and Pressey, R. 1966. “Studies with Soybean Protein and Fiber Formation,” Cereal Chem., 43:195–206. Kester, J.J. and Fennema, O.R. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Ko, S., Janes, M.E., Hettiarachchy, N.S. and Johnson, M.G. 2000. “Effects of Hydrophobicity/Hydrophilicity of Edible Films on Control of Listeria monocytogenes by Nisin,” J. Food Sci., submitted. Krochta, J.M and De Mulder-Johnston, C. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Kunte, L.A., Gennadios, A., Cuppett, S.L., Hanna, M.A. and Weller, C.L. 1997. “Cast Films from Soy Protein Isolates and Fractions,” Cereal Chem., 74:115–118. Lieberman, E.R. and Gilbert, S.G. 1973. “Gas Permeation of Collagen Films as Affected by Cross-Linkage, Moisture, and Plasticizer Content,” J. Polym. Sci., 41:33–43. Liu, K. 1997. Soybeans: Chemistry, Technology, and Utilization. New York: Chapman and Hall, pp. 198–202. Liu, K. 2000. “Expanding Soybean Food Utilization,” Food Technol., 54(7):46–48, 50, 52, 54, 56, 58. McHugh , T.H., Aujard, J.-F. and Krochta, J.M. 1994. “Plasticized Whey Protein Edible Films: Water Vapor Permeability Properties,” J. Food Sci., 59:416–419, 423. McKay, J.E., Stainsby, G. and Wilson, E.L. 1985. “A Comparison of the Reactivity of Alginate and Pectate Esters with Gelatin,” Carbohydr. Polym., 5:223–236. Miller, K.S. and Krochta, J.M. 1997. “Oxygen and Aroma Barrier Properties of Edible Films: A Review,” Trends Food Sci. Technol., 8:228–237. Miller, K.S., Chiang, M.T. and Krochta, J.M. 1997. “Heat Curing of Whey Protein Films,” J. Food Sci., 62:1189–1193.

136 Mohamed, S.B. and Stainsby, G. 1984. “Lysine Availability in Protein-Alginate Ester Gels,” Food Chem., 14:1–10. Motoki, M., Nio, N. and Takinami, K. 1987. “Functional Properties of Heterologous Polymer Prepared by Transglutaminase between Milk Casein and Soybean Globulin,” Agric. Biol. Chem., 51:237–239. Naga, M., Kirihara, S., Tokugawa, Y., Tsuda, F. and Hirotsuka, M. October 29, 1996. U.S. patent 5,569,482. Nielsen, P.M. 1995. “Reactions and Potential Industrial Applications of Transglutaminase. Review of Literature and Patents,” Food Biotechnol., 9:119–156. Ogale, A.A., Cunningham, P., Dawson, P.L. and Acton, J.C. 2000. “Viscoelastic, Thermal, and Microstructural Characterization of Soy Protein Isolate Films,” J. Food Sci., 65:672–679. Okamoto, S. 1978. “Factors Affecting Protein Film Formation,” Cereal Foods World, 23:256–262. Padgett, T., Han, I.Y. and Dawson, P.L. 1998. “Incorporation of Food-Grade Antimicrobial Compounds into Biodegradable Packaging Films,” J. Food Prot., 61:1330–1335. Park, S.K. and Hettiarachchy, N.S. 2000. Unpublished data. University of Arkansas, Fayetteville, AR, USA. Pérez-Gago, M.B., Nadaud, P. and Krochta, J.M. 1999. “Water Vapor Permeability, Solubility, and Tensile Properties of Heat-Denatured versus Native Whey Protein Films,” J. Food Sci., 64:1034–1037. Petersen, K., Nielsen, V.P., Bertelsen, G., Lawther, M., Olsen, M.B., Nilsson, N.H. and Mortensen, G. 1999. “Potential of Biobased Materials for Food Packaging,” Trends Food Sci. Technol., 10:52–68. Pfeifer, V.F., Sohns, V.E., Conway, H.F., Lancaster, E.B., Dabic, S. and Griffin, E.L. Jr. 1960. “Two-Stage Process for Dialdehyde Starch Using Electrolytic Regeneration of Periodic Acid,” Industr. Engr. Chem., 52:201–206. Pszczola, D.E. 1998. “Addressing Functional Problems in Fortified Foods,” Food Technol., 52(7):38, 40–41, 44, 46. Rampon, V., Robert, P., Nicolas, N. and Dufour, E. 1999. “Protein Structure and Network Orientation in Edible Films Prepared by Spinning Process,” J. Food Sci., 64:313–316. Ranganayaki, M.D. and Stainsby, G. 1978. “Thermostable Gels from Proteins-Alginate Ester Complexes,” Leather Sci., 25:441–442. Rangavajhyala, N., Ghorpade, V.M. and Hanna, M.A. 1997. “Solubility and Molecular Properties of Heat-Cured Soy Protein Films,” J. Agric. Food Chem., 45:4204–4208. Rayner, M., Ciolfi, V., Maves, B., Stedman, P. and Mittal, G.S. 2000. “Development and Application of Soy-Protein Films to Reduce Fat Intake in Deep-Fried Foods,” J. Sci. Food Agric., 80:777–782. Rhim, J.W. 1998. “Modification of Soy Protein Film by Formaldehyde,” Korean J. Food Sci. Technol., 30:372–378. Rhim, J.W., Gennadios, A., Handa, A., Weller, C.L. and Hanna, M.A. 2000. “Solubility, Tensile, and Color Properties of Modified Soy Protein Isolate Films,” J. Agric. Food Chem., 48:4937–4941. Rhim, J.W., Gennadios, A., Weller, C.L., Cezeirat, C. and Hanna, M.A. 1998. “Soy Protein Isolate-Dialdehyde Starch Films,” Industr. Crops Prod., 8:195–203. Rhim, J.W., Wu, Y., Weller, C.L. and Schnepf, M. 1999. “Physical Characteristics of a Composite Film of Soy Protein Isolate and Propyleneglycol Alginate,” J. Food Sci., 64:149–152. Roy, S., Weller, C.L., Gennadios, A., Zeece, M.G. and Testin, R.F. 1999. “Physical and Molecular Properties of Wheat Gluten Films Cast from Heated Film-Forming Solutions,” J. Food Sci., 64:57–60. Shih, F.F. 1994. “Interaction of Soy Isolate with Polysaccharide and Its Effect on Film Properties,” JAOCS, 71:1281–1285. Shimada, K. and Cheftel, J.C. 1988. “Determination of Sulfhydryl Groups and Disulfide Bonds in Heat-Induced Gels of Soy Protein Isolate,” J. Agric. Food Chem., 36:147–153.

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Shirai, M., Watanabe, K. and Okamoto, S. 1974. “Contribution of 11S and 7S Globulins of Soybean Protein to the Formation and Properties of Yuba-Film,” Nippon Shokuhin Kogyo Gakkaishi, 21:324–328. Sian, N.K. and Ishak, S. 1990a. “Effect of pH on Formation, Proximate Composition and Rehydration Capacity of Winged Bean and Soybean Protein-Lipid Film,” J. Food Sci., 55:261–262. Sian, N.K. and Ishak, S. 1990b. “Effect of pH on Yield, Chemical Composition and Boiling Resistance of Soybean Protein-Lipid Film,” Cereal Foods World, 35:748, 750, 752. Soy Protein Council. 1987. Soy Protein Products: Characteristics, Nutritional Aspects, and Utilization. Washington, DC. Soya & Oilseed Bluebook. 1999. Bar Harbor, ME: Soyatech Inc., pp. 225–228. Stuchell, Y.M. and Krochta, J.M. 1994. “Enzymatic Treatments and Thermal Effects on Edible Soy Protein Films,” J. Food Sci., 59:1332–1337. Subirade, M., Kelly, I., Guéguen, J. and Pézolet, M. 1998. “Molecular Basis of Film Formation from a Soybean Protein: Comparison between the Conformation of Glycinin in Aqueous Solution and in Films,” Biol. Macromol., 23:241–249. Tomihata, K., Burczak, K., Shiraki, K. and Ikada, Y. 1992. “Crosslinking and Biodegradation of Native and Denatured Collagen,” Polym. Preprints, 33:534–535. Torres, J.A. 1994. “Edible Films and Coatings from Proteins” in Protein Functionality in Food Systems, eds., N.S. Hettiarachchy and G.R. Ziegler, New York: Marcel Dekker, pp. 467–507. USB (United Soybean Board). 2000. http://www.unitedsoybean.org. USDA (United States Department of Agriculture). 1999. http://www.usda.gov. Weadock, K., Olson, R.M. and Siver, F.H. 1984. “Evaluation of Collagen Cross-Linking Techniques,” Biomater. Med. Dev. Art. Org., 11:293–318. Were, L., Hettiarachchy, N.S. and Coleman, M. 1999. “Properties of Cysteine-Added Soy Protein-Wheat Gluten Films,” J. Food Sci., 64:514–518. Witherly, S.A. 1990. “Soy Formulas Are Not Hypoallergenic,” Am. J. Clin. Nutr., 51:705–706. Wolf, W.J. and Cowan, J.C. 1975. Soybeans as a Food Source. Boca Raton, FL: CRC Press. Wu, L.C. and Bates, R.P. 1972a. “Soy Protein-Lipid Films. 1. Studies on the Film Formation Phenomenon,” J. Food Sci., 37:36–39. Wu, L.C. and Bates, R.P. 1972b. “Soy Protein-Lipid Films. 2. Optimization of Film Formation,” J. Food Sci., 37:40–44. Yannas, I.V. and Tobolsky, A.V. 1967. “Cross-Linking of Gelatine by Dehydration,” Nature, 215:509–510. Yildirim, M. and Hettiarachchy, N.S. 1997. “Biopolymers Produced by Cross-Linking Soybean 11S Globulin with Whey Proteins Using Transglutaminase,” J. Food Sci., 62:270–275. Yildirim, M. and Hettiarachchy, N.S. 1998. “Properties of Films Produced by Cross-Linking Whey Proteins and 11S Globulin Using Transglutaminase,” J. Food Sci., 63:248–252. Yildirim, M., Hettiarachchy, N.S. and Kalapathy, U. 1996. “Properties of Biopolymers from Cross-Linking Whey Protein Isolate and Soybean 11S Globulin,” J. Food Sci., 61:1129–1131, 1164.

Formation and Properties of Cottonseed Protein Films and Coatings

C

OTTON is a plant that has been cultivated worldwide for centuries as a fiber

crop. It is currently grown for fiber in more than 70 different countries, but there are also market outlets for its oil. Cottonseed accounts for about 13% of world oilseed production. Oil production, although more than double that of fiber production, accounts for only 15% of the commercial value of the cottonseed crop. To optimize usage of this low-cost, oil-rich, and protein-rich product, the film-forming properties of cottonseed proteins were studied to produce biodegradable materials of economic and environmental interest (Wu and Bates, 1973; Marquié et al., 1996, 1997a, b). This chapter discusses preparation of lipoprotein and protein films from cottonseed flours and their chemical modifications by gossypol, formaldehyde, glutaraldehyde, and glyoxal. Mechanical properties and biodegradability of such films are also discussed in light of potential applications.

With worldwide production estimated at about 33 million metric tons, cottonseed is the most important source of plant proteins after soybean. Generally, cottonseed is crushed to extract oil and produce cake that is chiefly used for ruminant livestock feed (Tacher et al., 1985; Zongo and Coulibaly, 1993). As it 139

140

contains gossypol, a toxic compound, this cake cannot be used for human consumption. Gossypol is present in seeds of most cotton cultivars grown in the world (glanded varieties). A few new glandless varieties do not produce this toxic compound. Cottonseed proteins make up 30 to 40% (w/w) of the cottonseed kernel (Table 5.1). Other important cottonseed components include lipids, soluble carbohydrates, cellulose, minerals, phytates, and polyphenolic pigments. The protein components are mainly globulins (60%) and albumins (30%) with lesser proportions of prolamins (8.6%) and glutenins (0.5%) (Saroso, 1989). Globulins include two protein fractions, the 11S and 7S types called gossypin and congossypin, respectively (Zarins and Cherry, 1981; Dumay et al., 1986; Reddy and Rao, 1988; Cho et al., 1992). They are insoluble in water at pH 6.8 and can be readily extracted by the salting-in process (Martinez et al.,

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Figure 5.1 Effect of pH on solubility of cottonseed proteins (% w/w). (Adapted from Marquié, 1996.)

1970; Bérot and Guéguen, 1985; Dumay et al., 1986) using the fractionation techniques described by Osborne. Gossypin is a hexameric protein, with an structure of 180 to 300 kDa molecular weight (Mw), depending on the dissociation conditions of the protein (Dumay et al., 1986; Cho et al., 1992). Congossypin is also an oligomer of 127 to 180 kDa molecular weight composed of three subunits (58 and 52 kDa Mw) partly modified by post-translational cleavage, leading to lower constitutive polypeptides (35, 23, and 18 kDa Mw) (Rossi-Fanelli et al., 1964; Martinez et al., 1970; Martinez, 1979; Youle and Huang, 1979; Zarins and Cherry, 1981; Zarins et al., 1984; Reddy and Rao, 1988; Cho et al., 1992). Albumins are grouped within a protein fraction with a sedimentation coefficient of 2S. They are composed of low Mw (10 to 25 kDa Mw) proteins that are water soluble at pH 6.8 (Martinez et al., 1970) and rich in lysine and sulfur amino acids, contrary to reserve globulins (Dumay et al., 1986). Cottonseed proteins have a high content of ionizable amino acids (aspartic and glutamic acids, arginine, histidine, and lysine) and a low content of sulfur amino acids (Table 5.2). These proteins are more soluble at basic pH than at acidic pH (Figure 5.1) with an isoelectric pH of around 5 (Bérot and Guéguen,

142 1985; Dumay et al., 1986). Cottonseed proteins are readily denatured by thermal treatment above 80 C, which leads to loss of solubility and nutritional value (Besançon et al., 1985). The temperature increase during the crushing of cottonseed leads to chemical interactions between gossypol and lysine and also to Maillard reactions (Marquié, 1987; Bourély, 1990).

Utilization of cottonseed proteins for film formation was first reported by Wu and Bates (1973). Their process involved soaking cottonseed kernels in hot water to prepare an “oilseed milk.” Films were successively formed on the surface of the heated (90 5 C) liquid, similarly to traditional formation of soy protein-lipid films (“yuba”) in East Asia (Wu and Bates, 1973). Due to their poor mechanical properties, films made with this process may be used as edible items but not as packaging materials. Cottonseed protein-based films were also obtained directly from cottonseed flours using a casting process (Figure 5.2) (Marquié, 1995, 1996; Marquié et al., 1995, 1997a, b, 1998). The first step involved solubilizing cottonseed flour proteins under appropriate conditions (solvent; pH; temperature; and addition of salts, plasticizers, and dissociating agents) to minimize protein-protein interactions. Changes in the protein structure prevented “remelting” of macromolecules and revealed potentially interactive zones. The dispersions were centrifuged to remove insoluble substances, and the supernatants were then homogenized. Plasticizer, such as glycerol, was then added to solutions (10–40% w/w, dry basis). Drying the cast film-forming solutions eliminated solvents and allowed the proteins to form three-dimensional networks through new inter- and intra-molecular linkages.

Film cohesiveness depends on various parameters such as polymer structure, film-forming process, temperature, pressure, solvent type, solids/solvent ratio (w/v), film application technique, and presence of plasticizers, bulking agents, and cross-linking agents (Gontard and Guilbert, 1994; Marquié, 1996). In contrast to protein isolates, the conditions required to obtain films from cottonseed flour-based solutions are difficult to determine because of the complexity of the raw material, which contains proteins, lipids, ash, cellulose, and carbohydrates. Based on preliminary experiments, we determined the feasibility range of cottonseed flour films, i.e., optimal pH (8 to 12), temperature (20 to 60 C), solid/solvent ratio (10 to 50% w/v), use of dispersive agents, and plasticizer content (10 to 50% w/w, dry basis). The combined effects of cottonseed flour/water ratio (CFWR), dispersion pH, and temperature during preparation of film-forming solutions on the ex-

143

Figure 5.2 Preparation of cottonseed protein films and schematic diagram of film network formation. (Adapted from Marquié et al., 1997a.)

144 traction efficiency of dry matter, proteins, and lipids were evaluated using response surface methodology (Marquié, 1996). On a laboratory scale, under optimal conditions (CFWR of 10%; pH of 9 to 10; temperature from 50 to 70 C), the dry matter extraction efficiency did not exceed 70% for glandless or glanded flours. Based on the amino acid composition of cottonseed proteins, protein extraction efficiency is enhanced at alkaline pH. However, pH had an opposite effect on lipid content in film dry matter obtained from glandless and glanded flours. An increase in pH dispersion led to a decrease in dry matter lipid content of films obtained from glandless flour, whereas the dry matter lipid content of glanded flour-based films increased. With glanded flours, the pH effect on protein dispersion could be offset by temperature, permitting film formation with both high protein and low lipid content. Protein and lipid contents of films obtained from non-delipidated glandless and glanded flours should range within 40 to 60% (w/w, dry matter basis) and 2 to 15% (w/w, dry matter basis), respectively. Examples of cottonseed flour film compositions are given in Table 5.3 (Marquié et al., 1995). In this case, protein and lipid contents of films made from glandless or glanded flours were nearly the same (i.e., protein/lipid ratio close to 1.4) as soybean films obtained by the yuba-like process (Wu and Bates, 1972a,b). The main difference between films from glanded and glandless flours was the presence of gossypol in films from glanded cottonseed flour. Films obtained from defatted glandless flour were richer in protein than those obtained from non-defatted flour, and they contained less than 2% lipids. Characteristics of films from cottonseed flour (delipidated or not) are shown in Table 5.4 (Marquié et al., 1995). Films containing lipids were weaker than those from delipidated flour as evidenced by their lower puncture strength and deformation values (Table 5.4). This could be explained by the protein content, which was lower in lipoprotein films than in films made from delipidated cottonseed flour. Gossypol in glanded flour led to less soluble films. Lipoprotein films from glanded cottonseed flour were opaque. This could have been due to the lipid distribution within the protein matrix with some lipid droplets immobilized inside the protein network and others exuded to the film surface. Film

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color varied from light yellow (in the absence of gossypol) to brown (in the presence of gossypol). Films from glandless delipidated cottonseed flour were smooth and transparent.

Cottonseed flour films were excessively brittle when the glycerol content was below 10% (w/w, dry basis), and they became sticky when the glycerol content was higher than 30% (Marquié et al., 1995). From 10 to 30% (w/w, dry basis) glycerol, film puncture strength decreased rapidly (Figure 5.3). Glycerol decreases intermolecular forces between polymer chains, with a decrease in cohesion, tensile strength, and glass transition temperature (Guilbert, 1986). Films from glandless flour (delipidated or not) completely dissolved or dispersed in water within a few minutes, regardless of glycerol content. Films from glanded flour were less soluble (Figure 5.4) due to the effect of gossypol. Gossypol is oxidized into quinones at alkaline pH forming Schiff bases with -amino groups of lysine. In protein films, gossypol strengthens the protein network (Lyman et al., 1959; Tchiegang and Bourély, 1990; Marquié et al., 1995), entrapping lipids inside the protein structure, and thus reducing film solubility (Figure 5.5). Lipoprotein glanded films became less soluble as glycerol content increased (solubility was only 12.7% at 30% glycerol content). Total nitrogen contents of lipoprotein glanded films before and after solubility testing indicated that no proteins or peptides were lost in the solution. The observed solubility was thus due to leaching of non-protein film components, such as carbohydrates, ash, and pigments.

Figure 5.3 Effect of glycerol content on puncture strength (N per 100 m film thickness) of films from glandless, glanded, or delipidated glandless cottonseed flours. Reprinted with permission from Marquié et al., 1995. Copyright 1995, American Chemical Society.

Figure 5.4 Effect of glycerol content on solubility (% w/w, dry matter without accounting for glycerol content in film dry matter) of films from glandless, glanded, or delipidated glandless cottonseed flours. Reprinted with permission from Marquié et al., 1995. Copyright 1995, American Chemical Society.

146

Film Formation Techniques and Mechanisms

147

Figure 5.5 Effect of gossypol added to film-forming solution on solubility and puncture strength (N per 100 m film thickness) of films from glandless cottonseed flour. Reprinted with permission from Marquié et al., 1995. Copyright 1995, American Chemical Society.

Films can be strengthened by cross-linking agents that chemically modify proteins during preparation of film-forming solutions. Cross-linking agents are natural or synthetic molecules containing at least two reactive groups that are able to form covalent inter- and/or intra-molecular links between protein chains. These agents, when used to prepare protein-based films, strengthen the material through formation of new covalent bonds, while reducing film elasticity and solubility in water (Gennadios and Weller, 1992; Marquié et al., 1995; Marquié, 1996). Mechanical properties of cottonseed films were improved through protein cross-linking by formaldehyde, glyoxal, or glutaraldehyde (Marquié et al., 1995, 1997a). Protein cross-linking was monitored by HPLC determination of available lysine that did not react with the cross-linking agent (Marquié et al., 1997b, 1998). Chemical treatment of cottonseed proteins with formaldehyde, using a formaldehyde/reactive lysine molar ratio in the film-forming solution greater than 5, produced films with greater puncture strength than using glyoxal or glutaraldehyde (Figure 5.6). In alkaline solution, protein cross-linking by formaldehyde [Figure 5.7(a)] formed short methylene cross-links between

148 lysine amino groups (Fraenkel-Conrat and Olcott, 1946; Fraenkel-Conrat et al., 1947; Means and Feeney, 1968; Bizzini and Raynaud, 1974), i.e., guanidinyle, amide, and phenol. Formation of the first methylene cross-links involving some of the numerous reactive groups on cottonseed proteins likely changed the protein conformation, thus dramatically reducing molecular mobility and limiting accessibility of remaining lysine groups to formaldehyde for further reactions. Under mild alkaline conditions, glyoxal is known to condense with the guanidinyle group in an initial reaction very similar to Schiff base formation, which undergoes further rearrangement to form cyclic products [Figure 5.7(b )] unable to cross-link proteins (Shan, 1991). In another type of reaction, glyoxal may also react with lysine [Figure 5.7(b)], and this reaction was important under our experimental conditions (Figure 5.8). The increase in film puncture strength at a glyoxal/reactive lysine molar ratio of 8 (Figure 5.6) could have resulted from protein cross-linking involving lysyl groups. At higher molar concentrations of glyoxal in film-forming solutions, film puncture strength decreased despite the decrease in reactive lysine content within films. In the presence of excess glyoxal in the film-forming solution, it could be assumed that only part of this reagent led to protein cross-linking. The glyoxal molecules that reacted with a lysyl group only or remained free in the protein network may have decreased film cohesion by reducing intermolecular forces within the protein network. In addition, chemical modification of the protein through reaction

Figure 5.6 Effect of cross-linking agent/lysine molar ratio on puncture strength of films from delipidated glandless cottonseed flour that were cross-linked with formaldehyde, glyoxal, or glutaraldehyde. Reprinted with permission from Marquié et al., 1997b. Copyright 1997, American Chemical Society.

Figure 5.7 Mechanisms of protein cross-linking by formaldehyde, glyoxal, or glutaraldehyde; (a) methylene cross-links formed with formaldehyde reaction; (b) cross-links and (b ) cyclic compounds formed with glyoxal; and (c) polyglutaraldehyde cross-links. Reprinted with permission from Marquié et al., 1997b. Copyright 1997, American Chemical Society.

Figure 5.8 Effect of cross-linking agent/lysine molar ratio on reactive lysine content in delipidated glandless cottonseed flour films cross-linked by formaldehyde, glyoxal, or glutaraldehyde. Reprinted with permission from Marquié et al., 1997b. Copyright 1997, American Chemical Society.

149

150 between arginyl groups and glyoxal, without cross-linking, may have led to the same phenomenon. Unlike formaldehyde and glyoxal, glutaraldehyde reacts only with lysyl groups (Monsan et al., 1975). Moreover, this cross-linking agent forms long unsaturated polymers in basic solutions (Monsan et al., 1975), which may include up to eight elementary glutaraldehyde molecules (Korn et al., 1972). These polymers, which possess many aldehyde functional groups, probably produced long flexible cross-links [Figure 5.7(c)] with protein chains that increased molecular mobility and enhanced the accessibility of remaining lysyl groups. The molecular structure of long bridges formed between glutaraldehyde cross-linked proteins decreased intermolecular forces among polymer chains, resulting in decreased film puncture strength compared with films cross-linked by glyoxal or formaldehyde. As a result, films cross-linked by glutaraldehyde had lower puncture strength than those cross-linked by glyoxal. When higher molar concentrations of glutaraldehyde were added to the film-forming solution, film puncture strength decreased (Figure 5.6), probably due to a plasticizing effect of excess glutaraldehyde molecules in the protein network. Of the three cross-linking agents used in our experiment, formaldehyde was the most effective in increasing puncture strength of films from glandless delipidated flour. These results are illustrated in Figure 5.9 showing film puncture strength versus modified reactive lysine percentage. There clearly was a positive correlation between modified reactive lysine content in films and film puncture strength, regardless of the type of cross-linking agent used.

Figure 5.9 Effect of percentage of modified reactive lysine in delipidated glandless cottonseed flour films cross-linked by formaldehyde, glyoxal, or glutaraldehyde. Reprinted with permission from Marquié et al., 1997b. Copyright 1997, American Chemical Society.

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It was possible to incorporate carded cotton fibers (mean length 20 mm) into cottonseed flour films and plastic-like materials (Marquié, 1996; Marquié et al., 1996). The resulting films resembled bilayer films in combining the properties of each polymer component (cellulose and protein). Plastic-like materials with proteins that were cross-linked by glutaraldehyde and contained 8% carded cotton fibers (w/w, dry basis) were fivefold more water resistant with solubility values ranging from 20 to 30% (w/w, dry basis), depending on the cottonseed flour used (Marquié et al., 1996). Therefore, cottonseed flour films may be strengthened with cotton fibers derived from spinning mill carding waste.

Films from cottonseed flour are highly hydrophilic due to their protein content. The water vapor adsorption isotherms of films from glandless flour that were cross-linked by formaldehyde or glutaraldehyde were represented by exponential curves for water activities up to 0.6 (Figure 5.10). These curve patterns were similar to those obtained for other protein-based films (Gontard et al., 1993; Gennadios and Weller, 1994).

Protein-based film mechanical properties are affected by both temperature and relative humidity (Marquié, 1996). Water acted as a plasticizer in cottonseed flour films, i.e., increasing molecular mobility in the protein network and decreasing the temperature at which film mechanical properties changed (apparent glass transition temperature) (Figure 5.11). Protein cross-linking hindered complete collapse of the protein structure when the films became highly hydrated (20% w/w moisture content). With added moisture, the films remained cohesive and dilated without dissolving. Protein cross-linking increased the loss and storage moduli (Figure 5.12). With a molar excess (more than threefold) of formaldehyde relative to reactive amino acids, the loss and storage moduli tended to level off, indicating a near optimal effect of cross-linking on the solid characteristics of the films (Marquié, 1996). Puncture strength of films from glandless and glanded cottonseed flours varied as a function of film moisture content or water activity (Figure 5.13). At low water activity (0.1), films with low moisture content (2–3% w/w, dry basis) were very brittle. Then, puncture strength increased sharply for film moisture contents within 3–7% (w/w, dry basis). The increase in film puncture strength

Figure 5.10 Moisture adsorption isotherm (20 C) of films made from glandless cottonseed flour and cross-linked by formaldehyde or glutaraldehyde. (Adapted from Marquié et al., 1997a.)

Figure 5.11 Effect of moisture content on apparent glass transition temperature (Tg), determined with dynamic mechanical thermal analysis, of glanded cottonseed flour films. (Adapted from Marquié, 1996.)

152

Figure 5.12 Effect of formaldehyde/(lysine + tyrosine) molar ratio on storage and loss moduli (determined with dynamic mechanical thermal analysis) of glanded cottonseed flour films. (Adapted from Marquié, 1996.)

Figure 5.13 Effect of moisture content on puncture strength of films from glandless, glanded, or delipidated glandless cottonseed flours. (Adapted from Marquié, 1996.)

153

154 at low water content may be interpreted as an antiplasticizing effect (Guo, 1994a, b). At film moisture contents up to 7% (w/w, dry basis), water plasticized the film structure through extensive hydrogen bonding among protein chains, thus increasing molecular mobility with a concomitant decrease in puncture strength. For low film moisture contents (% w/w, dry basis), the mechanical behavior of films from glanded flour differed from those made with glandless flour. The behavior of films from glanded flour at low moisture content was perhaps due to the fact that no antiplasticizing effect of water could occur because the protein network was cross-linked. DMTA of films demonstrated that cottonseed flours could be thermoformed at near ambient temperature at moisture contents greater than 30% (w/w, dry basis) (Figure 5.11). Therefore, molding cottonseed flours into films using industrial techniques common to manufacturing of synthetic packaging appears feasible. Tensile strength of films from cottonseed flour compared with those of selected synthetic and other biodegradable films is shown in Table 5.5. Cottonseed flour films whose proteins were cross-linked by formaldehyde were fiveto tenfold weaker in terms of tensile strength than the other films. However, cottonseed flour films that contained cotton fibers were as strong as Novon (starch-based) and Biopol (copolymer of polyhydroxybutyrate and polyhydroxyvalerate) films and were slightly weaker than high-density polyethylene films (Table 5.5).

Measured water vapor permeability values of cottonseed flour (glanded, glandless, and glandless delipidated) films were about 0.52 g mm/m2 h kPa at 20 C and 0/100% RH gradient (Marquié, 1996). These values were similar to

Potential Uses

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those reported in the literature for other protein-based films, but 100-fold lower than that of polyethylene.

A modified version of Sturm’s test was used to determine the biodegradability of cottonseed flour films in aerobic liquid medium (OCDE, 1981; Marquié, 1996). In this test, the assessed materials were used as a substrate (the only carbon source) by microorganisms in aerobic medium. The inoculum was obtained from activated sludge samples. Sample degradation was monitored indirectly by measuring the quantity of carbon dioxide produced by microorganisms during the test. The percentage of film biodegradation was determined in terms of the percentage of the initial theoretical carbon content in films converted to carbon dioxide. Results (Table 5.6) indicated that films from cottonseed flours were biodegradable under these conditions. Protein cross-linking, which only involves a few amino acids and does not affect peptide bonds, therefore does not hinder biodegradation. However, as it was not possible to analyze the quantity of carbon incorporated in the biomass or mineralized into carbonate by this test, we were unable to determine whether complete film biodegradation had occurred.

Cottonseed flour films may be suitable for certain applications in nonfood packaging (e.g., compostable waste bags) where good mechanical resistance and insolubility in water are required. They can also be used for agricultural packaging and mulching films to protect crops and fixate seeds, applications

156 where film color, porosity, and biodegradability are beneficial. Mulching films that progressively biodegrade after they have served their initial purpose can also function as natural fertilizers. Textural and adhesive properties of films from glanded cottonseed flours prompted us to assess their potential use in medical areas where biodegradability is necessary (e.g., prostheses and resorbable dressings). Preliminary results of in vivo studies in rats revealed good adherence and interesting healing properties of these materials as cutaneous dressings, without any allergenic effects. The ability of these films to absorb considerable amounts of moisture is a further advantage for exudate absorption (Marquié, 1995). Regardless of intended use, it should be possible during film preparation to introduce active compounds (e.g., insecticides, fungicides, and bactericides) that could migrate out of the films and offer a specific environmental action. Film hydration, which is accompanied by increased water vapor permeability and diffusion of small molecules, would make these materials suitable vectors for active substances in delicate environments.

Delipidated or non-delipidated cottonseed flours are interesting proteinaceous raw materials for formation of biodegradable films. Materials with satisfactory mechanical properties can be produced by cross-linking the cottonseed proteins and/or incorporating cotton fibers into the film. Gossypol, which is naturally present in glanded flours, has a specific effect during film formation, i.e., it promotes protein cross-linking as soon as the flour is dispersed in the film-forming solution. Their biodegradability and mechanical properties indicate that such films have potential applications in agricultural and biomedical fields.

Bérot, S. and Guéguen, J. 1985. “Fabrication de concentrats et d’isolats protéiques de coton glandless” in Proceedings of the Colloque IDESSA-CIDT-TRITURAF: Le Cotonnier sans Gossypol: Une Nouvelle Ressource Alimentaire, November 26–27, Abidjan, Côte d’Ivoire, pp. 186–210. Besançon, P., Henri, O. and Rouanet, J.M. 1985. “Valeur Nutritionnelle Comparée de Farines Délipidées de Coton Glandless et de Soja” in Proceedings of the Colloque IDESSA-CIDT-TRITURAF: Le Cotonnier sans Gossypol: Une Nouvelle Ressource Alimentaire, November 26-27, Abidjan, Côte d’Ivoire, pp. 63–79. Bizzini, B. and Raynaud, M. 1974. “Detoxification of Proteic Toxins by Formaldehyde: Possible Mechanisms and New Developments,” Biochim., 56:297–303. Bourély, J. 1990. “Développement des Cotonniers sans Gossypol comme Plante Vivrière: Aspects Technologiques et Nutritionnels,” Sci. des Aliments, 10:485–514. Cho, K.C., Park, P.S.W., Adams, E.T. and Rhee, K.C. 1992. “A Simple Rapid and Simultaneous Preparation of Glandless Cottonseed 7S and 11S Protein Fractions and Characterization of Some Physicochemical Properties,” J. Food Qual., 15:357–367.

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De Rham, O. 1982. “La Proportion d’Azote dans les Protéines et le Facteur de Calcul Protéine/Azote,” Lebensm.-Wis. u-Technol., 15:226–231. Dumay, E., Condet, F. and Cheftel, J.C. 1986. “Préparation Protéique de Graines de Coton Glandless: Caractérisation des Constituants Protéiques et Propriétés Fonctionnelles,” Sci. des Aliments, 6:623–656. Fraenkel-Conrat, H. and Olcott, H.S. 1946. “Reaction of Formaldehyde with Proteins. II. Participatin of the Guanidyl Groups and Evidence of Crosslinking,” JAOCS, 68:34–37. Fraenkel-Conrat, H., Brandon, B.A. and Olcott, H.S. 1947. “The Reaction of Formaldehyde with Proteins. IV. Participation of Indole Groups Gramicidin,” J. Biol. Chem., 168:99–118. Fritz, H.G., Seidenstücker, T. and Boölz, J.M. 1994. “Production of Thermo-Bioplastics and Fibres Based Mainly on Biological Materials,” Directorate-General XII, Science, Research and Development, EUR 16102 EN, p. 392. Gennadios, A. and Weller, C.L. 1992. “Tensile Strength Increase of Wheat Gluten Films,” ASAE Paper No. 92-6517, St. Joseph, MI: American Society of Agricultural Engineers, pp. 10. Gennadios, A. and Weller, C.L. 1994. “Moisture Adsorption by Grain Protein Films,” Trans. ASAE, 37:535–539. Gontard, N. and Guilbert, S. 1994. “Bio-Packaging: Technology and Properties of Edible and/or Biodegradable Material of Agricultural Origin” in Food Packaging and Preservation, ed., M. Mathlouthi, Glasgow, UK: Blackie Academic & Professional, pp. 159–181. Gontard, N., Guilbert, S. and Cuq, J.L. 1993. “Water and Glycerol as Plasticizers Affect Mechanical and Water Vapor Barrier Properties of an Edible Wheat Gluten Film,” J. Food Sci., 58:206–211. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation, ed., M. Mathlouthi, New York: Elsevier Applied Science Publishers, pp. 371–394. Guo, J. 1994a. “An Investigation into the Formation of Plasticizer Channels in Plasticized Polymer Films,” Drug Develop. Industr. Pharm., 20:1883–1893. Guo, J. 1994b. “A Theoretical and Experimental Study of Additive Effects of Physical Aging and Antiplasticization on the Water Permeability of Polymer Film Coating,” J. Pharm. Sci., 83:447–449. Korn, A.H., Feairheller, S.H. and Filachione, E.M. 1972. “Glutaraldehyde, Nature of the Reagent,” J. Mol. Biol., 65:525–529. Lawhon, J.T., Cater, C.M. and Mattil, K.F. 1977. “Evaluation of the Food Use Potential of Sixteen Varieties of Cottonseed,” JAOCS, 54:75–80. Lyman, C.M., Baliga, B.P. and Slay, M.W. 1959. Reaction of Proteins with Gossypol,” Arch. Biochem. Biophys., 84:486–497. Marquié, C. 1987. “Utilisation Alimentaire des Dérivés des Cotonniers Glandless,” Cot. Fib. Trop., 42:65–73. Marquié, C. 1995. Procédé de Fabrication de Films Biodégradables et Films Obtenus Selon ce Procédé. Breve déposé au nom du Cirad, No. 95 02640 du 07 Mars. Marquié, C. 1996. Mise au Point et Étude de Films Biodégradables Réalisés avec des Farines de Graines de Cotonnier. Thèse de Doctorat, Université Montpellier II, Sciences et Techniques du Languedoc, Montpellier, France, p. 193. Marquié, C., Aymard, C., Cuq, J.L. and Guilbert, S. 1995. “Biodegradable Packaging Made from Cottonseed Flour: Formation and Improvement by Chemical Treatments with Gossypol, Formaldehyde, and Glutaraldehyde,” J. Agric. Food Chem., 43:2762–2767. Marquié, C., Héquet, E., Guilbert, S., Tessier, A.M. and Vialettes, V. 1996. “Biodegradable Material Made from Cottonseed Flour” in Proceedings of the 23rd International Cotton Conference, Bremen, Germany, March 6–9, eds., H. Harig and S.A. Heap, Bremen, Germany: Faserinstitut, pp. 145–156. Marquié, C., Héquet, E., Vialettes, V. and Tessier, A.M. 1997a. “Cottonseed Processing into Biodegradable Material for Potential Agricultural and Biomedical Uses,” Beltwide Cotton Conferences, Cotton Improvement Conference, New Orleans, LA, USA, pp. 469–473.

158 Marquié, C., Tessier, A.M., Aymard, C. and Guilbert, S. 1997b. “HPLC Determination of the Reactive Lysine Content of Cottonseed Protein Films to Monitor the Extent of Cross-Linking by Formaldehyde, Glutaraldehyde, and Glyoxal,” J. Agric. Food Chem., 45:922–926. Marquié, C., Tessier, A.M., Aymard, C. and Guilbert, S. 1998. “How to Monitor the Protein Cross-Linking by Formaldehyde, Glutaraldehyde or Glyoxal in Cottonseed Protein-based Films,” Nahrung, 42:264–265. Martinez, W.H. 1979. “Functionality of Vegetable Proteins Other Than Soy,” JAOCS, 56:280–284. Martinez, W.H., Berardi, L.C. and Goldblatt, L.A. 1970. “Cottonseed Protein Products,” J. Agric. Food Chem., 18:961–968. Means, G.E. and Feeney, R.E. 1968. “Reductive Alkylation of Amino Groups in Proteins,” Biochem., 7:2192–2201. Monsan, P., Puzo, G. and Mazarguil, H. 1975. “Etude du Mécanisme d’Établissement des Liaisons Glutaraldéhyde-Protéines,” Biochim., 57:1281–1292. OCDE 301 B. 1981. “Biodégradabilité dite Facile. Essai de Sturm Modifié,” Ligne Directrice pour les Essais de Produits Chimiques, 12 Mai. Reddy, J.M. and Rao, N.M. 1988. “Method for the Isolation of Gossypin (11S Protein) and Congossypin (7S Protein) of Glanded Cottonseed,” J. Agric. Food Chem., 36:241–245. Rossi-Fanelli, A., Antonini, E., Brunori, M., Bruzzeri, M.R., Caputo, A. and Satriani, F. 1964. “Isolation of a Monodisperse Protein Fraction from Cottonseed,” Biochem. Biophys. Res. Commun., 15:110–115. Saroso, B. 1989. “Chemical Properties of Protein in Cottonseed Kernels,” Industr. Crop Res. J., 1:60–65. Shan, S.H. 1991. “Reactive Groups of Proteins and Their Modifying Agents” in Chemistry of Protein Conjugation and Cross-Linking, ed. S.S. Wong, Boca Raton, FL: CRC Press, pp. 7–45. Tacher, G., Rivière, R. and Landry, C. 1985. “Utilisation du Tourteau de Coton sans Gossypol dans l’Alimentation des Volailles de Type Chair” in Proceedings of the Colloque IDESSA-CIDT-TRITURAF: Le Cotonnier sans Gossypol: Une Nouvelle Ressource Alimentaire, November 26–27, Abidjan, Côte d’Ivoire, pp. 97–112. Tchiegang, C. and Bourély, J. 1990. “Contribution à l’Étude de la Combinaison du Gossypol avec les Autres Constituants des Amandes des Graines de Cotonniers au Cours de Certains Traitements Technologiques,” Cot. Fib. Trop., 45:105–117. Wu, L.C. and Bates, R.P. 1972a. “Soy Protein-Lipid Films. 1. Studies on the Film Formation Phenomenon,” J. Food Sci., 37:36–39. Wu, L.C. and Bates, R.P. 1972b. “Soy Protein-Lipid Films. 2. Optimization of Film Formation,” J. Food Sci., 37:40–44. Wu, L.C. and Bates, R.P. 1973. “Influence of Ingredients upon Edible Protein-Lipid Film Characteristics,” J. Food Sci., 38:783–787. Youle, R.J. and Huang, A.H.C. 1979. “Albumin Storage Protein and Allergens in Cottonseeds,” J. Agric. Food Chem., 87:500–503. Zarins, Z.M. and Cherry, J.P. 1981. “Storage Proteins of Glandless Cottonseed Flour,” J. Food Sci., 46:1855–1862. Zarins, Z.M., Phillips, R.D. and Martinez, W.H. 1984. “A Rapid Procedure for Isolation of the 7S Storage Protein of Cottonseed and Its Characterization,” Cereal Chem., 61:471–474. Zongo, D. and Coulibaly, M. 1993. “Le Tourteau de Coton sans Gossypol: Une Importante Source de Protéines pour l’Élevage Porcin,” Tropicultura, 11:95–98.

Formation and Properties of Whey Protein Films and Coatings

DIBLE biopolymer films have been a subject of intense interest in recent years because of consumer demands for food quality, convenience, and safety, as well as the necessity of improving recyclability and biodegradability of packaging. Protective, edible coatings can simplify packaging systems, thus improving recyclability of conventional packaging. Edible films and coatings prevent quality changes in foods by acting as barriers to moisture, oxygen, oil, and aroma migration between adjacent food components and/or between the food and the environment (Guilbert, 1986; Kester and Fennema, 1986; Krochta, 1992). The overall packaging requirement could also be reduced whenever an edible film can act as an efficient barrier to mass transfer and/or enhance mechanical integrity of the food (Krochta, 1997a). In addition, edible films and coatings can carry food additives, such as antimicrobials, flavors, or antioxidants. Materials that have been used in edible films include proteins, polysaccharides, and lipids. In particular, whey proteins have been intensively investigated in the past decade. With the addition of plasticizer, heat-denatured whey proteins produce transparent, bland, and flexible water-based edible films with excellent oxygen, aroma, and oil barrier properties at low relative humidity (RH) (McHugh and Krochta, 1994a; Miller and Krochta, 1997a; De Mulder-Johnston, 1999). When film formation includes heat-induced protein

E

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160 denaturation, these films are also water insoluble and have enhanced mechanical properties (Pérez-Gago et al., 1999), which can be beneficial in maintaining food integrity when necessary. However, the hydrophilic nature of whey protein films causes them to be less effective moisture barriers. The moisture barrier ability of these films is greatly improved by incorporation of lipids (McHugh and Krochta, 1994b; Shellhammer and Krochta, 1997a; Pérez-Gago and Krochta, 1999). This chapter reviews existing literature on whey protein-based films and coatings. The specific objectives are to summarize information on (1) whey protein nature and recovery, (2) whey protein-based film formation and properties, (3) relationship of whey protein-based film properties to molecular structure and environmental factors, and (4) applications and future directions for these films.

Whey proteins are those proteins that remain in milk serum after pH/rennet coagulation of casein during cheese or casein manufacture (de Wit, 1989; Morr and Ha, 1993). Whey protein, which represents approximately 20% of total milk proteins (Brunner, 1977) is a mixture of proteins with diverse functional properties. The five main proteins are -lactalbumins, -lactoglobulins, bovine serum albumin, immunoglobulins, and proteose-peptones. Table 6.1 shows the approximate composition of proteins in whey, component molecular weight, and the number of disulfide (S-S) bonds and sulfhydryl (SH) groups per molecule.

-Lactoglobulin ( -Lg) comprises approximately 57% of the protein in whey (Dybing and Smith, 1991). Its primary structure consists of 162 amino ac-

Nature of Major Whey Proteins

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Figure 6.1 Primary structure of (A) -lactoglobulin B (* variant A) and (B) -lactalbumin B ( variant A). Reprinted with permission from Cayot and Lorient, 1997.

ids with a molecular weight of 18,300 Da. Among seven genetic variants, the A and B variants are the most abundant, differing from each other by two amino acid residues. The primary structure of -Lg corresponding to variant B is shown in Figure 6.1. The secondary and tertiary structures show high degrees of organization, with 43–50% of residues in -sheets, 10–15% in -helices, and 15–20% in -turns (Papiz et al., 1986). Monomeric -Lg contains one free SH group, which is normally buried internally in the native molecule, and two S-S bonds. S-S bonds are formed between Cys 160 and 66, and Cys 119 and 106, and the free thiol is in the Cys-121 position (Morr and Ha, 1993). Crystallization analysis has shown that -Lg exists in a globular form, with stabilizing hydrophobic and SH groups located in the interior. The thiol group is important because it appears to facilitate molecular thiol-disulfide interchange reactions that allow formation of intermolecular disulfide-bonded dimers and polymers

162 upon heating (Kinsella, 1984). The functional properties of these groups can be exploited to form effective edible films. The conformation and association of -Lg are pH and temperature sensitive. In the pH range of 5.1–7.5, the molecule exists as a dimer of approximately 36,700 Da. At acidic conditions, especially below pH 3, the dimers dissociate into monomers due to strong electrostatic repulsive forces; but dissociation is not accompanied by changes in molecular shape or conformation of the monomers. Dissociation also occurs in the alkaline region above pH 8. The molecule undergoes reversible conformational changes, exposing carboxyl and thiol groups, which become more reactive (Fox, 1989). In the pH range of 3.5–5.2, dimers tetramerize to form octamers of approximately 147,000 Da (Morr and Ha, 1993). -Lg undergoes time- and temperature-dependent denaturation reactions at temperatures above 65 C, which result in a general molecular expansion, exposure of the internal SH group, and hydrophobic and -NH2 groups (Brunner, 1977; Kinsella, 1984).

-Lactalbumin ( -La) is the second most abundant whey protein, accounting for about 19% of the total whey protein (Dybing and Smith, 1991). It is a globular protein that contains 123 amino acid residues with a molecular weight of 14,000 Da and four S-S bonds (Table 6.1 and Figure 6.1). This molecule has a low content of organized secondary structure, i.e., 30% -helix, 9% -sheet, and 61% unordered structure (Alexandrescu et al., 1993), and, as a result, it has great flexibility. However, bound calcium and the four S-S bridges maintain the globular structure of -La and stabilize it against denaturation (Kinsella and Whitehead, 1989). Conformational changes occur in -La at pH 4, where the molecule loses the Ca+2 that is tightly bound at higher pH. At pH values between 4 and 5, -La appears to occur as two forms with different thermal stabilities. At pH 6.5, -La begins unfolding at 62 C; but on cooling, the molecule reverts to its native configuration. This reversibility is lost if the native S-S bonds are broken, for example, by heat-induced thiol-disulfide interchange reactions between -La and -Lg (de Wit, 1989).

Bovine serum albumin (BSA) is a large globular protein with 582 amino acid residues and a molecular weight of 66,000 Da. It represents approximately 7% of the proteins in whey (Dybing and Smith, 1991). BSA contains 17 S-S bonds and one free thiol group, which make it highly structured (Morr and Ha, 1993). BSA binds different ligands such as free fatty acids, other lipids, and fla-

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vors, that stabilize the molecule against thermal denaturation (Kinsella and Whitehead, 1989).

The immunoglobulins (Igs) and proteose-peptone (PP) fractions represent the remainder of the whey protein. The Igs are a heterogeneous family of glycoproteins of 150–1000 kDa with antibody activity. Up to 80% of the Ig in whey is Ig-G. These molecules are composed of four polypeptide chains joined by S-S bridges and denature at a higher temperature than -Lg or -La (de Wit, 1989). However, they are thermally unstable in the presence of other whey proteins, which may be related to the activity of thiol groups of -Lg and BSA (Morr and Ha, 1993). The PP fraction consists of proteolytic fragments of -casein-containing surface-active phospho-(glyco)peptides (Andrews, 1979).

Production of dry whey in 1999 was estimated at 1.6 million metric tons worldwide (USDA-NASS, 1999), containing approximately 200,000 metric tons of whey protein. U.S. production of dry whey is approximately one-third of word production. Increased production of whey because of expanding cheese manufacture has made identifying new uses for whey proteins a goal of the dairy industry. Improvements in membrane and ion-exchange technology have made possible the manufacture of quality whey proteins with useful functional properties, such as acid stability, gelation, film formation, aeration, and emulsification (Huffman, 1996; de Wit and Klarenbeek, 1984). The industrial processes used for whey protein recovery are ultrafiltration, reverse osmosis, gel filtration, electrodialysis, and ion exchange chromatography (de Boer et al., 1977; Matthews, 1984; Glover, 1985). Fractionated whey constituents of various degrees of concentration can be obtained by combining two or more of the above recovery processes (Sienkiewicz and Riedel, 1990). Whey protein products can be classified according to their composition. In particular, they are divided according to their protein content. Whey protein concentrate (WPC) contains 25–80% protein. Whey protein isolate (WPI) is nearly all protein (>90%). The wide range of protein concentration in these products depends on the membrane used and the pretreatment prior to membrane separation. Generally, ultrafiltration and diafiltration processes, using membranes with different molecular weight cut-off values, are used to separate lactose, minerals, and other small molecular weight components from high molecular weight molecules, producing undenatured WPC (Sienkiewicz and Riedel, 1990). The manufacturing process of undenatured WPI uses ion ex-

164 change chromatography after the whey is initially concentrated by ultrafiltration to reduce the volume of liquid processed (Dybing and Smith, 1991). This technique enables further reduction in mineral content. In general, the broad range in composition limits the final application, because some functional properties are modified.

Protein-protein interactions determine the characteristics of protein-based films. Film-forming ability may be influenced by amino acid composition, distribution, and polarity; conditions affecting formation of ionic cross-links between amino and carboxyl groups; presence of hydrogen bonding groups; and intramolecular and intermolecular S-S bonds (Gennadios and Weller, 1991). Native whey proteins are globular proteins containing most of the hydrophobic and SH groups hidden in the interior of the molecule. Formation of whey protein films has mainly involved heat denaturation of whey proteins in aqueous solutions. Heating modifies the three-dimensional structure of the protein, exposing internal SH and hydrophobic groups (Shimada and Cheftel, 1998), which promote intermolecular S-S bonding and hydrophobic interactions upon drying (McHugh and Krochta, 1994b). McHugh et al. (1994) produced plasticized whey protein isolate films by heat-treating 8–12% (w/w) solutions of whey proteins at temperatures between 75 C and 100 C. WPI solution concentrations greater than 12% (w/w) gelled upon heating. Optimal conditions for preparing these films included the heating of 10% (w/w) WPI solutions at 90 C for 30 min. These conditions formed films with a consistent structure, and X-ray diffraction indicated that whey proteins were irreversibly denatured (Lent et al., 1998). Lower WPI solution concentrations (down to 5% WPI (w/w)) have also been shown to form good whey protein-based films, whenever the amount of solids deposited per casting surface unit was kept constant (Pérez-Gago et al., 1999). Pérez-Gago et al. (1999) also observed that native whey protein (i.e., whey protein that has not undergone heat treatment) had good film-forming ability. Because native whey proteins maintain their globular structure with most of the hydrophobic and SH groups buried in the interior of the molecule, native protein films have a more random structure in which cohesion relies mainly on hydrogen bonding. In contrast, the intermolecular forces that promote cohesion in heat-denatured films also involve intermolecular S-S bonds and hydrophobic interactions among the unfolded protein strands. Both native and heat-denatured whey protein films are transparent, having similar water vapor permeabilities when made from protein solutions at their natural pH; however, they possess different solubility and mechanical properties. The unfolded structure of heat-denatured whey proteins and the covalent S-S bonding during drying lead to film insolubility in water and produce films that are stronger and that can withstand higher deformations.

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The low energy bondings and the globular structure of native whey proteins account for complete solubility in water and poor mechanical properties of native whey protein films compared to heat-denatured whey protein films (Pérez-Gago et al., 1999). Banerjee and Chen (1995) tested various denaturing temperature and pH conditions to optimize film-forming conditions for WPCs. They reported that heating WPC solutions at 75 C for 30 minutes after pH adjustment to 6.6 using 2 M NaOH produced solutions that ensured uniform film production. Transglutaminase has also been used to polymerize whey proteins in the production of films (Mahmoud and Savello, 1992, 1993; Yildirim and Hettiarachchy, 1998). Transglutaminase is a Ca+2-dependent enzyme that catalyzes the introduction of -( -glutamyl)-lysine cross-links into proteins via an acyl transfer reaction. This reaction introduces covalent cross-linkages between whey proteins, as well as between peptides and primary amines (Mahmoud and Savello, 1993). Plasticizer addition is necessary to reduce brittleness and increase flexibility of whey protein films. Plasticizers achieve these changes by interacting with polymer chains, thus reducing intermolecular forces among them. Plasticizers act mainly by disrupting hydrogen bonding between neighboring polymer chains, thus increasing chain mobility (Guilbert, 1986; Kester and Fennema, 1986). Glycerol (Gly), sorbitol (Sor), and polyethylene glycol (PEG) have been commonly used to plasticize whey protein films. The increase in film flexibility is also accompanied by an increase in film permeability, which depends on the type of plasticizer (McHugh et al., 1994). Plasticized whey protein films are transparent, bland, and flexible and have excellent oxygen, aroma, and oil barrier characteristics. However, these films are poor moisture barriers due to their hydrophilic character. Incorporation of lipid materials improves film moisture barrier properties by increasing hydrophobicity. The lipid materials are homogenized at temperatures above their melting points into the aqueous film-forming solutions of heat-denatured whey protein and plasticizer. The barrier efficiency of resulting composite films strongly depends on polarity of film components and distribution of lipid material in the film matrix (Kamper and Fennema, 1985; Debeaufort et al., 1993). Lipids that have been added to whey protein films include acetylated monoglycerides, waxes, fatty alcohols, and fatty acids. All protein-plasticizer solutions, either alone or emulsified with a lipid, are degassed until no air bubbles are observed, to ensure accurate permeability values.

Barrier properties commonly studied to determine the ability of edible films to protect foods from the environment and from adjacent ingredients are film water vapor permeability (WVP) and oxygen permeability (OP). Aroma and oil

166 permeability are equally important for many foods but have received less attention. On the other hand, Young’s Modulus (YM), tensile strength (TS), and elongation (E) are the most commonly investigated mechanical properties for assessing the ability of edible films and coatings to protect foods against mechanical abuse (Krochta, 1997a). Both barrier and mechanical properties are affected by film composition and by test conditions, specifically temperature and RH. These effects make comparisons among films difficult; nonetheless, such comparisons are important and are made when possible.

WVP is the most commonly studied transport property of edible polymer


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films. Table 6.2 compares WVP of whey protein films and synthetic polymer films. In general, the hydrophilic nature of proteins limits their ability to form films with good moisture barrier ability compared to synthetic polymer films such as low-density polyethylene (LDPE), polyvinylidene chloride (PVDC), and polyvinyl chloride (PVC). For the same reason, RH and plasticizer type and amount significantly affect the moisture barrier ability of protein films. Plasticized whey protein films have WVP nearly one order of magnitude greater than cellophane and nearly four orders of magnitude greater than LDPE under comparable test conditions and with appropriate level use of plasticizer. Even at low RH and low amount of plasticizer, WVP of WPI films is three orders of magnitude greater than that of LDPE. Increasing film hydrophobicity can decrease WVP of protein films. This can be accomplished by laminating the protein film with a lipid layer. However, these bilayer composite films tend to delaminate because of the high surface energy existing between the two components (Shellhammer and Krochta, 1997a). Alternatively, lipids can be homogenized into the protein-plasticizer film-forming solutions to form “emulsion composite films.” The emulsifying ability of whey protein makes this procedure particularly useful for obtaining films with lower WVP and good mechanical properties. McHugh and Krochta (1994b) investigated the effect of incorporating selected lipids into WPI film-forming solutions. Various acetylated monoglycerides, waxes, fatty alcohols, and fatty acids were examined for their effect on film WVP (Table 6.3). Similar to WPI films, an exponential effect of RH on WVP was observed when beeswax was present in the films. However, the effect of RH on WVP was reduced through addition of lipids to emulsion films. This is explained by the increased hydrophobicity imparted by the lipid. WVP measurements revealed some emulsion separation during drying, because different WVP values were obtained depending on whether the film upper (lipid-enriched) side was oriented toward the high or the low RH environment. Lipid type and content affected the final moisture barrier ability of films. Increasing lipid content of films decreased WVP, but the rate at which it decreased depended on the lipid type. As the chain length of both fatty acids and fatty alcohols increased, significantly lower WVPs were obtained. Beeswax and fatty acids were more effective at reducing WVP of WPI-based emulsion films than fatty acid alcohols. These results are consistent with the degree of lipid non-polarity. Sherwin et al. (1998) reported that increasing fatty acid chain length increased the particle size in the emulsion films. Another study indicated that, given a constant volume fraction of beeswax, decreasing the beeswax particle size in an emulsion film significantly correlated with a linear decrease in film WVP (McHugh and Krochta, 1994c). This was attributed to an increase in protein chain immobilization due to increased beeswax-protein interface area as the beeswax particle size decreased. Shellhammer and Krochta (1997a) studied how lipid type and amount affected the WVPs of dispersed-lipid films prepared with WPI and glycerol. Very

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low plasticizer level was used (WPI:Gly ratio of 15:1) to minimize the effect that the plasticizer would have on the final result. Emulsion films were prepared with candelilla wax (CanW), carnauba wax (CarW), beeswax (BW), and a hard anhydrous milkfat fraction (HAMFF). Film WVP decreased with increasing lipid concentration. However, it did not increase in a linear fashion with the increase in lipid content, but dropped suddenly at 40–50% content. Further increase in lipid content did not affect WVP. Films could be made with up to 70–80% BW and HAMFF, but with greater than 40% CanW or CarW, films did not form. Films formed with BW or HAMFF had up to 90% lower WVPs than films with no lipid content. CanW and CarW, the materials with the lower WVPs, gave emulsion films with the highest WVPs. In contrast, BW and HAMFF, the materials with the highest WVPs, gave emulsion films with the lower WVPs (Table 6.3). Interestingly, these results correspond with the viscoelastic properties of the lipids and suggest that BW and HAMFF may have yielded more easily to the internal forces related to shrinkage of the drying protein structure. Furthermore, the large drop in film WVP at 40–50% BW or HAMFF content could be explained by formation of an interconnecting lipid network internal to the film. Such a network could be made possible by deformation of BW and HAMFF due to film internal stress as water evaporates from the film-forming solution (Krochta, 1997b). McHugh and Krochta (1994c) showed that lipid particle size affected emulsion film WVP. It is unclear whether this was due to interactions at the pro-


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tein-lipid interface or whether it is a function of the disperse phase, such as localized concentration gradients or lipid particle shape (Shellhammer, 1996). When a lipid is laminating a hydrophilic film to form a bilayer, the barrier ability against water vapor transfer is higher than when a stable emulsion film is formed (Kamper and Fennema, 1984; Kester and Fennema, 1989). Therefore, lipid distribution in the matrix is very important. Lamination requires more steps than forming a composite film in which both hydrophilic and hydrophobic components are dispersed in the solvent. Furthermore, film delamination due to the high surface energy between polar and nonpolar materials can be a problem (Kamper and Fennema, 1984). However, because of the emulsifying ability of proteins, it is difficult to obtain complete bilayer films from protein-lipid emulsions. Pérez-Gago and Krochta (1999) examined the role of emulsion stability, as affected by pH, on the final morphology and WVP of WPI-lipid emulsion films. The films were cast from aqueous solutions of heat-denatured WPI (5% w/w), BW (lipid phase), and glycerol (plasticizer). In general, there was some phase separation and no significant difference among WVPs of emulsion films prepared at pH 6, 7, or 8. Rather than enhanced phase separation due to lipid particle coalescence at the pI (pH 4–5), phase separation was inhibited, and the film WVP was significantly higher. This occurred because emulsion viscosity increased at the pI and a weak gel was formed due to protein-protein aggregation. This lowered lipid mobility, thus prohibiting any phase separation and resulting in higher film WVP.

OP is the next most commonly studied transport property of edible polymer films after WVP. Table 6.4 compares OP values of biopolymer films and conventional synthetic polymer films. Compared to non-ionic polysaccharide films, protein films have lower OPs (Miller and Krochta, 1997a). This may be related to their more polar nature and more linear (non-ring) structure, leading to a higher cohesive energy density and lower free volume among polymer chains (Miller and Krochta, 1997a). The OPs of whey protein and other protein films are lower than those of high-density polyethylene (HDPE) and low-density polyethylene (LDPE) and are comparable to those of ethylene vinyl alcohol films (EVOH) under similar RH conditions and proper selection of plasticizer (Table 6.4).

The aroma barrier properties of edible films have received little attention (Miller and Krochta, 1997b). Miller et al. (1998) reported that WPI films were excellent barriers to d-limonene. WPI films containing 25% glycerol (dry basis) were comparable to EVOH films as a barrier to d-limonene under similar

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temperature and RH conditions (Miller et al., 1998). Additionally, temperature and RH significantly affected (exponentially) d-limonene permeability, but not by permeant concentration in the range of 62–226 ppm (mol/mol). Permeability data for d-limonene in WPI films are compared to EVOH and vinylidene chloride copolymer (co-VDC) films in Table 6.5. Even at high RH (80%), WPI films had d-limonene permeability values lower by three orders of magnitude than the industrial standard for hydrophobic aroma barriers, co-VDC, and comparable values to EVOH, the industry standard for hydrophilic aroma barriers.

Whey protein films have been proposed as excellent oil barriers, due to their hydrophilic nature, but limited relevant information has been published. De Mulder-Johnston (1999) developed a gravimetric method to quantify oil migration through WPI films. WPI films (with a constant WPI:Gly ratio of 2:1) containing 0–60% beeswax (BW) were studied. No visible oil permeability was detected for any of the films after 19 weeks at various RHs, which indicated that WPI films are excellent lipid barriers, even when large amounts of BW are added to improve the moisture barrier ability.


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Plasticizer and RH have the most significant effects on permeability and mechanical properties of WPI films. As mentioned, plasticizers reduce internal hydrogen bonding in films, thereby increasing film flexibility while also increasing film permeability. For this reason, simultaneous comparison of film permeability and mechanical properties is necessary for complete assessment of the effect of plasticizer type and amount (Krochta, 1998). McHugh et al. (1994) studied the effect of various plasticizers, including Gly, Sor, and PEG of different chain lengths, on WVP of WPI films (Table 6.2). Sor provided the highest flexibility increase per unit increase in WVP among the studied plasticizers at comparable concentrations and RH conditions. In a later study, McHugh and Krochta (1994a) compared the OP (Table 6.4) and mechanical properties (Table 6.6) of WPI films containing different amounts of Gly and Sor. Sor was more effective than Gly as a plasticizer, as films of equal TS, YM, or E had lower OP when plasticized with Sor. These results suggested that at the same mass content levels in films, lower molecular weight plasticizers might result in films with higher permeability, flexibility, and elongation, with lower strength. This is understandable considering that the molar content of the lower molecular weight plasticizer in the film is greater at constant plasticizer mass content (Krochta, 1998). Maté and Krochta (1996a) studied the effect of Gly amount on WVP (Table 6.2) and OP (Figure 6.2) for WPI and -Lg films. Again, increased Gly levels gave higher permeability values. However, WVP and OP of WPI and -Lg films were not significantly different at each Gly content studied. Despite the different protein molecular structures in WPI films compared to -Lg films and the resulting differences in hydrogen bonds, hydrophobic interactions, and S-S

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bonds (Table 6.1), no differences between WPI and -Lg WVP or between WPI and -Lg OP were detected. Similarly, Anker et al. (1998) reported no significant differences in WVP and mechanical properties between WPI and -Lg films using multivariate analysis. Due to the hydrophilic nature of whey proteins, moisture plasticizes whey protein films. Therefore, increased permeability as film moisture content increases is expected. McHugh et al. (1994) studied the effect of RH on WVP and OP of WPI films. The results (Tables 6.2 and 6.4) suggested an exponential-type relationship between WVP and RH and between OP and RH for WPI films. Interestingly, the effect of RH on WVP was more pronounced for Gly-plasticized films compared to Sor-plasticized films, in spite of higher WPI:Gly ratio compared to WPI:Sor ratio.

Figure 6.2 Effect of glycerol content and temperature on oxygen permeability of whey protein isolate (WPI) and -lactoglobulin ( -Lg) edible films. Reprinted with permission from Maté and Krochta, 1996a. Copyright 1996, American Chemical Society.


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Permeability properties of edible films are temperature-dependent, following the Arrhenius relationship. Maté and Krochta (1996a) confirmed an exponential temperature effect on OP for WPI and -Lg films (Figure 6.2). The Arrhenius model fit for OP has also been confirmed for other protein films (Gennadios et al., 1993), as well as for synthetic films.

Fairley et al. (1996a) studied the ability of sodium dodecyl sulfate (SDS) to plasticize WPI films and concluded that the primary effect of SDS was reduction of intermolecular S-S bond formation. Overall, the results with SDS suggested that film mechanical properties were affected both by limiting intermolecular S-S bond formation allowed to occur upon denaturation and by adding plasticizers to compete for intermolecular chain-to-chain hydrogen bonding after denaturing was complete (Table 6.7) (Krochta, 1998).

174 To better understand cross-linking in WPI films, Fairley et al. (1996b) studied the effect of adding N-ethylmaleimide (NEM) and cysteine to heat-denatured WPI solutions. NEM blocks free thiol groups, thus preventing sulfhydryl/disulfide interchange. Cysteine reduces S-S bonds under certain conditions, also reducing intermolecular S-S bonds. Neither NEM nor cysteine affected WVP of resulting films, indicating that the nature and extent of cross-linking had no effect on WVP (Table 6.7). However, blocking free SH groups with NEM had an unexpected effect on mechanical properties and film solubility (Table 6.7). Mechanical properties were not modified as NEM increased, except for the E that actually decreased. Film solubility, on the other hand, decreased as NEM concentration increased. The authors hypothesized that prevention of sulfhydryl/disulfide interchange enabled the protein to adopt conformations that resulted in modified hydrogen bonding, hydrophobicity, disulfide-disulfide interchange, or sulfhydryl-sulfhydryl reaction. Nevertheless, it appears that S-S bonds, whether intra- or intermolecular, play a very small role in determining the moisture barrier properties of WPI-based films. Pérez-Gago et al. (1999) compared the WVP, film solubility, and tensile properties of native versus heat-denatured whey protein films. Native whey protein films were totally water soluble, whereas heat-denatured films were insoluble. Heat-denatured whey protein films also had greater TS, YM, and E than native whey protein films. However, native and heat-denatured films had similar WVP. The unfolded structure due to heating and the covalent S-S bonding during drying led to film insolubility in water and produced stronger, stiffer films that could also endure greater deformations. The low energy bonds and the globular structure of whey proteins in native films are accountable for their complete solubility in water and poor mechanical properties compared to heat-denatured films. These results suggested that covalent cross-linking due to heat denaturation of whey protein was accountable for film water insolubility and greater tensile properties, but did not affect film WVP. Pérez-Gago et al. (1999) also reported that the pH of the film-forming solution did not significantly affect film solubility, mechanical properties, and WVP at pH away from the pI. The increase of WVP observed at the pI was likely due to incomplete removal of air bubbles because of increased film-forming solution viscosity. Cross-linking agents have been used to improve barrier properties and mechanical strength of different materials. Cross-linking by transglutaminase in the presence of Ca+2 has been used in -Lg, -La, and WPI film formation. Mahmoud and Savello (1992) reported that water vapor transmission rate and moisture content of films were not significantly different among -Lg, -La, and WPI films. These films were insoluble in SDS and -mercaptoethanol, but when films were incubated with proteolytic enzymes, they were hydrolyzed at various degrees depending on incubation time. Hydrolysis of whey protein films by proteolytic enzymes indicated that they are digestible and that food


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surfaces carrying proteolytic enzymes would reduce the stability of whey protein coatings on those foods (Mahmoud and Savello, 1992). Yildirim and Hettiarachchy (1998) compared the properties of transglutaminase-cross-linked WPI films with control WPI films. Solubility of transglutaminase-cross-linked films was lower than that of control films at pH 3, 4, 6, or 8. These results also correlated with film tensile strength, because values were twofold greater than those of the control. However, enzymatic hydrolysis of control and cross-linked films was similar. The WVP of transglutaminase cross-linked films, on the other hand, was lower than that for control films. Mahmoud and Savello (1992) stated that cross-linking polymerization of whey protein solutions induced gelation and that films were produced by casting the gelled whey protein solutions followed by air drying. Such films prepared from gelled solutions might contain large pores that could be the cause of the higher WVP. Galietta et al. (1998) investigated the plasticizing effect of Gly and the cross-linking effects of formaldehyde and CaCl2 on mechanical and thermomechanical properties of WPC (80% protein content) films. Increased plasticizer content increased film solubility in water and decreased the force at break, YM, and glass transition temperature. Formaldehyde, as a cross-linking agent, enhanced film mechanical properties and insolubility. Addition of a low concentration of CaCl2 in the film formulation did not affect film functional properties, perhaps because the extent of ionic bonding in films after CaCl2 addition was not sufficient to affect the mechanical behavior of the films. Heat curing is commonly used to cross-link synthetic polymers and to improve mechanical and barrier properties (El-Hibri and Paul, 1985; Perkins, 1988). Miller et al. (1997) heat-cured WPI films to study the effect of increased intermolecular S-S bonding on mechanical and water vapor barrier properties. They prepared films from WPI solutions that had been heated at 90 C for 30 minutes before adding Gly (WPI:Gly ratio of 3:1). Films dried at ambient conditions were heat-cured at several temperatures and RHs for different lengths of time and were tested after conditioning at the same temperature and RH. Results showed that curing time affected TS and YM linearly, while influencing E and TS exponentially. Increased cure temperature and reduced cure RH yielded stronger, less extendible films with diminished flexibility, but having more effective water vapor barrier properties. These data suggested that heat-curing may elicit additional cross-linking of protein and/or cause protein molecules to rearrange or relax, eliminating or lessening internal stress and producing more efficient protein chain packing with smaller free volume.

Common practice in film formation has been to dry films at ambient condi-

176 tions, normally at approximately 21 C and 50% RH. However, controlling the drying behavior is important to applying edible coatings on foods. Understanding the effect of drying conditions on WVP and mechanical properties of WPI-based films was the goal of Alcantara et al. (1998). In their study, higher drying rates significantly affected film thickness, tensile properties, and WVP. More rapid drying resulted in stiffer, less flexible films, but had smaller apparent effect on film TS and E. WVP results showed that WPI:Gly films were better moisture barriers when dried at a faster drying rate. This was attributed to the change in film morphology with subsequent thinner film thickness achieved at a faster drying rate.

Films based on WPC have also been developed and compared with films based on other milk proteins (Chen, 1995; Banerjee and Chen, 1995; Banerjee et al., 1996). In WPC films, S-S cross-linking induced by thermal denaturation (75 C for 30 min after the solution pH was adjusted to 6.6) was an essential step in film formation. Banerjee and Chen (1995) reported that the WVP of WPC (protein content 76.6%) film was 14% lower than that of WPI (protein content 93.6%) film. Transmission electron microscopy revealed the presence of residual milk fat embedded in the protein matrix in WPC films (6.8% fat) that might have increased film hydrophobicity and improved the moisture barrier properties. In later studies, microfluidization and ultrasound technologies were applied to improve mechanical strength and water vapor barrier ability of WPC films (Banerjee et al., 1996). These technologies reduced the particle size of milk fat present in WPC. Film E, WVP, and moisture content were not affected by ultrasonic treatment. However, resistance to puncture increased as the acoustic power increased.

While research continues to document and improve whey protein film barrier and mechanical properties, applications research dealing with forming films as coatings on various food surfaces has begun. The main focus of such research is to fully understand the surface chemistry of the food-coating interface (Trezza, 1999). Whey protein coatings significantly reduced oxygen uptake and rancidity of roasted peanuts at low RH (Maté and Krochta, 1996b; Maté et al., 1996). Coating fragile foods, such as freeze-dried chicken, with whey protein improved food integrity and reduced loss due to damage (Alcantara and Krochta, 1996). In frozen king salmon, lipid oxidation was delayed, and peak peroxide values were reduced in samples coated with WPI solution followed by an antioxidant overspray (Stuchell and Krochta, 1995). However, whey protein coatings did not affect respiration and moisture loss of

References

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green bell peppers stored at high RH, which is consistent with the fact that RH has a substantial effect on increasing permeability of hydrophilic film materials (Lerdthanangkul and Krochta, 1996).

Every food product has problems with moisture change, oxygen migration, aroma loss or gain, oil migration, or mechanical integrity. Often, several of these problems exist at the same time. Until now, research has focused on investigating whey protein film formation and properties. From the results obtained, it can be concluded that whey proteins can form films with excellent oxygen, aroma, and oil barrier properties at low RH, but they have poor moisture barrier properties. Incorporation of lipids has improved the moisture barrier ability of films, but the effect on their oxygen barrier properties remains unknown. In addition, a more effective incorporation of lipids, examination of lipids that have not been explored, and the effect of lipid particle size and shape may lead to breakthroughs in lowering the WVP of whey protein-lipid composite films. It is well known that plasticizer type and amount significantly affect barrier performance and film flexibility. However, there is a need to better understand how plasticizer composition, structure, and molecular size affect film properties. Incorporation of food ingredients, such as flavorings, antimicrobials, and antioxidants, has not yet been explored in whey protein films, and their use could improve the final food quality and safety. The ultimate goal of determining the film-forming ability and film properties of whey proteins is application of these films as food coatings. Even though some information is available, data are still lacking on forming such films as coatings on food products and assessing their effectiveness by using instrumental and sensory quality analysis. Furthermore, limited research has been conducted on efficient formation of whey protein films for food wraps or pouches. Such wraps and pouches would still require conventional secondary packaging. However, by providing protection and becoming part of the food, they reduce packaging requirements and can increase package recyclability (Krochta and De Mulder-Johnston, 1997).

Alcantara, C.R. and Krochta, J.M. 1996. “Mechanical Properties of Whey Protein-Isolate-Coated Freeze-Dried Chicken White Meat Dice,” in Book of Abstracts, 1996 IFT Annual Meeting, New Orleans, LA. Chicago, IL: Institute of Food Technologists, p. 179. Alcantara, C.R., Rumsey, T.R. and Krochta, J.M. 1998. “Drying Rate Effect on the Properties of Whey Protein Films,” J. Food Process. Engr., 21:387–405. Alexandrescu, A.T., Evans, P.A., Pitkeathly, M., Baum, J. and Dobson, C.M. 1993. “Structure and Dynamics of the Acid-Denatured Molten Globule State of -Lactalbumin: A Two-Dimensional NMR Study,” Biochem., 32:1707–1718.

178 Andrews, A.T. 1979. “The Formation and the Structure of Proteose Peptone Components,” J. Dairy Res., 46:215–218. Anker, M., Stading, M. and Hermansson, A-M. 1998. “Mechanical Properties, Water Vapor Permeability, and Moisture Contents of -Lactoglobulin and Whey Protein Films Using Multivariate Analysis,” J. Agric. Food Chem., 46:1820–1829. Banerjee, R. and Chen, H. 1995. “Functional Properties of Edible Films Using Whey Protein Concentrates,” J. Dairy Sci., 78:1673–1683. Banerjee, R., Chen, H. and Wu, J. 1996. “Milk Protein-Based Edible Film Mechanical Strength Changes Due to Ultrasound Process,” J. Food Sci., 61:824–828. Brunner, J. R. 1977. “Milk Proteins” in Food Proteins, eds., J.R. Whitaker and S.R. Tannenbaum, Westport, CT: AVI Publishers, Inc., pp. 175–208. Cayot, P. and Lorient, D. 1997. “Structure-Function Relationships of Whey Proteins” in Food Proteins and Their Applications, eds., S. Damodaran and A. Paraf, New York: Marcel Dekker, Inc., pp. 225–256. Chen, H. 1995. “Functional Properties and Applications of Edible Films Made of Milk Proteins,” J. Dairy Sci., 78:2563–2583. Debeaufort, F., Martin-Polo, M. and Voilley, A. 1993. “Polarity Homogeneity and Structure Affect Water Vapor Permeability of Model Edible Films,” J. Food Sci., 58:426–429, 434. de Boer, R., de Wit, J.N. and Hiddink, J. 1977. “Processing of Whey by Means of Membranes and Some Applications of Whey Protein Concentrate,” J. Soc. Dairy Tech., 30:112–120. De Mulder-Johnston, C. 1999. “Thermal Analysis of, and Oil Migration through Films from Whey Protein Isolate,” Ph.D. Dissertation, University of California, Davis, pp. 33–57. de Wit, J.N. 1989. “Functional Properties of Whey Proteins” in Developments in Dairy Chemistry, ed., P.F. Fox, New York: Elsevier Applied Science, pp. 285–321. de Wit, J.N. and Klarenbeek, G. 1984. “Effects of Various Heat Treatments on Structure and Solubility of Whey Proteins,” J. Dairy Sci., 67: 2701–2710. Dybing, S.T. and Smith, D.E. 1991. “Relation of Chemistry and Processing Procedures to Whey Protein Functionality: A Review,” Cultured Dairy Products J., 26:4–12. El-Hibri, M.J. and Paul, D.R. 1985. “Effects of Uniaxial Drawing and Heat-Treatment on Gas Sorption and Transport in PVC,” J. Appl. Polym. Sci., 30:3649–3678. Fairley, P., Monahan, F.J., German, J.B. and Krochta, J.M. 1996a. “Mechanical Properties and Water Vapor Permeability of Edible Films from Whey Protein Isolate and Sodium Dodecyl Sulfate,” J. Agric. Food Chem., 44:438–443. Fairley, P., Monahan, F.J., German, J.B. and Krochta, J.M. 1996b. “Mechanical Properties and Water Vapor Permeability of Edible Films from Whey Protein Isolate and N-Ethylmaleimide or Cysteine,” J. Agric. Food Chem., 44:3789–3792. Fox, P.F. 1989. “The Milk Protein System” in Developments in Dairy Chemistry—4, ed., P.F. Fox, New York: Elsevier Applied Science, pp. 1–53. Galietta, G., Di Gioia, L., Guilbert, S. and Cuq, B. 1998. “Mechanical and Thermomechanical Properties of Films Based on Whey Proteins as Affected by Plasticizer and Crosslinking Agents,” J. Dairy Sci., 81:3123–3130. Gennadios, A. and Weller, C.L. 1991. “Edible Films and Coatings from Soymilk and Soy Protein,” Cereal Foods World, 36:1004–1009. Gennadios, A., Weller, C. L. and Testin, R. F. 1993. “Temperature Effect on Oxygen Permeability of Edible Protein-Based Films,” J. Food Sci., 58: 212–214, 219. Glover, F.A. 1985. “Ultrafiltration and Reverse Osmosis for the Dairy Industry,” Tech. Bull. 5. NIRD. Reading, England. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation—Theory and Practice, ed., M. Mathlouthi, New York: Elsevier Applied Science Publishers, pp. 371–394. Hanlon, J.F. 1992. “Films and Foils” in Handbook of Package Engineering, ed., J.F. Hanlon, Lancaster, PA: Technomic, Publishing Co., Inc., pp. 3.1–3.59.

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Huffman, L.M. 1996. “Processing Whey Protein for Use as a Food Ingredient,” Food Technol., 50(2):49–52. Kamper, S.L. and Fennema, O. 1984. “Water Vapor Permeability of Edible Bilayer Films,” J. Food Sci., 49:1478–1481, 1485. Kamper, S.L. and Fennema, O. 1985. “Use of an Edible Film to Maintain Water Vapor Gradients in Food,” J. Food Sci., 50:382–384. Kester, J.J. and Fennema, O.R. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Kester, J.J. and Fennema, O. 1989. “An Edible Film of Lipids and Cellulose Ethers: Barrier Properties to Moisture Vapor Transmission and Structural Evaluation,” J. Food Sci., 54:1383–1389. Kinsella, J.E. 1984. “Milk Proteins: Physicochemical and Functional Properties,” CRC Crit. Rev. Food Sci. Nutr., 21:197–262. Kinsella, J.E. and Whitehead, D.M. 1989. “Proteins in Whey: Chemical, Physical and Functional Properties,” Adv. Food Nutr. Res., 33:343–438. Krochta, J.M. 1992. “Control of Mass Transfer in Foods with Edible Coatings and Films” in Advances in Food Engineering, eds., R.P. Singh and M.A. Wirakartakasumah, Boca Raton, FL: CRC Press, Inc., pp. 517–538. Krochta, J.M. 1997a. “Edible Protein Films and Coatings” in Food Proteins and Their Applications, eds., S. Damodaran and A. Paraf, New York: Marcel Dekker, Inc., pp. 529–549. Krochta, J.M. 1997b. “Edible Composite Moisture-Barrier Films” in Packaging Yearbook: 1997, ed., B. Blakistone. Washington, DC: National Food Processors Association, pp. 38–51. Krochta, J.M. 1998. “Whey Protein Interactions: Effects on Edible Film Properties” in Functional Properties of Proteins and Lipids, eds., J.R. Whitaker, F. Shahidi, A.L. Munguia, R.Y. Yada and G. Fuller, Washington, DC: American Chemical Society, pp. 158–167. Krochta, J.M. and De Mulder-Johnston, C. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Lent, L.E., Vanasupa, L.S. and Tong, P.S. 1998. “Whey Protein Edible Film Structures Determined by Atomic Force Microscope,” J. Food Sci., 63:824–827. Lerdthanangkul S. and Krochta J.M. 1996. “Edible Coating Effects on Postharvest Quality of Green Bell Peppers,” J. Food Sci., 61:176–179. Mahmoud, R. and Savello, P.A. 1992. “Mechanical Properties of and Water Vapor Transferability through Whey Protein Films,” J. Dairy Sci., 75:942–946. Mahmoud, R. and Savello, P.A. 1993. “Solubility and Hydrolyzability of Films Produced by Transglutaminase Catalytic Crosslinking of Whey Protein,” J. Dairy Sci., 76:29–35. Maté, J.I. and Krochta, J.M. 1996a. “Comparison of Oxygen and Water Vapor Permeabilities of Whey Protein Isolate and -Lactoglobulin Edible Films,” J. Agric. Food Chem., 44:3001–3004. Maté, J.I. and Krochta, J.M. 1996b. “Whey Protein Coating Effect on the Oxygen Uptake of Dry Roasted Peanuts,” J. Food Sci., 61:1202–1206, 1210. Maté, J.I., Saltveit, M.E. and Krochta, J.M. 1996. “Peanut and Walnut Rancidity: Effects of Oxygen Concentration and Relative Humidity,” J. Food Sci., 61:465–467, 472. Matthews, M.E. 1984. “Whey Protein Recovery Processes and Products,” J. Dairy Sci., 67:2680–2692. McHugh, T.H. and Krochta, J.M. 1994a. “Sorbitol- vs Glycerol-Plasticized Whey Protein Edible Films: Integrated Oxygen Permeability and Tensile Property Evaluation,” J. Agric. Food Chem., 52:841–845. McHugh, T.H. and Krochta, J.M. 1994b. “Water Vapor Permeability Properties of Edible Whey Protein-Lipid Emulsion Films,” JAOCS, 71:307–312. McHugh, T.H. and Krochta, J.M. 1994c. “Dispersed Phase Particle Size Effects on Water Vapor Permeability of Whey Protein-Beeswax Edible Emulsion Films,” J. Food Process. Preserv., 18:173–188.

180 McHugh, T.H, Aujard, J-F. and Krochta, J.M. 1994. “Plasticized Whey Protein Edible Films: Water Vapor Permeability Properties,” J. Food Sci., 59:416–419, 423. Miller, K.S. and Krochta, J.M. 1997a. “Oxygen and Aroma Barrier Properties of Edible Films: A Review,” Trends Food Sci. Technol., 8:228–237. Miller, K.S. and Krochta, J.M. 1997b. “Measuring Aroma Transport in Polymer Films,” Trans. ASAE, 41:427–433. Miller, K.S., Chiang, M.T. and Krochta, J.M. 1997. “Heat Curing of Whey Protein Films,” J. Food Sci., 62:1189–1193. Miller, K.S., Upadhyaya, S.K. and Krochta, J.M. 1998. “Permeability of d-Limonene in Whey Protein Films,” J. Food Sci., 63:244–247. Morr, C.V. and Ha, E.Y.W. 1993. “Whey Protein Concentrates and Isolates: Processing and Functional Properties,” CRC Crit. Rev. Food Sci. Nutr., 33:431–476. Papiz, M.Z., Sawyer, L., Eliopoulos, E.E., North, A.C.T., Findlay, J.B.C., Sivaprasadarao, R., Jones, T.A., Newcomer, M.E. and Kraulis, P.J. 1986. “The Structure of -Lactoglobulin and Its Similarity to Plasma Retinol-Binding Protein,” Nature, 324:383–385. Pérez-Gago, M.B. and Krochta, J.M. 1999. “Water Vapor Permeability of Whey Protein Emulsion Films as Affected by pH,” J. Food Sci., 64:695–698. Pérez-Gago, M.B., Nadaud, P. and Krochta, J.M. 1999. “Water Vapor Permeability, Solubility and Tensile Properties of Heat-Denatured versus Native Whey Protein Films,” J. Food. Sci., 64:1034–1037. Perkins, W. 1988. “Effect of Molecular Weight and Annealing Temperature on the Oxygen Barrier Properties of Oriented PET Film,” Polym. Bull., 19:397–401. Salame, M. 1986. “Barrier Polymers” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker. New York: John Wiley & Sons, pp. 48–54. Shellhammer, T.H. 1996. “Characterization and Optimization of Water Vapor Transport through Whey Protein-Lipid Edible Emulsion Films,” Ph.D. Dissertation, University of California, Davis. Shellhammer, T.H. and Krochta, J.M. 1997a. “Whey Protein Emulsion Film Performance as Affected by Lipid Type and Amount,” J. Food Sci., 62:390–394. Shellhammer, T.H. and Krochta, J.M. 1997b. “Water Vapor Barrier and Rheological Properties of Simulated and Industrial Milkfat Fractions,” Trans. ASAE, 40:1119–1127. Sherwin, P., Smith, D.E. and Fulcher, R.G. 1998. “Effect of Fatty Acid Type on Dispersed Phase Particle Size Distributions in Emulsion Edible Films,” J. Agric. Food Chem., 46:4534–4538. Shimada, K. and Cheftel, J.C. 1998. “Sulfhydryl Group Disulfide Bond Interchange during Heat Induced Gelation of Whey Protein Isolate,” J. Agric. Food Chem., 37:161–168. Sienkiewicz, T. and Riedel, C.-L. 1990. “Utilization of Whey” in Whey and Whey Utilization, Second Edition, eds., T. Sienkiewicz and C-L. Riedel, Gelsenkirchen-Buer, Germany: Verlag Th. Mann, pp. 92–130. Stuchell, Y.M. and Krochta, J.M. 1995. “Edible Coatings on Frozen King Salmon: Effect of Whey Protein Isolate and Acetylated Monoglycerides on Moisture Loss and Lipid Oxidation,” J. Food Sci., 60:28–31. Taylor, C.C. 1986. “Cellophane” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker, New York: John Wiley & Sons, pp. 159–163. Trezza, T.A. 1999. “Surface Properties of Edible, Biopolymer Coatings for Foods: Color, Gloss, Surface Energy and Adhesion,” Ph.D. Dissertation, University of California, Davis. U.S. Department of Agriculture. NASS. 1999. “Dairy Products: 1999 Summary,” Agr. Stat. Board. Washington, D.C. Yildirim, M. and Hettiarachchy, N.S. 1998. “Properties of Films Produced by Cross-Linking Whey Proteins and 11S Globulin Using Transglutaminase,” J. Food Sci., 63:248–252.

Formation and Properties of Casein Films and Coatings 1

ASEIN-BASED edible films are attractive for food applications due to their high nutritional quality, excellent sensory properties, and good potential to adequately protect food products from their surrounding environment. Milk protein-based films and coatings were extensively reviewed (Gennadios et al., 1994; McHugh and Krochta, 1994; Chen, 1995). This chapter focuses on caseinand caseinate-based films, and recent developments on such films are discussed. Casein is a unique protein because it is only synthesized in the mammary gland and is found nowhere else in nature (Fox and McSweeney, 1998). There are more than 4,000 species of mammal animals. Milk from only five or six of these species is used for human consumption. Discussion in this chapter is limited to bovine milk only because it is of most significant industrial and commercial value.

C

Average bovine milk contains about 30–35 g protein per liter, despite the 1Mention of a trademark or proprietary product is for identification only and does not imply a guarantee or warranty of the product by the U.S. Department of Agriculture. Mention of brand or firm names does not constitute an endorsement by the U.S. Department of Agriculture over others of a similar nature not mentioned.

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182 fact that a significant variation in concentration exists depending on the stage of lactation and season (Fox and McSweeney, 1998). Milk protein accounts for about 20% of the calories in bovine milk and casein accounts for about 80% of the total bovine milk proteins. Casein is easily digestible, which ensures its nutritive accessibility, due to its unique open structure in the natural state. Casein is of relatively high nutritional quality. Its average biological value (ABV), defined as body protein that can be replaced by 100 g of dietary protein, is 80, based on the ABV of egg albumen being 100 (Fox and McSweeney, 1998). Crop proteins generally have lower ABV values (e.g., 45 for wheat proteins and 69 for potato proteins) (Werner, 1981). Casein, as the major milk protein, is naturally designed to meet a young animal’s basic nutritional needs. It is rich in lysine, an essential amino acid in which many plant proteins are deficient. Use of casein with plant proteins can significantly enhance the nutritional value of the mixed diet. Casein also is a rich source of calcium. The factor limiting the ABV of casein is its low content of sulfurous essential amino acids (i.e., cysteine or cystine).

Chemical and physical properties of casein have been well characterized (Brunner, 1981; Swaisgood, 1982; Fox and McSweeney, 1998). Casein has physical properties different from those of either typical globular proteins or fibrous proteins, such as myosin. Several principal chemical and physicochemical properties of casein are summarized in Table 7.1. This section briefly discusses some aspects of this knowledge matrix most pertinent to film formation and properties. Casein is not a homogenous protein. Four different types of casein, namely

Chemical and Physical Characteristics of Casein and Caseinates

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s1, s2, , and , are found in bovine milk in the approximate ratio of 4:1:4:1 (Swaisgood, 1982; Dalgleish, 1997). Their molecular weights range from 19 to 25 kDa. Small variations of the casein molecules occur due to varying degrees of phosphorylation of the four casein species, varying degrees of glycosylation of -casein, and genetic polymorphism (Fox and McSweeney, 1998). The following discussion briefly distinguishes the differences among the four major caseins. At secretion, casein exists in the form of micelles containing all four casein species complexed with colloidal calcium phosphate. Each micelle consists of an average of 104 peptide chains with molecular weights of about 105 kDa. The casein micelles are stable to most common milk processes such as heating, compacting, and homogenization. Micellar integrity is preserved by extensive electrostatic and hydrogen bonding, and hydrophobic interactions. The micellar casein complexes disintegrate and the protein precipitates upon acidification of milk to pH 4.6 at around 30 C, as calcium phosphate dissolves under these conditions. This property is used as the operational definition of casein and is utilized in preparing industrial casein (acid casein) and in manufacturing certain cheese varieties (e.g., cottage cheese and cream cheese). Acid casein finds application in coffee creamers and in meat extenders. Caseins are quite soluble in water despite their high content of non-polar amino acids (35 to 45% of total amino acid residues). Aqueous solutions with casein concentrations up to 20% can be prepared at elevated temperatures. The high solubility is due to the high content of phosphate groups, the low level of sulfur-containing amino acids, and the high carbohydrate content in -casein. Casein also has a high water binding capacity. About 2.5 g of water can be bound to 1 g of casein. This high water-binding capacity results in casein films that are sensitive to environmental humidity. Caseins are phosphoproteins containing about 0.85% phosphorus (Fox and McSweeney, 1998). The phosphate in casein is essentially esterified to serine as monoester (Ser-P) and is referred to as organic phosphate. The organic phosphate residues in casein form “phosphate centers” that possess a considerable negative charge around neutral pH and bind calcium cations strongly (Dalgleish, 1997). This structure is also believed to contribute to the thermal stability of the caseins. Casein is low in sulfur (0.8%) as compared to whey proteins (1.7%). The major caseins ( s1- and -caseins) contain neither cysteine nor cystine. The other two casein fractions contain only two half-cystine residues. Thus, the caseins have good thermal stability. Differential scanning calorimetry showed no heat of denaturation when caseins were subjected to thermal treatments (Paulsson and Dejmek, 1990). They cannot form extensive covalent inter- or intramolecular disulfide bonds to render water-insoluble films. The primary structures of the caseins reveal that they contain many hydrophobic amino acid residues with non-polar side chains (35 to 45% of total resi-

184 dues). Uneven distribution of these amino acids results in hydrophobic ends and patches. The - and -caseins are amphipathic proteins having hydrophobic and hydrophilic ends and, thus, are especially suitable for use as emulsifiers. This feature helps the formation of stable composite protein-lipid emulsions for coating wet surfaces. However, caseins are generally considered hydrophilic because their hydrophobicity values (Table 7.1) are lower than that of valine (7.05 kJ/residue). Among the casein fractions, -casein is the most hydrophobic, and s2-casein is the most hydrophilic. Caseins have little secondary or tertiary structure, probably due to their high content of proline, which disrupts -helices and -sheets (Fox and McSweeney, 1998). The casein molecules have an open, flexible, and mobile conformation that is often referred to as “random coil” structure. Such a structure facilitates film formation through intermolecular hydrogen and electrostatic bonds, and hydrophobic interactions. The open structure also makes the caseins more accessible to enzymatic reactions, such as transglutaminase catalyzed cross-linking. The casein micelles are stabilized by -casein, which buries its hydrophobic N-terminal into the micelle while protruding its hydrophilic C-terminal into the surrounding aqueous environment. Rennet enzymes, such as chymosin, cleave the Phe105-Met106 bond producing para- -casein and glycomacropeptides. Removal of the hydrophilic macropeptides from the surface of the casein micelles reduces their surface (zeta) potential by half and removes the steric stabilizing layer, which eventually leads to casein coagulation in the presence of calcium cations. The coagulability of casein is utilized in manufacturing most cheese varieties as well as rennet casein. Rennet casein has been used for making cheese analogs. Casein molecules have a strong tendency to self-associate either with the same species or inter-species, and thus their separation is rather difficult without using dissociating agents, such as high concentrations of urea (Dalgleish, 1997). Common casein products in the market today are mixtures of the four protein species. It is currently cost prohibitive to produce individual casein fractions for commercial food consumption, even though they may have different film-forming properties due to their differing chemical and physico-chemical properties.

Its relative simple isolation and the useful properties of casein as an industrial material and food ingredient have led to commercial production of casein and caseinates since the 19th century (Muller, 1982). The first significant industrial application of casein as glue occurred in Europe in the 1850s (Southward, 1989). Expanded uses in industrial applications (especially in glues, pa-

Manufacture of Casein and Caseinates

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per coatings, paints, leather finishing, textile fibers, and plastics), and in various food products, pushed the world production of casein to around 150,000 metric tons in the mid-1960s (Muller, 1982). Subsequently, production slowly increased to about 205,000 metric tons over the past three decades (Anonymous, 1997). The end-uses of casein and caseinates have gradually shifted from industrial to food applications. About 70 to 80% of the casein produced worldwide in the 1980s was used as a food ingredient (Muller, 1982). New Zealand is presently the major casein producer in the world. Currently, there is no significant commercial casein production in the United States. About 94,647 metric tons of casein and casein mixtures (about 46% of the worldwide production) were imported by the United States in 1996 (Anonymous, 1997). Different manufacturing processes affect the physico-chemical properties of casein products. Two principal methods have been established for the production of casein on commercial scale, i.e., isoelectric precipitation (acid casein) and enzymatic coagulation (rennet casein) (Muller, 1982).

The preparation of acid casein from skim milk is quite simple. The essential steps involve acidification to about pH 4.6 (isoelectric point of casein) to induce the coagulation of casein, adjustment of temperature to between 30 and 40 C for better handling properties of the product, washing, pressing or centrifuging the curd to remove excessive water, and finally drying and grinding. Hydrochloric acid is usually used for both laboratory scale and industrial preparation of casein. Lactic acid produced in situ by a culture of mesophilic lactic acid bacteria, usually Streptococcus lactis and/or S. cremoris, is also widely used, especially in New Zealand. Sulfuric acid is only used occasionally because the resulting magnesium sulfate in the whey is a laxative. Acid casein is insoluble in water. Innovative techniques for producing acid casein have recently been investigated. Processes such as ion exchange, electrodialysis, and bipolar-membrane electroacidification have been tested and yielded acid casein with different chemical and physical properties (Bazinet et al., 2000). The implications of these new casein products on edible film properties have not been fully investigated.

The key difference in producing rennet casein from producing acid casein is the means of coagulation. A proteolytic enzyme, such as chymosin (rennin), cleaves the -casein fraction to release a glycomacropeptide, thus destabilizing the casein micelles and promoting coagulation of casein in the presence of calcium cations. Rennet casein is insoluble in water or alkali, but can be dissolved

186 by adding calcium-sequestering agents such as polyphosphates. It is suitable for making cheese analogs and has good flavor stability (Muller, 1982).

Though water-insoluble casein has some applications, most food application would require casein with high water solubility. This is achieved by dispersing the casein in water and adjusting the pH to between 6.5 and 7.0 with an alkali (Fox and McSweeney, 1998). The most commonly used soluble caseinate is sodium caseinate. It is normally manufactured by dissolving fresh acid casein curd in sodium hydroxide followed by spray drying. Other soluble caseinates prepared in a similar manner include potassium, calcium, magnesium, and ammonium caseinates.

A new process for precipitating casein by injecting high-pressure carbon dioxide (CO2) into skim milk at elevated temperature has been developed (Tomasula et al., 1995, 1997). It is a variation of acid precipitation that takes place at a pH value (6.0) higher than the isoelectric point of casein. The resulting curd and whey are free of precipitant because CO2 is removed upon depressurization. The casein produced has a high calcium content (about 1.5%) equivalent to that of calcium caseinate, because nearly intact casein micelles are precipitated. The CO2-precipitated casein also contains more phosphorus, which is an indication that the integrity of the casein micelles is preserved. The chemical and physical properties of CO2-precipitated casein are more similar to micellar casein.

The commercial production of casein and whey protein co-precipitates was developed to capture most of the proteins in skim milk and to improve the nutritive value of casein. The process involves a modification of acid casein production by adding calcium chloride into heated milk at 90 to 95 C. Heating denatures the whey proteins. The level of calcium in the co-precipitate has a significant effect on its functional properties (Muller, 1982). However, this process has not achieved commercial success.

Casein can easily form films due to its open secondary structure. It is gener-

Film Formation Mechanisms

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ally agreed in the literature that extensive hydrogen and electrostatic bonds, and hydrophobic associations facilitate the formation of intermolecular interactions that increase interchain cohesion to form film (McHugh and Krochta, 1994). The chemical and physical forces that may change the balances of the intermolecular interactions can perceivably modify the film properties. Adjusting the pH, changing the drying rate, and adding functional additives such as plasticizers, hydrophobic ingredients, and cross-linking ions, are examples of approaches used by investigators.

The protein matrix formed in casein films by the casting and natural drying process is random and less compact. Input of mechanical energy may help to promote intermolecular interactions, thus improving the mechanical and barrier properties of the films. Two techniques—ultrasound and microfluidization—have been attempted (Banerjee et al., 1994, 1996). Ultrasonic energy and acoustic cavitation can cause particle size reduction, rupture of biological cells, and homogenization of emulsions. Microfluidization, an ultra-homogenization process, subjects film-forming formulations to a combination of high shear, cavitation, and unique head-on-head collision impact, thus resulting in stable, very fine emulsions and dispersions. Such high intensity of mechanical energy input is perceived to promote protein conformational changes and intermolecular interactions. The increased molecular interactions lead to higher molecular order. An orderly polymeric matrix could result in rigid films exhibiting higher film strength (Guilbert, 1986). An additional feature of the ultrasonic technique is production of sterile films because there is no direct mechanical contact with the film-forming formulation.

Strong covalent bonds are perceived to promote tighter intermolecular interaction, thus increasing film strength and resistance to mass transfer. Enzymatic and physical (irradiation) treatments have also been explored in forming casein films.

Transglutaminase (EC 2.3.2.13, TGase) is an enzyme that catalyzes the formation of intra- and intermolecular -( -glutamyl) lysyl cross-links of protein molecules (Nielsen, 1995). The reaction occurs at mild temperature and within a wide pH range, and proceeds rapidly (Motoki et al., 1987a). Recently, a new industrial grade TGase produced by microbial fermentation became commer-

188 cially available at a lower cost than animal-derived TGase. The microbial-derived TGase also functions independently of calcium cations. Casein is an ideal substrate for TGase due to its less ordered conformation and relatively high content of glutamine and lysine (Table 7.1).

Lacroix et al. (1998) discussed their studies of cross-linking casein molecules by -irradiation. The irradiation of aqueous casein solution resulted in a chain reaction, in which hydroxyl radicals (.OH) were first generated as a product of water radiolysis (Brault et al., 1997). The hydroxyl radicals then reacted with aromatic amino acids, such as tyrosine and phenylalanine, to produce tyrosyl (phenoxyl) radicals, which then formed several stable biphenolic compounds through covalent bonding. The 2 ,2-biphenol bityrosine was the major product due to the strong directing effect of the hydroxyl group, although other products were also present. The concentration of bityrosine can be easily detected due to its characteristic fluorescence. It may be more likely to form intermolecularly than intramolecularly. The cross-linked caseinates have a molecular size up to 100 times that of monomer casein as confirmed by size exclusion chromatography (Lacroix et al., 1999). Due to their relatively high content of tyrosine and phenylalanine (Table 7.1), caseins are good candidates for cross-linking by irradiation. Another benefit of the irradiation treatment is the production of sterile films that may find special uses in medical or food applications.

Film preparation by casting solutions on flat surfaces and subsequently drying the castings is adequate for laboratory use and, perhaps, is also suitable for batch production of films similar to the traditional papermaking process. However, more efficient techniques are needed for commercial film production. Frinault et al. (1997) developed a wet spinning process to prepare films from acid casein and investigated the film properties as affected by the process and pH. The process involved extrusion of acid casein solution (17% protein content) into a coagulating bath containing sodium hydroxide, followed by hardening, rinsing, and drying. This process can be easily modified into a continuous film process. The pH of the film-forming solution was important to the film properties. Use of harsh chemicals in coagulation and hardening rendered the films questionable for edible applications. The prepared films had properties (e.g., tensile strength, elongation, and water vapor permeability) comparable to

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those of cast casein films. A similar process for producing casein films with mild chemicals was described in a patent (Metzger, 1997).

Edible films for food packaging must meet a number of functional needs to preserve food quality and to enhance sensory characteristics of foods. Important functional properties of edible films include resistance to mass transfers and mechanical protection.

Casein- and caseinate-based films, similar to most protein-based films, have limited moisture barrier ability due to their hydrophilic nature compared to commonly used synthetic plastic films (Table 7.2). The water vapor permeability (WVP) of casein films is two to four orders of magnitude higher than the WVP of low-density polyethylene and Saran wrapping films. The effects of structural materials, additives, and processing conditions on WVP of casein films are discussed below. Guilbert (1986) characterized casein films formed in sodium hydroxide solutions (pH 8) as poor water barriers. However, no quantitative information regarding WVP was reported. Krochta et al. (1990) compared relative water loss of simple sodium caseinate films plasticized with glycerol (protein to plasticizer ratio of 2:1) under a presumed relative humidity (RH) gradient of 0/100%. The sodium caseinate films reduced water permeation by only 7 to 14%. Chen (1995) reported a more substantial reduction in water vapor loss (76%) by a simple whey protein concentrate film, which had a similar WVP value to a sodium caseinate film (Banerjee and Chen, 1995). The noticeable difference between these two reports, i.e., Krochta et al. (1990) and Chen (1995), was the RH gradient. The permeability test cells were placed in a constant 55% RH environment in the study by Chen (1995), which was close to most practical food storage conditions. Thus, it is perceivable that casein films may be able to substantially protect the wet surface of foods, such as minimally processed vegetables and fruits, meat, and fish products. The water barrier ability of films is better characterized by the WVP value at certain test conditions and film thicknesses. The most popular method currently used for determining WVP of hydrophilic films is a version of Standard ASTM Method E 96-95 (ASTM, 1995) modified by McHugh et al. (1993) to account for a partial water vapor pressure gradient across the stagnant air-vapor layer underside of the film being tested. The first set of such WVP data for sodium caseinate films was reported by McHugh et al. (1993) (Table 7.2). Two

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194 other critical conditions were imposed by the modified WVP measurement method. First, an air speed greater than 152 m/min over the film surface outside of the testing cell is necessary to ensure that the assumption of constant external RH is valid. Second, a stagnant air gap less than 14 mm between the film sample and the liquid surface inside the testing cup should be maintained to avoid a convection effect.

WVP values of casein and caseinate films from various studies are compiled in Table 7.2. Different casein products may result in films of different water barrier properties. Ho (1992) compared the WVP of simple protein films plasticized with glycerol (4 to 1 ratio) prepared with several commercial casein products and determined that film WVP values ascended in the following order: magnesium caseinate < calcium caseinate < micellar casein < sodium caseinate < potassium caseinate < rennet casein. The WVP value increased by 28% from the least to the most permeable film (Table 7.2). Two independent studies from our laboratory confirmed Ho’s results, i.e., the WVP of caseinate salt films increased from calcium to sodium to potassium caseinate (Banerjee and Chen, 1995; Li and Chen, 2000). Chick and Ustunol (1998) also reported that lactic acid casein films had slightly lower WVP than rennet casein films. However, Tomasula et al. (1998) showed that films made from CO2-precipitated casein, which is chemically similar to micellar casein and is structurally more rigid, had lower WVP than calcium caseinate films. Unlike calcium caseinate films, the CO2-precipitated casein films did not swell. This indicates greater resistance to moisture vapor transfer.

The significant difference in WVP values between sodium and calcium caseinate films is attributed to electrostatic differences and to calcium cross-links. Avena-Bustillos and Krochta (1993) reported that the WVP value of calcium caseinate films was lower than that of sodium caseinate films by 24%. Similarly, Banerjee and Chen (1995) observed that the WVP of calcium caseinate films was lower compared to that of sodium caseinate films. Recently, Li and Chen (2000) again confirmed this trend (Table 7.2) and attributed it to the cross-linking effect of the divalent calcium cations. Divalent ions, such as calcium and magnesium cations, promote intermolecular cross-linking, which tightens the protein matrix by limiting biopolymer segmental mobility. This produces films with reduced WVP (Ho, 1992; Banerjee and Chen, 1995). Treating sodium caseinate films with calcium


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chloride at pH 9.6 reduced film WVP by 42% (Avena-Bustillos and Krochta, 1993). The treated films were also thinner by 31% due to the tighter molecular structure. There appears to be a correlation between the WVP values of simple caseinate films and the size of the cations. The smaller ions occupy less space between protein strands, thus resulting in tighter structures and greater resistance to mass transfer. WVP data in the literature are consistent with this postulation; namely, the WVP of magnesium caseinate films is lower than that of calcium caseinate films. Similarly, the WVP of sodium caseinate films is lower than that of potassium caseinate films (Ho, 1992; Li and Chen, 2000). Vigorous mechanical agitation of film-forming formulations is believed to facilitate a tighter intermolecular interaction, thus resulting in a more closely knitted protein matrix, which is perceivably more resistant to mass diffusion. Caseinate films treated by microfluidization at 34.5 MPa did show increased water resistance (Banerjee et al., 1994). The most noticeable difference for microfluidized simple protein formulations was with sodium caseinate films where the WVP was 29% lower compared to that of untreated control films. Among the casein-acetylated monoglyceride composite films, sodium caseinate films showed the most significant improvement (lower WVP by 43%) due to microfluidization (Table 7.2). Covalent cross-linking improves film resistance to moisture transfer. Intermolecular bityrosine cross-linking induced by -irradiation effectively improved the water vapor barrier properties of casein films. Lacroix et al. (1999) reported a reduction of 65% in WVP from 1.25 g mm/m2 h kPa for non-irradiated calcium caseinate films to 0.43 g mm/m2 h kPa for irradiated films.

Protein molecules tend to form tighter interactions when the environmental pH is adjusted toward the isoelectric point (pI) of the protein. This is the principle of acid coagulation of casein mentioned earlier. The resulting proteins become insoluble in water. However, the enhanced protein interaction within the protein film matrix by pH adjustment may or may not result in improved barrier properties. Krochta et al. (1990) reported that sodium caseinate films treated with sodium acetate buffer at the average isoelectric point of casein (pH 4.6) rendered the films insoluble, but had little effect on improving water barrier ability. The slight decrease of water vapor transmission, if any, may be attributed to the loss of glycerol in the soaking process. The hygroscopic nature of glycerol in the films compromises the water barrier ability. In the same study, treatment of sodium caseinate-acetylated monoglyceride emulsion films with the same buffer

196 solution (pH 4.6), followed by soaking in water for two minutes, effectively reduced water transmission by about 50% (Krochta et al., 1990). No difference in water transmission was observed among films treated with three different buffers, namely, sodium acetate, sodium ascorbate, and calcium ascorbate, all at pH 4.6. This indicated that pH adjustment to the pI, rather than cross-linking by calcium cations, was the predominant factor in modifying film permeability. Avena-Bustillos and Krochta (1993) reported that sodium caseinate films treated in various buffers at the pI reduced WVP by 32 to 43% compared to control films. Also, the treated films were about 14% thinner than control films. The underside of the treated films had a higher RH level indicating improved water vapor resistance. It was attributed to a tightened molecular structure as the result of pH treatment.

Incorporation of hydrophobic materials can effectively modify casein film properties. Various lipids, waxes, and fatty acids (including acetylated monoglycerides, beeswax, carnauba wax, and paraffin wax) have been tested for their effectiveness in improving the moisture barrier characteristics of casein films. Krochta et al. (1990) reported that films made from aqueous emulsions of sodium caseinate and acetylated monoglycerides had lower water vapor transmission rate by 41% compared to control films without lipids. The lowest moisture transmission was found among the films made of sodium caseinate emulsions with monoglycerides that have lower acetylation and fewer unsaturated fatty acids. Doubling the amount of acetylated monoglyceride further reduced water vapor permeation by about 50%. Treating the emulsion films with buffer at the pI followed by soaking in distilled water for two minutes reduced water transmission by another 50%. Water vapor transmission through the films made of sodium caseinate emulsion with paraffin, carnauba, or beeswax is quite similar to that of acetylated monoglyceride (Krochta et al., 1990). Acetylated monoglycerides are considered better film additives than waxes because they have lower melting temperatures and are easier to produce emulsions. Adding acetylated monoglyceride to various caseinate films in 1:2 ratio noticeably reduced film WVP by 23 to 36% (Li and Chen, 2000). The most significant reduction was noticed with potassium caseinate films (Banerjee and Chen, 1995; Li and Chen, 2000). More lipids in sodium caseinate films resulted in lower WVP values (Avena-Bustillos and Krochta, 1993). When the acetylated monoglyceride to protein ratio increased from 0:5 to 4:1, WVP decreased by 57%. The WVP of films of acetylated monoglyceride to sodium caseinate ratio of 1:4 was 0.66 g mm/m2 h kPa. In a different study, Avena-Bustillos and Krochta (1993) concluded that


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beeswax was more effective than stearic acid or acetylated monoglyceride in improving the water vapor barrier ability of sodium caseinate films. With a ratio of beeswax to sodium caseinate of 1:3, the film WVP was 0.41 g mm/m2 h kPa (73% lower than that of simple sodium caseinate film). The beeswax-sodium caseinate film had about half of the permeability of the acetylated monoglyceride-sodium caseinate film. The greater hydrophobicity and crystalline structure of beeswax suppressed moisture diffusion more effectively. However, adding beeswax at more than 25% of total solids did not further reduce the WVP of sodium caseinate films due to poor dispersion in the film system (Avena-Bustillos and Krochta, 1993).

Water vapor transfer rate increased with increasing moisture content and plasticizer concentration of caseinate films (Arvanitoyannis et al., 1996; Siew et al., 1999). Chick and Ustunol (1998) reported an increase in WVP (by 8 to 32%) when the protein to plasticizer ratio decreased from 1.4:1 to 1:1 for films made from lactic acid casein or rennet casein with either glycerol or sorbitol as plasticizer. Different plasticizers may impart different effects on film water barrier properties. Glycerol-plasticized casein films had lower WVP values than films plasticized with polyethylene glycol (PEG) (Siew et al., 1999). A correlation between protein conformation and WVP was found. The caseinate films plasticized with glycerol had a higher percentage of -helical structure and a lower percentage of random coil structure than films plasticized with PEG. It was perceived that the random coil structure formed a more “open” molecular network that facilitated water transmission. The higher WVP of the PEG-plasticized films compared to the glycerol-plasticized films was also attributed to the relatively higher hydration rate of PEG. Sorbitol-plasticized casein films were more effective moisture barriers than glycerol-plasticized films (Chick and Ustunol, 1998). Sorbitol is less hygroscopic than glycerol.

While there is no significant difference in WVP between either side of simple protein films facing the higher RH environment during testing, there usually is a notable difference for protein-lipid composite films. The orientation of protein-lipid composite films is an important factor in determining WVP. When the film side rich in hydrophobic material faces the higher water vapor partial pressure, a smaller WVP value is generally obtained. The WVP of a 72.5% calcium caseinate/27.5% beeswax film differed by nearly 100% from

198 one side to the other (Avena-Bustillos and Krochta, 1993). The lowest WVP (0.15 g mm/m2 h kPa) was observed for a 62.5% calcium caseinate/37.5% beeswax emulsion film that was measured with the hydrophobic-rich side facing the higher RH (Avena-Bustillos and Krochta, 1993).

Corn and wheat starches, being inexpensive, have been investigated for preparing films in blends with sodium caseinate (Arvanitoyannis et al., 1996). The starches were compatible with casein in forming uniform films with sufficient strength, acceptable gas barrier properties, and no phase separation at moisture contents lower than 15%. A higher protein content in the films ensured lower moisture and oxygen permeability due to the less hydrophilic nature of sodium caseinate compared to starch. Parris et al. (1995) reported that the WVP of composite films from 50% alginate/30% sodium caseinate/20% glycerol was reduced by 34% (Table 7.2) compared to control alginate only films while maintaining similar tensile strength (20 to 30 MPa). Sodium caseinate performed equally well to other milk protein products, including whole milk powder, whey proteins, and non-fat dry milk powder in such composite films with alginate.

A noticeable issue related to film WVP determination is that an ideal, but far from realistic, RH gradient is frequently employed. Subjecting films to 0% RH on one side and 100% RH on the other side does not simulate the actual conditions of most food applications. Some researchers measured the WVP at 50–65%/100% RH gradients. Such data are more applicable to most normal food environments, yet are still limited as point measurements. There is a gap in methodology of reliable prediction or extrapolation of WVP data of any edible films. These data are crucial in packaging design. The research community must address this issue to attract broader commercial interest in the use of edible films.

Generally, casein films, similar to other protein-based films, are brittle without the addition of plasticizers. Non-plasticized films are of limited practical use (Chen, 1995). Insertion of hydrogen bonding breakers, such as various plasticizers, relaxes the structure and improves the flexibility of protein films. Tomasula et al. (1998) reported that a minimal glycerol concentration of 20% was necessary to prepare casein films suitable for mechanical testing. Film thickness also directly affects the ease of film handling. Casein films cast from


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solutions with total solids contents of 2 or 4% were brittle and difficult to peel from the casting surface (Tomasula et al., 1998).

Tensile testing provides several important film mechanical parameters such as tensile strength (TS), elongation at break (E), and Young’s modulus (YM). TS represents film resistance to tension. E is a measure of film stretchability. YM indicates the stiffness of the material. While TS and E are extensively reported in the literature, YM has not been extensively studied. TS is an important parameter in packaging design. It frequently determines the thickness required for the wall of a package, thus relating to the cost of the package (Jenkins and Harrington, 1991). TS is defined as the maximum load during a tension test carried to rupture divided by the original cross-sectional area of the specimen (Mohsenin, 1986). It measures the maximum tensile stress that a material is capable of sustaining. For protein films, the maximum stress frequently occurs sooner than film rupture (Chen et al., 1993). The first set of tensile testing data of caseinate films was reported at the Food Preservation 2000 conference sponsored by the U.S. Army Natick Research, Development, and Engineering Center in October 1993 and subsequently published in the conference proceedings (Chen et al., 1993). The TS and E of sodium caseinate films plasticized with glycerol were determined at different strain rates (Table 7.3). TS increased and E decreased, both exponentially, as strain rate increased. This showed that the values of mechanical properties of biopolymer films are strain rate dependent. Ferry (1980) pointed out that the rupture properties of polymer materials depend greatly on the rate of deformation. Therefore, it is important to include the strain rate when reporting tensile data. TS values of casein films reported in the literature ranged from about 1 to 50 MPa (Table 7.3). The high end values are in the vicinity of commonly used polymeric packaging films such as low-density polyethylene (8 to 17 MPa) and high-density polyethylene (17 to 34 MPa) (Salame, 1986; Chen, 1995; Siew et al., 1999). A rather large variation of tensile property values of caseinate films is found in the literature (Table 7.3). For example, the TS and E of sodium caseinate films (Chen et al. (1993): TS 36.9 MPa, E 18.0%; Banerjee and Chen (1995): TS 2.98 MPa, E 29.9%) varied although the same strain rate of 5 mm/s was used during measurement. More recently, another independent test of sodium caseinate films in our laboratory determined TS and E values of 3.80 MPa and 204.05%, respectively (Li and Chen, 2000). These discrepancies indicated that the sensitive nature of film mechanical properties demands a standard method of using the most careful and consistent procedures of sample preparation and test conditions. Differences in film preparation techniques,

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202 moisture equilibrium, specimen cutting, testing temperature, and RH control, in addition to film formulation and tensile rate are critical to accurately and precisely measure protein film mechanical properties.

Different casein products may have an effect on film TS. Calcium caseinate films, which have significantly higher TS and lower E, are characterized as harder and more brittle than sodium or potassium caseinate films (Banerjee and Chen, 1995; Li and Chen, 2000). Divalent calcium ionic bridges result in extensive inter- and intramolecular cross-linking, thus enhancing film strength and limiting film flexibility. Chick and Ustunol (1998) reported that rennet casein films had higher TS (by 27 to 98% at various protein to plasticizer ratios) than lactic acid casein films. They attributed the difference to the higher ash content in rennet casein than lactic acid casein, which provided for additional electrostatic cross-linking between divalent cations and negatively charged groups along protein chains. The more rigid structure of rennet casein molecules may have also contributed to the greater TS of rennet casein films. The TS of CO2-precipitated casein films was greater (by more than 30%) than that of calcium caseinate films at low glycerol content (20% w/w), but the difference diminished when the glycerol content increased (Tomasula et al., 1998). Similarly, the YM value of CO2-precipitated casein films (40.9 MPa) was much greater than that of calcium caseinate films (8.9 MPa). The stronger and stiffer nature of CO2-precipitated casein films was attributed to the fact that most of the micellar calcium and phosphate linkages remained intact as casein precipitation occurred at pH 5.4, rather than at the pI where most of the micellar calcium phosphate dissolved. E values were not significantly different between CO2-precipitated casein films and calcium caseinate films (Tomasula et al., 1998).

The mechanical strength of casein films is directly proportional to the ratio of structural materials and fillers. The higher the ratio of protein to plasticizer, the greater the TS (Chick and Ustunol, 1998; Tomasula et al., 1998). Siew et al. (1999) reported that plasticizers were effective within a range, i.e., too little plasticizer resulted in brittle films and too much plasticizer rendered the films too tacky to yield reliable mechanical data. The TS of sodium caseinate films decreased with increasing plasticizer content (Table 7.3) within the effective range of the plasticizers. The molecular characteristics of plasticizers affect film strength significantly. Casein films plasticized with sorbitol had higher TS than those


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plasticized with glycerol (Chick and Ustunol, 1998). This was attributed to the less hydrophilic nature of sorbitol compared to glycerol, therefore giving casein films with lower affinity to bind water at equivalent RH. However, no equilibrium moisture content data were reported to support this statement. The higher TS of sorbitol-plasticized films might have also been partially due to the fact that the number of glycerol molecules was about double the number of sorbitol molecules at the same plasticizer weight as that used in the film-forming formulations. Glycerol is a much smaller molecule than sorbitol, and, therefore, it interrupts more hydrogen bonds within the protein matrix of casein films where an equal weight of plasticizer is applied. With less hydrogen bonding between protein strands, films have lower TS. Siew et al. (1999) compared the effects of glycerol and PEG 400 plasticizers on the TS of sodium caseinate films and noticed a “cross-over” phenomenon. Specifically, films plasticized with glycerol had higher TS at low plasticizer concentration, but lower TS at higher plasticizer concentration than films plasticized with PEG. Glycerol caused a larger change in film TS (about 20 MPa) while PEG resulted in a smaller change (about 10 MPa). The transition from glassy to rubbery state was observed at lower plasticizer concentration for glycerol-plasticized films compared to PEG-plasticized films. Furthermore, the transition in the glycerol films occurred over a broader plasticizer range, suggesting that the PEG system had a more homogeneous bonding distribution. Smaller plasticizer molecules, such as glycerol, can access the hydrophilic sites on the caseinate chain more easily than larger plasticizer molecules. Glycerol can be easily inserted between the protein strands through hydrogen bonding, thus reducing intermolecular protein interactions, increasing intermolecular spacing, and lowering the mechanical strength of casein films. The E of caseinate films generally showed successively an increase, plateau, and then decrease with increasing moisture and plasticizer content (Chick and Ustunol, 1998; Tomasula et al., 1998; Siew et al., 1999). Water molecules have a plasticizing effect on protein films. Insertion of plasticizer molecules as lubricants facilitates the movement of protein chains, thus increasing film flexibility and stretchability. However, too much plasticizer dilutes the cohesion of the protein network as the number of plasticizer-plasticizer bonds increases. Depending on the type of proteins and plasticizers, as well as on the range of protein to plasticizer ratio, one, two, or all three phases may be visible. The data reported by Chick and Ustunol (1998) serve as the best examples, that is (a) film E only increases with increasing plasticizer content (lactic acid casein with sorbitol); (b) film E increases to a peak value and then declines with increasing plasticizer content (lactic acid casein with glycerol); and (c) film E decreases with increasing plasticizer content (rennet casein with either glycerol or sorbitol). Addition of different plasticizers to the casein film matrix resulted in different E values. For example, glycerol-plasticized films were generally more

204 stretchable than sorbitol-plasticized films (Chick and Ustunol, 1998). Glycerol, being more hygroscopic, absorbs more moisture, which serves as an additional plasticizer.

Motoki et al. (1987a) reported that films made from TGase cross-linked casein had good mechanical strength when casein concentration in the film-forming solutions was greater than 3%. However, no film TS data were reported. The E of the TGase-treated films was 72%, which is comparable to that of other casein films.

Mechanical treatment of casein solutions or emulsions at high-energy intensity enhances molecular interactions within the protein film network, thus improving mechanical strength. Banerjee et al. (1996) showed that ultrasound treatment significantly increased TS (by up to 224%) and puncture strength (by up to 120%) of sodium caseinate films. E, WVP, and moisture content of the films were not affected by the ultrasound treatment. Similarly, caseinate films treated by microfluidization at 34.5 MPa showed drastic improvement in mechanical strength (Banerjee et al., 1994). The increase in TS of simple caseinate films ranged from 218 to 364% while the increase in TS of caseinate-acetylated monoglyceride emulsion films ranged from 72 to 653% compared to the TS of the respective untreated control films (Table 7.3).

Blending casein with other structural materials, such as starch, may alter the mechanical strength of resulting films. At 15% moisture content, films made from sodium caseinate and corn starch decreased in TS from 38.3 to 13.3 MPa, and increased in E from 6.2 to 28.0% and in YM from 23.8 to 29.5 MPa (Arvanitoyannis et al., 1996). Furthermore, film TS slowly decreased during storage for 60 days.

Puncture testing is a precise and reliable method to measure film mechanical strength (Chen et al., 1993). Measured values of film puncture strength (PS) depend on puncture speed and film thickness (Chen et al., 1993). Higher puncture rates generally result in higher PS (Table 7.3). It is noted that the PS value at the highest puncture rate in Table 7.3 was off the trend due to the limitation in the


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data acquisition rate of the instrument at that time. Also, the thicker the protein films, the greater the PS. Therefore, when comparing film PS values from the literature, the rate of puncture head speed and film thickness should be considered. Banerjee and Chen (1995) reported that calcium caseinate films had much greater PS than sodium or potassium caseinate films (Table 7.3). This was again attributed to cross-linking between protein polypeptides by calcium cations. Addition of acetylated monoglyceride in the presence of glycerol diminished differences in PS among calcium, sodium, and potassium caseinate films. The PS values of caseinate-acetylated monoglyceride films were much lower than those of simple caseinate films (Table 7.3), although the composite films were thicker than the simple protein films. This trend was recently reconfirmed in our laboratory (Table 7.3) (Li and Chen, 2000). Brault et al. (1997) reported that the PS of sodium caseinate and calcium caseinate films was not affected by -irradiation dose up to 12 kGy. However, the calcium caseinate films were stronger than the sodium caseinate films, especially at the highest irradiation dose at which the PS of the calcium caseinate films was greater by 17% than that of the sodium caseinate films.

Casein and caseinate films dissolve nearly instantaneously in water (Krochta et al., 1990; Chen, 1995; Tomasula et al., 1998). Casein films formed at pH 8 were soluble in both cold and hot water (Guilbert, 1986). The high solubility of casein and caseinate films is desirable for many food applications. For example, water-soluble spice and flavoring pouches can be used in preparation of soups. Also, water-soluble pouches can be used in precision manufacturing for adding pre-measured additives (e.g., yeast in dough making) to ensure product consistency. The solubility of casein films can be modified by pH treatment and cross-linking by divalent cations. Krochta et al. (1990) reported that sodium caseinate films remained undissolved in water for more than 24 hours after the films were soaked in sodium acetate buffer at the pI (pH 4.6). Soaking caseinate films in calcium salt solutions also decreased film solubility (Krochta et al., 1990). Casein films treated with hardening agents had low solubility in water and were opaque and whitish in appearance (Frinault et al., 1997). CO2-precipitated casein films retained their integrity in water, but became translucent and tacky (Tomasula et al., 1998). The water insolubility of such films was attributed to the tight protein interaction of the casein resulting from the unique precipitation process (Tomasula et al., 1998). Caseinate films with covalent intermolecular cross-links are not soluble in water. Films from both sodium caseinate and calcium caseinate that were irradiated at 4 and 12 kGy did not dissolve in water (Brault et al., 1997). Film insol-

206 ubility increased with irradiation doses up to 32 kGy where 70% of the caseinate films remained insoluble after boiling in water for 30 minutes followed by soaking in water at ambient temperature for 24 hours (Lacroix et al., 1999). TGase cross-linked s1-casein films were insoluble in boiling water, 10% 2-mercaptoethanol, 6.6 M urea, 10% SDS, or 6 M guanidine, and were digestible by common gastric enzymes, such as chymotrypsin (Motoki et al., 1987a).

Simple protein films prepared from casein and caseinates are transparent, colorless, tasteless, odorless, and smooth (McHugh and Krochta, 1994; Chen, 1995; Chick and Ustunol, 1998). Cross-linking, such as irradiation treatment, does not change the sensory appeal of caseinate films (Brault et al., 1997). Composite caseinate-lipid films are usually translucent. The extent of translucency depends on the type and amount of hydrophobic additives. The hydrophobic ingredients largely modify the flavor profile and texture of composite films. Films prepared from CO2-precipated casein had a slightly milky appearance due to the presence of intact casein micelles (Tomasula et al., 1998). The film surface had a texturized-like appearance, presumably due to localized phase separation during film drying. Comparison of scanning electron micrographs of the CO2-precipitated casein and calcium caseinate films indicated that the casein micelle structure was largely preserved during the CO2 precipitation process (Tomasula et al., 1998).

Films made from naturally occurring macromolecules are considered inherently biodegradable (Chen, 1995; Krochta and De Mulder-Johnston, 1997; Debeaufort et al., 1998; Arvanitoyannis and Gorris, 1999). However, limited data are available with respect to the degradation of such films. Some recent film preparation techniques employing covalent cross-linking polymerization warrant a careful checking of this degradability assertion. Biodegradability of caseinate films cross-linked by -irradiation was investigated (Mezgheni et al., 1998; Ressouany et al., 2000). Cross-linked caseinate films were more resistant to spoilage microorganisms (i.e., Pseudomonas aeruginosa) than control caseinate films, but still were susceptible to microbial degradation. The bacterial population in the medium containing the cross-linked caseinate films reached its maximum density much slower than that of the control films. Soluble nitrogen content, an indication of protein network degradation, agreed with the biodegradation trends of the microbial study. Conversion of carbon to CO2 by biodegradation also ensured the

Uses of Casein Films and Coatings

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degradability of the cross-linked caseinate films as well as the difference in degradation rate between the cross-linked films and the control films. This observation shows a possibility of customizing the shelf stability of protein films by controlling the number of cross-links.

Applications of casein films, and protein films in general, are discussed in detail elsewhere in this book. Therefore, only a brief discussion is presented below. The early exploration of casein films revealed several useful concepts, including extending the shelf life of coated bakery products and chocolate candies; reducing moisture loss from raisins to breakfast cereals, whole fruits and vegetables, chicken eggs, and frozen fish; preventing lipid oxidation of frozen fish; and improving microbial stability of intermediate moisture fruits (McHugh and Krochta, 1994). Some of these applications employed casein-lipid composite formulations to improve resistance to moisture loss. Krochta et al. (1990) proposed to extend the shelf life of lightly processed fruits and vegetables using films made from aqueous emulsions of casein with acetylated monoglycerides. Avena-Bustillos et al. (1993) studied the effect of various ratios and total solids contents of sodium caseinate-stearic acid emulsion coatings on the white blush and water vapor loss of peeled baby carrots using response surface methodology. They reported that an optimized coating formulation (about 1.5% sodium caseinate and 0.2% stearic acid) effectively retarded the white blush while increasing water vapor resistance by about 80% compared to uncoated control samples. In another study, Avena-Bustillos et al. (1994a) reported the effectiveness of calcium or sodium caseinate-lipid (1:1 ratio; 2% total solids) emulsion coatings on water vapor resistance, white blush, and respiration rate of peeled carrots. All the coatings, except for the sodium caseinate-acetylated monoglyceride coating, increased the water vapor resistance significantly. A calcium caseinate-beeswax coating offered the highest water vapor resistance, but had an undesirable whitish appearance, presumably due to wax crystallization. A sodium caseinate-stearic acid coating was the most effective in retarding white blush of carrots during storage. Some coating treatments also significantly reduced the respiration rate of the carrots. Avena-Bustillos et al. (1994b) attempted to optimize calcium caseinate-acetylated monoglyceride formulations to coat zucchini to reduce water loss and improve the quality. They determined that maximum water vapor resistance could be achieved with relatively high sodium caseinate and low acetylated monoglyceride contents in the edible coatings. The calcium caseinate-based coatings did not affect internal carbon dioxide and ethylene concentrations, or surface optical properties. Caseinate-acetylated monoglyceride emulsions also were applied on whole

208 fruit apples and celery sticks (Avena-Bustillos et al., 1997). The coatings improved the water vapor resistance of celery sticks with an optimal formulation of 1.5% calcium caseinate and 1.5% acetylated monoglyceride. However, the coatings did not affect water vapor resistance, respiration rate, or ethylene production of whole apples. It was also reported that water vapor resistance was not affected by the type of caseinates, calcium cross-linking, or addition of potassium sorbate (Avena-Bustillos et al., 1997). Other novel applications of casein coatings have also been considered. Japanese researchers demonstrated that casein-coated supplementary amino acids in fish diets improved plasma amino acid balance, which was significantly correlated with average weight gain, feed efficiency, and protein efficiency ratio (Murai et al., 1982a, b). In this case, coating amino acids with casein minimized variation in absorption rates of certain amino acids and resulted in simultaneous presentation. Another interesting application was immobilizing enzymes in transglutaminase cross-linked casein films (Motoki et al., 1987b). Four selected enzymes (i.e., -glucosidase, -mannosidase, -galactosidase, and glucose oxidase) were simultaneously entrapped in the casein films. The immobilized enzymes maintained their activities on repeated usage, washing, and drying. The thermal resistance of the enzymes was improved, and their suitable pH range was widened and shifted slightly toward the alkaline end for some of the enzymes. Enzymatic activity was similar to that of soluble reaction systems. Proteins have many side chain groups that serve as affinity moieties to carry functional additives such as antimicrobials and antioxidants. The protein matrix can also physically entrap functional ingredients and form a concentrated zone to boost efficiency. Guilbert (1988) reported that casein films containing sorbic acid significantly reduced the surface microbial load of intermediate moisture fruit cubes (apricots or papaya) inoculated with Aspergillus niger or Saccharomyces rouxii even at water activity values as high as 0.903. In this manner, it is possible to produce stable intermediate moisture foods with desirable soft texture, fresh appearance, and low dose of preservatives.

Casein and caseinate films can find broad application in food packaging. Their diverse functional properties are suitable for many applications. Their high nutritional value and superior sensory characteristics give them an edge over other protein films. Their mechanical properties are generally suitable for edible purposes, neither too tough nor too fragile. Though more permeable to water vapor than plastic films, they are capable of retarding moisture loss to a reasonable degree. To meet more stringent requirements, casein can be used with other materials in multi-layer packaging systems. Research efforts on ca-

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sein and protein films should continue to focus on some critical issues including, but not limited to (1) standardization of film testing methods; (2) development of mathematical models for predicting film functional properties for packaging design; (3) use of new composite film systems; (4) determination of film gas permeability values; and (5) development of new, continuous film-forming technologies. Researchers have showcased the potential of several interesting applications. The food industry should be encouraged to develop more product-specific applications based on the numerous concepts suggested by the research community. Only close cooperation between the industry and the research institutions can elevate the uses of casein films to their full potential.

Anonymous. 1997. Dry Milk Products Utilization & Production Trends—1996. Chicago, IL: American Dairy Products Institute, p. 16. Arvanitoyannis, I. and Gorris, L.G.M. 1999. “Edible and Biodegradable Polymeric Materials for Food Packaging or Coating” in Processing Foods: Quality Optimization and Process Assessment, eds., F.A.R. Oliveira and J.C. Oliveira, Boca Raton, FL: CRC Press, pp. 357–371. Arvanitoyannis, I., Psomiadou, E. and Nakayama, A. 1996. “Edible Films Made from Sodium Caseinate, Starches, Sugars or Glycerol. Part 1,” Carbohydrate Polym., 31:179–192. ASTM. 1995. “Designation E 96-95: Standard Test Methods for Water Vapor Transmission of Materials” in Annual Book of ASTM Standards, vol. 4.06, West Conshohocken, PA: American Society for Testing and Materials, pp. 697–704. Avena-Bustillos, R.J. and Krochta, J.M. 1993. “Water Vapor Permeability of Caseinate-Based Edible Films as Affected by pH, Calcium Crosslinking and Lipid Content,” J. Food Sci., 58:904–907. Avena-Bustillos, R.J., Cisneros-Zevallos, L.A., Krochta, J.M. and Saltveit, M.E. 1993. “Optimization of Edible Coatings on Minimally Processed Carrots Using Response Surface Methodology,” Trans. ASAE, 36:801– 805. Avena-Bustillos, R.J., Cisneros-Zevallos, L.A., Krochta, J.M. and Saltveit, Jr., M.E. 1994a. “Application of Casein-Lipid Edible Film Emulsions to Reduce White Blush on Minimally Processed Carrots,” Postharvest Biol. Technol., 4:319–329. Avena-Bustillos, R.J., Krochta, J.M., Saltveit, M.E., Rojas-Villegas, R.J. and Sauceda-Pérez, J.A. 1994b. “Optimization of Edible Coating Formulations on Zucchini to Reduce Water Loss,” J. Food Engr., 21:197–214. Avena-Bustillos, R.J., Krochta, J.M. and Saltveit, M.E. 1997. “Water Vapor Resistance of Red Delicious Apples and Celery Sticks Coated with Edible Caseinate-Acetylated Monoglyceride Films,” J. Food Sci., 62:351–354. Banerjee, R. and Chen, H. 1995. “Functional Properties of Edible Films Using Whey Protein Concentrate,” J. Dairy Sci., 78:1673–1683. Banerjee, R., Chen, H., Hendricks, G. and Levis, J.E. 1994. “Impact of Microfluidization on Mechanical and Water Vapor Barrier Properties of Milk Protein Based Edible Films,” J. Dairy Sci., 77(Suppl. 1):24. Banerjee, R., Chen, H. and Wu, J. 1996. “Milk Protein-Based Edible Film Mechanical Strength Changes Due to Ultrasound Process,” J. Food Sci., 61:824–828. Bazinet, L., Lamarche, F., Ippersiel, D., Gendron, C., Mahdavi, B. and Amiot, J. 2000. “Comparison of Electrochemical and Chemical Acidification of Skim Milk,” J. Food Sci., 65:1303–1307.

210 Brault, D., D’Aprano, G. and Lacroix, M. 1997. “Formation of Free-Standing Sterilized Edible Films from Irradiated Caseinates,” J. Agric. Food Chem., 45:2964–2969. Brunner, J.R. 1981. “Cow Milk Proteins: Twenty-five Years of Progress,” J. Dairy Sci., 64:1038–1054. Chen, H. 1995. “Functional Properties and Applications of Edible Films Made of Milk Proteins,” J. Dairy Sci., 78:2563–2583. Chen, H., Banerjee, R. and Limoge, E.J. 1993. “Testing Mechanical Strengths of Edible Films” in Proceedings of Food Preservation 2000, eds., I.A. Taub and R. Bell, Hampton, VA: Science and Technology Corporation, pp. 863–875. Chick, J. and Ustunol, Z. 1998. “Mechanical and Barrier Properties of Lactic Acid and Rennet Precipitated Casein-Based Edible Films,” J. Food Sci., 63:1024–1027. Dalgleish, D.G. 1997. “Structure-Function Relationships of Caseins” in Food Proteins and Their Applications, eds., S. Damodaran and A. Paraf, New York: Marcel Dekker, pp. 199–223. Debeaufort, F., Quezada-Gallo, J.A. and Voilley, A. 1998. “Edible Films and Coatings; Tomorrow’s Packagings: A Review,” Critical Rev. Food Sci., 38:299–313. Ferry, J.D. 1980. Viscoelastic Properties of Polymers. New York: John Wiley & Sons, Inc. Fox, P.F. and McSweeney, P.L.H. 1998. Dairy Chemistry and Biochemistry. London: Blackie Academic & Professional. Frinault, A., Gallant, D.J., Bouchet, B. and Dumont, J.P. 1997. “Preparation of Casein Films by a Modified Wet Spinning Process,” J. Food Sci., 62:744–747. Gennadios, A., McHugh, T.H., Weller, C.L. and Krochta, J.M. 1994. “Edible Coatings and Films Based on Proteins” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 201–277. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation. Theory and Practice, ed., M. Mathlouthi, London: Elsevier Applied Science, pp. 371–394. Guilbert, S. 1988. “Use of Superficial Edible Layer to Protect Intermediate Moisture Foods: Application to the Protection of Tropical Fruits Dehydrated by Osmosis” in Food Preservation by Moisture Control, ed., C.C. Seow, London: Elsevier Applied Science, pp. 119–219. Ho, B. 1992. Water Vapor Permeabilities and Structural Characteristics of Casein Films and Casein-Lipid Emulsion Films. M.S. Thesis, University of California, Davis, CA, USA. Jenkins W.A. and Harrington, J.P. 1991. Packaging Foods with Plastics. Lancaster, PA: Technomic Publishing Company, Inc. Krochta, J.M. and De Mulder-Johnston, C. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Krochta, J.M., Pavlath, A.E. and Goodman, N. 1990. “Edible Films from Casein-Lipid Emulsions for Lightly-Processed Fruits and Vegetables” in Engineering and Food, Vol. 2, Preservation Processes and Related Techniques, eds., W.E.L. Spiess and H. Schubert, London: Elsevier Applied Science, pp. 329–340. Lacroix, M., Jobin, M., Mezgheni, E., Srour, M. and Boileau, S. 1998. “Polymerization of Calcium Caseinate Solutions Induced by Gamma Irradiation,” Radiat. Phys. Chem., 52:223–227. Lacroix, M., Ressouany, M., Ouattara, B., Yu, H. and Vachon, C. 1999. “Radiation Cross-Linked Protein Films and Their Physicochemical Properties” in Proceedings of 5th Conference of Food Engineering, Dallas, TX, pp. 602–607. Li, J. and Chen, H. 2000. Unpublished data. University of Vermont, Burlington, VT, USA. McHugh, T.H. and Krochta, J.M. 1994. “Milk-Protein-Based Edible Films and Coatings,” Food Technol., 48(1):97–103. McHugh, T.H., Avena-Bustillos, R.J. and Krochta, J.M. 1993. “Hydrophilic Edible Films: Modified Procedure for Water Vapor Permeability and Explanation of Thickness Effects,” J. Food Sci., 58:899–903. Metzger, W. October 28, 1997. U.S. patent 5,681,517.

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Mezgheni, E., Vachon, C. and Lacroix, M. 1998. “Biodegradability Behavior of Cross-Linked Calcium Caseinate Films,” Biotech. Progress, 14:534–536. Mohsenin, N.N. 1986. Physical Properties of Plant and Animal Materials. New York: Gordon and Breach Science Publishers. Motoki, M., Aso, H., Seguro, K. and Nio, N. 1987a. “ s1-Casein Film Prepared Using Transglutaminase,” Agric. Biol. Chem., 51:993–996. Motoki, M., Aso, H., Seguro, K. and Nio, N. 1987b. “Immobilization of Enzymes in Protein Films Prepared Using Transglutaminase,” Agric. Biol. Chem., 51:997–1002. Muller, L.L. 1982. “Manufacture of Casein, Caseinates and Coprecipitates” in Developments in Dairy Chemistry—1. Proteins, ed., P.F. Fox, London: Applied Science Publishers, pp. 315–337. Murai, T., Akiyama, T., Ogata, H., Hirasawa, Y. and Nose, T. 1982a. “Effect of Coating Amino Acids with Casein Supplemented to Gelatin Diet on Plasma Free Amino Acids of Carp,” Bull. Jap. Soc. Sci. Fisheries, 48:703–710. Murai, T., Akiyama, T. and Nose, T. 1982b. “Effects of Casein Coating on Utilization of Dietary Amino Acids by Fingerling Carp and Channel Catfish,” Bull. Jap. Soc. Sci. Fisheries, 48:787–792. Nielsen, P.M. 1995. “Reactions and Potential Industrial Applications of Transglutaminase. Review of Literature and Patents,” Food Biotechnol., 9:119–156. Parris, N., Coffin, D.R., Joubran, R.F. and Pessen, H. 1995. “Composition Factors Affecting the Water Vapor Permeability and Tensile Properties of Hydrophilic Films,” J. Agric. Food Chem., 43:1432–1435. Paulsson, M. and Dejmek, P. 1990. “Thermal Denaturation of Whey Proteins in Mixtures with Caseins Studied by Differential Scanning Calorimetry,” J. Dairy Sci., 73:590–600. Ressouany, M., Vachon, C. and Lacroix, M. 2000. “Microbial Resistance of Caseinate Films Crosslinked by Gamma Irradiation,” J. Dairy Res., 67:119–124. Salame, M. 1986. “Barrier Polymers” in The Wiley Encyclopedia of Packaging Technology, ed., M. Bakker, New York: John Wiley & Sons, pp. 48–54. Siew, D.C.W., Heilmann, C., Easteal, A.J. and Cooney, R.P. 1999. “Solution and Film Properties of Sodium Caseinate/Glycerol and Sodium Caseinate/Polyethylene Glycol Edible Coating Systems,” J. Agric. Food Chem., 47:3432–3440. Southward, C.R. 1989. “Uses of Casein and Caseinates” in Developments in Dairy Chemistry—4. Proteins, ed., P.F. Fox, London: Applied Science Publishers, pp. 173–244. Swaisgood, H.E. 1982. “Chemistry of Milk Protein” in Developments in Dairy Chemistry—1. Proteins, ed., P.F. Fox, London: Applied Science Publishers, pp. 1–59. Tomasula, P.M., Craig, Jr., J.C., Boswell, R.T., Cook, R.D., Kurantz, M.J. and Maxwell, M. 1995. “Preparation of Casein Using Carbon Dioxide,” J. Dairy Sci., 78:506–514. Tomasula, P.M., Craig, Jr., J.C. and Boswell, R.T. 1997. “A Continuous Process for Casein Production Using High-Pressure Carbon Dioxide,” J. Food Engr., 33:405–419. Tomasula, P.M., Parris, N., Yee, W. and Coffin, D. 1998. “Properties of Films Made from CO2-Precipitated Casein,” J. Agric. Food Chem., 46:4470–4474. Werner, H. 1981. “Whey Protein,” Dairy Industr. Int., 46(9):33.

Formation and Properties of Fish Myofibrillar Protein Films and Coatings

F

OR many years, various raw materials from agricultural resources have been

used to produce renewable, biodegradable, or edible “agricultural-materials” or “agro-packaging.” Among agricultural resources, some of them rich in proteins have long been used empirically as raw material. For instance, buttons were made from formaldehyde-treated caseins, and Henry Ford manufactured automobile parts from mixtures of soy protein and phenol-formaldehyde resins. Numerous cereal and vegetable proteins (e.g., corn zein, wheat gluten, and soy protein) and animal proteins (e.g., milk proteins, collagen, gelatin, keratin, and myofibrillar proteins) are commonly used or have been tested to form films and coatings (Cuq et al., 1998a). Whereas the first reports describing film-forming properties of proteins (mainly corn zein, wheat gluten, soy proteins, collagen, gelatin, and milk proteins) are relatively old, the film-forming applications of myofibrillar proteins were first mentioned by Okamoto (1978), who described the transformation of fish meats for preparing edible coatings. Recently, the film-forming properties of myofibrillar proteins were extensively studied, and the main functional properties of films were evaluated (Cuq et al., 1995, 1998b; Souza et al., 1997). The potential use of myofibrillar proteins for the preparation of films and coatings was proposed to identify new practical applications for meat and fish animal proteins (particularly from fish surimi); to test new natural film-forming 213

214 polymers; and to assess the development of edible and/or biodegradable films and coatings for packaging fish or meat products (Cuq et al., 1998b). In this chapter, the structure, main sources, and physicochemical properties of myofibrillar proteins in relationship with their film-forming properties are discussed. Also, the formation methods, principal functional properties, and potential applications of fish myofibrillar protein films are reviewed.

Myofibrillar proteins are found in meats (e.g., beef, sheep, pork, and poultry) and fish. These are particularly important food products that represent significant sources of proteins rich in essential amino acids.

Myofibrillar proteins represent the main component of muscles (more than 50% of total muscle weight). Mammal muscles consist of giant cells, whereas fish muscles are divided into encased layers. Muscles have a high number of parallel fibers containing filamentous bodies called myofibrils that are formed by myofibrillar proteins. Myofibrillar proteins are structured with contractile proteins, contraction regulatory proteins, and structural proteins (Table 8.1). The association of myosin and actin forms contractile proteins. Myosin associates two sub-units known as heavy chains (200 kDa) and two pairs of light chains (16 to 20 kDa). There is a cylindrical part of the two heavy chains entangled in a double helix, and a globular part is formed by the entanglement of

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Figure 8.1 Schematic representation of the structure of myofibrillar proteins.

heavy chains associated with light chains (Figure 8.1). Actin is the second main protein in muscles. It exists in a globular form, i.e., G-actin (42 kDa), and in a fibrous form, i.e., F-actin (14,000 kDa) formed by linear polymerization of G-actin. The actomyosin term is often used to indicate the protein fraction formed by the association of actin and myosin. Actin and myosin are directly involved in the contraction-relaxation cycle of muscular cells. Contraction regulatory proteins play a key role in the regulation of muscular contraction. Tropomyosin is closely associated with actin and troponin filaments. This association contributes to mechanical stabilization of muscular fibers (Figure 8.1). Troponins are located on the F-actin filaments, along tropomyosin chains. Both - and -actinin are globular proteins that have regulating functions in muscle. M-proteins have enzymatic and structural function. C-protein consists of only one polypeptide chain and can form cylindrical rings to join the tails of myosin chains. Titin and nebulin constitute the structural proteins. Titin represents one-third of the total weight of myofibrillar proteins (Trinick, 1991) and is characterized by an exceptionally high molecular mass (2,800 kDa). Titin filaments are parallel to actin and myosin filaments and contribute to formation and stabilization of their organization. Nebulin is one of the main structural proteins of the muscle, accounting for 3–4% of all myofibrillar proteins. Nebulin and F-actin are directly associated over their whole lengths.

Myofibrillar proteins can only be used for film-forming applications after purification and concentration from meat or fish. This involves successive

216 washing treatments to remove undesirable compounds such as blood components, sarcoplasmic proteins (myoglobin), extracellular proteins (collagen), and lipids (Cuq et al., 1995; Monterrey-Q, 1998). The process is known as surimi (or fish and meat mince) process. Surimi production technology relies on water extraction of low molecular weight components (e.g., lipids and soluble proteins) from mechanically separated fish muscles by successive washing. This results in semi-purified protein fractions containing high concentrations of myofibrillar proteins. Addition of cryoprotective agents (e.g., sucrose and sorbitol) to the resulting concentrate of myofibrillar proteins permits long-term storage at 20 to 30 C without changes in protein functional properties. Surimi preparation from white (lean) fish (e.g., hake, cod, and Alaska pollock) is described below (Lee, 1986; Pigott, 1986; Venugopal, 1992). After the reception of fresh fish preserved in ice, resolution of rigor mortis (5 to 20 h) is necessary prior to any processing. Fish cleaning, head removal, and evisceration must be rapid to prevent intestinal proteases from penetrating into the muscle. It is often better to scale fish and remove their central bones. Several washing steps to remove undesirable components follow the mechanical separation of bones and skins. Washing, which is repeated two or three times, involves gentle agitation of the flesh mince in two to three volumes of cool water for five minutes, and then drying by pressing or centrifugation. Sodium chloride (0.1 to 0.3 g per 100 g of fish flesh mince) is added to the last washing bath to facilitate final drying. Sieving of the washed fish pulp eliminates residual connective tissues, skin, and small bones. Final drying in a screw press decreases the water content of fish mince (about 80 g of water/100 g of fish mince). The washed fish mince is homogenized in a grinder with cryoprotective agents (for example, 5 g sucrose, 5 g sorbitol, and 0.3 g polyphosphates per 100 g of fish mince). Following conditioning in blocks packed in polyethylene bags, the surimi is frozen at 25°C. Surimi from white fish is colorless and odorless and has interesting functional properties, particularly gelling properties (Lanier, 1986). The heat-induced gelation characteristics of surimi have been exploited to produce a wide range of fabricated seafood products. Close to 2–3 million tons of fish are used annually for surimi production. The cost of surimi varies from $2 to $5/kg depending on fish species and quality. Preparing surimi from fatty pelagic fish (e.g., sardine, mackerel, anchovy, and herring) requires addition of sodium bicarbonate (0.5 g/L) to the first washing batch to improve elimination of fat components. Surimi from fatty pelagic fish is much less neutral than surimi from white fish. For instance, sardine-based surimis have good gelling properties but also have a slight grayish color and typical fishy odor (Roussel and Cheftel, 1988; Trevino et al., 1990). Surimi-like products from muscles of other animal species have similar properties to fish surimi. However, there is limited information available on preparation and properties of surimi-like products from beef, pork, or chicken (Park et al., 1996; Smyth and O’Neill, 1997). Using red meat as a starting mate-

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rial presents challenges (i.e., high contents of fat, pigments, connective tissue, and collagen). However, such a process could increase the value of underutilized meat cuts or animal by-products, increasing profitability. Aqueous washing and sieving have been used to remove fat, heme pigments, connective tissue, and other water-soluble components from mechanically separated chicken meat (Smyth and O’Neill, 1997). The resulting preparation was a wet concentrate of meat myofibrillar proteins, which may be described as meat surimi.

Preparation of protein-based films requires formation of a continuous, low-moisture, and more or less ordered macromolecular network containing numerous and uniformly distributed interactions between polymer chains. The probability of forming intermolecular bonds mainly depends on protein shape (fibrous versus globular) and on physicochemical conditions during processing. High molecular weight proteins (e.g., myosin) and fibrous proteins (e.g., myosin and F-actin) generally can form films with good mechanical properties (Cuq et al., 1998a), while globular or pseudo-globular proteins (e.g., G-actin) need to be unfolded before film formation. Knowledge of the myofibrillar protein structure and more particularly of the amino acid composition (Table 8.2) can help indicate the physicochemical conditions required for preparing film-forming solutions. Solubilization and gel formation of myofibrillar proteins have been extensively studied (Acton et al., 1983; Acton and Dick, 1984; Regenstein, 1984; Whiting, 1988; Lavelle and Foegeding, 1993). According to the Osborne classification (Table 8.2), myofibrillar proteins are soluble in dilute salt solutions. It is also possible to solubilize myofibrillar proteins into very low ionic strength solutions at neutral pH (Wu et al., 1991; Stanley et al., 1994; Stefansson and Hultin, 1994;

218 Monterrey-Q, 1998). Below 0.001 ionic strength at pH 7, repulsive forces among negative charges on side chains are sufficient to solubilize most myofibrillar proteins (Figure 8.2). Solubility decreases quickly to a minimum at 0.025–0.15 ionic strength. Above 0.15 ionic strength, a salting-in effect is observed that peaks around 0.75 ionic strength. Protein solubility then decreases with higher salt contents through the salting-out effect. Myofibrillar proteins are sensitive to pH variations because of their particularly high contents of ionized polar amino acids (27–31%) (Table 8.2). Close to the isoelectric pH (pI near 5), electrostatic repulsions are minimal, and myofibrillar proteins are not soluble (Orban et al., 1992; Stefansson and Hultin, 1994). Protein solubility increases significantly when the pH is lower than 4.5 or higher than 6.5 (Figure 8.2). The dissociation and solubilization of myofibrillar proteins are then sufficient to expect film formation. Below pH 4, myofibrillar protein dispersions in water-acetic acid gel at ambient temperature (Fretheim et al., 1985; Hermansson et al., 1986; Orban et al., 1992; Shahidi and Venugopal, 1994; Venugopal et al., 1994). Solubilization of myofibrillar proteins induces probable changes in conformation and polymerization-depolymerization reactions (Skaara and Regenstein, 1991). Electrophoretic analysis of myofibrillar protein dispersions revealed reduction of band intensity for myosin heavy chains due to some proteolysis or formation of covalent intermolecular bonds (Kamat et al., 1992). Myosin heavy chains, initially fibrous, are the main components involved in gelling of myofibrillar proteins (Samejima et al., 1981; Roussel and Cheftel, 1990; Sano et al., 1990). Myosin sub-units also participate in network formation by stabilizing the myosin heavy chains (Samejima et al., 1988).

Figure 8.2 Solubility in water of myofibrillar proteins as a function of ionic strength at pH 7 (Stefansson and Hultin, 1994) and as a function of pH (Orban et al., 1992).

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Tropomyosin and G-actin specifically inhibit myosin-myosin interactions while being intercalated between molecules (Samejima et al., 1982; Sano et al., 1989a, b, c). The relative contribution of interactions and bonds involved during gelation of myofibrillar proteins is not yet completely elucidated (Table 8.3). Myosin chains contain approximately 40 sulfhydryl groups, but apparently no disulfide bonds. However, intermolecular disulfide bonds are certainly formed during aggregation of myofibrillar proteins (Roussel and Cheftel, 1990). The contribution of hydrophobic interactions and hydrogen bonds during network formation has been demonstrated by fixating specific probes on myosin chains (Niwa, 1975; Samejima et al., 1981; Niwa et al., 1982). The role of electrostatic interactions has been shown by the positive effect of calcium chloride on the rigidity of myofibrillar protein-based gels (Roussel and Cheftel, 1990). Structured networks of myofibrillar proteins could also be formed in the presence of transglutaminases, which catalyze formation of intermolecular isopeptidic bonds (Kim et al., 1993). These covalent bonds strengthen the mechanical resistance and elasticity of gels (Joseph et al., 1994).

Preparation of films and coatings based on myofibrillar proteins has mainly been studied using fish surimi from sardines (Cuq et al., 1995) or tilapia (Monterrey-Q, 1998). Some data are also available on films based on surimi-like material from beef meat (Souza et al., 1997). Surimi could be used directly after thawing to prepare films or coatings (Cuq et al., 1995). It is also possible to dry surimi (or fish mince) before using it as raw material for film-forming applications. For instance, fish minces were lyophilized (Monterrey-Q, 1998) or air-dried (Cuq et al., 1997d) and ground into powders that were used to prepare films.

220

Protein film macromolecular networks are formed in three stages: (1) rupture of intermolecular bonds that stabilize proteins in the native state (by solubilization or thermal treatment); (2) arrangement and orientation of polymer chains (shaping); and (3) formation of a three-dimensional network stabilized by new interactions and bonds after the agent that initially ruptured the intermolecular bonds is removed (e.g., solvent removal or cooling). Two methods have been widely used to prepare protein-based films and coatings. The solvent process involves dispersion or solubilization of proteins in film-forming solutions. This process has been extensively studied and applied to produce films and coatings from various proteins, particularly from myofibrillar proteins (Cuq et al., 1995; Monterrey-Q, 1998). On the other hand, the dry process is based on the thermoplastic properties of proteins at low moisture contents. Thermoplastic properties of proteins are defined in terms of the glass transition theory. Heating amorphous thermoplastic proteins above the glass transition temperature (Tg) produces soft rubbery materials that can be shaped into specific forms such as films. Cooling at ambient temperature reconverts the rubbery product into a glassy material, giving a more or less rigid form to the desired structure. The dry process was successfully applied to produce films and/or rigid materials using common melt processing technologies (e.g., extrusion, molding, and rolling mill process), but few studies have been published on proteins as thermoplastic raw materials (Fritz et al., 1994). The dry process was tested by Cuq et al. (1997d) with myofibrillar proteins.

Film formation by the solvent process requires dissolution of myofibrillar proteins by adjusting the pH of the film-forming solution (with acetic acid, lactic acid, or ammonium hydroxide). The physicochemical conditions have been optimized to obtain homogeneous film-forming solutions (i.e., myofibrillar proteins do not precipitate out of solution) of low viscosity (i.e., solutions can be easily spread in thin layers) (Cuq et al., 1995; Ocuno and Sobral, 1998; Ocuno et al., 1998). The viscosity of film-forming solutions is greatly modified by pH and protein content. Low protein contents are required to obtain low viscosity film-forming solutions. The viscosity is at minimum between pH 4.5 and 6.5, but film formation is inhibited by myofibrillar protein precipitation. The viscosity is adequate at pH 2.5–4.0 and at pH 9.0. Alkaline conditions (pH between 10.5 and 12) were reportedly suitable for film formation from fish meat (Okamoto, 1978). However, films formed at highly alkaline conditions had a persistent, undesirable ammonia odor (Cuq et al., 1995). Optimal conditions

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for preparing film-forming solutions based on myofibrillar proteins were determined at pH 3 and 2 g protein/100 g solution by Cuq et al. (1995), and at pH 2.5 and 1.25 g protein/100 g solution by Monterrey-Q (1998). Glycerol, sorbitol, and sucrose were tested as plasticizers and were incorporated in the film-forming solutions. Lactic acid used to adjust the pH of film-forming solutions (instead of acetic acid) had a plasticizing effect on films (Souza et al., 1997, 1998). Lactic acid does not evaporate during film drying and thus participates in plasticization of the protein film network. The film-forming solutions were cast on poly(vinylidene chloride) plates and then air-dried. Transparent, easily handled films were formed. Temperature, pH, and protein content of film-forming solution did not significantly affect film functional properties. For instance, film mechanical properties did not change by increasing film-forming solution pH from 2 to 3 (Monterrey-Q, 1998; Monterrey-Q and Sobral, 1998).

The dry process, based on the thermoplastic properties of myofibrillar proteins, was tested by Cuq et al. (1997d) to produce films and rigid packagings. Thermoplastic properties of myofibrillar proteins were studied by dynamic mechanical thermal analysis (Figure 8.3). Sharp changes in mechanical behavior between 215 and 250°C are associated with the glass transition of unplasticized myofibrillar proteins. The Tg range for unplasticized myofibrillar proteins is close to the Tg level typically observed for collagen, gelatin, or starch. Thermal degradation of myofibrillar proteins occurred up to 250 C. It is noted that the glass transition of dry proteins (between 215 and 250 C) occurred just before their thermal degradation, so that the workable temperature range for thermomolding processing is very restricted. Incorporation of plasticizers (such as sucrose and sorbitol) in myofibrillar proteins markedly decreased (>75 C) the Tg (Figure 8.3). The depressive effect of water on the Tg of myofibrillar proteins is also very important. The thermodynamic theory of glass transition (i.e., the Couchman-Karasz equation) is suitable to partially describe the plasticizing effect of water on myofibrillar proteins (Cuq et al., 1997d). Thermomolding of myofibrillar proteins was only possible when the process temperature was higher than the Tg value at the corresponding moisture content. Materials were prepared using a pilot thermomolding process apparatus from myofibrillar protein-based powders hydrated at various levels by water addition (Cuq et al., 1997d). The structures of the materials depended on processing conditions (e.g., translucent glass or foamed structures such as polystyrene).

222

Figure 8.3 Thermomechanical scans (storage modulus and loss modulus) for unplasticized or plasticized (1:1 mixture of sucrose and sorbitol at 35 g/100 g dry matter) myofibrillar proteins (at 2.2 g water/100 g dry matter). (Adapted from Cuq et al., 1997d.)

Irrespective of plasticizer type and content, visual observations clearly showed that myofibrillar protein-based films did not disperse and did not lose their integrity after being immersed in water for 24 hours (Cuq et al., 1997b). High molecular weight proteins are generally insoluble or only slightly soluble in water and thus have potential for forming water-resistant films (Cuq et al., 1998b). However, irrespective of plasticizer type (glycerol, sucrose, or sorbitol), increasing the plasticizer content in the film increased the water-soluble dry matter content (Figure 8.4). In general, hydrophilic plasticizers enhance water solubility.

Properties of Myofibrillar Protein-Based Films

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Variations in the non-protein dry matter content in films (essentially plasticizers) as a function of plasticizer content appeared to be close to experimental variations in water-soluble dry matter content (Figure 8.4). The observed slight difference between solubilized dry matter content and non-protein dry matter content in films was relatively constant and close to 3–5 g/100 g initial dry matter in films. Low molecular weight protein chains (i.e., monomers and small peptides) formed during conditioning of the film-forming solution and immobilized in the film network could thus constitute the water-soluble proteinic components of the films (Cuq et al., 1995). Most likely, the plasticizer will mainly comprise the film dry matter solubilized in water. The myofibrillar protein-based network will therefore not likely solubilize or disperse in water. High density of molecular interactions and certainly the presence of intermolecular covalent bonds or physical knots (chain entanglements) are responsible for the partial insolubility of these films. This water solubility behavior cannot be generalized, and understanding film solubility remains a complex issue. Monterrey-Q (1998) reported that a significant part of glycerol remained water-insoluble in films, suggesting occurrence of glycerol-protein interactions.

Mechanical properties of protein-based films are usually evaluated by extension tests in isothermal and controlled RH conditions. A procedure described by Bourne (1968) and modified by Gontard et al. (1992) was success-

Figure 8.4 Effect of plasticizer content (glycerol (䊊), sorbitol (䊏), or sucrose (䉭) on the watersoluble dry matter content of myofibrillar protein-based films. The dashed line (– – –) represents the non-proteinic dry matter contents in films (g/100 g dry matter). (Adapted from Cuq et al., 1997b.)

224 fully applied to characterize the mechanical properties of various films containing proteins, particularly myofibrillar protein-based films (Cuq et al., 1996a). Force and strain at break are recorded after displacement of a cylindrical probe perpendicular to the film surface. A rheological model was proposed by Cuq et al. (1996b) to describe the mechanical properties of protein-based films as a function of film thickness. Tensile strength and elongation at break values of fish myofibrillar protein-based films and various other films were compared (Table 8.4). Myofibrillar protein-based films had greater tensile strength and lower elongation values, by an order of magnitude, than values reported in the literature for soy protein, wheat gluten, and corn zein films. Also, fish myofibrillar protein films had substantially lower tensile strength values than synthetic polymer films such as polyester, poly(vinylidene chloride), and cellulose acetate, but similar to those of high- and low-density polyethylene (Table 8.4). The distribution and concentration of inter- and intra-molecular interactions allowed by primary and spatial structures most likely affect the mechanical properties of myofibrillar protein-based films.

Water vapor permeability (WVP) values of fish myofibrillar protein-based films and of other biopolymer or synthetic films are presented in Table 8.5. Fish myofibrillar protein-based films had high WVP (3.5 10 12 mol m/m2 s Pa),


225

similar to other hydrocolloid-based films (Cuq et al., 1995). In general, resistance of protein-based films to water vapor transmission is limited due to the inherent hydrophilicity of proteins. Hydrophilic plasticizers, such as glycerol, also facilitate transmission of water vapor through protein-based films. WVP values determined for fish myofibrillar protein-based films were greater by two orders of magnitude than those of low- and high-density polyethylene (Table 8.5).

Oxygen and carbon dioxide permeability values of fish myofibrillar protein-based films and of other biopolymer or synthetic films are shown in Table 8.6. Fish myofibrillar protein-based films have excellent gas barrier properties, especially against oxygen, when they are dry. For instance, the oxygen permeability of a dry fish myofibrillar protein-based film was lower by three orders of magnitude than that of low-density polyethylene (Table 8.6). Macromolecular chains become more mobile when moisture is present, which leads to a substantial increase in gas permeability. Development of fish myofibrillar protein-based films with selective gas permeability could be promising for controlling respiratory exchange and preserving fresh or minimally processed

226 fruits and vegetables (Barron et al., 1999) or for preserving fresh fish that is sensitive to oxidative degradation.

Similar to other hydrophilic polymers, functional properties of fish myofibrillar protein-based films are substantially influenced by plasticization from incorporated plasticizers (e.g., glycerol, sorbitol, and sucrose) and from moisture. Lactic acid used to adjust the pH of film-forming solutions also plasticized myofibrillar protein-based films (Souza et al., 1998). Plasticization of myofibrillar protein-based films with glycerol, sorbitol, or sucrose markedly decreased film strength and apparent elastic modulus, and increased extendability and water vapor permeability (Cuq et al., 1997b; Monterrey-Q, 1998). Film functional properties did not vary significantly as a function of plasticizer type when plasticizers were added at the same molecular contents, due to structural similarities between glycerol, sorbitol, and sucrose (Cuq et al., 1997b). Functional properties of myofibrillar protein-based films also strongly depend on their water content and therefore on surrounding RH. Water interactions with proteins could explain physical property modifications induced by sorbed moisture. Relationships between equilibrium RH and moisture contents of myofibrillar protein-based films were investigated by determining water sorption isotherms at different temperatures (Cuq et al., 1997a; Monterrey-Q,


227

Figure 8.5 Effect of mean water activity on water vapor transmission rate ( 10 12 mol m/m2 s) of myofibrillar protein-based films at 5 C (䉭), 20 C (䊉), or 50 C (ⵧ). (Adapted from Cuq et al., 1997a.)

1998). Increasing the water content generally changed barrier (Figure 8.5) and mechanical properties. For example, the strain at failure of myofibrillar protein films was 100-fold higher when ambient RH increased from 11 to 95% (at 20 C) and 12-fold higher when glycerol content increased from 0 to 0.44 mol/100 g protein (Cuq et al., 1997a). These changes were observed at temperature-dependent RHs (RH was reduced by increasing temperature). The combined effect of temperature and hydration on mechanical and barrier properties of myofibrillar protein-based films could be interpreted in terms of disruptive water-polymer hydrogen bonding. The role of water as a plasticizer of amorphous materials could be therefore associated with glass transition. Thermal properties of myofibrillar protein-based films as a function of their water contents were characterized by dynamic mechanical thermal analysis and by differential scanning calorimetry. During a temperature increase, the films showed sudden changes in mechanical properties and specific heats, which are typically associated with glass-to-rubber transition of amorphous materials. Increasing film water content caused a non-linear decrease in glass transition temperature (Cuq et al., 1997c; Souza et al., 1998). The thermodynamic theory of glass transition (i.e., the Couchman-Karasz equation) is partially suitable for describing the plasticizing effect of water on films (Cuq et al., 1997c).

228

Figure 8.6 Strength-deformation curves during the cutting of a piece of meat with or without edible coating, where (T) was the uncoated control (Cuq, 1996). Coating thickness ranged from 10 to 20 µm (a), 20 to 30 µm (b), 30 to 40 µm (c), and 40 to 50 µm (d).

Films from myofibrillar proteins can be used as edible, protective food coatings. Such films should have mechanical properties that are sufficient to withstand abuse arising during food manufacturing, transport, and marketing, but not too resistant to be detected during their consumption. As an integral part of food, edible films and coatings must have sensory properties that are neutral or compatible with the coated food. Also, it is preferable that the films are not visible on the food surface because edible films are consumed at the same time as the product. Application of myofibrillar protein-based edible coatings was tested on meat pieces (Cuq, 1996). The mechanical properties of meat specimens with and without edible coatings were assessed using cutting tests and were characterized by strength-deformation rheograms (Figure 8.6). The coatings slightly modified the texture profiles of the meat specimens, mainly by increasing mechanical resistance. Generally, mechanical resistance of the coated meat increased as coating thickness increased (Figure 8.6). The tested edible coatings seemed to be too resistant for such applications and would have to be weakened (Cuq, 1996).

Use of myofibrillar proteins for preparation of films and coatings by casting or thermomolding appears possible. Myofibrillar protein-based films are trans-

References

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parent and water-insoluble, and have poor water barrier ability, a slight fishy odor, and good mechanical properties. The moisture vapor and gas barrier properties of such films are similar to those of films from other proteins. Edible films and coatings containing myofibrillar proteins could thus be produced on a large scale as myofibrillar protein concentrates (i.e., surimis) are commercially available and could be used directly for preparing film-forming solutions. The high cost of the raw materials ($2–5/kg for surimi with 70–80% water content) should be noted especially in comparison to the low cost of vegetable proteins (e.g., $1/kg for wheat gluten with 10% water content). However, using underutilized fish species (e.g., some fatty fish species) or by-products of meat or fishing industries could significantly reduce the costs to around $1–3/kg. The principal applications that could be considered for myofibrillar protein-based films are related to coating meat or fish products. In such cases, using an edible coating derived from the same source as the coated food constitutes a particularly interesting characteristic. Applications may include protecting fish or meat pieces from oxidation or dehydration during storage or being used on processed meat or fish products (e.g., sausages and kamaboko) to replace currently used cellulose coatings or plastic films.

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230 Cuq, B., Aymard, C., Cuq, J.L. and Guilbert, S. 1995. “Edible Packaging Films Based on Fish Myofibrillar Proteins: Formulation and Functional Properties,” J. Food Sci., 60:1369–1374. Cuq, B., Gontard, N., Cuq, J.L. and Guilbert, S. 1996a. “Functional Properties of Myofibrillar Protein-Based Biopackagings as Affected by Film Thickness,” J. Food Sci., 61:580–584. Cuq, B., Gontard, N., Cuq, J.L. and Guilbert, S. 1996b. “Rheological Model for the Mechanical Properties of Myofibrillar Protein-Based Films,” J. Agric. Food Chem., 44:1116–1122. Cuq, B., Gontard, N., Aymard, C. and Guilbert, S. 1997a. “Relative Humidity and Temperature Effects on Mechanical and Water Vapor Barrier Properties of Myofibrillar Protein-Based Films,” Polymer Gels & Networks, 5:1–15. Cuq, B., Gontard, N., Cuq, J.L. and Guilbert, S. 1997b. “Selected Functional Properties of Fish Myofibrillar Protein-Based Films as Affected by Hydrophilic Plasticizers,” J. Agric. Food Chem., 45:622–626. Cuq, B., Gontard, N. and Guilbert, S. 1997c. “Thermal Properties of Fish Myofibrillar Protein-Based Films as Affected by Moisture Content,” Polymer, 38:2399–2405. Cuq, B., Gontard, N. and Guilbert, S. 1997d. “Thermoplastic Properties of Fish Myofibrillar Proteins: Application to the Biopackaging Fabrication,” Polymer, 38:4071–4078. Cuq, B., Gontard, N. and Guilbert, S. 1998a. “Proteins as Agricultural Polymers for Packaging Production,” Cereal Chem., 75:1–9. Cuq, B., Gontard, N., Cuq, J.L. and Guilbert, S. 1998b. “Packaging Films Based on Myofibrillar Proteins : Fabrication, Properties and Applications,” Nahrung/Food, 42:260–263. Fretheim, K., Egelandsdal, B., Harbitz, O. and Samejima, K. 1985. “Slow Lowering of pH Induces Gel Formation of Myosin,” Food Chem., 18:169–178. Fritz, H.G., Seidenstücker, T., Bölz, U., Juza, M., Schroeter, J. and Endres, H.J. 1994. Study on Production of Thermoplastics and Fibers Based Mainly on Biological Materials. Science Research Development, European Commission, EUR 16102 EN. Gennadios, A., Brandenburg, A.H., Weller, C.L. and Testin, R.F. 1993. “Effect of pH on Properties of Wheat Gluten and Soy Protein Isolate Films,” J. Agric. Food Chem., 41:1835–1839. Goll, D.E., Robson, R.M. and Stromer, M.H. 1977. “Muscle Proteins” in Food Proteins, eds., J.R. Whitaker and S.R. Tannebaum, Westport, CT: AVI Publishing Co., pp. 121–174. Gontard, N., Guilbert, S. and Cuq, J.L. 1992. “Edible Wheat Gluten Films: Influence of the Main Process Variables on Film Properties Using Response Surface Methodology,” J. Food Sci., 57:190–195, 199. Gontard, N., Duchez, C., Cuq, J.L. and Guilbert, S. 1994. “Edible Composite Films of Wheat Gluten and Lipids: Water Vapour Permeability and Other Physical Properties,” Int. J. Food Sci. Technol., 29:39–50. Gontard, N., Thibault, R., Cuq, B. and Guilbert, S. 1996. “Influence of Relative Humidity and Film Composition on Oxygen and Carbon Dioxide Permeabilities of Edible Films,” J. Agric. Food Chem., 44:1064–1069. Herald, T.J., Hachmeister, K.A., Huang, S. and Bowers, J.R. 1996. Corn zein packaging materials for cooked turkey. J. Food Sci., 61:415–417, 421. Hermansson, A.M., Harbitz, O. and Langton, M. 1986. “Formation of Two Types of Gels from Bovine Myosin,” J. Sci. Food Agric., 37:69–84. Joseph, D., Lanier, T.C. and Hamann, D.D. 1994. “Temperature and pH Affect Transglutaminase-catalyzed ‘Setting’ of Crude Fish Actomyosin,” J. Food Sci., 59:1018–1023, 1036. Kamat, G.G., Lanier, T.C., Foegeding, E.A. and Hamann, D.D. 1992. “Non-disulfide Covalent Cross-Linking of Myosin Heavy Chain in ‘Setting’ of Alaska Pollock and Atlantic Croaker Surimi,” J. Food Biochem., 16:151–172. Kim, S.H., Carpenter, J.A., Lanier, T.C. and Wicker, L. 1993. “Polymerization of Beef Actomyosin Induced by Transglutaminase,” J. Food Sci., 58:473–474, 491. Lanier, T.C. 1986. “Functional Properties of Surimi,” Food Technol., 40(3):107–114, 124.

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Formation and Properties of Egg White Films and Coatings

N recent years, there has been increased interest in edible film research, partic-

I

ularly in the development of free-standing films and coatings derived from proteins (Gennadios et al., 1994). The driving forces behind this interest can be attributed to elevated demands for high-quality food products, scientific advances in new functionalities of food components, and increased public concern for environmental conservation and protection (Chen, 1995; Miller and Krochta, 1997). The promise of edible protein materials in food packaging applications arises from their capability to supplement, and possibly improve, the performance of existing synthetic packaging polymers while reducing environmental impacts (Krochta and De Mulder-Johnston, 1997). Egg white is a complex protein system made up of a solution of globular proteins containing ovomucin fibers (Woodward, 1990). Major protein components of egg white are summarized in Table 9.1. Ovalbumin, which constitutes more than half of egg white protein by weight, is the only fraction that contains free sulfhydryl (SH) groups. Other proteins, such as ovotransferrin, ovomucoid, and lyzozyme contain disulfide (S-S) bonds (Mine, 1995). The S-S bonds are considered important in film formation for proteins containing cysteine and/or cystine amino acids (Okamoto, 1978; Gennadios et al., 1994). Egg yolk has a greater number of applications in the food industry as compared to egg white, leading to a surplus of egg albumen in the egg-breaking in233

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dustry of North America (Gennadios et al., 1996a). In an effort to develop further uses for this surplus product, several studies have explored the feasibility of using egg white as the base material for forming edible films and coatings. This chapter provides an overview of formation methods and properties of films and coatings based on egg white protein.

Most methods for preparing egg white protein films involve denaturation of egg white protein in aqueous solution, followed by a casting process whereby the film-forming solution is spread on a level surface and then dried to produce films. Gennadios et al. (1996b) prepared films from alkaline (pH 11.25) aqueous egg albumen solutions (9 g/100 mL water) containing glycerol (30, 40, or 50% w/w of protein), polyethylene glycol (PEG, 50, or 60% w/w of protein), or sorbitol (50 or 60% w/w of protein) as film plasticizers. The egg white solutions were heated at 45 C for 20 minutes, cast on leveled Teflon -coated glass plates, and then allowed to solidify under ambient conditions for five to six hours before placing them in an environmental chamber to complete the drying process at 25 C and 50% relative humidity (RH). Gennadios et al. (1996b) reported that a pH range of 10.5 to 12.0 was necessary to obtain smooth and homogeneous egg white films. A similar pH range (10.5 to 11.5) was suggested by Okamoto (1978) for film formation on the surface of heated egg white solutions (similar to film formation on the surface of heated soymilk). Other proteins that require alkaline conditions for optimal film formation process include peanut, wheat, and fish myofibrillar proteins (Okamoto, 1978). For egg white protein, Gennadios et al. (1996b) noted that pH above 12 should be avoided as the

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film-forming solutions would become too viscous, or would coagulate during thermal treatment such that the solutions could not be easily cast. On the other extreme, although self-supporting films could be obtained with solution pH as low as 3, the surfaces of such films were rough and gritty due to insoluble protein particles. Recently, Handa et al. (1999) utilized a similar technique for preparing egg white films. In this study, spray-dried egg white (9% w/v), silicone oil (0.1% w/w of egg white), and PEG 400 (60% w/w of egg white) were dissolved in distilled water, with silicone oil acting as an anti-foaming agent. The solution was adjusted to pH 10.5 to 11.5, using 2N NaOH, heated at 40 C for 30 minutes, and then was cast on a level surface to form a film. In general, the mechanism of film formation is hypothesized to involve inter- and intramolecular S-S bonds. At alkaline pH, the S-S bonds in protein molecules are reduced to SH groups, thereby facilitating dispersion of the protein. The heating treatment further unfolds the protein chains, exposing more SH and hydrophobic groups. It has been suggested that, during the gelation and drying steps, SH groups are converted to inter- and intramolecular S-S covalent cross-links through oxidation and sulfhydryl-disulfide interchange reactions (Mine, 1992; Gennadios et al., 1996b). This causes the formation of three-dimensional networks. In addition to S-S bonding, hydrogen bonding as well as hydrophobic and electrostatic interactions are important in dictating the physical properties of egg white films.

Functional properties of protein films can be improved by cross-linking through physical, chemical, or enzymatic treatments. Transglutaminase (TGase, protein-glutamine -glutamyltransferase, EC 2.3.2.13) is an enzyme that catalyzes acyl-transfer reactions in proteins resulting in formation of -( -glutaminyl)lysine inter- and intramolecular cross-links (Figure 9.1) (Nielsen, 1995). While flexible proteins (e.g., casein and gelatin) are good substrates for this enzyme (Nio et al., 1985, 1986; Sakamoto et al., 1994; Lim et al., 1999), globular proteins (e.g., -lactoglobulin, -lactalbumin, and bovine serum albumin) are poor substrates when they are in their native state. The compact structure of these globular proteins limits the accessibility of TGase to the target glutamine and lysine residues (Motoki and Nio, 1983; Mahmoud and Savello, 1992; Dickinson and Yamamoto, 1996; Matsumura et al., 1996). Susceptibility to the TGase-catalyzed cross-linking reaction can be improved by cleaving S-S bonds and unfolding the globular protein with protein denaturants such as dithiothreitol or 2-mercaptoethanol (Mahmoud and Savello, 1992, 1993; Nielsen, 1995; Færgemand et al., 1997; Yildirim and Hettiarachchy, 1998).

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Figure 9.1 Transglutaminase-catalyzed cross-linking reaction in proteins.

TGase is suitable for cross-linking various proteins, including s1-casein (Motoki et al., 1987), -lactalbumin and -lactoglobulin (Mahmoud and Savello, 1992, 1993), and whey protein isolate and soybean 11S globulin (Yildirim and Hettiarachchy, 1998) to form biopolymer films. In these referenced studies, dithiothreitol was used as a denaturant to promote the enzymatic cross-linking reaction. However, classifying films treated with dithiothreitol as “edible” is highly questionable due to the toxicity of this compound (Nielsen, 1995). Because of their globular nature, egg white proteins are not readily polymerizable by TGase without a denaturation step. To circumvent the dithiothreitol toxicity issue, Lim et al. (1998) employed a combined alkaline and thermal treatment to denature the egg white. In their study, dialyzed, freeze-dried egg white protein powder was dissolved in an alkaline solution (pH 10.5, using NaOH) and was thermally treated at 80 C for 20 minutes before the enzymatic (TGase) treatment. Thermal denaturation of egg white proteins without forming coagula is possible at low ionic strength and at pH levels distinct from pI, due to the electrostatic repulsion forces that hinder the formation of random aggregates (Doi et al., 1994). Under these conditions, the proteins are partially unfolded and are more flexible than in their native form (Mine et al., 1990). After thermal treatment, the egg white solution was adjusted to pH 8.2 using 0.1 N HCl, and TGase (0.003%) was added to allow for the enzymatic cross-linking reaction of the egg white protein. To form films, the egg white protein solution was poured onto a leveled glass plate fitted with a rim around the edge and incubated at 50 C for eight hours, which allowed formation of a layer of translucent gel. The casting tray was covered with a lid during incubation to prevent drying. Due to the high RH of the headspace above the

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gel-forming solution, condensate droplets tended to form on the inner side of the lid and drip onto the gelling solution, resulting in films with uneven surfaces. To overcome this problem, a fiberglass screen was fitted underneath the glass lid to entrap the condensate droplets (Figure 9.2). Subsequent drying of the gel at the same temperature for 12–14 hours with the lid removed produced transparent, colorless egg white protein films. In contrast to -lactoglobulin, which has been shown by Færgemand et al. (1997) to be polymerizable by TGase at elevated pH conditions (pH 8.5–9.0), high pH alone would not induce sufficient conformational changes in egg white proteins for the TGase cross-linking reaction to occur. Electrophoretic analysis by Lim et al. (1998) detected only minimal amounts of aggregates when egg white protein samples were pre-heated at 60 C (Figure 9.3). However, when the pre-heating temperature increased to 80 C, the aggregate bands became progressively more intensified. The formation of aggregates was accompanied by reductions in band intensities of the constituent proteins of the egg

Figure 9.2 Schematic diagram of a casting tray for preparing TGase cross-linked egg white films.

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Figure 9.3 SDS-PAGE patterns of egg white proteins subjected to pre-heating treatments at various temperatures (pH 10.5 for 20 min), followed by incubation in the presence of TGase (pH 8.2, 50 C). A, B, C, D, and E represent samples incubated for 0, 30, 60, 120, or 300 min, respectively. Identified bands: 1 polymerized protein; 2 ovotransferrin; 3 ovalbumin; 4 ovomucoid; 5 lysozyme (and hydrolytic products for samples subjected to 80 C treatment). Reprinted with permission from Lim et al., 1998. Copyright 1998, American Chemical Society.

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white. Because the buffer systems used for electrophoresis included 2-mercaptoethanol (which cleaves S-S bonds), this reduction in band intensities provided evidence for the formation of higher molecular weight polymers through TGase-induced cross-linking, rather than to intermolecular S-S bonds between protein chains. The gelation process appeared to be essential for forming cohesive egg white protein films (Lim et al., 1998). Without enzymatic treatment, the protein solutions remained in a liquid state even after prolonged periods of incubation. Subsequent drying of the solutions resulted in fragile films that readily cracked when peeled from the casting tray. In contrast, films produced from egg white protein solutions that went through the gelation process remained intact upon peeling from the casting surface and showed good integrity.

Another cross-linking agent that merits discussion is dialdehyde starch (DAS), a polymeric aldehyde obtained by reacting native starch with periodic acid (molecular weight 300,000–5,000,000). Aldehydes such as formaldehyde, glutaraldehyde, and glyoxal promote inter- and intramolecular cross-linking in proteins (Habeeb and Hiramoto, 1968). However, due to the inherent toxicity of these aldehydes, their use in improving the physical properties of protein films is limited. In contrast, DAS has shown low toxicity to rats by oral, dermal, and respiratory routes (Wilson, 1959). Gennadios et al. (1998a) reported that DAS could be used as an effective cross-linking agent for polymerizing egg white during film preparation. In their study, egg white films were prepared as described previously (alkaline treatment and heating method; Gennadios et al., 1996b) except that DAS was added after a pH adjustment to 11.25, followed by heating at 40 C for 20 minutes. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) patterns, which displayed increased aggregate band intensity with increasing amounts of DAS, indicated the formation of DAS-induced cross-links in the egg white protein. The formation of cross-links was also reflected by a 15% reduction in total soluble matter in distilled water, as well as 66 and 75% reductions in protein solubility after immersing the film in buffer solutions containing urea/2-mercaptoethanol and urea, respectively. Because urea disrupts hydrophobic interactions and hydrogen bonds, while 2-mercaptoethanol reduces S-S bonds, the increased resistance to solubilization was attributed to formation of other types of intermolecular cross-links induced by DAS. DAS cross-linked to the egg white proteins may also act as an effective internal plasticizer. Potentially, egg white films exhibiting different degrees of flexibility and stretchability may be achieved by cross-linking egg white with DAS of different molecular weights.

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It has long been known that physical properties of synthetic polymers can be improved through ionizing radiation (Pascat, 1986). In proteins, ultraviolet (UV) radiation is absorbed by aromatic amino acid residues resulting in conformational modifications and, if the energy levels are high enough, in cleavage of S-S bonds (Cheftel et al., 1985). Individual proteins would be expected to exhibit different responses to UV radiation because of the varying amino acid compositions and molecular structures. Rhim et al. (1999) investigated the mechanical and barrier properties of UV-irradiated protein films derived from egg albumen and other proteins (wheat gluten, corn zein, and sodium caseinate). Film specimens, prepared according to the method used by Gennadios et al. (1996b), were treated with 253.7 nm UV radiation for 24 h (51.8 J/m2 radiation dosage). Egg albumen, wheat gluten, and sodium caseinate films all increased in yellowness as a result of UV exposure. The egg albumen films had slightly but significantly (P < 0.05) lower mean total soluble matter contents than untreated films, which was attributed to formation of cross-links in the UV-treated films. Rhim et al. (1999) suggested that the UV-induced cross-linking in these films involved aromatic amino acids, such as phenylalanine and tyrosine, present in egg white proteins.

Coatings may be differentiated from free-standing films in that they are applied directly onto the surface of food products. The formation of protective coatings in situ is the main advantage of coatings versus pre-formed films, in that the coating can adapt itself to the multiple shapes and surfaces of a product. Accordingly, good interfacial adhesion between the protective layer and surface of the item being coated is imperative to ensure proper surface coverage. Wong et al. (1996) evaluated several proteins (i.e., egg albumen, soy protein isolate, wheat gluten, and corn zein) as coating materials for reducing quality loss in hen shell eggs. Egg albumen coating solutions were prepared as described by Gennadios et al. (1996b). Eggs were then immersed in the protein solutions at room temperature for one minute and were allowed to dry at room temperature. The coatings reduced moisture loss and increased shell strength. Bolin (1976) reported that egg albumen coatings notably retarded moisture loss from raisins in cereal/raisin mixtures. In that study, protein coatings were applied by dipping the raisins in egg white protein solutions (40% w/v), allowing them to drain for one minute, and then heating the coated raisins in a dehydrating oven at 115 C for four to five minutes to denature the egg white protein. The heat denaturation step was essential to remove excess water from the coating and to provide a rigid barrier structure (Bolin, 1976).

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Several patents discussed methods for coating food products using egg white protein. Reutimann et al. (1996) described a formulation and process for coating meat products to reduce fluid losses after cooking. The coating mixture consisted of egg albumen (at least 5%), milk protein (5%), unmodified starch (25%), and water (40%). The egg albumen and milk protein components provided the primary fluid retention function of the coating while the starch absorbed any moisture escaping from the coated food. The starch also contributed elasticity to prevent the coating from cracking. Heating the coating mixture (preferably at 70 C) was necessary to achieve optimal fluid retention properties via coagulation, denaturation, and gelation of the albumen, milk protein, and starch fractions, respectively. Berberat and Wissgott (1993) reduced moisture transfer among components in food products using a coating prepared by mixing egg white (3–12%), milk protein (3–18%), and pre-gelatinized starch (7–12%) in water. A thin layer of this suspension was applied to a pizza dough surface by brushing or spraying, followed by heat treatment (15 to 120 s at 70 to 100 C) to solidify the layer. After the heat treatment, the moist pizza topping components (e.g., ham, cheese, vegetables, and tomato sauce) were placed on this barrier layer on the dough. They emphasized that the heat treatment was critical to the development of desirable hydrophobic and impermeable characteristics of the coating (Berberat and Wissgott, 1993).

Due to the inherent hydrophilicity of most proteins, films derived from these proteins are moisture-sensitive. At low RH conditions, protein films are brittle and susceptible to cracking due to the strong cohesive energy density of the biopolymer. To overcome this problem, compatible plasticizers (e.g., polyols) are added to impart flexibility and stretchability to the films for practical end-use applications. Because polyols are hydrophilic, their addition as plasticizers increases the hydrophilicity of the protein films. Moisture sorption isotherms of TGase cross-linked egg white films, as affected by plasticizer content, were determined by Lim et al. (1998) (Figure 9.4). At any given water activity (aw), equilibrium moisture contents were higher for films containing higher glycerol contents, which was attributed to the additional polar hydroxyl groups introduced by glycerol, resulting in films with higher hydrophilicity. Moisture sensitivity of the films was also reflected by changes in tensile strength (TS) and elongation at break (E) values with RH. Increasing glycerol content and RH notably decreased TS of the TGase cross-linked egg white films (Figure 9.5), indicating that both water and glycerol were capable of plasticizing the cross-linked egg

Figure 9.4 Moisture sorption isotherms for TGase cross-linked egg white films as affected by temperature and glycerol content. Reprinted with permission from Lim et al., 1998. Copyright 1998, American Chemical Society.

Figure 9.5 Ultimate tensile strength of TGase cross-linked egg white films as affected by relative humidity and glycerol content at 22 C. Reprinted with permission from Lim et al., 1998. Copyright 1998, American Chemical Society.

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white protein films. McHugh and Krochta (1994) reported similar effects of plasticizers on mechanical properties of whey protein films. For TGase cross-linked egg white films, TS values were higher and more sensitive to RH variation when the plasticizer content of the film was low, but became weaker and less sensitive to RH at higher glycerol levels (Lim et al., 1998). At RH above 70%, films were weak (TS < 2 MPa) with minimal change in TS, regardless of the amount of glycerol incorporated, due to extensive plasticization of the polymer matrix caused by the sorbed water. Hydrogen bonds are considered important in contributing to the TS of protein films. Plasticizers appear to function by masking polar sites along polymer molecules, thereby reducing interactions between adjacent molecular chains (Paton, 1972; Meier, 1990). As plasticizers “force” the chains further apart, the freedom of movement of the chains is increased, and van der Waals forces between the chains are reduced (Coulson, 1972), thus weakening the physical strength of the polymer. Plasticizers can also be envisioned as “lubricants” that exist between the polymer chains, allowing the neighboring molecules to slide past each other more readily, thus increasing the extensibility of the polymer. In general, egg white films with higher glycerol contents have greater E values (Figure 9.6) (Lim et al., 1998).

Figure 9.6 Elongation at break of TGase cross-linked egg white films as affected by relative humidity and glycerol content at 22 C. Reprinted with permission from Lim et al., 1998. Copyright 1998, American Chemical Society.

244 Both the level of plasticizer and the type of plasticizer added during film formation can affect the mechanical properties of egg albumen and other protein films (McHugh and Krochta, 1994). Gennadios et al. (1996b) showed that regardless of the added plasticizer (glycerol, sorbitol, or PEG 400), TS decreased and E increased with increasing plasticizer content in egg white films (Table 9.2). Moreover, PEG 400 gave films with greater TS and greater E than films obtained using glycerol or sorbitol. Higher amounts of sorbitol than glycerol were required for films to achieve similar tensile properties (Gennadios et al., 1996b). The smaller size of the glycerol molecule likely enabled it to influence film mechanical properties more readily than sorbitol. Ultimate TS represents the maximal tensile stress a film can sustain before breaking, while E reflects the extensibility of the material. A stand-alone film should possess not only reasonably high TS, but also moderate E to stretch and comply with points of load application. Although stronger films would be expected when lower amounts of glycerol are used (Figure 9.5), the corresponding films would possess very poor flexibility (Figure 9.6), which would limit their packaging applications. Under these conditions, samples tended to crack or fracture when subjected to stress. In view of the water-sensitive nature of egg white films, RH conditions during the end-use application should also be considered to better predict the performance of such materials.

In addition to the effects of moisture and plasticizers, different film formation methodologies can result in egg white films of varying mechanical properties. Rhim et al. (1999) demonstrated an increase in mean TS from 1.7 MPa to 2.9 MPa for UV-irradiated egg albumen films compared to untreated films. In contrast, E values for protein films tend to decrease with increasing UV irradia-


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tion, as shown by Gennadios et al. (1998b) who reported a linear decrease in E for soy protein films with UV dosage. Table 9.3 summarizes reported mechanical properties of several protein films. Although direct comparisons among film samples was difficult due to different test conditions and substrates used, egg white films compared favorably with other protein films. Moreover, TGase cross-linked egg white films appeared to have improved mechanical properties compared to egg white films prepared using an alkaline/thermal method. Under the same test conditions (protein:glycerol 1:0.5, 50% RH), TGase cross-linked films had much higher TS and E values (4.4 MPa TS and 74% E; Lim et al., 1998) than untreated films (1.3 MPa TS and 32% E; Gennadios et al., 1996b). The improved mechanical properties of the TGase-treated films were likely due to formation of covalent -( -glutaminyl)lysine cross-links between the protein molecules. Other cross-linking agents, such as DAS, can be used to effectively enhance the mechanical properties of egg white films. Gennadios et al. (1998a) reported that TS and E increased significantly (P < 0.05) with increasing amounts of DAS added to egg white films (optimal DAS level at 5% w/w of egg white), suggesting the formation of covalent cross-links between egg white proteins and DAS. Usually, an increase in TS of protein films due to cross-linking tends to reduce film extensibility (i.e., lower E values); however, this effect was not observed by Gennadios et al. (1998a). These authors suggested that cross-links involving DAS, which is a long chain biopolymer, are unlike cross-links involving low molecular weight compounds that increase rigidity and “tightness” of a film matrix. Instead, by “fixing” the macromolecular DAS chains within the protein film structure, the flexibility of the polymer matrix is not severely restricted. However, in spite of exhibiting improved mechanical properties, DAS cross-linked films tend to develop a yellow hue, which may limit applications where colorless films are required.

Exposing an egg white protein solution to heat under alkaline conditions is essential for forming films with good physical integrity. Surface SH group concentration is reported to increase significantly with increasing pH. Moreover, at a given alkaline pH, heated egg white film-forming solutions had greater concentrations of surface SH groups than unheated solutions (Table 9.4) (Handa et al., 1999). This effect was previously attributed to reduction of S-S bonds, as well as to unfolding of previously “buried” SH groups of the egg white protein (Beveridge and Arntfield, 1979; Mine et al., 1990; Mine, 1992). It is noteworthy that thermal and alkaline treatments appeared to exert a synergistic effect in increasing the surface SH concentration of the egg white protein solutions (Table 9.4); a greater difference in surface SH concentration was detected between heated and unheated protein solutions as pH increased (Handa et al., 1999).

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The increased surface SH concentration is important for egg white film formation, because, upon exposure to air, the SH groups are converted to intermolecular S-S bonds through oxidation and sulfhydryl-disulfide interchange reactions (Hayakawa and Nakai, 1985; Mine, 1992). The number of surface SH groups formed through thermal and alkaline denaturation will determine the number of intermolecular S-S cross-links in the cast egg white films and their mechanical properties. Handa et al. (1999) correlated the mechanical properties of egg white films with the surface concentrations of SH groups in egg white film-forming solutions. Films cast from heated solutions had slightly greater TS than those from unheated solutions, but in most cases film TS did not increase with pH. Conversely, films prepared from heated solutions had greater E than unheated solutions only at the most alkaline test condition (pH 11.5), but film E values increased with pH in most cases. Concentration of surface SH groups correlated positively with E (P < 0.05, r 0.86). It was concluded that the increase in surface SH groups due to heat and alkaline treatments resulted in increased S-S bonding in the egg white films, which rendered the films more stretchable (Handa et al., 1999).

Due to the inherent hydrophilic nature of proteins, films derived from proteins are poor water vapor barriers. To improve this inadequacy, various techniques have been used to reduce the moisture sensitivity of protein films. More than three decades ago, Durst (1967) described a method of preparing protec-

248 tive coatings made up of egg albumen and sodium caseinate dispersed in immiscible oil to reduce the hydrophilicity of protein film matrices. Gennadios et al. (1996b) reported that the mean water vapor permeability (WVP) of egg white films containing egg yolk solids (50% w/w of total egg solids) decreased from 6.21 (no egg yolk) to 5.62 g mm/m2 h kPa. It was suggested that the hydrophobic egg yolk lipids (32–36% on a wet basis in egg yolk) were responsible for the observed reduction in WVP. O2 permeability coefficients of TGase cross-linked egg white protein films have been shown to increase substantially with increasing RH (Figure 9.7) (Lim et al., 1998). Similar observations on the effect of RH on O2 permeability were reported for other protein-based films (McHugh and Krochta, 1994; Gontard et al., 1996; Krochta and De Mulder-Johnston, 1997; Miller and Krochta, 1997). Lower O2 permeability values were detected by Lim et al. (1998) for films containing a lower amount of glycerol. Such films also exhibited less sensitivity to RH below intermediate RH levels compared to films with higher glycerol content. However, as RH approached saturation, permeability values of the glycerol-plasticized films became similar, regardless of the amount of glycerol incorporated. The increase in O2 permeability of egg white films as a function of RH is related to structural plasticization of the protein matrix caused by the sorbed water. Water is believed to increase the free volume of the polymer system, causing a drop in the glass transition temperature (Tg), the temperature above which the macromolecules would possess sufficient energy to allow large-scale vibrational motion and undergo a glassy-to-rubbery state transition (Tager, 1978). Permeation of gases can be considered a two-step process: the thermodynamic dissolution of a permeant in the matrix and the kinetic diffusion of the permeant through the matrix; that is, P DS, where P, D, and S are permeability, diffusivity, and solubility coefficients, respectively (Robertson, 1993). The increase in polymer segmental movement due to the plasticization effects of water would increase the diffusivity of the permeant molecules. Moreover, formation of larger water clusters may also favor the solubility of oxygen (Gavara and Hernandez, 1994; Gontard et al., 1996), resulting in an overall increase in O2 permeability. When O2 permeability of egg white films was plotted against RH (Figure 9.7A), changes in slope were observed near the 50% RH region, particularly for the 35% glycerol films. According to free volume theory, the magnitude of Tg depression would depend upon the volume fraction of the plasticizer in the polymer (Cowie, 1991). Below 50% RH, with lower amounts of water sorbed, the polymer is assumed to exist in a glassy state, i.e., with the test temperature < Tg. The limited segmental movements of the polymer would thus reduce the influence of RH upon O2 permeability. Accordingly, if the test temperature were elevated, increased segmental motion would be expected to favor diffu-

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Figure 9.7 Oxygen permeability coefficients for TGase cross-linked egg white protein films with 35 and 45% glycerol content, as related to relative humidity and temperature. Symbols represent means of two repeated tests. Note logarithmic scale on ordinates. Reprinted with permission from Lim et al., 1998. Copyright 1998, American Chemical Society.

sion. This behavior is evident in Figure 9.7A, where the change in slope of permeability versus RH plots became less prominent with increased temperature. Furthermore, it could be concluded that the less conspicuous changes in slope for the 45% glycerol samples were caused by more extensive plasticization of the protein films (T < Tg) by the additional glycerol.

Although egg white films can be formed at a pH range from 10.5 to 12, in combination with thermal treatments, the sensory qualities of the resulting alkaline films may be undesirable if the films are intended for edible packaging applications. Alternatively, TGase may be used for polymerizing the egg white protein if high pH is undesirable. A combined high pH and heat treatment is important for egg white film formation. This treatment is also effective for enhancing the susceptibility of egg white proteins to the cross-linking reaction catalyzed by TGase and may be used for denaturing other globular proteins to achieve similar enzymatic polymerization. Varying the amount of added plasticizers can form egg white films with a spectrum of mechanical and oxygen barrier properties. In view of the water-sensitive nature of these films, RH conditions during end-use applications should be taken into account during film development. The potential of egg white films and coatings for commercial applications to extend the shelf life of food products appears promising.

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Krochta, J.M. and De Mulder-Johnston, C.D. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Lieberman, E.R. and Gilbert, S.G. 1973. “Gas Permeation of Collagen Films as Affected by Cross-Linkage, Moisture, and Plasticizer Content,” J. Polym. Sci.: Symposium, 41:33–43. Lim, L.-T., Mine, Y. and Tung, M.A. 1998. “Transglutaminase Cross-Linked Egg White Protein Films: Tensile Properties and Oxygen Permeability,” J. Agric. Food Chem., 46:4022–4029. Lim, L.-T., Mine, Y. and Tung, M.A. 1999. “Barrier and Tensile Properties of Transglutaminase Cross-Linked Gelatin Films as Affected by Relative Humidity, Temperature, and Glycerol Content,” J. Food Sci., 64:616–622. Mahmoud, R. and Savello, P.A. 1992. “Mechanical Properties and Water Vapor Transferability through Whey Protein Films,” J. Dairy Sci., 75:942–946. Mahmoud, R. and Savello, P.A. 1993. “Solubility and Hydrolyzability of Films Produced by Transglutaminase Catalytic Cross-Linking of Whey Protein,” J. Dairy Sci., 76:29–35. Matsumura, Y., Chanyongvorakul, Y., Kumazawa, Y., Ohtsuka, T. and Mori, T. 1996. “Enhanced Susceptibility to Transglutaminase Reaction of a-Lactalbumin in the Molten Globule State,” Biochim. Biophys. Acta, 1292:69–76. McHugh, T.H. and Krochta, J.M. 1994. “Sorbitol- vs Glycerol-Plasticized Whey Protein Edible Films: Integrated Oxygen Permeability and Tensile Property Evaluation,” J. Agric. Food Chem., 42:841–845. McHugh, T.H., Aujard, J-.F. and Krochta, J.M. 1994. “Plasticized Whey Protein Edible Films: Water Vapor Permeability Properties,” J. Food Sci., 59:416–419, 423. Meier, L. 1990. “Plasticizers” in Plastics Additives, 8th Edition, eds., R. Gächter and H. Müller, New York: Hanser Publishers, pp. 327–420. Miller, K.S. and Krochta, J.M. 1997. “Oxygen and Aroma Barrier Properties of Edible Films: A Review,” Trends Food Sci. Technol., 8:228–237. Mine, Y. 1992. “Sulfhydryl Groups Changes in Heat-Induced Soluble Egg White Aggregates in Relation to Molecular Size,” J. Food Sci., 58:254–255. Mine, Y. 1995. “Recent Advances in the Understanding of Egg White Protein Functionality,” Trends Food Sci. Technol., 6:225–232. Mine, Y., Noutomi, T. and Haga, N. 1990. “Thermally Induced Changes in Egg White Proteins,” J. Agric. Food Chem., 38:2122–2125. Motoki, M. and Nio, N. 1983. “Cross-Linking between Different Food Proteins by Transglutaminase,” J. Food Sci., 48:561–566. Motoki, M., Aso, H., Seguro, K. and Nio, N. 1987. “ s1-Casein Film Prepared Using Transglutaminase,” Agric. Biol. Chem., 51:993–996. Nielsen, P.M. 1995. “Reactions and Potential Industrial Applications of Transglutaminase. Review of Literature and Patents,” Food Biotechnol., 9:119–156. Nio, N., Motoki, M. and Takinami, K. 1985. “Gelation of Casein and Soybean Globulins by Transglutaminase,” Agric. Biol. Chem., 49:2283–2286. Nio, N., Motoki, M. and Takinami, K. 1986. “Gelation Mechanism of Protein Solution by Transglutaminase,” Agric Biol. Chem., 50:851–855. Okamoto, S. 1978. “Factors Affecting Protein Film Formation,” Cereal Foods World, 23:256–262. Pascat, B. 1986. “Study of Some Factors Affecting Permeability” in Food Packaging and Preservation. Theory and Practice, ed., M. Mathlouthi, London: Elsevier Applied Science, pp. 7–24. Paton, C. 1972. “Theories of Plasticizer Action” in Plasticizers, Stabilizers and Fillers, eds., P.D. Ritchie, S.W. Critchley and A. Hill, London: Iliffe Books Ltd., pp. 39–49. Reutimann, E.J., Vadehra, D.V. and Wedral, E.R. October 22, 1996. U.S. patent 5,567,453. Rhim, J.W., Gennadios, A., Fu, D., Weller, C.L. and Hanna, M.A. 1999. “Properties of Ultraviolet Irradiated Protein Films,” Lebensm. Wiss. Technol., 32:129–133. Robertson, G.L. 1993. Food Packaging: Principles and Practice. New York: Marcel Dekker, p. 95. Sakamoto, H., Kumazawa, Y. and Motoki, M. 1994. “Strength of Protein Gels Prepared with Microbial Transglutaminase as Related to Reaction Conditions,” J. Food Sci., 59:866–871.

252 Tager, A. 1978. Physical Chemistry of Polymers. Moscow, Russia: Mir Publishers, pp. 559–560. Wilson, R.H. 1959. “Utilization and Toxicity of Dialdehyde- and Dicarboxyl-Starches,” Proc. Soc. Exp. Biol. Med., 102:735–737. Wong, Y.C., Herald, T.J. and Hachmeister, K.A. 1996. “Evaluation of Mechanical and Barrier Properties of Protein Coatings on Shell Eggs,” Poultry Sci., 75:417–422. Woodward, S.A. 1990. “Egg Protein Gels” in Food Gels, ed., P. Harris, Essex, England: Elsevier Science Publishers Ltd., pp. 175–199. Yildirim, M. and Hettiarachchy, N.S. 1998. “Properties of Films Produced by Cross-Linking Whey Proteins and 11S Globulin Using Transglutaminase,” J. Food Sci., 63:248–252.

Formation and Properties of Wool Keratin Films and Coatings

K

ERATINS are proteins found in hair, wool, feathers, nail, horns, and other ep-

ithelial coverings (MacLaren and Milligan, 1981; Feughelman, 1985; Fraser et al., 1986; Husken, 1986; Heid et al., 1986). In wool, keratins are main constituents of long cortical cells (Figure 10.1) and weigh 30–60% of the total fiber. Keratins are distinguished from collagen, silk fibroin, and other structural proteins by their high content of cysteine and half-cystine residues (7–20 mol% of all amino acid residues). The ratio of cysteine to half-cystine in wool is about 0.2 to 0.8 (Arai et al., 1996). Because the aforementioned biological materials are constructed from keratins interconnected three-dimensionally through cystine disulfide (S-S) bonds, they may be disassembled into basic fragments by chemically reducing S-S bonds to sulfhydryl (SH) groups (of cysteine residues). Then, SH groups in the blocks may be re-associated by oxidative S-S bond formation to produce new polymeric proteinaceous materials, which may be useful as biomaterials. This recycling reduction/oxidation approach is not feasible with proteins, such as collagen and silk fibroin, that lack cysteine and cystine residues. Here, we discuss keratin films produced by oxidation of a reduced form of keratins (referred to hereafter as “reduced keratins”). This chapter deals mainly with the preparation and characteristics of the reduced keratins and the properties and potential uses of their films. 253

254

Figure 10.1 Schematic representation of a wool hair fiber showing the major cellular components.

Figure 10.2 displays an amino acid analysis of keratins extracted from wool, where amino acids are arranged from the left in the order they were eluted from an anion-exchange resin. The amino acid analysis was conducted after completely hydrolyzing the reduced keratins in 6 N hydrochloric acid at 120°C for 6 h. Under these conditions, asparagine (Asn) and glutamine (Gln) were converted to aspartic acid (Asp) and glutamic acid (Glu), respectively. Asn/Asp and Gln/Glu ratios of 0.96–1.86 and 0.75–0.93, respectively, have been reported for Merino 64s wool (MacLaren and Milligan, 1981). Keratins consist of a wide variety of amino acids (Figure 10.2) and, hence, should react with various chemical reagents. Also, the first half series of amino acid residues is more abundant than the later half (Figure 10.2), resulting in keratins possessing a low isoelectric point (pI) of 5–6. Therefore, keratins have anionic properties at around neutral pH. Keratins also are unique in their location of cysteine and/or half-cystine residues. In the case of component 8c-1 of wool keratin microfibrils (Figure 10.3), most S-containing amino acid residues are localized in both amino- and carboxy acid-terminal regions (Dowling et al., 1986). Keratin fibrils may thus be considered as polymers, which are formed by intra- and intermolecular oxidative S-S bonds of cysteine residues between the

Molecular Characteristics of Keratins

255

Figure 10.2 Amino acid residues in reduced keratins (prepared by method a) and in keratin films (amino acid residues are arranged from the left in the order they were eluted from an anion exchange resin; glutamic acid and aspartic acid residues include glutamine and asparagine, respectively; cysteine residues were quantified upon conversion into the S-carboxymethylated [CmCys] derivative).

monomeric keratins. Thus, one may draw a simple model for reduced keratin as a proteinaceous monomer having many reactive SH groups in both ends. Although amino acid sequences of matrix keratins have not been studied, they contain even more cysteine/half-cystine residues (13–25 mol%) than microfibril keratins (5–10 mol%). Hair is similar to wool in structure. A classification of keratins is listed in Table 10.1.

Figure 10.3 Distribution of half-cystine in component 8c-1 of wool keratins. The strings and the broad solid bars are non-helical and helical regions, respectively. Most cysteine/half-cystine residues are localized in the amino (N)- and carboxy acid (C)- terminals.

256

Several keratin extraction methods have been reported. Rhodes et al. (1967) described an oxidative fission of S-S bonds in keratins with performic acid or peracetic acid to obtain the water-soluble proteins (keratoses) that possess [ NH CH( CH2 SO3 ) CO ] residues of cysteic acid in place of cystine residues. Similar to the cupra-rayon producing process, Harrap and Woods (1964) dissolved keratins in a solution of bisulfite and cupric ammonia to furnish the Bunte’s salt [ NH CH( CH2 S SO3 Na + ) CO ] of keratins. Reportedly, the cupric ions associated with amide bonds, thus disrupting hydrogen bonding among keratin fibrils. Oxidative sulfitolysis to obtain the Bunte’s salt of keratins by a combination of NaHSO3 and Na2S2O4 and subsequent conversion of the S-sulfo groups into free SH groups by a treatment with 2-mercaptoethanol to obtain the reduced keratins has been discussed (Thomas and Spei, 1984; Thomas et al., 1986; Van de Locht, 1987). Finally, reductive extraction with an aqueous mixture of urea and 2-mercaptoethanol followed by a reaction with methyl iodide or -iodoacetic acid to produce the corresponding S-alkylated keratins has been proposed (Ito et al., 1988; O’Donnell and Thompson, 1964; Miyamoto et al., 1987). In addition, several reports, including patents, discuss degraded keratins produced by partial hydrolysis (with acid, alkali, or enzymes) of wool, hair, feather, and other keratinous materials. The small keratin fragments obtained by hydrolysis are used in the manufacture of cosmetics, artificial leather, and filaments.

Preparation of Reduced Keratins

257

This procedure (Figure 10.4) is referred to hereafter as method a (Yamauchi et al., 1996). Wool was washed with water, dried, and defatted by Soxhlet extraction using a 1:1 v/v mixture of hexane and dichloromethane. The cleaned wool (10 g) was mixed with 8 M urea (180 mL), sodium dodecyl sulfate (SDS) (6 g), and 2-mercaptoethanol (15 mL) in a 300-mL round bottom flask and shaken at 50 C for 6 h. The aqueous phase was maintained around neutral pH during the process. The resulting mixture was quickly filtered (50 mesh), and the filtrate was subsequently dialyzed (two to three times) using a Union Carbide cellulose tubing ( 8,000 Da molecular weight cut-off) against degassed water (3.5 L) containing 0.2% (w/w) 2-mercaptoethanol to obtain a colorless, clear solution of reduced keratins (2.1% w/w, 230 mL; 48% yield); [ ] 2 dL/g at 20 C). Keratins in the aqueous solution consisted mainly of proteins that had molecular weights of 43 to 57 kDa and 14 to 29 kDa (Figure 10.5) with an overall cysteine content of 8–10 mol% of total amino acids (Figure 10.2). Such features typify reduced keratins from wool microfibrils (Dowling et al., 1986; Thomas et al., 1986). Chemically untreated woolen cloth or yarn may be used as starting materials with the described process.

Figure 10.4 Extraction of reduced keratins using urea/reducing agent/surfactant system (method a).

258

Figure 10.5 SDS-PAGE (10–15% gradient polyacrylamide gel; Pharmacia PhastGel; code 17-0517-01; pH 8.1/tris-alanine). Lane R molecular weight markers (kDa); lane L reduced keratins prepared by extraction method a; and lane M reduced keratins prepared by extraction method b.

This procedure is referred to hereafter as method b (Yamauchi et al., 1996). A mixture of woolen cloth (10 g), urea (65 g), 2-mercaptoethanol (15 mL), and water (80 g) was deoxygenated, warmed at 60 C for 24 h with mechanical shaking, and filtered (50 mesh). The filtrate was dialyzed using the aforementioned cellulose tubing against 2-mercaptoethanol (0.2% w/w) in water and was separated into the supernatant and the precipitates. The supernatant mainly contained reduced keratins from microfibrils as determined from amino acid analysis (for a S-carboxymethylated sample and a performic acid-treated sample) and SDS-PAGE. The precipitates contained reduced proteins primarily from the matrix keratins.

Keratins Films

259

Urea and 2-mercaptoethanol in the two extraction methods previously described cleaved hydrogen bonds and covalent S-S bonds between proteins, respectively, causing keratin fibril bundles to disassociate and be reduced simultaneously into monomeric reduced keratins. Method a uses SDS where the aqueous solution contains the surfactant up to an amount of 5–17% (w/w) of the proteins. SDS accelerated the extraction rate and stabilized the aqueous solution. Anionic surfactants, such as SDS, are more effective for this purpose than cationic or neutral ones. Although experimental conditions have not been optimized, keratins can be extracted rapidly when the concentrations of urea, 2-mercaptoethanol, and the surfactant are greater than 5 M, 0.3 g/g wool, and 0.2 g/g wool, respectively. 2-Mercaptoethanol may be substituted by thioglycolic acid and other water-soluble reducing reagents, including sodium bisulfite (which is less effective). Temperature and pH of the solution may be set at 50–70 C and 7–8, respectively. Extraction at about 100 C and/or at extremely alkaline pH causes decomposition of reduced keratins. The yield of reduced keratins ranges from 30 to 70% (w/w) based on the starting wool material. Most of the insoluble remainder comes from the cuticle. Aqueous solutions of reduced keratins were stored in glass bottles at ambient temperature for more than a year without precipitation and rotting. A 1H-NMR spectrum of the reduced keratin solution and an elemental analysis of the freeze-dried powder suggested that, even after repeated dialysis, SDS remained associated with keratins in amounts corresponding to 5–17% (w/w) of the proteins. Light scattering of the solution indicated the presence of particles with a diameter of 30 to 50 nm. Although the stability of keratin solutions has not been fully explained, we speculate that keratins are solubilized as giant micelles separated by electrostatic repulsion. Such repulsive forces retard aggregation of protein molecules, thus preventing oxidative linking of adjacent SH groups to form S-S bonds.

Aqueous solutions of reduced keratins (2.1% w/w, 10 mL) obtained by method a were mixed with glycerol (50% w/w of protein), spread to a circular area (about 40 cm2) on a horizontal polypropylene sheet, and dried in a desiccator over granular silica gel (Figure 10.6). The dried films were heated for 15 min at 80 C and peeled off from the sheet. When necessary, the films were washed with distilled water, laid on a stainless-steel ring, and allowed to dry at ambient temperature. At 65% relative humidity (RH), the films were 30–40 m in thickness and had surface densities of 4 mg/cm2.

260

Figure 10.6 Preparation of a keratin film from an aqueous solution of reduced keratins.

We mainly studied films of reduced keratins obtained by extraction method a. Under the scanning electron microscope (magnification of 1000–5000), the films had smooth surfaces and dense cross sections with light transmittances of >99% at wavelength of 400–700 nm, 92–99% at 330–400 nm, and 50% at 300 nm (film thickness 50 m). In contrast, films from reduced keratins obtained by method b had “frosted” surfaces. The films were insoluble in organic solvents, including DMF and DMSO, at 100 C. They swelled by 30–60% length-wise in water and shrank to 90% in boiling water. However, the films were dissolved or broken to fragments upon heating in an aqueous solution of 2-mercaptoethanol, presumably due to reductive cleavage of S-S bridges. Amino acid analysis indicated that (1) the cysteine residues accounted for 7–9 mol% of total amino acid residues of the reduced keratins and (2) about 40–60% of the cysteine residues were converted to half-cystine residues when the reduced proteins were cast into films (Figure 10.2).

Tensile properties of glycerol-plasticized cast films are presented in Table 10.2. Films did not melt upon heating. Gradual decomposition above 160 C was evidenced by thermogravimetric analysis [Figure 10.7(a)]. Differential scanning calorimetric analysis showed an endothermic peak at around 270 C, although the peak did not re-appear in the second run [Figure 10.7(b)]. The heat

Keratin Films

261

absorption may have been caused by melting or decomposition of the -structure of keratins in films.

Apparent permeability values of various organic and inorganic substances across keratin films were determined according to Nakagaki and Yonese (1971). Film specimens with an effective diffusion area of 0.785 cm2 were placed between two aqueous compartments. One compartment was filled with permeators and the other with an aqueous ethylene glycol solution of the same osmotic pressure. Generally, apparent permeability through keratin films decreased with increasing molecular weight of the permeating species (Table 10.3). The keratin films were less permeable to the tested substances than collagen films cross-linked by glutaraldehyde (Table 10.3). Collagen films for apparent permeability testing (Table 10.3) were prepared by mixing 1% w/w aqueous solutions of collagen (type I, from calf skin) with 5% w/w aqueous glutaraldehyde (0.1 mL/1 mL of protein solution) below 10 C for 30 min. Then, the solutions were cast into films similar to the keratin film preparation

Figure 10.7 Thermal analysis of films from reduced keratins prepared by extraction method a: (a) thermogravimetric analysis and (b) DSC, first run (solid line) and second run (dotted line).

262

Keratin Films

263

(Figure 10.6). In terms of dissolved oxygen, the keratin membranes were as permeable (4 2 cm3 (STP) cm/cm2 s cmHg at 37 C) as membranes prepared from silk fibroin and other polypeptides (Minoura et al., 1979, 1990; Aiba et al., 1985).

The keratin films were biologically degradable. As displayed in Figure 10.8 (lines 䊊 and 䊉), hydrolysis of films (40 m thick) by trypsin (from porcine pancreas) at 37 C (in 0.05 M Tris-HCl of pH 7.6) proceeded rapidly at the beginning of incubation to a degradation extent of 45–55% and then almost stopped. Degradation extent was defined as [(initial film weight degraded film weight)/(initial film weight)] 100. The incomplete decomposition was attributed to the substrate specificity of the enzyme (preferred links are the carboxyl groups of lysine and arginine) (Dixon et al., 1979) and to the

Figure 10.8 Biological degradation of films from reduced keratins prepared by extraction method a by trypsin (䊉 and 䊊 are 520 and 2600 USP units of trypsin/mL, respectively) or subcutaneously in mice (䉱). Biological degradation of type I collagen films subcutaneously in mice is also shown (䊏). The degradation extent did not increase appreciably when trypsin (same amount as first dosage) was added again to the assay system at the incubation time point marked by ✳. Results suggested that keratin films did not have any remaining trypsin susceptible bonds after about 45–55% degradation.

264 three-dimensional bridged film structure (possible steric hindrances). Keratin films also degraded subcutaneously in mice (Figure 10.8, line 䉱) (Yamauchi et al., 1996). Their in vivo degradation rate was slower than that of type I collagen films (Figure 10.8, line 䊏) and poly(lactic acid). The films did not adhere strongly to the body. No abnormal scars were observed in mice around the embedded sites during the experimental period of about 18 weeks.

Cell substratum properties of the keratin films were compared with those of collagen films using L929 fibroblast cells from mouse connective tissue (Yamauchi et al., 1998). The substratum was the glass and polystyrene dishes, which were coated with reduced keratins obtained by extraction methods a and b. As anticipated, when the culture media did not contain fetal bovine serum (FBS), cells did not grow either on the cell-culture dish or on the dishes coated with the reduced keratins. In contrast, cells grew smoothly on reduced keratin-coated dishes in bipolar and sometimes multipod elongation in the presence of FBS. Most likely, certain mediator proteins in FBS, including those for cell adhesion and growth, overlayered first the protein coating, enabling the seeded fibroblast cells to scaffold and proliferate (presumably through interactions between fibroblast receptors and the corresponding ligands of the mediator). Cell growth was virtually independent of the amount of keratins applied per area. For example, coatings of 0–1 mg/cm2 (0–10 m and 0–14 m thick when dry and wet, respectively) of the reduced keratins (extraction method b) supported cell adhesion and proliferation almost similarly as shown in Figure 10.9(a) and 10.9(b) (line F). Figure 10.9(a) suggests that there was better cell attachment on the keratin substratum than on the glass (control) and collagen substrata. Figure 10.9(b) may be considered to display the proliferation specificity of the substrata (the culture period of 24 h is a doubling time). It appeared that the keratin coating (to about 1 mg/cm2) was more effective than both the glass itself and the collagen coating [Figure 10.9(b), lines 䊉 and 䊊 vs. line 䊏]. When reduced keratins (by extraction method a) were used for coatings that were thick and washing with phosphate buffered solution (PBS) (8.00 g NaCl, 0.20 g KCl, 1.15 g Na2HPO4, and 0.20 g KH2PO4 in total volume of 1 L) was brief, cell growth was suppressed [Figure10.10(b), line ✳ for 3 h washing]. The phenomenon was ascribed to the toxicity of the residual SDS used with extraction method a (Gloxhuber, 1974). Indeed, cell growth and morphology returned to the levels of reduced keratins prepared by the extraction method b (which did not make use of SDS) after repeated washing with PBS( ) [Figure 10.10(b), lines 䊊 and 䊉 for 3 + 24 and 3 + 48 h, respectively]. In contrast, the cells attached well onto coatings of reduced keratins without washing with PBS( ) [Figure 10(a)]. Also, it was noticed that cell attachment on collagen coatings required a time-lag of about 1 h after planting. The enhanced attach-

(a)

(b) Figure 10.9 Relationship between quantity of the substrata coated on glass with the cell number-ratios (the number of cells on the substratum divided by the number of cells on the glass); 䊉 and 䊊, coatings with reduced keratins (extraction methods a and b, respectively) washed with ( )PBS for 3 + 48 h; 䊏, collagen. Conditions: 1 105 cells/cm2 were plated at 37 C in minimum essential medium (MEM) containing 10% FBS. Culture periods of 1 h [Figure 10.9(a)] and 24 h [Figure 10.9(b)], respectively. Cell populations determined on glass at 1 and 24 h-periods were 1.4 104 and 3.0 104, respectively.

265

(a)

(b) Figure 10.10 Effect of washing with PBS( ) on the adhesion and growth of L929 cells. The abscissa and the ordinate are defined similarly as in Figure 10.9; ✳, 䊊, and 䊉, coatings with reduced keratins (extraction method a) were washed for 3, 3 + 24 and 3 + 48 h, respectively. Incubation periods of 1 h [Figure 10.10(a)] and 24 h [Figure 10.10(b)], respectively. Cell populations determined on glass at 1 and 24 h-periods were 1.4 104 and 3.0 104, respectively.

266

Keratin Microcapsules

267

ment of the L929 cells on reduced keratin coatings compared to collagen coatings has not been fully understood. It may be ascribed to a fast covering of mediator proteins (from FBS) because the keratins consist of a wide variety of amino acids and have a greater amount of polar amino acids than collagen ( 27 mol% vs. 19 mol% of Arg, Lys, His, Asp, and Glu). In addition, a study on the microfibril of wool keratins (8c-1 component) revealed the presence of an Arg328-Gly-Asp sequence in the 2B region (Dowling et al., 1986). Reportedly, the tripeptide sequence in fibrinogen, fibronectin, vitronectin, von Willebrand factor, and silk fibroin is considered the cell-binding site (Pierschbacher and Ruoslahti, 1984; Hynes, 1986; Ruoslahti and Pierschbacher, 1987; Minoura et al., 1995).

Microcapsules are used as carriers of fine chemicals such as dyes, flavors, fragrances, and drugs. Encapsulation may sustain the duration of activity of the substances and alter the releasing mode into aimed sites (Kondo, 1979; Pitt et al., 1980; Sun et al., 1984; Weiner, 1990). Suslick and Grinstaff (1990) developed an ultrasound method to prepare aqueous suspensions of protein microcapsules filled with water-insoluble liquids. Their proposed microencapsulation process involved both emulsification and chemical cross-linking of protein molecules (mainly through S-S bond formation by sonically generated superoxide). We applied this sonication method to aqueous solutions of reduced keratins prepared by extraction method a (Yamauchi and Khoda, 1997). The procedure is as follows (Figure 10.11). A mixture of the

Figure 10.11 Preparation of keratin microcapsules.

268 aqueous solution of reduced keratins (1.8% w/w, 4 mL) and a toluene solution (2 mL) of a substance to be encapsulated, such as Sudan II (0.09 mg/mL), was irradiated by a probe-type sonicator at 22 C for 3 min under mechanical stirring (output power of 165 W/cm2 of the probe cross-sectional area). The resulting milky suspension was centrifuged at 1000 g for 15 min. The upper layer containing microcapsules was stirred with water and centrifuged to separate the vesicles again. This washing process was repeated a few more times until the lower aqueous phase became almost clear. Figure 10.12(a) (optical micrograph) displays the resulting vesicles (toluene and water as internal and external phases, respectively), while Figure 10.12(b) (transmission electron micrograph) displays empty vesicles formed by solvent evaporation under reduced pressure. The vesicles were estimated at 6–10 m in diameter by light scattering (Figure 10.13). Membrane (shell) thickness was estimated from the electron micrograph at 0.05–0.1 m. The keratin vesicles were superior in encapsulating properties to protein vesicles prepared similarly from egg yolk albumin, bovine serum albumin (BSA), soy protein (11S fraction), and myoglobin (Yamauchi and Khoda, 1997). Generally, all these encapsulating agents retained the hydrophobic compounds (vide infra) within the capsules when water or methanol were employed as external solvents. Microcapsules containing polymers such as poly(methyl methacrylate) and polystyrene were stable even in potent solvents (e.g., toluene and diethyl ether) for the polymers. In a hydrophobic solvent, however, small organic compounds leaked out of the protein capsules. Leakage from capsules varied with the protein used as the microencapsulating agent (Figure 10.14). Although the experiment offered only a qualitative insight on permeability of the capsule membranes, it appeared that the barrier ability of the membranes increased with increasing cysteine content in the proteins. Specifically, the protein membranes were more effective barriers in the order of keratins (112) > BSA (49) > soy protein 11S (17) > myoglobin (0) (where the numbers in parenthesis correspond to cysteine plus cystine residues per 1000 amino acid residues of the proteins). These results suggested that the protein membranes were developed through intermolecular S-S bonds between cysteine residues of protein molecules. In fact, the keratin capsules lost their shape upon heating in a dilute aqueous solution of a mercaptanol, such as 2-mercaptoethanol or dithiothreitol, presumably because of cleavage of S-S bonds by the mercaptanols. Myoglobin, which lacks both cysteine and cystine residues, resulted in leaky and fragile vesicles, perhaps because the membrane was formed merely from aggregates of the denatured protein. Toluene, xylenes, and o-isopropylphenol were suitable water-immiscible organic solvents for the sonication-promoted microencapsulation. Various hydrophobic materials including n-C10H21OH, C11H23COOH, glycerides [a mixture of various CH2(OCOR)CH(OCOR )CH2(OCOR )] and dyes were encapsulated in yields greater than 95% within keratin microcapsules. However,

269

Figure 10.12 (a) Optical micrograph of an aqueous suspension of keratin microcapsules filled with toluene. The capsules were stained by Coomassie brilliant blue R. Unstained white spots are toluene suspensions. The bar is 20 m. (b) Transmission electron micrograph of empty keratin microcapsules; negatively stained by phosphotungstic acid/NaOH (pH 7). The bar is 10 m.

Figure 10.13 Average size distribution (%) of keratin microcapsules filled with toluene in an aqueous suspension measured by light scattering (assay temperature 20 C).

Figure 10.14 Leakage extent of Sudan II from various protein microcapsules ( 7–9 m mean diameter) over time (䊉 wool keratins; 䊏 bovine serum albumin; 䉱 myoglobin).

270

References

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microencapsulation of antioxidants such as vitamins C and E in keratins was not successful. In fact, antioxidants rather inhibited microcapsule formation presumably by quenching the radical-mediated oxidative intermolecular cross-linking of SH groups in keratins (Suslick and Grinstaff, 1990).

Stable aqueous solutions of reduced keratins can readily be prepared from wool and hair with a mixture of urea, mercaptanol, surfactant, and water at 40–60 C. SDS was especially effective as a surfactant not only in promoting extraction but also in stabilizing the aqueous protein solutions. The reduced keratins had molecular weights of 14–57 kDa, cysteine contents of 8–10 mol%, and pI of about 6.7. The aqueous solutions were cast into clear films, which were insoluble in water and in most organic solvents (including dimethyl sulfoxide). The keratin films were mechanically stronger than similarly prepared collagen (Type I) films and were less permeable than the latter to glucose, urea, sodium chloride, and proteins such as BSA. The keratin films degraded both in vitro (by trypsin) and in vivo (subcutaneous embedding in mice). They supported attachment and growth of mouse L929 fibroblast cells. The comparative culture assay on keratins, collagen (type I), and glass revealed that the keratins were more adhesive to the cells and more supportive for cell proliferation than collagen and glass. The results were explained by an enhanced initial adsorption of mediator proteins from fetal bovine serum onto the keratin substrata. Upon sonication, reduced keratins formed spherical microcapsules (2–10 m in diameter and 0.05–0.1 m in membrane thickness). When oil, fat, and dyes were dissolved in the solvent, they were trapped within the keratin vesicles in yields greater than 95%. The keratin capsules were more effective in retaining encapsulating substances than microcapsules made from BSA, soy protein (11S fraction), or myoglobin.

Aiba, S., Minoura, N., Fujiwara, Y., Yamada, S. and Nakagawa, T. 1985. “Laminates Composed of Polypeptides and Elastomers as a Burn Wound Covering. Physicochemical Properties,” Biomat., 6:290–296. Arai, K., Naito, S., Dang, V.B., Nagasawa, N. and Hirano, M. 1996. “Crosslinking Structure of Keratin. VI. Number, Type, and Location of Disulfide Crossslinkages in Low-Sulfur Protein of Wool Fiber and Their Relation to Permanent Set,” J. Appl. Polym. Sci., 60:169–179. Dixon, M., Webb, E.C., Thorne, C.J.R. and Tipton, K.F. 1979. Enzymes, 3rd edition, London: Longman Group Ltd., pp. 261, 302. Dowling, L.M., Crewther, W.G. and Parry, D.A.D. 1986. “Secondary Structure of Component 8c-1 of -Keratin: Analysis of the Amino Acid Sequence,” Biochem. J., 236:705–712.

272 Feughelman, M. 1985. “Keratin” in Encyclopedia of Polymer Science and Engineering, Vol. 8, eds., H.F. Mark, N.M. Bikales, C.G. Overberger, G. Menges, and J.I. Kroschwitz, New York: John Wiley & Sons, pp. 566–600. Fraser, R.D.B., MacRae, T.P., Parry, D.A.D. and Suzuki, E. 1986. “Intermediate Filaments in -Keratins,” Proc. Natl. Acad. Sci. USA, 83:1179–1183.Gloxhuber, C. 1974. “Toxicological Properties of Surfactants,” Arch. Toxicol., 32:245–270. Harrap, B.S. and Woods, E.F. 1964. “Soluble Derivatives of Feather Keratin,” Biochem. J., 92:8-18. Heid, H.W., Werner, E. and Franke, W. 1986. “The Complement of Native Alpha-Keratin Polypeptides of Hair-Forming Cells: A Subset of Eight Polypeptides That Differ from Epithelical Cytokeratins,” Differentiation, 32:101–119. Husken, P. 1986. “Isolierung von Mikrofibrillen und Matrix-Proteinen aus Humanhaar in der Disulfidform,” Parfumerie und Kosmeitik, 67:434–442. Hynes, R.O. 1986. “Fibronectins,” Scient. Amer., 254(6):32–41. Ito, M., Tazawa, T., Shimizu, N., Ito, K., Katsumi, K. and Sato,Y. 1988 “Cell Differentiation in Human Anagen Hair Follicles Studied with Anti-Hair Keratin Monoclonal Antibodies,” J. Invest. Dermatol., 86:563–569. Kondo, A. 1979. Microcapsule Process and Technology. New York: Marcel Dekker. MacLaren, J.A. and Milligan, B. 1981. Wool Science. The Chemical Reactivity of the Wool Fibre. Marrickville, NSW, Australia: Science Press, pp. 1–18. Minoura, N., Fujiwara,Y. and Nakagawa, T. 1979. “Permeability of Synthetic Poly( -Amino Acid) Membranes to Oxygen Dissolved in Water,” J. Appl. Polym. Sci., 24:965–973. Minoura, N., Tsukuba, M. and Nagura, M. 1990. “Physico-chemical Properties of Silk Fibroin Membrane as a Biomaterial,” Biomat., 11:430–434. Minoura, N., Aiba, S., Gotoh, Y., Tsukuda, M. and Imai, Y. 1995. “Attachment and Growth of Cultured Fibroblast Cells on Silk Protein Matrices,” J. Biomed. Mat. Res., 29:1215-1221. Miyamoto, T., Sakabe, H. and Inagaki, H. 1987. “Ordered Structure of High-Glycine Proteins from Reduced Merino Wool,” Bull. Inst. Chem. Res. Kyoto Univ., 65:109–119. Nakagaki, S. and Yonese, M. 1971. “Permeability and Its Temperature Dependency on Gelatin, Collagen Membranes and Bovine Eye Lens Capsule,” J. Pharm. Soc. Jpn., 91:1211–1216. O’Donnell, I.J. and Thompson, E.O.P. 1964. “Reduced Wool. IV. The Isolation of a Major Component,” Austr. J. Biol. Sci., 17:973–979. Pierschbacher, M. D. and Ruoslahti, E. 1984. “Variants of the Cell Recognition Site of Fibronectin That Retain Attachment-Promoting Activity,” Proc. Natl. Acad. Sci., 81:5985–5988. Pitt, C.G., Marks, T.A. and Schindler, A. 1980. “Biodegradable Drug Delivery Systems Based on Aliphatic Polyesters: Applications to Contraceptives and Narcotic Antagonists” in Controlled Release of Bioactive Biomaterials, eds., R.W. Barker and W. Richards, New York: Academic Press, pp. 19–43. Rhodes, H.J., Potter, B. and Widra, A. 1967. “Characteristics of the Alpha-Keratose Fraction of Hair Inducing Ascosporogenesis in Nannizzia Grubyia,” Mycopathologia et Mycologia Applicata, 33:345–348. Ruoslahti, E. and Pierschbacher, M.D. 1987. “New Perspectives in Cell Adhesion: RGD and Integrins,” Science, 238:491–497. Sun, A.M., O’Shea, G.M. and Goosen, M.F.A. 1984. “Injectable Microcapsulated Islet Cells as a Bioartificial Pancreas,” Appl. Biochem. Biotechnol., 10:87–99. Suslick, K.S. and Grinstaff, M.W. 1990. “Protein Microcapsulation of Nonaqueous Liquid,” J. Am. Chem. Soc., 112:7807–7809. Thomas, T. and Spei, M. 1984. “Isolierung der Mikrofibrillenproteine von Wolle in der Disulfidform,” Melliand Textilber, 65:208–209. Thomas, H., Conrads, A., Phan, K-H., van de Locht, M. and Zahn, H. 1986. “In vitro Reconstitution of Wool Intermediate Filaments,” Int. J. Biol. Macromol., 8:258–264. Van de Locht, M. 1987. “Rekonstitution von Mikrofibrillen aus Wolle und Filamenten aus Epidermisproteinen,” Melliand Textilber, 68:780–786.

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Weiner, A.L. 1990. “Developing Lipid-Based Vesicles and Protein Drugs. Part I: Selection and Analysis Issues,” Biopharm., 3:27–32. Yamauchi, K. and Khoda, A. 1997. “Novel Proteinous Microcapsules from Wool Keratins,” Colloids and Surfaces B: Biointerfaces, 9:117–119. Yamauchi, K., Yamauchi, A., Kusunoki, T., Khoda, A. and Konishi,Y. 1996. “Preparation of Stable Aqueous Solution of Keratins, and Physicochemical and Biodegradational Properties of Films,” J. Biomed. Mat. Res., 31:439–444. Yamauchi, K., Maniwa, M. and Mori, T. 1998. “Cultivation of Fibroblast Cells on Keratin-Coated Substrata,” J. Biomater. Sci. Polym. Edn., 9:259–270.

Formation and Properties of Collagen and Gelatin Films and Coatings

OLLAGEN is an abundant protein constituent of connective tissue in vertebrate (about 50% of total human protein) and invertebrate animals (Johnston-Banks, 1990). Similar to cellulose in plants, collagen molecules support mechanical stresses transferred to them by a low-modulus matrix (Yannas, 1972). Thermal or chemical dissociation of collagen polypeptide chains forms products known as gelatin. Insoluble collagen is converted to soluble gelatin by acid or alkaline/lime (mild and slow) processing. Two processes are mainly used for commercial production of gelatin. In the first process, the collagen in hide or demineralized bone is partly depolymerized by prolonged liming that breaks down covalent cross-links. The occurring hydrolysis results in extensive release of collagenous material, which is solubilized at near neutral pH at temperatures of 60–90 C (Type B gelatin). The acid process (Type A gelatin) involves soaking skin or bone in a dilute acid followed by extraction at acid pH. Gelatin manufacturing has been discussed extensively (Robinson, 1953; Rose, 1987; Johnston-Banks, 1990; GMIA, 1993). The properties of collagen and gelatin are of great interest to various fields, such as surgery (implantations; wound dressings), leather chemistry (tanning), pharmacy (capsule production; tablet binding), and food science (gels; edible films) (Gustavson, 1956; Harrington and Von Hippel, 1961; Von Hippel, 1967; Johnston-Banks, 1990; Arvanitoyannis et al., 1997, 1998a, b). Reportedly,

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276 about 65% of gelatin manufactured worldwide is used in foods, 20% in photographic applications, 10% in pharmaceutical products, and 5% in other specialized and industrial applications (Slade and Levine, 1987).

Because of the importance of collagen, its structure has been extensively studied. The primary structure (amino acid sequence) of collagen and gelatin consists of 18 different amino acids (Table 11.1). Collagen is rich in glycine (~1/3 of total residues), hydroxyproline, and proline. Also, it is one of few proteins containing hydroxylysine and is devoid of tryptophan. Mammalian collagen is richer in hydroxyproline than fish collagen (Foegeding et al., 1996). Collagen is a hydrophilic protein because of the greater content of acidic, basic, and hydroxylated amino acid residues than lipophilic residues. Therefore, it swells in polar liquids with high solubility parameters. The secondary structure (local configuration of chains) does not favor formation of -helical chain segments because of the high number of prolyl and hydroxyprolyl residues. Collagen chains are helical macromolecules with a tendency mainly for interchain, rather than intrachain, hydrogen bonding (Carver and Blout, 1967). The tertiary struc-

Figure 11.1 Order in collagen: (a) primary, (b) secondary, (c) and (d) tertiary, and (e) quaternary structure (Yannas, 1972; Georgatsos, 1982; Foegeding et al., 1996).

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278 ture standing for large-scale folding and helicity of collagen is better understood through the fundamental unit tropocollagen. Tropocollagen is a right hand superhelix with a repeat unit of about 100 Å consisting of three left-hand strands (Ramachandran and Kartha, 1954; Rich and Crick, 1955, 1961; Yannas, 1972). Two or three of these chains can be covalently linked to form a -chain or a -chain, respectively. The quaternary structure refers to formation of small to medium size aggregates of tropocollagen molecules, i.e., complex crystallites first identified by small angle X-ray diffraction (Highberger et al., 1950; Hodge and Schmitt, 1960) (Figure 11.1). Collagen molecules link end to end and adjacently to form collagen fibers. There is a periodicity in the cross-striations of collagen at about 640–700 Å intervals. The fibers are sometimes arranged in parallel having great strength, as in tendons, or they may be highly branched and disordered, as in skin. The primary structure of gelatin is almost identical to that of parent collagen but for some small differences due to the pretreatment and extraction processes. These differences include removal of amide groups of asparagine and glutamine, conversion of arginine to ornithine, and (partial) removal of telopeptides (Johnston-Banks, 1990). The precise macromolecular constitution of gelatin resulting from a melting process depends on collagen source and on the extraction method (Oth, 1959; Yannas, 1972). Gelatin chains can be intertwined back into the collagen helix through an appropriate technique, such as cooling or annealing in solution. Films “cold cast” from gelatin solutions at temperatures below 35 C provided concluding evidence of tertiary structure but not of quaternary structure while “hot cast” gelatin films were amorphous (Bradbury and Martin, 1952; Robinson, 1953).

The lability of peptide bonds and cross-linkages depends upon temperature, pH, and raw material source. The number average molecular weight (Mn) and the weight average molecular weight (Mw) of non-hydrolyzed gelatins ranged from 5 104 to 1 105 Da and from 105 to 106 Da, respectively (Rose, 1987). Average molecular weight and molecular weight distributions can be determined by osmometry, viscometry, sedimentation, light scattering, filtration chromatography, and gel electrophoresis (Yannas, 1972; Rose, 1987). Gelatin distributions were polydisperse and were resolved into many peaks depending primarily on the measurement method. The proportion of and chains or cross-linked aggregates is mainly affected by the pretreatment method (Figure 11.2).

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Figure 11.2 Molecular weight distributions of high Bloom gelatins: – – – , type A (pigskin); – – –, limed calfskin; ––––, limed ossein; and , thermally denatured soluble calfskin collagen. From Rose, 1987, in Encyclopedia of Polymer Science and Engineering, 2nd edition, Volume 7, Mark et al., eds. Copyright 1987. Reprinted by permission of John Wiley & Sons, Inc.

Early infrared (IR) work on proteins was promising and resulted in several proposed models. The IR spectra of all proteins resemble each other closely because protein molecules consist of varying amounts of 20 amino acid residues. The varying abundance of these amino acid residues is reflected in the IR spectrum of each protein. In fact, molecular configuration affects the IR spectrum of a polymer molecule, and different forms of the same protein may have different spectra. Blood proteins, muscle proteins, plant proteins, and scleroproteins belong to class A, whereas collagen, gelatin, and silk fibroin belong to class B. Because the variability of protein IR spectra is rather limited, proteins can fairly easily be distinguished from other biological molecules, such as carbohydrates, lipids, or nucleic acids (Beer et al., 1959). IR spectra of collagen and gelatin have been extensively investigated over the past 50 years, and several of the initial resolution problems have been resolved. Several changes appearing in the IR spectrum by denaturation of collagen to hot cast gelatin were identified (Figure 11.3) (Yannas, 1972). Collagen in the solid state was also studied with IR spectroscopy by using deuterated samples (Beer et al., 1959). Structures of gelatin gels, formed by cooling down sufficiently concentrated

280

Figure 11.3 Infrared absorption spectra of collagen (from acetic acid) and hot-cast gelatin (cast at 65 C from aqueous solution). Reprinted from Yannas, 1972, courtesy of Marcel Dekker, Inc.

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gelatin solutions, were investigated by holographic relaxation spectroscopy (HRS) that revealed two peaks, presumably corresponding to “coarse” and “fine” network (Wu et al., 1991). The former consisted of aggregations of a collagen-like triple stranded helix, whereas the latter consisted of entanglements between polymer chains. Predominance of the former over the latter depended on cooling rate. This model could possibly be expanded to account for structural differences occurring during gelatin film formation (Wu et al., 1991).

Melting of molecularly dispersed collagen (tropocollagen) in heated, salt-free water solutions proceeded with an extremely intensive heat absorption (Privalov, 1982). The high specific heat of collagen denaturation facilitates the thermal study of collagen intramolecular melting, which should be carried out by sensitive instruments because of the artifacts that have occasionally occurred. Flory and Garrett (1958) derived the first order transition (Tm 145 C) occurring in collagen and gelatin from an extrapolation of dilatometric data. Subsequent studies on collagen/gelatin samples with or without plasticizers led to substantial deviations. Table 11.2 summarizes reported melting temperature (Tm) and glass transition temperature (Tg) values of collagen and gelatin. Several deviations arise from insufficient documentation of measurement conditions or sample treatment, thermal history of samples, or presence of water. Even in small amounts, water depresses the Tg of gelatin by as much as 5 C per 1% (w/w) water in the region of low moisture content. Furthermore, removal of moisture below 1% (w/w) causes covalent cross-linking, thus increasing Tg due to network formation (Yannas, 1972). Figure 11.4 shows the state diagram of the gelatin-water system. The considerable deviations among reported results (Hirai, 1955; Luescher et al., 1974; Borchard et al., 1980; Marshall and Petrie, 1980; Reutner et al., 1985; Slade and Levine, 1987; Nishinari et al., 1997; Arvanitoyannis, 1999) should be attributed to differences in origin of gelatin samples, preparation methods (hot vs. cold casting), heating rates, and thermal history. The values reported by Borchard et al. (1980) were obtained by extrapolation of experimental differential scanning calorimetry (DSC) data (10 C/min) with quenching of samples to homogeneous glass. Tg data by Hirai (1955) were obtained from dielectric (DETA) measurements, which may be the reason for their deviation from Tgs obtained by calorimetric measurements. The Tgs reported by Nishinari et al. (1997) tended to increase at higher water contents. This was attributed to gelatin molecules being restricted by the ice due to difficulties in completely solubilizing gelatin at concentrations higher than 60%. The Tg of pure water (138 C), as determined by Sugisaki et al. (1968), was used to extrapolate Tg data to very dilute gelatin solutions. The data from Marshall and Petrie (1980) showed the effect of different

282

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equilibrium water contents on the transition temperatures of a cold-dried gelatin film. At higher water contents, the transition temperatures were lower, and the intervals between Tms and Tgs increased so that the first and second order transitions were well defined and separated. Some representative first and second run heatings of dynamic thermal analysis (DTA) and dynamic mechanical thermal analysis (DMTA) traces of plasticized gelatin are shown in Figure 11.5. Slade and Levine (1987) also produced state diagrams for commercial calfskin and pigskin gelatins. Their observation that both transition temperatures (Tm and Tg) were independent of moisture content in the presence of excess moisture did not agree with other studies (Borchard et al., 1980; Marshall and Petrie, 1980; Reutner et al., 1985). However, there is general agreement that both Tm and Tg increase with decreasing moisture content, finally converging or extrapolating to a high temperature range of 210 to 240 C at very low/zero moisture contents (Yannas and Tobolsky, 1964, 1968; Koleske and Faucher, 1965; Jolley, 1970; Marshall and Petrie, 1980; Slade and Levine, 1987). Recently, Tg of gelatin films was indirectly determined from microhardness measurements vs. temperature (Vassileva et al., 1998). The first heating of gelatin samples showed a softening point at around 200 C, whereas extrapolation of the two linear parts of the fourth cycle resulted in a Tg of 215 C (Vassileva et al., 1998). This was in satis-

Figure 11.4 State diagram of the gelatin-water system.

Figure 11.5 Thermal properties of plasticized gelatin: (a) Representative traces of 1st and 2nd run heatings of gelatin-water systems and (b) DMTA traces of glycerol plasticized gelatin (Arvanitoyannis, 1999).

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factory agreement with reported Tg values (Table 11.2) of 220 C (Marshall and Petrie, 1980), 217 C (Rose, 1987), and 212 C (Arvanitoyannis, 1999). Nishinari et al. (1992, 1997) studied rheologically and thermally the effect of sucrose and glucose on gelatin gels. Sucrose raised the gel to sol transition temperature and increased the Young’s modulus by increasing the number of elastically active network chains. An increase in the amount of unfreezable water, which exists in junction zones, in the presence of sucrose was related to an increase in the number of junction zones. Sucrose produced unfreezable water by itself and also interacted directly with gelatin molecules, thus forming additional junction zones capable of holding even more unfreezable water (Nishinari et al., 1992). The Tg values of 40% (w/w) gelatin gels containing glucose moved to lower temperatures contrary to the gels with sucrose. This different behavior could be possibly attributed to glucose having a lower molecular weight than sucrose and to the fact that a decrease in freezable water, counterbalances the unfreezable water, which acts as a plasticizer shifting the Tg to lower temperatures (Nishinari et al., 1997). Arvanitoyannis et al. (1997, 1998a, b) and Arvanitoyannis (1999) studied with DTA and DMTA the effects of plasticizers (i.e., water, sugars, and glycerol) on gelatin, gelatin/starch (unmodified and modified), and gelatin/chitosan blends. A second plasticizer (sugar or polyol) in addition to water resulted in a much more pronounced depression of Tg values for gelatin/chitosan blends (Figure 11.6). Several gelatin products are based on the lyotropic effect of the Hofmeister series of ionic salts, acids, and denaturants such as urea (Von Hippel and Schleich, 1969). The Tms of gelatin solutions were depressed below 20 C in the presence of urea, thus rendering gelatin concentrates liquids at ambient temperature (Slade and Levine, 1987).

Thin gelatin films are rich in colloid material and behave similarly to rigid gels exhibiting elastic moduli and time-dependent phenomena. Commercially, gelatin is mainly characterized by Bloom strength and viscosity. However, tensile strength and elongation are equally important physical properties for edible film applications. The main parameters affecting film-forming properties of gelatin are raw material source, extraction method, molecular weight, film preparation method (hot vs. cold casting), and degree of hydration or presence of plasticizer. Table 11.3 presents measured mechanical properties of gelatin films. Healey et al. (1974) determined the mechanical properties of films made from several binders used in tableting. Gelatin films showed a slight yielding before fracture and had higher values of Young’s modulus and tensile strength than methylhydroxyethyl cellulose, starch, acacia gum, or polyvinylpyrrolidone.

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Figure 11.6 Representative traces of chitosan/gelatin/water/glycerol blends prepared by the lowtemperature process: DTA and DMTA. Reprinted from Carbohydrate Polymers, Volume 37, Arvanitoyannis et al., “Chitosan and Gelatin Based Edible Films,” pp. 371–382. Copyright 1998, with permission from Elsevier Science.

High elastic modulus, low plasticity, and relative brittleness were possible limitations of gelatin as a tablet binder (Healey et al., 1974). Gelatin and collagen films dried over phosphorus pentoxide (both equilibrated and non-equilibrated) showed a two-dimensional contraction of up to 10%, known as “hydrothermal shrinkage,” for samples containing greater than 15% moisture. The strength of alkaline, acid, and alkaline-acid (1:1) films was sensitive to the amount of water present, reaching a maximum in the range of 14–16%

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288 moisture (Kellaway et al., 1978b). The effects of dyes (i.e., amaranth, erythrosine, brilliant blue, and indigo carmine) and titanium dioxide (TiO2), which are commonly added to hard and soft gelatin capsule shells, were studied by Kellaway et al. (1978b). Both TiO2 and the dyes had minor effects on tensile strength and Young’s modulus of acid and alkaline gelatin films. However, the dyes affected stress contraction of films by interacting with gelatin and thus interfering with the strain-induced orientation of gelatin molecules. Bradbury and Martin (1952) showed the importance of hot vs. cold casting on mechanical properties of gelatin films. Cold cast (20 C) films had greater tensile strength than hot cast (56–60 C) films (89.3 vs. 64.2 MPa at 45% RH during testing) (Bradbury and Martin, 1952). The greater tensile strength of cold cast films was due to their higher crystallinity and degree of orientation of the crystallites. Tensile strength of both cold and hot cast films decreased with increasing RH because intermolecular bonds were weakened by absorbed water and lower tension was required for rupturing the bonded regions within gelatin films. Also, cold cast films had slightly greater extension at break than hot cast films at 45–65% RH during testing (Bradbury and Martin, 1952). However, at higher RH (75–85%) during testing, hot cast films had substantially greater extension at break, presumably due to an almost simultaneous failure of a high number of bonded regions. Similar observations were reported for films from gelatin/starch (modified and unmodified) and gelatin/chitosan blends (Arvanitoyannis et al., 1997, 1998a, b), which were prepared by both hot and cold casting. The plasticizing effect of polyols/sugars on gelatin/starch films was also studied. Percent elongation at break showed a pronounced increase proportional to plasticizer content except from sucrose, which had the opposite effect (Arvanitoyannis et al., 1997, 1998a, b) (Table 11.4). Eliassaf and Eirich (1960) studied creep behavior of gelatin films at 100% RH. Equilibrium stress-strain curves differed considerably with temperature. At 26.5 C, gelatin films were rubber-like (8 GPa elastic modulus; 6 MPa stress; 300% elongation) because of high (27%) water uptake (Table 11.3). At higher temperatures, gelatin films became stiffer (4 GPa elastic modulus; 10 MPa stress; 230% elongation), and at 46.5 C, the films became highly brittle (5.2 MPa stress; 20% elongation) due to very low water contents (15%) that cannot affect the glass-like behavior of gelatin. The major disadvantage of gelatin as a food packaging material is its poor mechanical properties. Therefore, segmental orientation induced by hot and cold and rapid zone drawing, and cross-linking were used to improve mechanical properties of gelatin films (Fakirov et al., 1996, 1997). Gelatin cross-linking, which is particularly important for the medical and photographic industries, hardens gelatin, making possible the use of higher solution temperatures (Honda et al., 1997). The Young’s modulus for non-cross-linked gelatin was 0.8 GPa increasing to 1.0, 1.1, or 1.2 GPa for cross-linked gelatin subjected

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to draw ratios ( ) of 0, 1, or 4, respectively. Similarly, stress at break increased from 20% for non-cross-linked gelatin films to 40, 100, or 200% for cross-linked gelatin films with values of 0, 1, or 4, respectively. Increases up to 4 in modulus and stress at break of gelatin films as increased, during drying, were attributed to effective orientation within the film network. However, at higher values, film modulus and stress at break decreased due to extensive changes in the network affecting contacts or joints between crystallites and microfibrils, thus resulting in chain scission or disassociation (Fakirov et al., 1996, 1997) (Figure 11.7). Hood (1987) determined tensile properties of collagen casings prepared

290

Figure 11.7 Stress-strain curves for cross-linked gelatin films at different draw ratios ( ). Reprinted with permission from Fakirov et al., 1996. Copyright 1996, Springer-Verlag.

both by the wet and dry process and containing glycerol or sorbitol, respectively. Although casings prepared by the dry process had higher tensile strength and percent elongation, it is difficult to draw valid conclusions because of the complexity of interactions among ingredients (e.g., cellulose, fat, and carboxy methylcellulose) present in casings. Yannas (1972) discussed properties of collagen and gelatin films. Glycerol-plasticized collagen films had lower moduli than gelatin films. The effect of gelatin plasticization was more pronounced (3.5 times higher moduli values than non-plasticized gelatin films) when plasticizer content varied within 30–50% (w/w). In the diluent range of 60 to 75% (w/w) plasticizer content, gelatin films were tough rubbers extending up to 700%. When collagen films were subjected to stress, tropocollagen molecules slid past each other with negligible mutual entanglement excluding any recovery of original shape upon removal of the load. Collagen fibers were strong under tension, having a tensile modulus of 100 to 1,000 MPa (Farquhar et al., 1990; Fung, 1993; Schwartz et al., 1994). Recently, particularly high microhardness values, reaching up to 200 MPa, were reported for gelatin (Vassileva et al., 1998). Moreover, as mentioned previously, after four heating cycles, microhardness measurements were used to indirectly determine the Tg of gelatin.

The viscoelastic properties of gelatin have been investigated (Tobolsky,

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1960; Castello and Goyan, 1964; Smith and Goyan, 1965; Genot et al., 1989; Borchard and Burg, 1990) (Table 11.5). Gelatin gels are widely used in photographic, pharmaceutical, and food applications (Melnichenko and Klepko, 1992). Such applications usually include stages of gelatin swelling in water or mixed solvent systems. Observed differences in viscoelastic behavior among various gelatin films were readily assigned to characteristics of the tertiary structure and to parameters such as extraction methods, temperature, diluent content, and density of cross-links (Yannas, 1972; Oakenfull, 1984). In dilute systems (1–5% w/w), the rate of gelatin renaturation into collagen-like structures appears to increase as temperature and concentration increase. Although the heat of reaction of the renaturation process is high, the heat of activation of the rate-determining step is very small (Te Nijenhuis, 1981). Gelatin has the typical properties of an amphoteric polyelectrolyte (pH-dependent net charge and an isoelectric point) and is widely used with cationic or anionic surfactants (Wustneck et al., 1989; Johnston-Banks, 1990; Fruhner and Kretzschmar, 1992; Dreja et al., 1996). At pH 6, Type B gelatin molecules already exhibit negative net charges and are loosely coiled and rather flexible. Due to the high concentration of hydrophobic regions along gel-

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Figure 11.8 Possible structures of gelatin-surfactant complexes formed in various concentration regimens: (a) a salt complex is formed by specific binding of surfactants to the gelatin, (b) micellar regions aggregated along the gelatin chain lead to unfolding, and (c) free micelles disturb the gelatin-micelle complexes by electrostatic repulsion. Reprinted with permission from Dreja et al., 1996. Copyright 1996, Springer-Verlag.

atin chains, micellar structures along these hydrophobic regions nucleate, forming micellar gelatin-surfactant complexes. These micellar gelatin-surfactant complexes increase the hydrophilicity of gelatin molecules. As a result, chains uncoil and tend to be more soluble, leading to increased Mw for each gelatin chain due to association of surfactant molecules (Dreja et al., 1996) (Figure 11.8).

There is strong demand for minimally processed foods (Shewfelt, 1987). Edible coatings and films have been applied to processed foods to retard and prevent movement of water and gases. Although commercially available synthetic polymers, such as polyethylene and polypropylene, effectively control gas permeability (GP) and water vapor permeability (WVP), concerns over their non-renewable nature generates interest in more environmentally friendly, renewable biopolymers.

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Collagen and gelatin coatings have been used on meats and sausages to reduce GP and/or WVP (Hood, 1987). Gelatin is a good gas barrier, but is highly hydrophilic. Cross-linking of gelatin membranes reduces their solubility in water. Another approach to increasing moisture resistance is the use of laminated membranes. GP of films changes considerably below and above the glass transition zone (Psomiadou et al., 1996; Arvanitoyannis et al., 1997, 1998a, b). Similar behavior was observed for gelatin films (Figure 11.9) with activation

Figure 11.9 Permeability of oxygen in gelatin/water and gelatin/water/glycerol blends vs. inverse temperature (1/T). Presented values are means and standard deviations of at least six replicates (Arvanitoyannis, 1999).

294 energies for oxygen permeability being higher at temperatures above Tg than below Tg. At temperatures above Tg, gas molecules have to create their own interstitial spaces by separating interchain polymer contacts. The permeant then diffuses through the polymer matrix along cylindrical voids created by the synchronized rotation of polymer chain segments. On the contrary, at temperatures below Tg, gas molecules can diffuse through existing interstitial spaces, thus requiring lower activation energy (Biliaderis et al., 1999). Activation energies for oxygen permeability through gelatin films varied from 30 to 90 kJ/mol (Arvanitoyannis et al., 1998a). Similar values (20–120 kJ/mol) have been reported for other types of biopolymer films (Mujica-Paz and Gontard, 1997; Arvanitoyannis et al., 1998b; Biliaderis et al., 1999).

Many transparent substances lacking symmetry in their molecular or crystalline structure can rotate the plane of polarized radiation (Djerassi, 1960). Such materials are said to be optically active. The wavelength dependence of optical activity, known as optical rotatory dispersion (ORD), is a more useful source of structural information about asymmetric compounds than is the specific rotation at a single wavelength (Ewing, 1981). It is closely related to the phenomenon called circular dichroism (CD). Combination of ORD and CD results in what is known as the cotton effect, which appears in connection with chromophoric absorptions in optically active compounds. The maxima of CD and absorption and the inflection of the ORD curve theoretically coincide in wavelength but the superimposed effects of other chromophores may cause shifts in the maxima. CD curves often disclose “hidden” absorption maxima, not easily detected by ORD, whereas ORD curves can provide more specific information about characteristic chromophoric bands. In general, both CD and ORD provide valuable information about the stereochemical characteristics of optically active monomers or polymers (Abe and Inomata, 1989). In addition to its unique amino acid composition and wide angle X-ray diffraction pattern, collagen is also characterized by unusual optical rotatory (OR) properties. Initially, OR (Harrington and Von Hippel, 1961) and, only recently, CD have been extensively used to study gelatin and collagen structures in solutions/gels and in films. OR and CD values for collagen and gelatin are summarized in Tables 11.6 and 11.7, respectively. Collagen (regardless of source) had greater negative [ ]D values, close to those obtained for poly-L-proline II, than its respective gelatin. Therefore, these two structures are similar, whereas the observed variation in gelatin values was attributed to inherent properties of the helix (Harrington and Von Hippel, 1961). Robinson and Bott (1951) reported that specific rotation was identical for dissolved gelatin and gelatin films at 35 C, whereas cold cast gelatin films

295

296

had considerably higher rotation due to formation of collagen helix. Yannas (1972) noticed distinct differences among OR spectra of collagen, hot cast (amorphous) gelatin, and cold cast (semi-crystalline) gelatin, thus proposing use of this method for differentiating states. Although OR spectra for gelatin films did not differ substantially, cross-linking agents (e.g., formaldehyde) considerably decreased OR values in proportion to the amount of cross-linking agent (OR values of 610 , 570 , and 480 for 1, 5, and 10% formaldehyde in pigskin gelatin, respectively) (Coopes, 1968). According to Macsuga (1972), the OR for cold-dried gelatin at 570 nm was 660 , a much higher value than that of the gel ( 275 ). Various explanations for this observation have been suggested, such as increased triple helix formation, formation of anisotropic structure, random orientation of crystallites (triple helix segments), and orientation of molecules in the plane of the film. The latter explanation was more strongly supported by experimental evidence (Coopes, 1968; Macsuga, 1972). Several surprisingly high OR values for collagen and for cold cast gelatin or gelatin (no exact conditions mentioned) have been reported (Robinson and Bott, 1951; Robinson, 1953; Yannas, 1972) without any valid explanations. Achet and He (1995) monitored the renaturation level (total fraction of gelatin strands having helical conformation) in gelatin films (limed bone, pigskin, and modified gelatins). Negative [ ]D values increased with increasing film moisture caused by placing dehydrated films in a 50% RH chamber (20 C). This was attributed to the slow rate of helical growth due to higher Tg ( 45 C for 16% mc) and to subsequent stabilization of gelatin strands by absorbed water molecules (Achet and He, 1995). Melting of collagen or gelatin substantially affects OR (Figure 11.10). CD is an established method for studying structural changes of biological macromolecules both in solutions and in films (Young and Pysh, 1975). The triple helical conformation of collagen and gelatin exhibits in solution a positive CD band at 222 nm and a stronger negative band at 196 nm (Nishio and Hayashi, 1985; Wetzel et al., 1987). CD spectra for gelatin films were similar to

Optical Rotation and Circular Dichroism of Gelatin Solutions/Films

297

those for gelatin solutions, but the 222- and 196-nm peaks were slightly shifted toward the red (Wetzel et al., 1987). For all gelatin films, the intensity of the 222-nm band decreases with decreasing RH. In particular, the decrease in triple-helix content at RHs below 20% is highly pronounced. The decrease in the intensity of the 222-nm peak is followed by an increase in the negative CD 238 nm. However, film dehydration increases peak intensity at spectrum at 224 nm, whereas it decreases peak intensity at 238 nm. In general, the content of the triple-stranded structure decreases with increasing temperature of gelatin solution and with isothermal dehydration. Removal of water is associated with a trans to cis transition of peptide bonds in separated chains (Ohsaku et al., 1984). Although generally accepted that helix stabilization in gelatin systems is exclusively due to intermolecular triple-chain associations, other means of helix stabilization such as electrostatic and steric have been suggested (Harrington and Rao, 1967). De Wolf and Keller (1996) used CD to elucidate the relation

Figure 11.10 Optical rotation changes caused by thermal denaturation (melting) of native soluble collagen (——) and partially denatured gelatin (– – –). (Adapted from Rose, 1987.)

298 between junction properties and junctions/cross-links. They showed that by applying temperature-dependent CD, it is possible to discern multiple and single helical structures occurring at high and low gelatin concentrations. Moreover, the widely applied random coil reference spectra do not really represent unordered structures while the random coil reference spectra based on the cationic form of poly(lysine) reflected a clear helicity (De Wolf and Keller, 1996).

Proteins and polysaccharides are good film formers and have occasionally been used for food preservation (Keil et al., 1960; Keil, 1961). Although they exhibit satisfactory gas barrier behavior for coating applications, their water vapor permeability values are usually high (Ghosh and Gyani, 1953; Baldwin, 1994; Psomiadou et al., 1996; Arvanitoyannis et al., 1997, 1998a, b). In general, the behavior of non-chemically interacting, mixed polymer systems is dictated by the enthalpy of endothermic segment-segment interactions leading to repulsive forces and mutual exclusion of each polymer from the domain of the other (Ledward, 1994). Protein-polysaccharide interactions have been exploited to recover protein from dilute effluent streams, to fractionate purified proteins, to inhibit protein precipitation, to inactivate enzymes for food preservation, and to stabilize foams and emulsions. However, most interest resides in combining proteins and polysaccharides to prepare novel textured products such as fat replacers (Ledward, 1993). Binary, ternary, or quaternary single-phase blends consisting of gelatin, polysaccharide (i.e., soluble starch, modified starch, or chitosan), and plasticizer (i.e., water, sugars, or polyols) have recently been prepared (Arvanitoyannis et al., 1997, 1998a, b). Plasticizers of low molecular weights lowered both Tm and Tg and increased gas permeation (proportionally to plasticizer content) (Figures 11.6 and 11.11). Cohesion of starch/gelatin or chitosan/gelatin blends increased in the presence of self-associated molecules (i.e., water, polyols). The storage modulus (E ) was clearly observed passing through the glassy, leathery, and rubbery states. A shoulder-like -transition at high sorbitol contents was attributed to the pure sorbitol. Similar behavior was observed for glycerol. The mechanical properties of gelatin-based films depended considerably on film preparation method. Films obtained by evaporation at high temperature (~60 C) were weaker (lower tensile strength and percent elongation) than those prepared by the low temperature process for total plasticizer content (TPC) of up to 25%. At TPC 25%, percent elongation increased considerably. This increase was more pronounced for films of low crystallinity (high temperature process) because such films were more easily plasticized and were more extendable than the respective films of high crystallinity (low temperature process). The more crystalline films had more constraints to elongation because they contained

Gelatin-Based Films

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Figure 11.11 Permeability of carbon dioxide in chitosan/gelatin/water blends vs. inverse temperature (1/T). Presented values are means and standard deviations of at least six replicates. Reprinted from Carbohydrate Polymers, Volume 37, Arvanitoyannis et al., “Chitosan and Gelatin Based Edible Films,” pp. 371–382. Copyright 1998, with permission from Elsevier Science.

higher numbers of triple-stranded crystallites (Table 11.4) (Bradbury and Martin, 1952; Kellaway et al., 1978a, b; Arvanitoyannis et al., 1998a, b). Graft copolymers of gelatin and vinyl monomers, butyl acrylate, and N-vinylpyrrolidone have been studied (Croome, 1982; Gao et al., 1998). Non-specific ionic polysaccharide-protein interactions have been utilized in food applications (e.g., binding fish protein, sausage casings, gums, stabilizers for frozen dairy products, gelatin-phospholipid gels, and carriers for enzymatic systems) (Keeney and Kroger, 1974; Rangayanaki and Stainsby, 1978; Galant, 1983; McKay et al., 1985; Genot et al., 1989; Gennadios et al., 1994). Propylene glycol alginate or pectate esters interacted with gelatin, forming covalently

300 bonded gels with high thermal stability (Tm > 100 C) (McKay et al., 1985). Also, polysaccharide-protein interactions have been applied to microencapsulation (gelatin-gellan), drug release (gelatin-dextran), and tissue adhesion (gelatin-poly[carboxylic acid]) (Chilvers and Morris, 1987; Kurisawa and Yui, 1998; Otani et al., 1998). Quick-setting materials from gelatin both for edible (Shank, 1982, 1984, 1985) and photographic uses (Borchard et al., 1976) have also been discussed.

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304 Shewfelt, R.L. 1987. “Quality of Minimally Processed Fruits and Vegetables,” J. Food Qual., 10:143–156. Slade, L. and Levine, H. 1987. “Polymer-Chemical Properties of Gelatin in Foods” in Advances in Meat Research, Vol. 4, eds., A.M. Pearson, T.R. Dutson and A.J. Bailey, New York: Van Nostrand Reinhold Company, pp. 251–266. Smith, H.L. and Goyan, J.E. 1965. “Method for Studying Rheology of Gelatin Melts,” J. Pharm. Sci., 54:545–548. Sugisaki, M., Suga, H. and Seki, S. 1968. “Calorimetric Study of the Glassy State. IV. Heat Capacities of Glassy Water and Cubic Ice,” Bull. Chem. Soc. Jp., 41:2591–2599. Te Nijenhuis, K. 1981. “Investigation into the Ageing Process in Gels of Gelatin/Water Systems by the Measurements of Their Dynamic Moduli,” Colloid Polym. Sci., 259:1017–1026. Tobolsky, A.V. 1960. Properties and Structure of Polymers. New York: John Wiley & Sons. Vassileva, E., Balta-Calleja, F.J., Esperanza Cagiao, M. and Fakirov, S. 1998. “Gelatin Films with Very High Surface Hardness,” Macromol. Rapid Commun., 19:451–454. Veis, A. 1964. The Macromolecular Chemistry of Gelatin. New York: Academic Press. Von Hippel, P.H. 1967. “Gelatin” in Treatise on Collagen, Vol. 1, ed., G.N. Ramachandran, New York: Academic Press, pp. 38–67. Von Hippel, P.H. and Schleich, T. 1969. “The Effects of Neutral Salts on the Structure and Conformational Stability of Macromolecules in Solution” in Structure and Stability of Biological Macromolecules, eds., S.N. Timasheff and G.D. Fasman, New York: Marcel Dekker, pp. 436–438. Wetzel, R., Buder, E., Hermel, H. and Huttner, A. 1987. “Conformations of Different Gelatins in Solutions and in Films; an Analysis of Circular Dichroism (CD) Measurements,” Colloid Polym. Sci., 265:1036–1045. Witnauer, L.P. and Fee, J.G. 1957. “Effect of Diluents on Fusion Temperature of the Crystalline Regions in Plain and Tanned Cowhide,” J. Polym. Sci., 26:141–150. Wu, C.W., Schrof, W., Lilge, D., Luddecke, E. and Horn, D. 1991. “Holographic Relaxation Spectroscopic Study on the Structure of Gelatin Gel Doped with Fluorescein as a Tracer,” Colloid Polym. Sci., 269:682–688. Wustneck, R., Buder, E., Wetzel, R. and Hermel, H. 1989. “The Modification of the Triple Helical Structure of Gelatin in Aqueous Solution 3. The Influence of Cationic Surfactants,” Colloid Polym. Sci., 267:429–433. Yannas, I.V. 1972. “Collagen and Gelatin in the Solid State,” J. Macromol. Sci.-Rev. Macromol. Chem., C7(1):49–104. Yannas, I.V. and Tobolsky, A.V. 1964. “Viscoelastic Properties of Plasticized Gelatin Films,” J. Phys. Chem., 68:3880–3882. Yannas, I.V. and Tobolsky, A.V. 1968. “High-Temperature Transformations of Gelatin,” Eur. Polym. J., 4:257–264. Young, M.A. and Pysh, E.S. 1975. “Vacuum Ultraviolet Circular Dichroism of Poly(L-proline) I and II,” J. Amer. Chem. Soc., 97:5100–5103.

Films and Coatings from Proteins of Limited Availability

have been widely studied as film-forming materials due to their relative abundance, film-forming ability, and nutritional qualities. Prevalent types of plant and animal protein film-formers have been discussed in other chapters. Nevertheless, these are not the only film-forming proteins available. Other proteins, which are of limited availability, may be of interest due to a unique property they provide to films or an advantage they provide to film formation. Limited availability may be due to relatively low production of the protein source or limitations in recovering the protein as a co-product from a process. This chapter reviews eight additional vegetable proteins and one additional animal protein of limited availability, which have been reported in the literature as film-formers. Protein flours, concentrates, isolates, and solutions from vegetable and animal sources play an important role in film formulation. Acceptability of such protein preparations depends on their functional properties, which are affected by the specific preparation method. These are governed by the physicochemical properties that affect protein behavior in prepared films. The proteins that are reviewed in this chapter are those from peanuts, rice, peas, lupins, grain sorghum, winged beans, pistachios, cucumber pickle brine, and feather keratin. Nature, recovery, film formation, and film properties are discussed for each protein.

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The nutritional value of the peanut as a dietary protein source has been known for a great number of years. Peanut kernels, similar to other oilseeds, are comprised of four major components, i.e., lipids, protein, insoluble residue from protein extraction, and non-protein solubles (Natarajan, 1980). Genetics, growing environment and geographic location, handling, storage, and processing of peanuts can affect nutritional value. Young (1970) showed that amino acid composition of peanut kernels varied substantially among peanut varieties. Peanut kernels have a protein content between 22 and 30% (Ahmed and Young, 1982). The defatted peanut meal contains about 55–60% protein of good nutritional quality (Prakash and Rao, 1986). Fractionated peanut proteins are classified as albumins, arachin, and conarachin (or nonarachin) with the two later fractions composed mainly of complex, high molecular weight globulins (Natarajan, 1980; Prakash and Rao, 1986). Arachin and conarachin make up nearly 75% of the total peanut proteins. They differ mainly in content of sulfur-containing amino acids. Conarachin contains three times more total cysteine and methionine than arachin (Woodroof, 1983). Basha and Cherry (1976) reported that gel electrophoresis of sodium dodecyl sulfate-dissociated globulins showed isolates each containing five different components of molecular weights between 20,000 and 84,000 Da. Peanut protein concentrates and isolates are commercially produced from defatted peanut flour using several methods. Hydraulic pressing, screw pressing, solvent extraction, and pre-pressing followed by solvent extraction have been used for defatting (Natarajan, 1980; Woodroof, 1983). After oil removal, protein isolation and purification are necessary. Peanut concentrates (~70% protein) are produced from dehulled kernels by removing most of the oil and the water-soluble, non-protein components. Techniques for removing water-soluble components include water leaching at the isoelectric pH, aqueous alcohol leaching, air classification, liquid cyclone fractionation, and moist heat denaturation followed by water leaching (Natarajan, 1980). Peanut protein isolates (~90–95% protein) are produced by further processing concentrates to remove water-insoluble oligosaccharides and polysaccharides, and other minor constituents (Natarajan, 1980). The essential amino acid compositions of peanut kernels, flour, concentrate, and isolate are shown in Table 12.1. Peanut proteins are deficient in methionine, lysine, and, possibly, threonine and tryptophan. This limitation must be considered when peanut proteins are added to foods for nutritional purposes.

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Two different methods have been used to prepare peanut protein films. The first method involved formation of peanut protein-lipid films on the surface of heated peanut milk (Wu and Bates, 1973; Aboagye and Stanley, 1985; Del Rosario et al., 1992). This film production on the surface of peanut milk was similar to the traditional production of soy protein-lipid films (“yuba”) in East Asia (Okamoto, 1978). Wu and Bates (1973) prepared protein-lipid films from peanut milk containing 1.6% protein, 3.0% lipid, and 5.6% total solids. Films were successively formed and removed from the surface of peanut milk that was heated (95 5 C) in stainless-steel pans. The peanut films had lower strength than similar films produced from soymilk. Also, considerable excess lipid was evident on the surface of dried peanut films, and rancid odors developed rapidly. Addition of soy protein isolate (SPI) or SPI:sodium caseinate (1:1) effectively improved film strength. Aboagye and Stanley (1985) produced surface films on an aqueous extract of blanched, full-fat peanut by heating the dilute solution (2–5% protein, w/v) to 95 C in a stainless-steel pan. They investigated the effect of pH, heating temperature, and protein concentration on film formation and reported that process temperature was the most influential parameter on both film formation rate and film yield. They determined that peanut proteins underwent a sequential breakdown of higher molecular weight globulins into lower molecular weight fractions followed by reaggregation into an insoluble complex at the surface during film formation. The process was enhanced by alkaline pH conditions and higher protein concentration (up to 5% w/v). However, scanning electron mi-

308 croscopy revealed that elevated pH decreased film strength, and a pH around 7 was recommended for optimal film strength (Aboagye and Stanley, 1985). Del Rosario et al. (1992) prepared protein-lipid peanut films from full-fat and partially defatted peanut milk and reported that fat reduction of milk by creaming, starch removal, and boiling improved film quality but did not increase film yield. Also, film yield from full-fat peanut milk was higher than film yields from low-starch milk and milk low in both starch and fat. However, films from full-fat peanut milk were more susceptible to oxidative rancidity than those from partially defatted peanut milk (Del Rosario et al., 1992). The second method for production of peanut protein films involves casting of peanut protein concentrate or isolate solutions. Jangchud and Chinnan (1999a, b) prepared films from peanut protein concentrate solutions adjusted to pH 6.0, 7.5, or 9.0 and dried at 70, 80, or 90 C. Glycerin, sorbitol, polyethylene glycol, or propylene glycol were added as plasticizers at 0.67, 1.17, or 1.67 g/g protein. Films formed at higher pH (7.5 or 9.0) and higher drying temperature (80 or 90 C) were less moist and sticky at the surface when peeled than films formed at pH of 6.0 and drying temperature of 70 C. Glycerin was determined to be the best performing plasticizer. Barrier and mechanical properties of peanut protein concentrate films plasticized with glycerin (1.67 g/g protein) are shown in Table 12.2 (Jangchud and Chinnan, 1999a). Increasing film drying temperature decreased water vapor permeability (WVP), but pH had no effect. Jangchud and Chinnan (1999a) hypothesized that reduction of WVP with increased drying temperature was due to greater cross-linking, resulting in a tight and compact protein film structure. The effects of pH and drying temperature on oxygen permeability (OP) were similar

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to those on WVP. Mean OP decreased from 15.40 to 2.13 cm3 mm/m2 day kPa when temperature increased from 70 to 90 C (Jangchud and Chinnan, 1999a). Film tensile strength (TS) and elongation (E) increased when film drying temperature increased from 70 to 90 C. This was attributed to increased protein denaturation at the higher temperature, which also likely resulted in tighter, more compact protein film structures. Film color varied from pale-yellow to dark-yellow depending on pH and drying temperature while film solubility decreased with increasing drying temperature and increased with increasing solution pH (Jangchud and Chinnan, 1999a). The sorbic acid diffusivity of peanut 10 12 m2/s protein concentrate films was determined as 8.18–12.60 (Jangchud et al., 1999). The ability of peanut protein films to effectively carry and disperse sorbic acid was limited, and the films were judged to be highly hygroscopic (Jangchud et al., 1999).

Abrasive milling removes the outer tissue material from rice kernels, producing polished rice and the side-products of bran and polish. The content of protein, which is the second most abundant constituent in milled rice following starch, can vary substantially. For example, 14 selected milled rice samples had protein contents from 5.2 to 11.9% (Villareal and Juliano, 1978). Bran contains more of the pericarp, seed coat, nucellus, aleurone layer, and germ than the polish, which contains relatively more starchy endosperm (Juliano, 1985). Usually, 10% by weight of brown rice is removed during milling. Bran accounts for 5–8% of the rough rice weight, and polish accounts for a further 2–3% (Houston, 1972). Rice bran and polish contain 12.0–15.6% and 11.8–13.0% protein, respectively. Amino acid compositions of rice bran and polish are shown in Table 12.3. Although rice bran and polish are rich sources of vitamin B, oil, and good quality protein, they also are rich in phytin, silica, dietary fiber, trypsin inhibitor, and lectin. The outer tissues of the rice grain are rich in albumins (water-soluble proteins) and globulins (salt-soluble proteins), whereas milled rice (endosperm) is rich in glutelins (alkali-soluble proteins). Prolamins (alcohol-soluble proteins) comprise a minor constituent in all milling fractions of the rice grain. Ratios of albumins, globulins, prolamins, and glutelins were reported as 37:36:5:22 for bran, 30:14:5:51 for polish, and 5:9:3:83 for milled rice (Cagampang et al., 1966). Tashiro and Maki (1979) obtained a ratio of 24:44:5:27 for the mentioned rice bran protein fractions and determined lysine contents of 7.7, 5.1, 5.1, and 5.0 g/16 g of nitrogen for crude albumins, globulins, prolamins, and glutelins, respectively. Rice bran protein concentrates are prepared from commercially available

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unstabilized or heat-stabilized rice bran by alkaline extraction and isoelectric precipitation (Gnanasambandam and Hettiarachchy, 1995). Protein solubility and extractability have been reported in descending order: untreated > acid stabilized > parboiled > heat stabilized (Prakash and Ramanathan, 1994). In a concentrate preparation method described by Gnanasambandam and Hettiarachchy (1995), rice bran samples were dispersed (1:4) in technical grade hexane, defatted, and centrifuged. The sediment was extracted once again following the same procedure. Defatted bran samples were dispersed in water at pH 9.5 and were centrifuged to remove insoluble material. Then, the supernatant was adjusted to pH 4.5 to precipitate proteins and was centrifuged again. The precipitate was washed with distilled, deionized water at pH 4.5 and was dispersed in small amounts of distilled, deionized water at pH 7.0. Finally, the dispersions were freeze-dried and stored at 5 C. Recently, Wang et al. (1999) prepared rice bran protein isolates (~92% protein) from unstabilized and defatted rice bran using a combination of phytase and xylanase enzymes. Methods of rice bran stabilization are important when producing protein ingredients from rice bran. Chen and Houston (1970) reported that the yield of extracted protein increased linearly from pH 7.5 to pH 11.0, and then decreased at pH 12.0 due to solubilization of non-protein solids. Lew et al. (1975) prepared protein concentrate from rice bran by alkaline extraction at pH 11.0. It has also been reported that protein purity decreased when extracted at pH 12.0 (Juliano, 1985). The severe protein degradation induced by alkaline pH can reduce the

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nutritive value of the protein (De Groot and Slump, 1969). Also, a combination of high alkaline pH and high temperature resulted in protein degradation and undesirable flavor effects in rice bran protein concentrate (Lynn, 1969). Gnanasambandam and Hettiarachchy (1995) reported that a pH of 9.5 might be considered moderate but useful for preparing rice bran concentrates with >70% protein. Rice protein concentrate can be prepared by alkaline extraction of rice flour or of broken rice kernels, a side-product of rice milling. Alternatively, rice flour or kernels may be treated with enzymes to partially remove the starch component (Chen and Chang, 1984). Rice flour and broken rice kernels are also good sources of sugar and starch. Several functional polysaccharides can be readily generated by sugar or starch fermentation (Catley, 1979).

Gnanasambandam et al. (1997) prepared protein films from rice bran protein solutions (70% protein, dry basis). Glycerol was added at 2% w/v as plasticizer. The solutions were adjusted to either pH 9.5 or 3.0, heated to 80 C, poured onto polyethylene plates, and dried at 60 C. Films prepared at pH 9.5 were darker (had more reddish-yellow tan color) and more transparent than films prepared at pH 3.0. No substantial differences in WVP were observed between rice bran protein films at pH 9.5 and 3.0 (Table 12.4). Rice bran films had higher WVP than synthetic films because of the hydrophilic nature of the protein. The majority of proteins in rice bran are albumins and globulins, which are water- and salt-soluble. Film OP was affected by pH with films prepared at pH 3.0 having lower OP than films prepared at pH 9.5 (Table 12.4). The lower solubility of rice bran protein at pH 3.0 might have resulted in a tighter film structure and, thus, reduced OP (Gnanasambandam et al., 1997). Mechanical properties of protein films provide an indication of expected film integrity under conditions of stress that may occur during processing, handling, and storage. Puncture strength was greater for rice bran protein films prepared at pH 9.5 (1.67 N) than for those prepared at pH 3.0 (0.85 N) (Table 12.4).

312 Rice bran proteins were less soluble at pH 3.0, resulting in fewer protein-protein interactions and hence lower film puncture strength than films prepared at pH 9.5. A similar trend was observed with film TS where films prepared at pH 3.0 had lower TS (7.0 vs. 16.4 MPa) than films prepared at pH 9.5 (Gnanasambandam et al., 1997). Shih (1996) prepared films from mixtures of rice protein concentrate (53% protein content) and pullulan. TS and WVP of rice protein-pullulan films were a function of rice protein concentrate content. Pure pullulan films or those with a low content (50%) of rice protein (Shih, 1996). Addition of propylene glycol alginate (1.5% w/w of total solids) to the film-forming solutions at pH 8.5 increased film TS and water resistance through cross-linking with both amino groups in protein and hydroxyl groups in pullulan.

Peas of various genera, types, and varieties are closely related to beans. In some countries, certain kinds of peas may be called beans and vice versa. Generally, the majority of plants with their pods and seeds known as peas are of the genus Pisum while those called beans are of the genus Phaseolus. Common types of peas are green pea (Pisum sativum), chick pea (Cicer arietinum), lentil (Lens culinaris), and pigeon pea (Cajunus cajan) (Considine, 1982). Peas have long been regarded as a low-cost, healthful food that is rich in protein, vitamin A, and calcium. Pea flour contains nearly half the protein of soybean flour, but pea protein is richer in lysine than soy protein. Nickel (1981) separated pea proteins from starch and fiber by multi-step solubilization at a pH of 2.5–3.0, followed by centrifugation. Naczk et al. (1986) evaluated functional properties of pea protein prepared by Nickel’s method and compared them to those of wheat gluten, soy protein concentrate, and soy protein isolate. The pea protein preparations had a mean crude protein content of 85.3% and an ash content of 4.1–5.0% (Naczk et al., 1986). The pH (about 6.5) of a 10% pea protein dispersion was similar to that of a 10% soy protein isolate dispersion. Pea protein solubility (30.3–41.9%) was greater than that of soy protein isolate (22.2 1.0%) and wheat gluten (16.3 1.0%). The water absorption of pea protein (278–293%) was greater than that of wheat gluten (152 2%), but notably lower than those of soy protein concentrate (445 8%) and soy protein isolate (584 23%). The fat absorption of pea protein (90.1–94.5%) was comparable to that of gluten (96.3 2.2%) but substantially lower than those of soy protein concentrate (157.0 2.2%) and soy protein isolate (144.0 5.5%) (Naczk et al., 1986). The low fat absorption of pea protein

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suggested that there were more hydrophilic than hydrophobic groups on the surface of protein molecules.

Protein film preparation involves formation of a network in which interactions among polypeptide chains are modulated by a plasticizer. Because of their globular structure, pea proteins need unfolding, e.g., under alkaline conditions, prior to film formation. Gueguen et al. (1995, 1998) used an industrial pea (P. sativum L.) protein isolate (70.6% protein; nitrogen factor 5.7) to cast films plasticized with various polyols. Film-forming solutions were prepared by dispersing the isolate (13% w/w) in highly alkaline (pH 12.5) glycine (0.1 M) solutions. The final pH of the film-forming solutions decreased to about 10 due to the buffering action of the protein. Ethylene glycol, diethylene glycol, triethylene glycol, tetraethylene glycol, glycerol, or propane diol was added (100% w/w protein) as a plasticizer. Following centrifugation at low speed to remove air bubbles, the solutions were cast and dried at 60 C (Gueguen et al., 1995, 1998). Glycerol-plasticized pea protein films were sticky, while propane diol-plasticized films became brittle rapidly (Gueguen et al., 1995). Ethylene glycol gave films of greater TS and E and lower WVP than diethylene, triethylene, or tetraethylene glycol (Gueguen et al., 1998). Overall, pea protein isolate films had lower TS and E and greater (by one or two orders of magnitude) WVP than low-density and high-density polyethylene (Table 12.5). Cross-linking with formaldehyde substantially reduced water solubility and increased TS of pea protein isolate films (Gueguen et al., 1998). This was expected due to the high lysine content of pea protein. The -amino group of lysine is considered a primary reactive site between proteins and formaldehyde (Feeney et al., 1975). Succinylation (87% of protein) improved film transparency but did not change film properties (Gueguen et al., 1995).

314

Lupin or lupine are trivial names for plants of the genus Lupinus belonging to the Leguminosae family. The main lupin species are Lupinus albus, L. luteus, L. angustifolius, and L. mutabilis, with the latter being the species most often used for animal feed and human food (Chango et al., 1995). Lupin seeds are rich in protein, with L. luteus having the highest protein content (>42%) (Kiosseoglou et al., 1999). All lupin species contain alkaloids (quinolizidines) at 0–4% w/w of the seed, which impart bitterness or sweetness to a particular variety (Chango et al., 1995). These alkaloids can be removed by soaking seeds in water for several hours (Karara, 1989). Sathe et al. (1982) prepared flour and protein concentrate (alkali solubilization method) from L. mutabilis (cultivar H-6). Proximate compositions of the flour and protein concentrate are shown in Table 12.6. Sodium dodecyl sulfate polyacrylamide gel electrophoresis indicated the presence of 13 and 12 subunits in lupin flour protein and protein concentrate, respectively (Sathe et al., 1982). Lupin seed protein isolates have been prepared by extraction (optimum pH between 8 and 9), acid coagulation, ultrafiltration/ diafiltration, isoelectric precipitation (pH of 4.5), or dialysis (Ruiz and Hove, 1976; Duranti et al., 1983; Kiosseoglou et al., 1999). Lupin seed proteins are either albumins (13%) or globulins (87%) (Duranti et al., 1983). Electrophoresis has fractionated the albumins and globulins into 5 and 11 components, respectively (Kiosseoglou et al., 1999). Amino acid composition profiles of seed proteins from various lupin species can vary considerably (Table 12.7), but all lupin species are deficient in methionine, cystine, and tryptophan (Gross, 1988; Gross et al., 1989).

Protein-lipid films have been prepared on the surface of heated lupin

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seed milk (Karara, 1989; Chango et al., 1995), similar to film formation from soymilk and peanut milk. Karara (1989) prepared lupin seed (L. termis) milk by soaking seeds in water, draining, grinding, and filtering. Soaking at 22 C for 5 h resulted in protein extractability of 94% and was determined as optimal for subsequent milk and film production. According to Wu and Bates (1972), solids concentration, pH, and heating temperature of milk are the most important factors in protein-lipid film formation. Optimal conditions for maximizing yields of protein-lipid films from lupin seed milk were determined as solids concentration of 5–6%, pH of 7–8, and processing temperature of 80 C (Karara, 1989). Proximate compositions of similarly prepared lupin and soy protein-lipid films are shown in Table 12.8. Chango et al. (1995) investigated protein-lipid film formation on the surface of lupin seed (L. luteus) milk heated at 85 C for 1 h. At the natural pH (5.6) of lupin milk, proteins coagulated and did not form films. At acidic conditions (pH 2.5), only weak films were obtained. Film formation was more successful at slightly alkaline conditions (pH 8.5) (Chango et al., 1995), similar to earlier observations by Karara (1989).

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Grain sorghum (Sorghum bicolor L. Moench) ranks fifth among cereals produced in the world (Dendy, 1995). Serna-Saldivar and Rooney (1995) classified grain sorghum according to pericarp color (white, yellow, or red), presence or absence of a pigmented testa (with or without tannins), pericarp thickness (thin or thick pericarp), and endosperm color (normal, heterowaxy, or waxy). Grain sorghum composition varies significantly due to genetic and environmental influences, and is similar to that of maize. Starch (72.3–75.1%) is the major component, followed by protein (11.5–12.3%) and oil (3.6%). The main protein fraction in the sorghum kernel is prolamin, known as kafirin. Kafirin is similar to corn zein in molecular weight, solubility, structure, and amino acid composition (Shull et al., 1991). The amino acid composition of kafirin is shown in Table 12.9. Taylor et al. (1984) determined the average molecular weight of kafirin at 26,300 Da. Kafirin fractionation by SDS-PAGE revealed two major bands with molecular weights of 25,000 and 23,000 Da and four minor bands with molecular weights of 28,000, 20,000, 18,000, and 16,000 Da (Shull et al., 1991). In general, kafirin can be readily extracted with alcohol and/or a reducing

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agent such as 2-mercaptoethanol (Watterson et al., 1990; Shull et al., 1991; Hamaker et al., 1995). In a process described by Hamaker et al. (1995), 90–95% of total protein was extracted from sorghum whole grain flour or sorghum endosperm using a pH 10 buffer containing 1% sodium dodecyl sulfate and 2% 2-mercaptoethanol. Subsequently, t-butanol was added to the extracts at 60% to precipitate non-prolamin proteins. This method showed that kafirin contents in sorghum whole grain flour and in sorghum endosperm were 68–73% and 77–82%, respectively (Hamaker et al., 1995).

Buffo et al. (1997) prepared films from laboratory-extracted (Buffo, 1995) kafirin (89.0% protein, 6.7% crude free fat, and 0.04% ash). Film-forming solutions containing kafirin (25 g), 95% ethanol (105.5 mL), glycerin (6.08 g), and polyethylene glycol 400 (5.48 g) were heated to boiling and cast onto glass plates. The dry films (85 m thick) had similar TS, E, and WVP as films prepared from commercial zein (Table 12.10). The kafirin films were notably darker than the zein films, presumably due to a higher content of inherent pigments in the laboratory-extracted kafirin than in commercial zein (Buffo et al., 1997). Obuz et al. (1998) extruded mixtures of sorghum flour and low-density polyethylene into films. Prior to extrusion, sorghum flour was plasticized with sorbitol, palmitic acid, or stearic acid at weight ratios of 1:1 or 1:2 with sorbitol being the best plasticizer. Films containing sorghum flour were less extendable and weaker than films containing similar amounts of wheat gluten. Film strength decreased with increasing amounts of sorghum flour in the extruded blends (Obuz et al., 1998).

318

Despite the high protein content of its ripe seeds, tubers, and fresh leaves (ranging within 29.3–39.0%, 3.0–15.0%, and 5.0–7.6%, respectively) and the high nutritional quality of its protein, the winged bean (Psophocarpus tetragonolobus) remained an obscure food source until about 30 years ago. Recently, this legume has been receiving attention because of its potential use as a food protein source in the humid tropical regions. The similarity of winged bean seed and soybean in terms of protein and fat contents has been established. Hildebrand et al. (1981) evaluated 240 winged bean accessions obtained from 16 countries and determined seed oil and protein ranges of 7.2–21.5% and 20.7–45.9%, respectively. Of the total proteins in winged bean seed, globulins and albumins accounted for 29–33% and 15–22%, respectively. Electrophoresis on cellulose acetate membranes resolved three protein components labeled psophocarpins A, B, and C, in order of increasing mobility. Psophocarpin A is a single protein comparatively rich in sulfur-containing amino acids. The other two fractions, psophocarpin B and C, are each composed of several related compounds. Polyacrylamide gel electrophoretic studies revealed that winged beans contain predominantly low molecular weight ( anhydrides > esters

carboxylic acids > amides > carboxylates

The -amino group from the NH2 terminal amino acid and lysine side chains in proteins are potential nucleophiles in the reaction. The pH is an important factor because the amine must be non-ionized (pH should be neutral or basic) for the reaction to occur.

Many of the early solid particulate drug delivery investigations used non-biodegradable polymers such as polystyrene (Illum and Davis, 1984). Other examples include poly(ethylene glycol) and poly(vinylpyrrolidone) (Woiszwillo, 1994). Non-biodegradable polymers are difficult to use as implantable microparticulate delivery systems because they are nearly impossible to remove surgically. However, these polymers have potential for oral administration.

Improvements to existing methods are needed to optimize protein stability. Removal of denaturing conditions during encapsulation, such as organic solvents, elevated temperatures, and agitation, is required. Newer polymers that are being investigated do not require organic solvents or elevated temperatures that would inactivate the protein [e.g., poly(phosphazenes) and poly(amino acids)] (Schwendeman et al., 1996). However, these polymers have not been used in FDA-approved products nor have they been extensively tested for protein compatibility. A recent review by Ravi Kumar (2000) discusses novel processing of nano and microparticles. An example is a self-assembled method of preparing protein microspheres reported by Woiszwillo (1994). This method uses

Stabilization of Proteins with Cosolvents

357

no organic solvents, no agitation, low temperature, and polymers [poly(ethylene glycol), PEG and poly(vinylpyrrolidone), PVP] that have been used in FDA-approved products. Therefore, this method of protein microencapsulation has the advantage of milder processing conditions in comparison to other microencapsulation methodologies.

For protein delivery systems to be useful, cosolvents used in processing should act as stabilizers. The mechanism of stabilization for most cosolvents is preferential hydration, which may involve solute exclusion from the protein molecules or protein binding. The interaction between the protein and the stabilizer is a balance between protein binding and exclusion (Arakawa et al., 1990; Timasheff and Arakawa, 1989). Cosolvents can be preferentially excluded by two mechanisms. The first involves interactions with proteins that are determined by the solvent properties. The second mechanism involves repulsive or attractive interactions that are determined by the chemical properties of the protein surface. The potential also exists for both of these mechanisms to occur concurrently (Timasheff and Arakawa, 1989). Steric exclusion results from the size difference between the cosolvent molecules and water (Timasheff, 1995). This is shown schematically in Figure 14.3 (Timasheff, 1995). A shell is formed around the protein that is inaccessible to the cosolvent molecules from chemical inertness (protein and cosolvent) limited by the center to center distance between the protein and cosolvent (Rp + Rc). Water molecules, being much smaller in size, are able to access the protein shell, resulting in a zone of water around the protein molecule (preferential hydration) (Timasheff, 1995). This mechanism accounts for the exclusion of PEG from the protein surface (Lee and Lee, 1981, 1987; Arakawa and Timasheff, 1985b). Preferential exclusion may also occur from interactions between the cosolvents and chemical properties of the protein. Solvophobicity and repulsion from charges on the protein surface are two such mechanisms. The solvophobic effect occurs when contacts of non-polar regions of a protein and the water-cosolvent mixture are more entropically unfavorable than contact with water. Solvophobicity is possibly due to stronger hydrogen bonding than in pure water, and the cosolvent molecules move into the bulk solvent to maintain constant chemical potential (activity) of the cosolvent and water in the system. This results in a shell of water as described above (Gibbs, 1878). Glycerol is one such molecule that exhibits the solvophobic effect (Gekko and Timasheff, 1981; Cleland, 1997a, b). The net interaction of protein stabilization by cosolvents is preferential

358 hydration; however, this does not exclude the possibility that the cosolvent molecules may be excluded and bound to the protein surface. This situation has been reported for salts, which are excluded from the protein surface, evidenced from the increase in the interfacial tension of water, but can weakly bind to charges on the protein surface (Arakawa and Timasheff, 1984; Arakawa et al., 1990). Figure 14.4 lists cosolvents shown to bind non-specifically to several proteins (positive preferential cosolvent interaction) as well as cosolvents that preferentially hydrate proteins, resulting in a negative preferential interaction (Cleland et al., 1993, 1994a). The effect of the cosolvent on the unfolding of a protein is complex. Protein structural changes can alter the protein solvent interactions. These cosolvent effects depend on a balance of the interaction forces between the native and denatured protein and may involve a combination of exclusion and binding. The effect of a cosolvent on the native to denatured protein equilibrium requires knowledge of the preferential interactions in both the native and denatured state (Arakawa et al., 1990; Timasheff, 1995).

Figure 14.3 Schematic representation of varying degrees of preferential hydration from cosolvent molecules. (Adapted from Timasheff, 1995.)

Summary

359

Figure 14.4 Protein and cosolvent interaction diagram (Cleland et al., 1993, 1994a). The cosolvents with positive preferential interaction may bind non-specifically to proteins, and cosolvents with negative preferential interaction may preferentially hydrate proteins (Lee and Lee, 1981, 1987; Lee and Timasheff, 1981; Arakawa and Timasheff, 1982a, b, 1984, 1985a, b; Timasheff and Arakawa, 1989; Arakawa et al., 1990; Timasheff, 1995).

Proteins are used in microencapsulation of pharmaceuticals for encapsulating drugs and as the actual encapsulant material themselves for drug delivery. Presently, the latter has received more attention as the controlled delivery of proteins has benefits in many areas of therapeutics (e.g., cancer, AIDS, and diabetes). These benefits include targeted drug delivery; availability of many processing methods; wide variety of physicochemical properties; and FDA-approved synthetic and natural polymers. However, there are a finite number of microparticle formulations that are approved for human use. Currently available microsphere formulations that deliver proteins as the active agent include Lupron Depot (leuprolide acetate, Takeda-Abbott), Zoladex (goserelin acetate, AstraZeneca), Decapeptyl (triptorelin, Debiopharm), octreotide acetate (Sandostatin LAR Depot, Novartis), and somatropin (Nutropin Depot , Genentech). Microparticle formulations that deliver

360 non-protein drugs as the active agent include Pancrease (pancrelipase, McNeil), Ultrase (pancrelipase, Axcan Scandipharm), Cipro Oral Suspension (ciprofloxacin, Bayer), Naprelan (naproxen sodium, Carnrick), Retin-A MICRO (tretinoin gel using MICROSPONGE system, Ortho Dermatological), and Micro-K Entencaps (potassium chloride, Ther-Rx). The commercially available products are successful, and further advancements with microparticles will result in systems with improved encapsulation efficiency that prevent drug or protein inactivation.

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364 Latha, M.S. and Jayakrishnan, A. 1995. “A New Method for the Synthesis of Smooth, Round, Hydrophilic Protein Microspheres Using Low Concentrations of Polymeric Dispersing Agents,” J. Microencapsul., 12:7–12. Lavelle, E.C., Yeh, M.K., Coombes, A.G. and Davis, S.S. 1999. “The Stability and Immunogenicity of a Protein Antigen Encapsulated in Biodegradable Microparticles Based on Blends of Lactide Polymers and Polyethylene Glycol,” Vaccine, 17:512–529. Lee, J.C. and Lee, L.L. 1981. “Preferential Solvent Interactions between Proteins and Polyethylene Glycols,” J. Biol. Chem., 256:625–631. Lee, J.C. and Timasheff, S.N. 1981. “The Stabilization of Proteins by Sucrose,” J. Biol. Chem., 256:7193–7201. Lee, L.L. and Lee, J.C. 1987. “Thermal Stability of Proteins in the Presence of Poly(ethylene glycols),” Biochem., 26:7813–7819. Li, S., Schoneich, C. and Borchardt, R.T. 1995. “Chemical Instability of Proteins,” Pharm. News, 2(5):12–16. Li, J.K., Wang, N. and Wu, X.S. 1998. “Gelatin Nanoencapsulation of Protein/Peptide Drugs Using an Emulsifier-Free Emulsion Method,” J. Microencapsul., 15:163–172. Madan, P.L., Jani, R.K. and Bartilucci, A.J. 1978. “New Method of Preparing Gelatin Microcapsules of Soluble Pharmaceuticals,” J. Pharm. Sci., 67:409–411. Madan, P.L., Luzzi, L.A. and Price, J.C. 1972. “Factors Influencing Microencapsulation of a Waxy Solid by Complex Coacervation,” J. Pharm. Sci., 61:1586–1593. Magdassi, S. and Vinetsky, Y. 1996. “Microencapsulation of Oil-in-Water Emulsions by Proteins” in Microencapsulation Methods and Industrial Applications, ed., S. Benita, New York: Marcel Dekker, pp. 21–33. Manning, M.C., Patel, K. and Borchardt, R.T. 1989. “Stability of Protein Pharmaceuticals,” Pharm. Res., 6:903–913. Masters, K. 1985. Spray Drying Handbook. New York: John Wiley & Sons. Mathiowitz, E. and Cohen, M.D. 1989. “Polyamide Microcapsules for Controlled Release. V. Photochemical Release,” J. Membr. Sci., 40:67–86. Mathiowitz, E. and Langer, R. 1987. “Polyanhydride Microspheres as Drug Carriers I. Hot Melt Microencapsulation,” J. Contr. Rel., 5:13–22. Mathiowitz, E., Kreitz, M.R. and Brannon-Peppas, L. 1999. “Microencapsulation” in Encylopedia of Controlled Delivery, Vol. 2, ed., E. Mathiowitz, New York: John Wiley & Sons, pp. 493–546. Maysinger, D., Filipovic-Grcic, J. and Cuello, A.C. 1993. “Effects of Coencapsulated NGF and GM1 in Rats with Cortical Lesions,” Neuroreport, 4:971–974. McGinity, J.W. and O’Donnell, P.B. 1997. “Preparation of Microspheres by the Solvent Evaporation Technique,” Adv. Drug Deliv. Rev., 28:25–42. Morimoto, Y. and Fujimoto, S. 1985. “Albumin Microspheres as Drug Carriers,” Crit. Rev. Therap. Drug Carr. Systems, 2:19–63. Morlock, M., Kissel, T., Li, Y.X., Koll, H. and Winter, G. 1998. “Erythropoietin Loaded Microspheres Prepared from Biodegradable LPLG-PEO- LPLG Triblock Copolymers: Protein Stabilization and in-vitro Release Properties,” J. Contr. Rel., 56:105–115. Nellore, R.V., Pande, P.G., Young, D. and Bhagat, H.R. 1992. “Evaluation of Biodegradable Microspheres as Vaccine Adjuvant for Hepatitis B Surface Antigen,” Parenter. Sci. Technol., 46:176–180. Nixon, J.R., Khalil, S.A. and Carless, J.E. 1968. “Gelatin Coacervate Microcapsules Containing Sulphamerazine: Their Preparation and the in vitro Release of the Drug,” J. Pharm. Pharmacol., 20:528–538. Okada, J., Kusai, A. and Ueda, S. 1985a. “Core Treatment for Improving Microencapsulability in Simple Gelatin Coacervation Method,” J. Microencapsul., 2:175–182. Okada, J., Kusai, A. and Ueda, S. 1985b. “Factors Affecting Microencapsulability in Simple Gelatin Coacervation Method,” J. Microencapsul., 2:163–173. Palmieri, G.F., Lauri, D., Martelli, S. and Wehrle, P. 1999. “Methoxybutropate Microencapsulation by Gelatin-Acacia Complex Coacervation,” Drug Dev. Ind. Pharm., 25:399–407.

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Hard Gelatin Capsules

G

ELATIN capsules are small, usually oblong, containers used to enclose a va-

riety of substances. They are classified as solid dosage forms and are generally intended for oral use but may also be used vaginally and rectally. Gelatin is the main ingredient used to prepare capsules because it produces relatively strong films at room temperature and readily dissolves after ingestion or insertion. Depending upon their gelatin composition, capsules may be classified as soft or hard. Soft gelatin capsules (SGC) are one-piece containers, which contain plasticizers to make them flexible. On the other hand, hard gelatin capsules (HGC) are comprised of two pieces that combine to form a closed container and do not contain any plasticizers, which gives them a rigid structure. This chapter will cover the raw materials used to make HGC, the techniques used for both preparing and filling HGC, and the release of drugs from HGC (Ansel, 1985; Jones, 1990; Rudnic and Schwartz, 1990; Augsburger, 1996).

In 1833, Mothes invented the first gelatin capsule, which by today’s standards may be classified as an SGC. These capsules were developed primarily to 367

368 mask the poor taste of the medications being used at the time. It was not until 1846 when Lehuby patented what he called a medicinal envelope that the first two-piece HGC appeared. But in 1848, Murdock became the first person to actually describe a two-piece HGC in his patent and is often given credit for its invention (Jones and Turner, 1974). Initially, HGC were less popular than their “soft” gelatin counterparts because they were more expensive to produce, and, at this time, most medicines were either liquids or semi-solids, which required the use of a SGC (Jones, 1987a). However, because of the development of better equipment and the abundance of solid drug entities, HGC are now ten times more likely to be used in the manufacture of solid dosage forms than SGC (Delaney, 1982).

In addition to masking taste, HGC offer several advantages over other solid dosage forms, but they also have certain disadvantages depending upon the situation (Augsburger, 1996). HGC allow for greater flexibility during formulating and manufacturing because the compaction process used for tableting is not required. However, capsule products are generally more expensive to produce than tablets because of limited capsule suppliers and slower filling equipment. There is also the concern of interactions between the capsule fill and the gelatin capsule itself, especially in the presence of moisture. Many patients find it easier to swallow capsules than tablets because of their shape and slickness when wetted, but some individuals (such as children and the elderly) may still have difficulty and may not be able to use them. Powder-filled capsules are generally thought to have greater bioavailability (the relative amount of drug that reaches the bloodstream) than tablets because the gelatin shell rapidly dissolves, whereas tablets must first disintegrate or break up before dissolving (Augsburger, 1996). However, some capsule formulations have shown diminished capsule shell dissolution and subsequent drug release problems (Johnson et al., 1977; Chafetz et al., 1984; Martin et al., 1985; Mohamad et al., 1986; Dey et al., 1993). In addition, there may be cases where rapid dissolution of the capsule and quick drug release is not desirable. In these cases, capsules cannot be used unless they are first modified in such a way as to limit dissolution.

Gelatin is the major ingredient used to manufacture capsules and is prepared by partially hydrolyzing collagen, a naturally occurring fibrous protein found in animal skins, bones, and connective tissues. Hydrolysis may be done under acidic conditions to produce type A gelatin or under basic conditions to produce type B gelatin. Gelatin has a number of unique properties but the two most important to capsule manufactures are Bloom strength and viscosity. Materials

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such as starch have been used as gelatin extenders to help lower costs by reducing the amount of gelatin needed during manufacturing (Jones, 1987b). Recently, a method to prepare gelatin-free capsules made of a cellulose ether derivative has been developed (Grosswald et al., 1998); however, gelatin continues to be the predominant material used for capsule manufacturing. Along with gelatin, several other agents are used to prepare capsules. Coloring and opacifying agents are used for a number of reasons, including improving patient compliance and protecting capsule contents from light. Processing aids such as surfactants and preservatives may be incorporated into a gelatin formulation, and protective or enteric coatings may be used to improve product stability or alter capsule solubility in a specific medium.

Several of the amino acids in gelatin contain ionizable groups, which give gelatin a specific net charge depending upon the gelatin type and pH of the solution. The net charge of the gelatin affects molecular interactions and contributes to differences between the properties of each gelatin type (Table 15.1). Type B gelatin is prepared by basic hydrolysis, which causes a greater degree of deamidation of the asparagine and glutamine amino acids than acid hydrolysis. This gives type B gelatin a larger number of free carboxyl groups than type A and results in each having its own unique isoelectric point. The isoelectric point (IEP) of a gelatin solution is the pH at which there is an equal number of negative and positive charges so that the net charge on the gelatin molecule is zero. Type A gelatin has an isoelectric point of 7.0–9.0, whereas Type B gelatin has an isoelectric point of only 4.8–5.0 because of the greater degree of deamidation and the larger number of free carboxylic acid groups. Cooling sufficient concentrations of aqueous gelatin solutions below 40 C allows gelatin molecules to renature or partially reform themselves back into their original collagen structure (Veis, 1964a). This produces a three-dimensional network and causes gel formation, which is essential for capsule manufacturing. The strength or rigidity of the resulting gel is very important for capsule production and is related to the molecular weight of the gelatin. Several factors including concentration, pH, maturing time, and temperature also affect the gel strength. Capsule manufacturers often use Bloom strength as a quantitative measurement of gel rigidity. Bloom strength has usu-

370 ally been measured under standard conditions using a Bloom gelometer, which determines the gram weight needed to depress a standardized plunger 4 mm into the surface of a 6.67% w/w gel after maturing the gel at 10 C for 17 hours (AOAC, 1999). However, the Bloom gelometer has largely been replaced by the Stevens LFRA/Voland Texture Analyzer and the TA.XT2 Texture Analyzer. Gelatins ranging in Bloom strength from 150 to 280 g are most often used for capsule manufacturing (Augsburger, 1996). The viscosity of the gelatin solution used during manufacturing is also important for capsule production. During manufacturing, viscosity is used to control capsule wall thickness and ensure that the two pieces of the HGC fit together properly. The viscosity of gelatins used for capsule preparation is measured as a 6.67% (w/w) solution at 60 C and is usually determined with either an Ostwald or a pipette viscometer (Jones, 1987c). The values generally range from 30 to 60 milliPoise (Augsburger, 1996). The primary determinants of viscosity are concentration and molecular weight, but pH is also an important determining factor. The pH of the gelatin solution determines the net charge of the gelatin molecule and, therefore, the repulsion of ionizable groups. As the net charge increases, electrostatic repulsion increases and uncoils gelatin molecules, which increases the viscosity of the solution (Veis, 1964b). Because of this, gelatin viscosity has its highest value at approximately pH 3.0 and 10.5 and its lowest value around the IEP (Sheppard and Houck, 1930). At both high gelatin concentrations and extreme pH values, the effect of pH on viscosity is lessened due to the increasing presence of counter-ions, which swamp out the electrostatic repulsive effect and, therefore, reduce viscosity (Veis, 1964b).

Coloring agents are commonly used to identify specific drug products during manufacturing, dispensing, or patient usage. In addition to identification, inclusion of coloring agents into a capsule offers several advantages. Reportedly, color influenced both patient compliance and perceived effectiveness (Lucchelli et al., 1978; Bailie and Kesson, 1981; Buckalew and Coffield, 1982). Certain colors were associated with certain treatments such as lavender for hallucinogenic effects, white for analgesia, and orange/yellow for stimulants and antidepressants (Buckalew and Coffield, 1982). Colorants may also be used to maintain patient confidence in products that vary in appearance and prevent identification of products being used during clinical studies by concealing capsule contents. This ability to conceal can also be used to protect substances that are susceptible to light degradation by blocking the detrimental wavelengths of light (Jones, 1987b). Two types of colorants, water-soluble dyes and insoluble pigments, may be used to color capsules. Soluble dyes reveal their color by absorbing certain wavelengths of visible light, whereas insoluble pigments reveal their color by

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reflecting certain wavelengths. Water-soluble dyes are mainly synthetic and include materials from many different chemical classes with the most common class being the azo dyes. These dyes contain an azo-linkage, –N苷N–, which is responsible for their color production. A commonly used class of insoluble pigments is the iron oxides, of which black, red, and yellow are the most common. Titanium dioxide is a white pigment (opacifier) and may be the most frequently used colorant in all of manufacturing. In the past, several colorants such as amaranth (formally FD&C Red No. 2) have been reported to be toxic, so safety is a concern when manufacturing colored capsules (Jones, 1990). For this reason, use of colorants in the U.S. is under the control of the American Food and Drug Administration (FDA).

Several materials such as surfactants and preservatives may be used as processing aids during HGC manufacturing. Surfactants such as sodium lauryl sulfate are commonly added to the gelatin solution to lower its surface tension. This improves the interaction between the gelatin solution and the processing machinery, which ensures capsule wall uniformity. In addition to surfactants, preservatives such as sulfur dioxide and parabens are commonly added to the gelatin solution because gelatin is an ideal growth medium for bacteria and fungi, especially at the slightly elevated temperatures used during manufacturing. The United States Pharmacopoeia (USP) limits sulfur dioxide, added as either sodium sulfite or sodium metabisulfite, to a concentration of < 0.15% (USP XXIII, 1995). Parabens are used at a concentration of 0.2% (w/w) and, usually, in combinations (such as a 4:1 methyl/propyl paraben mixture).

After manufacturing, the capsules may be coated to protect them from moisture present during storage, thereby improving the stability of the final product. Also, an enteric coating may be applied to alter the capsule’s performance in the biological environment. Enteric coatings, such as cellulose acetate phthalate, resist dissolution in the acidic stomach environment but readily dissolve in the environment of the small intestine (Porter, 1990). Enteric coatings are used for multiple reasons, including protecting capsule contents from the acid or enzymes present in the stomach, protecting the stomach from drugs that may be irritating or harmful, and controlling the site of drug delivery in the gastrointestinal tract. Although coatings offer several advantages, their use creates additional problems. A major concern with applying coatings is that additional processing is required. This not only increases the time needed to make the final product but it also exposes capsules to additional forceful actions that may damage them or even cause them to separate (Jones, 1987b).

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Because HGC are rather expensive to manufacture, only a few companies such as Capsugel, a division of Warner-Lambert, and the Japanese company Shionogi Qualicaps manufacture them. However, some companies such as Eli Lilly market them after they are contract manufactured by another company. Capsules are manufactured using a process similar to that first described by Lehuby in his original patent more than 150 years ago although it has since been refined and automated. Today, HGC are manufactured primarily on Colton machines that were invented more than 50 years ago. The manufacturing process involves several steps (Norris, 1959, 1961; Martyn, 1974–75; Jones, 1982, 1987d; Hostetler, 1986). First, the proper raw materials are prepared and mixed to produce the desired capsule properties. Then, capsules of a certain size and shape are prepared using the appropriate equipment. After preparation, the capsules are sorted, and the defects are removed. Finally, the capsules are printed if needed and properly stored. The completed HGC must meet a number of standards after production set forth by both the USP and the American Federal Standards for Capsules.

The manufacturing process begins with the preparation of a 30–40% (w/w) gelatin solution by dissolving the gelatin directly into purified water at 60–70 C or by first hydrating the gelatin in cold purified water before dissolving (Jones, 1987d). At these elevated temperatures, the gelatin molecule may undergo partial hydrolysis, which changes its molecular weight. Therefore, Bloom strength and viscosity may be altered and the resulting capsule properties affected. To prevent significant hydrolysis from occurring, small batch sizes are prepared. In addition, the gelatin solution is highly concentrated and, therefore, highly viscous. To prevent air entrapment in the solution, a vacuum is usually applied during preparation. Once the gelatin solution is prepared, 20to 30-liter aliquots are mixed with the appropriate amounts of additives such as colorants and processing aids (Jones, 1990). The viscosity of the final solution is determined and adjusted to provide the desired capsule wall thickness. The final gelatin solutions are delivered to the capsule manufacturing machinery and stored in temperature-controlled holding tanks. The gelatin solution is fed from the holding tanks into a “dip pan” which is maintained at 45–55 C (Jones, 1987d). The level of solution within the pan is held constant using a level-sensing device (Jones, 1982). When the amount of gelatin solution in the pan becomes too low, more solution is pumped in from the holding tanks. Because the pan is maintained at an elevated temperature, water is continually being evaporated from the gelatin solution, which increases its viscosity. To maintain a consistent viscosity, the gelatin solution that is subsequently

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pumped into the pan to maintain a constant solution level has a lower viscosity than the initial solution (Jones, 1987d). In other words, a gelatin solution with a lower concentration is subsequently added to replace the water lost though evaporation to maintain a constant viscosity. Sets of lubricated mold pins on a steel plate form what is termed a pin bar and are used to prepare the capsules. The mold pins are slightly tapered, 0.1 to 0.3 mm/cm of length, toward the closed capsule end (Jones, 1987d). If the capsule walls were made parallel, a vacuum would be created upon capsule removal from the molds, possibly causing the capsule walls to collapse. The caps and bodies of the capsules are produced simultaneously on opposite sides of the machine, and the mold pins used for each are similar except the cap molds have a slightly larger diameter. The mold pins are at approximately 22 C when dipped into the heated gelatin solution, which causes the gelatin to immediately gel onto the molds (Augsburger, 1996). The molds remain in the gelatin solution for about 12 seconds, depending upon the capsule size being produced with larger capsules requiring longer dipping times (Augsburger, 1996). The molds are then slowly withdrawn, allowing the excess gelatin solution to run off (Figure 15.1). A higher viscosity causes a greater amount of gelatin to be picked up and remain on the molds after removal, which creates a thicker capsule wall (Jones, 1990). After dipping, the pin bars are transferred from the lower level to the upper level of the machine. As they do this, they are rotated horizontally to evenly spread the gelatin over the mold pins and are passed through a stream of cool air to harden the gelatin. Once the pin bars reach the upper level, the gelatin is com-

Figure 15.1 Illustration of the hard gelatin capsule dipping process.

374 pletely set, and the molds are now facing upward. To thoroughly dry the gelatin, the pin bars are passed through a series of ovens where humidity-controlled air at 22–28 C is passed over them (Jones, 1987d). Temperatures only slightly above ambient are used to prevent gelatin melting and/or overdrying. If overdrying occurs, the capsules may become too brittle for subsequent use and may even split on the pins due to shrinkage. After drying, the pin bars are cooled down to ambient temperatures. Sets of metal jaws remove the dried capsules from their mold pins and place them into a metal holder. The capsules are then rotated against a sharpened knife and trimmed to the desired length. The trimmings are removed by suction and recycled while the capsules are placed into a central joining block. The caps and bodies are fitted together to a pre-set length termed the unclosed join length, which is designed so that the capsules are closed tight enough to prevent separation upon handling but are still separable prior to filling (Jones, 1990). This entire process lasts about 45 minutes with capsule drying requiring about two-thirds of the time (Augsburger, 1996). Throughout the cycle, an operator is sampling the capsules to check both capsule length and wall thickness, and he or she may adjust the machinery based on this information. Once the capsules are joined, they are sorted and visually inspected for defects. Defects are categorized by their severity (Jones, 1987e). Critical defects such as improper sizes, dents, and holes could cause stoppage of filling machines. Major defects such as thin walls and splits could compromise dosage integrity and reduce capsule effectiveness. Minor defects such as small bubbles or specks within the film deter from the overall capsule appearance. The capsules may then be printed with a variety of information, which can be done either axially along the capsule length or radially around its circumference using specially designed equipment (Augsburger, 1996). The capsules are then evaluated for printing defects, packaged, and stored. The USP XXIII requires that empty HGC be stored in tight containers protected from potential microbial contamination. Capsules can usually be stored at 10–30 C and relative humidity (RH) of F% for extended periods of time (Jones, 1987d).

A variety of sizes and types of HGC are currently being manufactured. Capsules for human use are prepared in eight different sizes, 000, 00, 0, 1, 2, 3, 4, and 5, with 000 being the largest and 5 being the smallest (Figure 15.2). They differ in both length and diameter. Capsules larger than size 000 are also manufactured but they are mainly for veterinary use. Drug products are usually made with a capsule size that is small enough for the patient to easily swallow but large enough for the capsule to be easily processed and handled. Because of this, the capsule sizes most commonly used for drug products are 0, 1, and 2. Most capsules are prepared in the traditional oblong shape except for the prod-

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Figure 15.2 Comparison of hard gelatin capsule sizes for human use (from left to right, 000, 00, 0, 1, 2, 3, 4, and 5).

ucts manufactured by Eli Lilly and SmithKline Beecham. Lilly prepares its products using Pulvule capsules, with bodies that are tapered at the closed end to give them a bullet-like shape. SmithKline, which recently ended its capsule manufacturing, had prepared its products using Spansule capsules with ends are angular instead of rounded (Rudnic and Schwartz, 1990). A major concern with capsule products is the possible separation of the cap and body after filling, which not only causes product loss but also enables product tampering. Capsule manufactures have developed methods to prevent this separation. A number of capsules are prepared with a self-locking mechanism. Indentations are formed within the cap and body so that when the capsule is closed, these indentations fit snuggly together and prevent separation (Figure 15.1). Products of this type include Lok-Cap and Posilok capsules from Lilly and Snap-Fit and Coni-Snap capsules from Capsugel, who at the time of their invention was a division of Parke-Davis (Cole, 1987a). HGC may also be sealed after filling by spot welding or banding. Capsules are spot welded using hot melt pins, which press against the cap and fuse it to the body (Belthoff, 1986). Capsules are banded by applying a strip of melted gelatin around the bottom end of the cap, which cools and seals the cap to the body. Banding has been used often by Parke-Davis for its Kapseals products (Rudnic and Schwartz, 1990).

Empty HGC must meet several industrial standards set forth by the American Federal Standards for Capsules (Am. Fed. Std. No. 285A, 1976). In addition to being free of the mentioned defects, the capsules must meet standards for dimensions, solubility, moisture content, and odor. The most important capsule dimensions are the cap length and two particular diameters. The cap length must be exactly one-half the total capsule length after closing. The diameter of

376 the body opening influences the joining of the cap and body. The diameter of the cap at the point it meets the end of the body after capsule closing influences how well the capsule stays together. For optimal performance, the empty capsule must remain undissolved in water at 25 C for 15 minutes but completely dissolve or disintegrate within 15 minutes when placed in a simulated gastric environment such as 0.5% (w/w) HCl (~ pH 1.0) at 36–38 C. Although the regulation specifies dissolution in an acidic pH, gelatin capsules should dissolve in water at this temperature within this time. An appropriate moisture content for HGC is 13.0–16.0% (w/w). Lower moisture contents can make the capsules too brittle and cause splitting; whereas higher moisture contents could make the capsules deform and become sticky. To maintain patient acceptability, the capsules must not develop any foreign odors after storage in a sealed bottle for 24 hours at 30–40 C.

HGC are primarily filled with powder formulations but may also be used to encapsulate a variety of materials such as pellets, tablets, smaller capsules, and even liquids. The first powder filling techniques for HGC appeared in the late nineteenth century. These techniques were manual until the development of the first automatic capsule filling machines in the early twentieth century (Cole, 1987b). Today, there are several types of capsule-filling machines available (Cole, 1987c), but they all follow the same basic procedure. First, the capsules are oriented into the proper position, and the caps and bodies are separated. The capsule bodies are then filled with the appropriate materials, and the capsules are rejoined and closed. Finally, the capsules are cleaned, polished, and stored. The powder formulation’s characteristics are very influential in determining successful filling, and, like empty HGC, filled capsules are required to meet certain pharmacopoeial standards.

The most basic capsule-filling technique, often termed the punch method (Ansel, 1985), is to fill the capsules by hand. This is mostly done in pharmacies when specialized drug products that are not currently manufactured need to be dispensed. First, the powder formulation is prepared by uniformly mixing or triturating together the appropriate ingredients using a mortar and pestle (O’Connor et al., 1990). After mixing, the powder is placed on a clean surface and compressed into a “square” pile using a spatula. An appropriately sized empty capsule is separated into its cap and body halves, and to keep the capsule body clean, either an extra cap is placed over its closed end or gloves are worn. The body is then stamped or “punched” repeatedly into the previously com-

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pressed pile of powder. Once the body is full, the cap is loosely placed back on, and the capsule is weighed. To meet the desired capsule weight, powder is either removed or further packed into the body or cap. Complete filling of the cap is avoided to ensure a proper fit upon closure. Once the capsule is at the proper weight, it is securely closed and gently tapped onto a hard surface to evenly distribute the powder throughout the capsule. The capsule is finally “polished” or cleaned of excess powder before dispensing.

Preparing capsules one at a time can be very time consuming even on a small scale, so a number of hand-operated capsule-filling devices have been developed. These machines can prepare 24, 96, 100, or 144 capsules per batch and have an output ranging from 2000/day to 2000/hour (Ansel, 1985). Generally, all hand-operated devices operate in a similar fashion (Figure 15.3). Empty capsules of a particular size are placed into the device, body end down, either by hand or by using a capsule rectifier. The rectifier orients the capsule with its body end down by utilizing the difference between the capsule’s cap and body diameters. Once the capsules are properly filled into the machine, a lid is placed over them, and the bodies are clamped into position. The upper part of the device is then separated from the lower part, which separates the caps from the bodies. The clamp holding the bodies in place is loosened, allowing them to fall into the machine so that their open ends are flush with the top of the machine’s surface. The previously prepared powder formulation is then uniformly spread over the surface of the machine, and, as this is done, the powder falls into the bodies. For higher fill weights, the powder may need to be packed into the body using a tamping device. Once the appropriate amount of powder is filled into

Figure 15.3 Illustration of a hand-operated hard gelatin capsule-filling device (see text for description).

378 the bodies, the upper part of the machine is rejoined with the lower part, thereby placing the caps back onto the bodies. The capsules are then emptied from the device, securely closed, and cleaned before dispensing. On a large scale, HGC are filled using either semi-automated or automated filling machines. Semi-automated machines require the presence of an operator at all times and can fill 120,000–160,000 capsules per eight-hour shift, whereas the automated machines can fill the same number of capsules in about one hour (Augsburger, 1996). Semi-automated and automated machines use the same general filling process (Cole, 1987d; Augsburger, 1996; Jones, 1990; Rudnic and Schwartz, 1990). First, the capsules are oriented body-end down. The bodies are then separated from the caps by vacuum suction and are filled with the powder formulation according to a certain filling technique (Ridgway and Callow, 1973; Augsburger, 1974; Jones, 1985; Hostetler, 1986). Once the bodies are filled, the capsules are rejoined, securely closed, and removed from the machine. Automated machines may be classified as either intermittent or continuous depending upon their operation. Intermittent machines must start and stop at various steps in the filling process while continuous machines complete each step simultaneously and therefore have a faster filling rate. The four most common capsule filling techniques are the auger, vibratory, pisto-tamp, and vacuum fill methods (Augsburger, 1996). The auger and vibratory methods fill the powder formulation directly into the capsule body and are, therefore, considered direct techniques, whereas the pisto-tamp and vacuum fill methods use independent equipment to fill the capsule body and are, therefore, considered indirect techniques (Jones, 1990). The auger method (Reier et al., 1968; Ito et al., 1969) delivers the powder formulation from a holding container to the capsule body using a rotating auger while the vibratory method (Kurihara and Ichikawa, 1978) uses a vibrating plate to fluidize the powder formulation and initiate flow (Jones, 1990). The pisto-tamp method is used by both dosing-disk and dosator machines to first form a powder compact, which is then placed into the capsule body. Dosing-disk machines (Ridgway and Callow, 1973; Kurihara and Ichikawa, 1978; Shah et al., 1983) deliver the powder formulation onto disks containing particular sized cavities. A set of pistons compresses the powder into these cavities to form the compacts (Figure 15.4). On the other hand, dosator machines (Small and Augsburger, 1977; Jolliffe and Newton, 1978; Jolliffe et al., 1979) use a cylindrical dosing tube containing a movable piston. The tube is plunged into the powder formulation, and the piston is then compressed to form the compact (Figure 15.5). The vacuum fill technique (Ridgway and Callow, 1973) also uses a cylinder/piston system, but in this case a porous piston is preset within the cylinder to create a particular sized cavity. The tube is dipped into the powder bed where a vacuum, which is applied through the piston, holds the powder in the cavity. The powder is then emptied into the capsule body by removing the vacuum and forcing air back through the piston.

Figure 15.4 Illustration of the dosing-disk hard gelatin capsule filling method. The pistons are arranged circularly above the dosing-disk (see text for description).

Figure 15.5 Illustration of the dosator filling method (see text for description).

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The powder formulation itself is very influential in determining successful capsule filling. In most cases, the drug’s dose and physical characteristics determine both the appropriate capsule size to be used and the need for other formulation ingredients. The drug’s dose is usually lower than the amount needed to totally fill a conveniently sized capsule. In this case, the drug is first mixed with an appropriate amount of filler or diluent before the capsules are filled. These diluents can neither react with gelatin nor have a high moisture content, which will allow the gelatin capsule to absorb water, causing it to become distorted (Jones, 1990). The most commonly used diluents for capsule filling are starch, lactose, dicalcium phosphate, and microcrystalline cellulose (Augsburger, 1996). These diluents may also be used to improve the compactibility of poorly compressible drugs, thereby making capsule filling easier. The type of equipment used for capsule filling is also an important consideration when developing the powder formulation (Jones, 1988). To ensure rapid and uniform filling, especially when using direct filling techniques, the formulation must have good flowability. Also, to prevent the formulation from sticking to the machine, especially when using indirect filling techniques, the formulation must have minimal powder/machine interactions. Specially prepared diluents such as pregelatinized starch, spray processed lactose, and unmilled dicalcium phosphate dihydrate (Augsburger, 1996) may be used to help improve these formulation properties. In addition to diluents, glidants may be added to improve flowability, and lubricants may be used to prevent powder/machine interactions. Colloidal silica, cornstarch, and talc are the common glidants used and appear to work by coating the particles in the powder formulation (Augsburger and Shangraw, 1966; York, 1975). Commonly used lubricants are magnesium stearate and stearic acid, which reduce friction and adhesiveness between machine surfaces and formulation particles. A summary of formulation additives and their functions is given in Table 15.2.

Filled HGC are required to meet certain standards set forth by the USP XXIII. One of these standards is filled capsule uniformity, which can be demonstrated by either content uniformity or weight variation. The test for content uniformity may be required for all cases; however, the test for weight variation may be required only for products containing 50 mg or more of drug, which comprises 50% or more of the total dosage form’s weight. Content uniformity is determined by individually assaying 10 capsules as directed in the drug’s individual USP monograph. Weight variation is determined by individually weighing 10 filled capsules, removing each capsule’s contents, and individu-

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ally weighing each empty capsule. The net weight of each capsule is calculated by subtracting the empty capsule weight from the filled capsule weight. The amount of active in each capsule is then determined assuming homogenous distribution of drug. The criteria for content uniformity and weight variation are that the amount of active in nine of the 10 assayed capsules must lie within the range of 85.0–115.0%, and no capsule must lie outside the range of 75.0–125.0% of the label claim. If these criteria are not met, 20 more capsules must be assayed. In addition to these standards, disintegration and dissolution tests of filled HGC are also required and are discussed below.

The ability of HGC to release their contents and facilitate drug delivery to the bloodstream is dependent upon gelatin’s ability to dissolve in the gastrointestinal environment. To assess the competency of dosage forms, the FDA requires that the bioavailability, or relative amount of drug that reaches the bloodstream, be determined for all drug products before marketing. However, the true determination of bioavailability requires time-consuming and expensive in vivo experiments. Because of this, in vitro experiments, such as dosage form disintegration and drug dissolution, have been developed to help predict the capsule’s in vivo performance. Several factors can influence a capsule’s performance, including the formulation contained within the capsule, the process by which the capsules were filled, and the gelatin shell itself. Usually, these influences do not greatly affect the capsule product’s bioavailability, but in a few cases there have been problems.

Bioavailability is defined as “a measurement of the rate and extent (amount)

382 of therapeutically active drug that reaches the general circulation” (Shargel and Yu, 1993) and is influenced by many processes, such as disintegration of the dosage form, drug dissolution, and drug absorption. The bioavailability of a capsule product is determined by measuring the plasma concentration of its active ingredient over a specific period of time after ingestion. The area under the curve (AUC) of the plasma concentration vs. time plot is often used to indicate the amount of drug that reaches the systemic circulation (Figure 15.6). Bioavailability may be discussed in terms of absolute or relative bioavailability. Absolute bioavailability is determined by comparing the AUC of the capsule product to the AUC of an IV administration of the same dose of active ingredient. Relative bioavailability is determined by comparing the AUC of a capsule product to the AUC of an acknowledged reference product of the same dose. The reference may be a different route of administration other than IV such as intramuscular, different dosage form such as an oral solution, or different manufacture such as a currently marketed capsule product.

As mentioned above, in vitro tests, such as dosage form disintegration and drug dissolution, have been developed to predict in vivo performance. The standards for these tests are given in the USP XXIII. The standard disintegration test apparatus (Figure 15.7) consists of a basket containing six glass tubes, each having a wire mesh bottom. One capsule is placed into each tube, and the basket is repeatedly immersed into a fluid, usually water maintained at 35–39 C, to simulate movement within the stomach. Because capsules have a tendency to float, a wire mesh is placed over the top opening of the tubes to prevent the capsules from escaping during immersion. The qualitative test is conducted over a

Figure 15.6 Typical plasma concentration vs. time plot after oral administration of a hard gelatin capsule product (AUC area under the curve).


383

specified period of time, and the capsules are then observed for complete disintegration. The USP XXIII defines complete capsule disintegration as the state in which the capsule residue remaining on the bottom mesh of the test apparatus is a soft mass having no palpably firm core. If one or two of the capsules fail, the test is then repeated on 12 additional capsules and 16 of the 18 total capsules must disintegrate in the allotted time for the capsules to pass. Dissolution tests measure the rate at which the drug is released from the capsule product into solution. The standard dissolution test apparatus (Figure 15.8) uses either a rotating wire mesh basket or a rotating paddle to simulate the movement within the stomach. The baskets or paddles are immersed into a glass vessel containing a specified amount of dissolution medium at 37.0 C. The entire apparatus usually contains six dissolution vessels with one capsule tested in each. If the basket method is used, the capsules are simply placed within the basket but if the paddle method is used, the capsules must first be placed within a weighted cage to prevent them from floating. After the capsules are placed into the dissolution medium and rotation begins, the dissolution medium is sampled at specified times and analyzed for drug content. The use of a third apparatus has recently increased in prevalence (Bottom et al., 1997). In an attempt to better simulate stomach movement, the disintegration and dissolution apparatuses have been combined into a single method. The capsules are dipped into the dissolution media at a specified rate (dips/minute), and the dissolution medium is sampled and analyzed as described above.

Figure 15.7 Standard USP disintegration apparatus. Dipping motion introduces mixing in an aqueous medium maintained at 37 C with a water bath.

384

Figure 15.8 Standard USP dissolution apparatus. Apparatus II uses paddles, while Apparatus I uses baskets. Either is rotated for mixing in an aqueous medium at 37 C maintained with a water bath.

The USP XXIII uses the quantity Q as the dissolution test criterion. The Q value is the acceptable amount of drug dissolved within a specified period of time expressed as a percentage of the labeled contents. Each drug’s individual USP monograph specifies the acceptable amount to be dissolved for that particular drug. An example Q value would be 75% of the labeled drug content dissolved within 45 minutes. The USP XXIII acceptance standards are as follows. The amount of drug released from each of the six capsules in the specified time period is not less than Q 5%. If this is not met, six more capsules are tested, and the average drug release from the 12 capsules must be equal to or greater than Q and no one capsule can be less than Q 15%. Finally, if this is not met, 12 more capsules are tested, and the average drug release from the 24 capsules must be equal to or greater than Q and no more than two capsules can be less than Q 15%.

The biggest influence on bioavailability of HGC is from the powder formulation ingredients, particularly the drugs, diluents, and lubricants (Jones, 1972). The extent to which these affect bioavailability is ultimately dependent upon the absorption site of the drug. For optimum bioavailability, a capsule product must release its ingredients, and the drug must dissolve within the gastrointestinal tract before it reaches its site of absorption. The dissolution rate of many


385

drugs is directly proportional to their solubility (Hamlin et al., 1965). Therefore, drugs having a low solubility often have a low bioavailability. A common way to increase the dissolution rate of a drug is to reduce its particle size, which increases the surface area available for dissolution (Fincher et al., 1965; Paul et al., 1967). However, in some cases, this may actually decrease the dissolution rate by limiting drug/water interactions and lowering drug wettability (Newton and Rowley, 1970; Newton and Bader, 1980). Diluents have been shown to both increase and decrease drug dissolution under different circumstances (Withey and Mainville, 1969; Newton et al., 1971; Bastami and Groves, 1978). Generally, soluble diluents increase the dissolution of poorly soluble drugs (Davies and Fell, 1973). Lubricants usually have a detrimental effect on drug dissolution because of their hydrophobicity, which increases the overall hydrophobic nature of the capsule contents and decreases their wettability (Samyn and Jung, 1970; Murthy and Samyn, 1977). However, there have also been instances where the opposite effect occurred (Stewart et al., 1979; Mehta and Augsburger, 1981). Materials, such as disintegrants (Samyn and Jung, 1970; Goodhart et al., 1973), may be added to capsule powder formulations for the sole purpose of improving bioavailability. Disintegrants, which are mainly used to manufacture tablets, are compounds that absorb water and subsequently expand or swell. It is believed that their swelling may help disintegrate the gelatin shell and deaggregate the powder formulation, thereby increasing the rate of drug dissolution. The effectiveness of disintegrants is questionable because capsules are filled with little compression and provide little structure for the disintegrants to swell against. However, the increasing popularity of the pisto-tamp filling method, which somewhat compresses the powder formulation, and the development of improved disintegrants such as croscarmellose sodium type A, sodium starch glycolate, and crospovidone (Shangraw et al., 1980) may make their inclusion into capsule formulations beneficial (Augsburger, 1996). In addition to formulation ingredients, capsule product bioavailability is also influenced by the filling technique. Capsule-filling techniques affect the structure of the capsule’s contents and in particular their degree of compaction. The most desirable filling technique is one that prevents a large degree of compaction and, therefore, allows for a more rapid deaggregation of the capsule contents (Newton, 1987). It is generally accepted that a more compact capsule fill results in longer capsule disintegration and drug dissolution times (Newton and Rowley, 1970; Newton, 1972). Because of this, the pisto-tamp filling method has the greatest impact on bioavailability since it produces a powder compact prior to capsule filling. However, this may vary depending upon other formulation factors (McGee et al., 1970; O’Callaghan et al., 1971), and, as mentioned, bioavailability problems resulting from compaction during capsule filling may be overcome by inclusion of a disintegrant in the powder formulation.

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In most instances, the gelatin shell has little influence on bioavailability of capsule products. However, diminished capsule shell dissolution and subsequent drug release problems have recently received attention although problems were reported as early as 1974 for a chloramphenicol HGC product (Khalil et al., 1974). In most cases, there was no effect on bioavailability of the drug compared to the readily dissolving product (Johnson et al., 1977; Chafetz et al., 1984; Mohamad et al., 1986; Dey et al., 1993). One exception occurred with a HGC product containing phenytoin, an anticonvulsant drug, which was identified upon hospital admission for loss of seizure control (Martin et al., 1985). These dissolution problems were associated with adverse storage conditions of elevated temperature and humidity, or prolonged storage. An FDA drug recall of a nifedipine SGC product in 1993 due to these problems led to the formation of the FDA/Industry Gelatin Capsule Working Group to examine the problem. It is now generally accepted that these dissolution problems have only a small risk of reduced bioavailability because of gastric and intestinal enzymes that contribute to the physiological dissolution process but which have not been routinely used in dissolution tests. One result of the FDA scrutiny was to initiate the regulatory process to allow so-called Tier Two testing. This involves a second USP dissolution test containing a proteolytic enzyme if the product failed its original monograph dissolution test because of this problem (Capsule Working Group, 1998). The dissolution problem, nevertheless, remains a possible safety issue, and its causes are a concern. There is a strong body of evidence that this problem is caused by gelatin cross-linking from aldehyde impurities initially present in the formulation, or formed under adverse storage conditions. Gelatin cross-inking leads to an intricate network of high molecular weight protein chains, which produces a swellable hydrogel and substantially reduces, or even prevents, gelatin dissolution (Albert and Bayer, 1991; Bubnis and Ofner, 1994; Mwangi and Ofner, 1995; Ofner and Bubnis, 1996; Ofner et al., 1999). Such a cross-linked and swollen film, or pellicle, can prevent the capsule from dissolving and releasing its contents (Ofner, 1996), thereby lowering capsule bioavailability (Digenis et al., 1994; Adesunloye and Stach, 1998). The aldehyde impurities initiating the cross-linking can occur from a surprising range of sources. For example, incorporation of the surfactant polysorbate 80 into a gemfibrozil capsule formulation produced formaldehyde, insoluble films, and impaired capsule dissolution after one month storage at 37 C and 80% RH (Chafetz et al., 1984). Formaldehyde production was attributed to autoxidation of the end hydroxyl groups in the polyoxyethylene chains of polysorbate 80. In another case, cellulose in rayon coilers (as a “cotton” packer in capsule containers) underwent decomposition to furfural. This alde-

References

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hyde was proposed to react with gelatin leading to a poorly soluble gelatin product after storage at 40 C and 75% RH for two months (Schwier et al., 1993). Other non-aldehyde agents have also induced a similar problem. For example, the capsule dyes FD&C Red No. 3 and FD&C Blue No. 1 produced substantial dissolution reductions after exposure to high humidity and UV or fluorescent light (Murthy et al., 1989a, b).

HGC have changed little over the past 150 years, but they continue to offer several advantages over other dosage forms. Gelatin, due to its unique properties, remains as the most suitable material for capsule production, and additives such as colorants, preservatives, and surfactants have improved capsule manufacturing. Today, a few companies produce a variety of different capsules, some with special properties and functions. The evolution of capsule-filling machinery has made the manufacturing of capsule products much more efficient by utilizing either hand-operated devices on a small scale or fast automated machines on a large scale. A major reason for the continued use of HGC has been the high drug bioavailability compared to tablets. However, many factors may influence bioavailability and several variables must be considered when working with capsule products. Nevertheless, HGC continue to be one of the most commonly and successfully used pharmaceutical dosage forms.

Adesunloye, T.A. and Stach, P.E. 1998. “Effect of Glycine/Citric Acid on the Dissolution Stability of Hard Gelatin Capsules,” Drug Dev. Ind. Pharm., 24:493–500. Albert, K. and Bayer, E. 1991. “Investigation of the Hardening Reaction of Gelatin with 13C Labeled Formaldehyde by Solution and Solid State 13C NMR Spectroscopy,” Z. Naturforsch, 46b:385–389. American Federal Standard No. 285A. 1976. Capsules (for Medicinal Purposes). Washington, DC: General Services Administration. Ansel, H.C. 1985. Introduction to Pharmaceutical Dosage Forms. Philadelphia, PA: Lea & Febiger, pp. 126–136. AOAC. 1999. Official Methods of Analysis, 16th Ed., 5th Rev., ed., P. Cunniff, Gaithersburg, MD: Association of Official Analytical Chemists, p. 38–41. Augsburger, L.L. 1974. “Powdered Dosage Forms” in Sprowl’s American Pharmacy, 7th Ed., ed., L.W. Dittert, Philadelphia, PA: J.B. Lippincott Co., pp. 301–343. Augsburger, L.L. 1996. “Hard and Soft Gelatin Capsules” in Modern Pharmaceutics, 3rd Ed., eds., G.S. Banker and C.T. Rhodes, New York: Marcel Dekker, Inc., pp. 395–440. Augsburger, L.L. and Shangraw, R.F. 1966. “Effect of Glidants in Tableting,” J. Pharm. Sci., 55:418–423. Bailie, G.R. and Kesson, C.M. 1981. “The Effect of Capsule Color on Hypnotic Efficacy,” Drug Intell. Clin. Pharm., 15:492–493. Bastami, S.M. and Groves, M.J. 1978. “Some Factors Influencing the in vitro Release of Phenytoin from Formulations,” Int. J. Pharm., 1:151–164.

388 Belthoff, R.C. 1986. “Capsule Sealing Update: Combining Efficient Production with Tamper-Resistant Procedures,” Packag. Technol., 16(4):6–7. Bottom, C.B., Clark, M. and Carstensen, J.T. 1997. “Dissolution Testing of Soft Shell Capsules—Acetaminophen and Nifedipine,” J. Pharm. Sci., 86:1057–1061. Bubnis, W.A. and Ofner III, C.M. 1994. “Semi-Quantitative and Quantitative Determinations of the Extent of Amino Group Crosslinking in Gelatin and Modified Gelatin,” Pharm. Res., 11:S–161. Buckalew, L.W.and Coffield, K.E. 1982. “An Investigation of Drug Expectancy as a Function of Capsule Color and Size and Preparation Form,” J. Clin. Psycopharma., 2:245–248. Capsule Working Group. 1998. “Collaborative Development of Two-Tier Dissolution Testing for Gelatin Capsules and Gelatin-Coated Tablets Using Enzyme-Containing Media,” Pharmacopeial Forum, 24:7045–7050. Chafetz, L., Hong, W., Tsilifonis, D.C., Taylor, A.K. and Philip, J. 1984. “Decrease in the Rate of Capsule Dissolution Due to Formaldehyde from Polysorbate 80 Autoxidation,” J. Pharm. Sci., 8:1186–1187. Cole, G.C. 1987a. “Capsule Types, Filling Tests, and Formulation” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 165–175. Cole, G.C. 1987b. “The History of Capsule-filling Machinery” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 87–91. Cole, G.C. 1987c. “Capsule-filling Machinery” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 104–163. Cole, G.C. 1987d. “The Mechanical Operations of Filling Hard Capsules” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 92–103. Davies, J.E. and Fell, J.T. 1973. “The Influence of Starch and Lactose on the Release Rates of Drugs from Hard Gelatin Capsules,” J. Pharm. Pharmacol., 25:431–432. Delaney, R. 1982. “Surveying Consumer Preferences,” Pharm. Exec., 2(3):34–38. Dey, M., Enever, R., Kraml, M., Pruek D.G., Smith, D. and Weierstall, R. 1993. “The Dissolution and Bioavailability of Etodolac from Capsules Exposed to Conditions of High Relative Humidity and Temperatures,” Pharm. Res., 10:1295–1300. Digenis, G.A., Gold, T.B. and Shah, V.P. 1994. “Cross-Linking of Gelatin Capsules and Its Relevance to Their In-Vitro-In-Vivo Performance,” J. Pharm. Sci., 83:915–921. Fincher, J.H., Adams, J.G. and Beal, H.M. 1965. “Effect of Particle Size on Gastrointestinal Absorption of Sulfisoxazole in Dogs,” J. Pharm. Sci., 54:704–708. Goodhart, F.W., McCoy, R.H. and Ninger, F.C. 1973. “New In-Vitro Disintegration and Dissolution Test Method for Tablets and Capsules,” J. Pharm. Sci., 62:304–310. Grosswald, R.R., Anderson, J.B. and Andrew, S. May 12, 1998. U.S. patent 5,750,157. Hamlin, W.E., Northam, J.I. and Wagner, J.G. 1965. “Relationship Between In-Vitro Dissolution Rates and Solubilities of Numerous Compounds Representative of Various Chemical Species,” J. Pharm. Sci., 54:1651–1653. Hostetler, V. 1986. “Capsules, Part One: Hard Capsules” in The Theory and Practice of Industrial Pharmacy, eds., L. Lachman, H.A. Lieberman and J.L. Kanig, Philadelphia, PA: Lea & Febiger, pp. 374–398. Ito, K., Hitomi, M., Kaga, S.I. and Takeya, Y. 1969. “Studies on Hard Gelatin Capsules. II. The Capsule Filling of Powders and Effects of Glidant by Ring Filling Method-Machine,” Chem. Pharm. Bull., 17:1138–1145. Johnson, B.F., Maculey, P.V., Smith, P.M.and French, J.A.G. 1977. “The Effects of Storage Upon In-Vitro and In-Vivo Characteristics of Soft Gelatin Capsules Containing Digoxin,” J. Pharm. Pharmacol., 29:576–578. Jolliffe, I.G. and Newton, J.M. 1978. “Powder Retention within a Capsule Dosator Nozzle,” J. Pharm. Pharmacol., 30(Supp.):41P. Jolliffe, I.G., Newton, J.M. and Walters, J.K. 1979. “A Theoretical Approach to Optimizing Capsule Filling by a Dosator Nozzle,” J. Pharm. Pharmacol., 31(Supp.):70P.

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Jones, B.E. 1972. “Disintegration of Hard Gelatin Capsules,” Acta. Pharm. Suec., 9:261–263. Jones, B.E. 1982. “The Manufacture of Hard Gelatin Capsules,” Chem. Engr. (London), 380:174–177. Jones, B.E. 1985. “Hard Gelatin Capsules and the Pharmaceutical Formulator,” Pharm. Tech., 9(9):106–112. Jones, B.E. 1987a. “The History of the Gelatin Capsule” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 1–12. Jones, B.E. 1987b. “Gelatin Additives, Substitutes and Extenders” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 49–60. Jones, B.E. 1987c. “Gelatin: Physical and Chemical Properties” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 31–48. Jones, B.E. 1987d. “The Manufacture of Hard Gelatin Capsules,” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 68–79. Jones, B.E. 1987e. “Capsule Standards” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 61–67. Jones, B.E. 1988. “Powder Formulations for Capsule Filling,” Mfg. Chem., 59(7):28–30, 33. Jones, B.E. 1990. “Capsules, Hard” in Encyclopedia of Pharmaceutical Technology, Vol. II, eds., J. Swarbrick and J.C. Boylan, New York: Marcel Dekker, Inc., pp. 251-268. Jones, B.E. and Turner, T.D. 1974. “A Century of Commercial Hard Gelatin Capsules,” Pharm. J., 213:614–617. Khalil, S.A.H., Ali, L.M.M. and Abdel Khalek, M.M. 1974. “Effects of Ageing and Relative Humidity on Drug Release,” Pharmazie, 29(1):36–37. Kurihara, K. and Ichikawa, I. 1978. “Effect of Powder Flowability on Capsule Filling Weight Variation,” Chem. Pharm. Bull., 26:1250–1256. Lucchelli, P.E., Cattaneo, A.D. and Zattoni, J. 1978. “Effect of Capsule Colour and Order of Administration on Hypnotic Treatments,” Europ. J. Clin. Pharmacol., 13:153–155. Martin, E.D., Fraser, R.J.L. and Camens, I. 1985. “Storage of Phenytoin Capsules,” Med. J. Austral., 143:634–635. Martyn, G.W. 1974-75. “The People Computer Interface in a Capsule Molding Operation,” Drug Devel. Comm., 1(1):39–49. McGee, B.J., Kennedy, D.R. and Walker, G.C. 1970. “Some Factors Affecting Release and Availability of Drugs from Hard Gelatin Capsules,” J. Pharm. Sci., 59:1430–1433. Mehta, A.M. and Augsburger, L.L. 1981. “A Preliminary Study of the Effect of Slug Hardness on Drug Dissolution from Hard Gelatin Capsules Filled on an Automatic Capsule-Filling Machine,” Int. J. Pharm., 7:327–334. Mohamad, H., Renoux, R., Aiache, S., Aiache, J.-M., Sirot, J. and Kantelip, J.-P. 1986. “Investigation of Drug Biopharmaceutical Stability. Part 3. Study of Ampicillin Trihydrate Capsules,” S.T.P. Pharma. Pratiques, 2(Oct–Nov):912–917. Murthy, K.S. and Samyn, J.C. 1977. “Effect of Shear Mixing on In-Vitro Drug Release of Capsule Formulations Containing Lubricants,” J. Pharm. Sci., 66:1215–1219. Murthy, K.S., Enders, N.A. and Fawzi, M.B. 1989a. “Dissolution Stability of Hard-Shell Capsule Products, Part I: The Effect of Exaggerated Storage Conditions,” Pharm. Tech., 13(3):72–86. Murthy, K.S., Reisch Jr, R.G. and Fawzi, M.B. 1989b. “Dissolution Stability of Hard-Shell Capsule Products, Part II: The Effect of Dissolution Test Conditions on In Vitro Drug Release,” Pharm. Tech., 13(6):53–58. Mwangi, J.W. and Ofner III, C.M. 1995. “Modulation and Evaluation of the Degree of Crosslinking in Gelatin Hydrogel Matrices Using a Water Soluble Carbodiimide,” Pharm. Res., 12:S212. Newton, J.M. 1972. “The Release of Drugs from Hard Gelatin Capsules,” Pharm. Weekbl. Ned., 107:485–498. Newton, J.M. 1987. “Drug Release from Capsules” in Hard Capsules, Development and Technology, ed., K. Ridgeway, London: The Pharmaceutical Press, pp. 195–204.

390 Newton, J.M. and Bader, F. 1980. “The Influence of Drug and Diluent Particle Size on the In-Vitro Release of Drug from Hard Gelatin Capsules,” J. Pharm. Pharmacol., 32:167–171. Newton, J.M. and Rowley, G. 1970. “On the Release of Drug from Hard Gelatin Capsules,” J. Pharm. Pharmac., 22(Suppl.):163S–168S. Newton, J.M., Rowley, G. and Tornblom, J.F.V. 1971. “The Effect of Additives on the Release of Drug from Hard Gelatin Capsules,” J. Pharm. Pharmac., 23:452–453. Norris, W.G. 1959. “Hard Gelatin Capsules-How Eli Lilly Make 500 Million a Year,” Mfg. Chem., 30:233–236. Norris, W.G. 1961. “Output Replaces Imports and Provides Exports,” Mfg. Chem., 32:249–252, 258. O’Callaghan, C.H., Toothill, J.P.R. and Robinson, W.D. 1971. “A New Approach to the Study of Serum Concentrations of Orally Administered Cephalexin,” J. Pharm. Pharmac., 23:50–57. O’Connor, R.E., Rippie, E.G. and Schwartz, J.B. 1990. “Powders” in Remingtons, 18th Ed., ed., A.R. Gennaro, Easton, PA: Mack Publishing Company, pp. 1615–1632. Ofner III, C.M. 1996. “Evaluation of Gelatin Capsule Crosslinking Using Gelatin Dissolution and Chemical Assay,” Final Report to the Food and Drug Administration, Subcontract of Contract #223-95-3006; Philadelphia College of Pharmacy and Science, Philadelphia, PA. Ofner III, C.M. and Bubnis, W.A. 1996. “Chemical and Swelling Evaluations of Amino Group Crosslinking in Gelatin and Modified Gelatin Matrices,” Pharm. Res., 13:1821–1827. Ofner III, C.M., Zhang, Y.E., Jobeck, V. and Bowman, B.J. 1999. “Evaluation of Dissolution and Crosslinking in Gelatin Capsules Treated with Formaldehyde and Capsules Exposed to Elevated Temperature and Humidity,” submitted. Paul, H.E., Hayes, K.J., Paul, M.F. and Borgmann, A.R. 1967. “Laboratory Studies with Nitrofurantoin, Relationship between Crystal Size, Urinary Excretion in the Rat and Man, and Emesis in Dogs,” J. Pharm. Sci., 56:882–885. Porter, S.C. 1990. “Coating of Pharmaceutical Dosage Forms” in Remingtons, 18th Ed., ed., A.R. Gennaro, Easton, PA: Mack Publishing Company, pp. 1666–1675. Reier, G., Cohn, R., Rock, S. and Wagenblast, F. 1968. “Evaluation of Factors Affecting the Encapsulation of Powders in Hard Gelatin Capsules. I. Semi-automatic Machines,” J. Pharm. Sci., 57:660–666. Ridgway, K. and Callow, J.A.B. 1973. “Capsule-filling Machinery,” Pharm. J., 211:281–285. Rudnic, E. and Schwartz, J.B. 1990. “Oral Solid Dosage Forms” in Remingtons, 18th Ed., ed., A.R. Gennaro, Easton, PA: Mack Publishing Company, pp. 1633–1665. Samyn, J.C. and Jung, W.Y. 1970. “In-Vitro Dissolution from Several Experimental Capsule Formulations,” J. Pharm. Sci., 59:169–175. Schwier, J.R., Cooke, G.G., Hartauer, K.J. and Yu, L. 1993. “Rayon: Source of Furfural-Reactive Aldehyde Capable of Insolubilizing Gelatin Capsules,” Pharm. Tech., 17(5):78, 80. Shah, K., Augsburger, L.L., Small, L.E. and Polli, G.P. 1983. “Instrumentation of a Dosing Disc Automatic Capsule Filling Machine,” Pharm. Tech., 7(4):42–54. Shangraw, R.F., Mitrevej, A. and Shah, M. 1980. “A New Era of Tablet Disintegrants,” Pharm. Tech., 4(10):49–57. Shargel, L. and Yu, A.B.C. 1993. “Bioavailability and Bioequivalence” in Applied Biopharmaceutics and Pharmacokinetics, 3rd Ed., Norwalk, CT: Appleton & Lange, pp. 193–223. Sheppard, S.E. and Houck, R.C. 1930. “The Structure of Gelatin Sols and Gels. I. The Viscosity of Gelatin Solutions,” J. Phys. Chem., 34:273–298. Small, L.E. and Augsburger, L.L. 1977. “Instrumentation of an Automatic Capsule-Filling Machine,” J. Pharm. Sci., 66:504–509. Stewart, A.G., Grant, D.J.W. and Newton, J.M. 1979. “The Release of a Model Low-Dose Drug (Riboflavine) from Hard Gelatin Capsule Formulations,” J. Pharm. Pharmacol., 31:1–6. United States Pharmacopeial Convention, Inc. 1995. The United States Pharmacopeia 23/The National Formulary 18. Taunton, MA: Rand McNally.

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Soft Gelatin Capsules

OFT gelatin capsules are one-piece, hermetically sealed soft gelatin shells containing a liquid, a suspension, or a semi-solid (Figure 16.1) (Hom and Jimerson, 1990; Wilkinson and Hom, 1990). In contrast to the rigid two-piece hard gelatin capsule shells, soft gelatin capsule shells contain large amounts of plasticizers, which make them flexible. Similar to hard gelatin capsules, they are solid dosage forms intended mainly for oral administration, although they may also be used as rectal or vaginal suppositories. In the late 1980s, a trade association comprised of soft gelatin manufacturers in the U.S. introduced the name “softgels” to further distinguish this dosage form from hard gelatin capsules. Softgels are formed, filled, and sealed in a continuous operation, which has been most cost-effective for a few contract manufacturers (Hom and Jimerson, 1990). The list of companies operating softgel manufacturing facilities in North America in 2000 includes Banner Pharmacaps (High Point, NC), R.P. Scherer (Basking Ridge, NJ), Accucaps Limited (Windsor, Ontario, Canada), Soft Gel Technologies (Los Angeles, CA), Pharmavite (Mission Hills, CA), Nutricia Manufacturing USA (Greenville, SC), Goldcaps (Miami, FL), CapsuleWorks (Bayport, NY), Tishcon Corporation (Westbury, NY), IVC Industries (Freehold, NJ), Swiss Caps (Miami, FL), Gelcell Capsules Limited (Tecumseh, Ontario, Canada), Captek Softgel International (Cerritos, CA), and National Vitamin Company (Porterville, CA). At present, the number of in-

S

393

394

Figure 16.1 Softgels manufactured using the rotary die encapsulation process.

stalled softgel encapsulation lines/machines in North America is estimated at 250. This chapter discusses the advantages, limitations, uses, and manufacturing of softgels. Gelatin-enrobed and gelatin-coated tablets/caplets also are briefly discussed.

The French pharmacists Mothes and DuBlanc are credited with developing the softgel dosage form in the 1830s (Hom and Jimerson, 1990). They patented a method of preparing capsules by dipping a mercury-filled leather sac into molten gelatin. The gelatin coating was allowed to solidify, the sac was removed, and medications were added to the capsule with a pipette (Hom and Jimerson, 1990; Wilkinson and Hom, 1990). The capsule was then hand-sealed with molten gelatin. Although iron molds were later introduced, this tedious softgel preparation method had high fill variations and yield losses, and was not commercially viable.

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Later, a plate method was developed that made the commercialization of softgels viable. This batch process used two sets of metallic plates with matching cavities. A gelatin sheet was cast on the surface of the lower die plate, vacuum was applied to pull the sheet into the die pockets, medication was filled into the formed pockets, a second gelatin sheet was laid on top, and the two plates were pressed together to form and separate the capsules (Hom and Jimerson, 1990). The plate method was used for many years by The Upjohn Company (Kalamazoo, MI) until it was discontinued in 1989 (Wilkinson and Hom, 1990). In the early 1930s, Robert P. Scherer invented the continuous rotary die encapsulation process for large-scale manufacturing of softgels (Scherer, 1934). Over the years, this process has undergone various modifications and improvements in automation and has become the industry standard worldwide (Ebert, 1977). The concept of the Scherer process was the basis for two additional processes suitable for filling softgels with powders and pelleted formulations. One, the Accogel process, was developed by Lederle Laboratories in 1948, and the other, the reciprocating die process, was developed by the Norton Company in 1949 (Hom and Jimerson, 1990; Wilkinson and Hom, 1990). Today, the vast majority of encapsulating machines operating around the world are custom-manufactured based on the Scherer concept and are self-maintained by the softgel manufacturers. However, “turn-key” softgel manufacturing systems have become available in recent years, thus lowering the technological barrier for entry into the softgel business (at least for dietary supplements, such as oils and vitamin E, which require minimal fill formulation expertise).

The following are generally recognized as functional and commercial advantages of the softgel as a dosage form for administering pharmaceutical and dietary formulations: (1) Softgels generally exhibit enhanced dissolution rates of encapsulated biologically active compounds because they absorb water, open at the seams, disintegrate, and rapidly release their contents (Hom and Miskel, 1970, 1971). The elevated body temperature accelerates the rapid in vivo release of the softgel contents because some degree of gel melting occurs. (2) Biologically active compounds with poor water solubility can be solubilized or dispersed in oils or aqueous-miscible liquids within the softgels. Upon ingestion, the capsule shell disintegrates, and the fill formulation dissolves or emulsifies, yielding dispersions of high surface area and good bioavailability (Berry, 1983; Seager, 1985; Karunakar, 1998). The enhanced bioavailability of several pharmaceutical compounds administered within softgels compared to hard gelatin capsules and/or tablets has

396 been demonstrated (Mallis et al., 1975; Angelucci et al., 1976; Ghirardi et al., 1977; Lindenbaum, 1977; Lucchelli et al., 1978; Stella et al., 1978; Alvisi et al., 1979; Astorri et al., 1979; Nitsche and Mascher, 1982; Helqvist et al., 1991; Gumkowski et al., 1994). However, other studies reported no significant differences in the bioavailability of various pharmaceutical compounds administered within softgels versus hard gelatin capsules and/or tablets (Albert et al., 1974; Fuccella et al., 1977; Steinbach et al., 1980a, b; Pierce et al., 1984). (3) The improved bioavailability of compounds delivered within softgels allows for administering lower dosages, thus resulting in reduced raw material costs (Seager, 1985). (4) Compounds sensitive to oxidation can be protected through solubilization or dispersion in oils or aqueous-miscible liquids within the softgels (Seager, 1985). In addition, the gelatin shell is a potent oxygen barrier (Hom et al., 1975; Anonymous, 1992), as is generally the case with protein-based films (at least at low relative humidity conditions) (Gennadios et al., 1993). (5) The softgel manufacturing process often allows for higher dosage accuracy and content uniformity than other oral dosage forms (Berry, 1982). (6) Highly potent (e.g., cytotoxic) compounds present health and safety concerns with the resulting airborne particles during tableting. Such concerns can be alleviated by introducing the compounds into liquid formulations and encapsulating them into softgels. (7) Although not tamper-proof, softgels are both tamper-evident and tamper-resistant. Puncturing the softgel shell, introducing a contaminant, and resealing the shell without resultant leakage or signs of alteration is a highly difficult task (Berry, 1982; Hom and Jimerson, 1990). (8) Unpleasant tastes and odors of active compounds are masked by the capsule shell (Ebert, 1977; O’Brien, 2000). (9) There is a high degree of flexibility in selecting softgel sizes, shapes, and colors, which, combined with capsule printing capabilities, offers wide opportunities for product identification and differentiation (Stanley, 1986; Schofield, 1999). (10) As an oral dosage form, softgels typically rate high in consumer preference because of their elegance, ease of swallowing, and strong perceived effectiveness due to their liquid fill formulations (Berry, 1983; Schofield, 1999).

The following are often identified as limitations of the softgel dosage form and technology:

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(1) The softgel manufacturing process is slower than tableting. (2) Softgels require intensive inspection due to several potential defects, such as capsules that leak, have shape imperfections, or are stuck together. (3) The lengthy drying process substantially extends the manufacturing cycle of softgels. (4) Operation of softgel encapsulation machines requires experienced personnel. (5) The encapsulation process is not fully automated in terms of monitoring in-process parameters, such as capsule seal strength and wet shell thickness or weight. (6) Although the softgel encapsulation process allows for accurate dosing and thus economical use of the fill material, it results in a notable waste of shell formulations (about 30%). (7) Due to increased labor requirements, softgels are generally produced at a higher cost than directly compressed tablets. (8) Prior to drying, softgel shells have a high moisture content, which allows for increased interactions among shell and fill ingredients.

Both over-the-counter (OTC) and ethical (prescription; Rx) drugs are encapsulated and marketed in softgels. It is noted that few facilities worldwide have the necessary technical expertise and regulatory approvals for manufacturing softgels containing drugs, particularly ethical drugs. The categories of OTC drugs typically available in softgels include analgesics (e.g., acetaminophen); anti-inflammatory agents (e.g., ibuprofen); antihistamines (e.g., chlorpheniramine maleate, brompheniramine maleate, doxylamine succinate, and diphenhydramine hydrochloride); stool softeners (e.g., docusate salts); decongestants (e.g., pseudoephedrine hydrochloride); antitussive agents (e.g., dextromethorphan hydrochloride); expectorants (e.g., guaifenesin); and antiflatulents (e.g., simethicone). Combinations of two or more active compounds are quite common, particularly in formulating cough and cold medications. The ethical drugs that have been or are currently formulated within softgels cover a wide range of therapeutic indications and include nifedipine (antianginal), valproic acid (anticonvulsant), benzonatate (antitussive), isotretinoin (treatment of severe recalcitrant nodular acne), amantadine hydrochloride (antiviral and antiparkinsonian), calcitriol (hypocalcemia management), ergocalciferol (treatment of refractory rickets and hypoparathyroidism),

398 cephalexin (antibacterial), amoxycillin (antibacterial), etoposide (antineoplastic), cyclosporine (immunosuppressant), ritonavir (HIV protease inhibitor), ethosuximide (anticonvulsant), chloral hydrate (sedative), dronabinol (cannabinoid; complex effects on central nervous system), ethchlorvynol (hypnotic), and ranitidine hydrochloride (ulcer treatment).

A wide array of traditional dietary supplements and compounds associated with supplement-style structure/function claims—regulated in the U.S. by the Food and Drug Administration (FDA) under the Dietary Supplement Health and Education Act of 1994 (DSHEA)—are currently available in softgels including the following: (1) Vitamins (mainly oil-soluble such as vitamins A, D, and E), minerals (e.g., calcium as calcium carbonate and chromium as chromium picolinate), and multi-vitamin and multi-mineral combinations (2) Antioxidants (e.g., grape skin extract, alpha-lipoic acid, rosemary extract, astaxanthin, and coenzyme Q10) (3) Phospholipids (e.g., lecithins) (4) Carotenoids (e.g., lycopene and lutein) (5) Oils that are rich in essential fatty acids (e.g., flaxseed oil, borage oil, evening primrose oil, and black currant seed oil) or omega-3 fatty acids (e.g., marine oils) (6) Herbal supplements (e.g., saw palmetto, aloe vera, panax ginseng, Siberian ginseng, St. John’s wort, valerian, kava, maca, echinacea, cat’s claw, dong quai, elderberry, ginkgo biloba, goldenseal, black cohosh, horsechestnut, olive leaf, and milk thistle) (7) Enzymes (e.g., lactase) (8) Amino acids and protein hydrolyzates In addition to dietary supplements and herbals, which also are widely referred to as nutraceuticals, traditional food items and food processing ingredients (e.g., cooking oils, peanut butter, tallow, butter, sauces, and chocolate syrup) also have been encapsulated into softgels to form single-use, single-dosage packages (Yamada and Makino, 1986; Anonymous, 1992). However, such a use of softgels has remained a niche application with limited commercialization.

Bath oils are the most common personal care products marketed in softgels.

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The functional properties of gelatin are well suited for such products (bath beads) because the plasticized gelatin shells quickly swell and then dissolve in contact with hot water, thus releasing the aromatic oils. In addition, softgels also are used as single-dose packages for higher value cosmetic formulations intended for topical use. Typically, such softgels have a “twist-off” or “break-off” feature at one end for dispensing the fill material (Spellman et al., 1991; Rinaldi et al., 1999). For example, Melnik et al. (1992) described the encapsulation of cosmetic compositions (e.g., sun screens, tanning agents, skin care, and anti-dandruff agents) using silicone polymers as carriers. Punto et al. (1996) disclosed a skin-treating formulation incorporated into softgels in the form of an emulsion comprised of a water-soluble active ingredient (e.g., ascorbic acid), polyethylene glycol, and an oil (e.g, silicone, paraffin, or vegetable oil). Lambrechts (1996) described a shampoo/conditioner formulation in a softgel that included a concentrated surfactant, a cationic conditioner, and a carrier (e.g., polyethylene glycol). Morton et al. (1997) discussed fragrance-containing softgels intended primarily for dispensing as perfume testers or samples. Lambrechts (1997) described skin conditioning compositions that were comprised of hydroxy and/or keto acids, a thixotropic agent, and an emulsifying agent (e.g., glyceryl monoesters of long-chain fatty acids) and were suitable for encapsulation into softgels. Skin lotion compositions containing vitamin E and/or vitamin A palmitate that were encapsulated into softgels were described by Fishman (1998). Softgel fill formulations intended for skin care that contained retinol-impregnated microparticles and, optionally, ascorbic acid-impregnated microparticles were described by Rinaldi et al. (1999).

Over the past 25 years, paintballs have emerged as an important application for softgels. The paintballs are softgels containing dyes in an oil (Haman and Schmoke, 1987), polyoxyethylene sorbitol monolaureate (Skogg, 1987), or polyethylene glycol (Rouffer, 1995) vehicle. They are fired from compressed air guns, including rapid firing devices, during adult war games or training and target shooting. Upon impact, the paintballs readily crush, thus “splattering” the contained dyes and marking the hit target. The paintball sport or recreational activity started in the U.S. in the 1970s and has been growing in popularity, both in the U.S. and overseas, ever since. In addition to recreational products, other niche industrial applications of softgels have been commercialized over the years. Examples include tube-shaped softgels filled with glue or technical grade grease, and round-shaped softgels filled with starter fluid for trucks.

Hard gelatin capsules are mainly produced in traditional oblong shapes and

400 in eight different sizes. In contrast, softgels for oral administration are manufactured in oval, oblong, and round shapes (Figure 16.2) and are able to accommodate a wide range of fill volumes. The nominal fill volume in minims is traditionally used to indicate the size of a softgel. A minim is 1/60 of a fluid dram (1 fluid dram 1/8 fluid ounce). Thus, a 1 cm3 volume corresponds to approximately 16.2 minims. Overall, softgels can range in size from 1 to 480 minims. For oral consumption in particular, oval-, oblong-, and round-shaped softgels typically range in size from 2 to 16, 3 to 24, and 2 to 9 minims, respectively. Sample calculations used for determining the minimum fill volume of a softgel product based on the desired active dose and the necessary excipients were presented by Stanley (1986). Fill formulations should result in a softgel of the smallest possible size so that raw material usage, manufacturing output, and patient compliance are optimized. To some extent, the capsule shell can shrink to the volume of its contents without negatively affecting product appearance. This offers sufficient leeway for filling a capsule with a lesser than the nominal volume. According to Stanley (1970), this leeway for smaller volume filling is 10, 20, or 30% of the nominal capacity for oblong, oval, or round capsules, respectively. In contrast, overfilling is not recommended because it can affect product appearance and stress the capsule seams leading to leakage and possibly rupture. Also, overfilled products can cause problems in post-processing operations such as blister packaging. In addition to the traditional oval, oblong, and round shapes used for human consumption, softgels also are manufactured in a wide variety of shapes for personal care products. For example, bath oil softgels often are marketed in the shapes of animals, seashells, stars, hearts, teardrops, and triangles. Bath oil softgels with a partially or fully textured outer surface also are manufactured. This surface texture can be applied on the cast, moldable gelatin ribbons

Figure 16.2 Examples of differently shaped softgels. From left to right: oval, oblong, round, and a tube with the “twist-off” feature.

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through contact with a roller having a textured surface (Ratko et al., 1993). Another method for enhancing softgel appearance and differentiation was described by Schurig et al. (1997). They produced color-striped or marblelized softgels by using patterned gelatin ribbons. Stone (1998) described the manufacture of softgels having a filled and a non-filled portion with one or both portions carrying impressed graphical representations (Stone, 1998). Single-use softgels containing cosmetic formulations for topical application are marketed in the form of tubes (regular, oval, or round) with a “break-off” or “twist-off” feature (Figure 16.2). Finally, softgel suppositories are typically manufactured in bullet-like shapes.

Details on softgel fill formulations are beyond the scope of this chapter, and only a few general considerations are discussed here. A comprehensive discussion on the nature of softgel contents was presented by Stanley (1986). Furthermore, substantial information on softgel fill formulation approaches, often targeted to a specific active compound, is available in the patent literature (Grainger, 1980; Stoopak et al., 1982; Shah et al., 1984; Henmi et al., 1987; Brox, 1988a, b; Yu et al., 1991; Torosian, 1992; Makino et al., 1993; Shelley et al., 1996; Tanner and Shelley, 1996; Vàsquez, 1997; Cimiluca, 1997; Woo, 1997; Becourt et al., 1998; Cody et al., 1999; Devlin and Hoy, 1999; Goldman, 2000; Hong et al., 2000; Hoy, 2000; Lacy et al., 2000; Rouffer, 2000). Fill formulations intended specifically for chewable softgels also have been discussed in the patent literature (Steele and Montes, 1999; Lech, 2000).

With the rotary die encapsulation process, the capsule contents are typically a liquid or a combination of miscible liquids; a solution of a solid(s) dissolved in a liquid(s); or a suspension of a solid(s) in a liquid(s) (Stanley, 1986). Rotary die apparatuses for encapsulating solids into softgels have been developed (Rowe, 1998) but have found limited application thus far. A large number of liquids that are either actives themselves or function as solubilization excipients for solid actives can be encapsulated. Such liquids that can be encapsulated without any limitations include water-immiscible liquids (e.g., vegetable oils, aromatic oils, aromatic and aliphatic hydrocarbons, chlorinated hydrocarbons, ethers, esters, alcohols, and organic acids) and water-miscible, non-volatile liquids (mainly limited to polyethylene glycols and non-ionic surfactants such as polysorbate 80) (Ebert, 1977; Stanley, 1986). A few other water-miscible and relatively non-volatile liquids, such as glycerin and propylene glycol, can be included in fill formulations but only in small

402 amounts (not more than 5–10% of the total liquid in the fill) (Stanley, 1986). Typically, water itself cannot be present in the fill at more than 8% (Sundararajan et al., 1996). However, a method to encapsulate fills with up to a 20% water content was described by Miskel et al. (1974). They prepared fill formulations by incorporating active compounds into aqueous solutions of gelling proteins (e.g., collagen, gelatin, soy protein, egg albumin, and casein). The gelling proteins formed fluid macromolecular gel matrices, and, upon drying of the softgels, these matrices set into rigid gels that retained as much as 20% water (Miskel et al., 1974). Addition of colloidal silica into fill formulations (0.5 to 10% by weight) to immobilize water also was suggested for encapsulating fills having a high water content (Altmann, 1995). Solid compounds that are poorly soluble in the abovementioned liquids can be encapsulated by being formulated into stable, homogeneous suspensions. To achieve good content uniformity and stability, the particle size of suspended solids typically should not exceed 80 mesh (180 m) (Hom and Jimerson, 1990). The suspending medium (referred to as the base, carrier, or vehicle) is typically a vegetable oil (e.g., soybean oil), a combination of a vegetable oil and a surfactant, a non-ionic surfactant (e.g., polysorbate 80), or polyethylene glycol (PEG, 400 or 600 molecular weight) (Stanley, 1986). PEG having a lower molecular weight (e.g., 200) is avoided because it can easily migrate into the gelatin shell over time causing overplasticization (softening). To facilitate the complete wetting of solids by oil bases, a wetting agent (often lecithin) is added at 2–3% by weight of the oil (Stanley, 1986). Suspensions also require a suspending agent to ensure homogeneity (content uniformity) and good flow characteristics (Ebert, 1977). Typical suspending agents for oil suspensions are waxes (e.g., beeswax and paraffin wax), stearates, and cellulose ethers (Ebert, 1977; Stanley, 1986). For non-oil suspending mediums, PEGs of high molecular weight (e.g., 4000 and 6000), glycol esters, and acetylated monoglycerides are generally used as suspending agents (Ebert, 1977; Stanley, 1986).

There are several limitations in the types of compounds that are suitable for encapsulation into softgels. Aldehydes (e.g., formaldehyde, acetaldehyde, and glutaraldehyde) can cross-link gelatin, thus slowing capsule disintegration and dissolution (Digenis et al., 1994; Hakata et al., 1994; Bottom et al., 1997). In general, the cross-linking of proteins by aldehydes is well documented (Feeney et al., 1975). Formaldehyde-induced cross-linking of gelatin mainly involves the lysine and arginine amino acids (Taylor et al., 1978; Albert et al., 1986, 1991; Gold et al., 1996). Aldehydes may be directly present as impurities in fill or shell ingredients, or they may be generated by autoxidation of lipid excipients, such as polysorbate 80 (Chafetz et al., 1984; Doelker and

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Vial-Bernasconi, 1988; Singh et al., 2000). PEG can be particularly problematic because it tends to react with atmospheric oxygen to form aldehydes (Tanner and Shelley, 1996). To alleviate this problem, PEG is typically handled in an inert atmosphere, for example, under a nitrogen blanket (Tanner and Shelley, 1996). Even capsule packaging materials can function as a source of cross-linking aldehydes as was shown for furfural from the rayon fiber inserted into high-density polyethylene bottles (Schwier et al., 1993). Recently, the use of near-infrared spectrophotometry as a non-invasive and non-destructive method for assessing aldehyde-induced cross-linking in softgels was proposed (Gold et al., 1998). Reducing carbohydrates (e.g., glucose, fructose, lactose, maltodextrin, and corn syrup solids), which often are used as drug or dietary supplement excipients, also cross-link proteins through the Maillard reaction (non-enzymatic browning) (Cheftel et al., 1985; Ames, 1998) and can affect the gelatin shell (Tanner and Shelley, 1996). However, satisfactory in vitro dissolution of cross-linked softgels or hard gelatin capsules may be obtained by adding proteolytic enzymes to the dissolution medium (Hom et al., 1973; Doelker and Vial-Bernasconi, 1988; Murthy et al., 1989b; Digenis et al., 1994; Gelatin Capsule Working Group, 1998). Succinylated gelatin, which is not susceptible to aldehyde-induced cross-linking, can be used in capsule manufacturing (Kobayashi et al., 1986; Sato et al., 1986; Yamamoto et al., 1995). Acylation of proteins with succinic anhydride reduces the e-amino groups, which are the prime reactive sites for aldehydes (Cheftel et al., 1985). However, succinylated gelatin is not approved for use with ingestible softgels in the U.S. Nonetheless, personal care formulations containing high amounts of aldehydes have occasionally been encapsulated into softgels manufactured with succinylated gelatin. Organic compounds of low molecular weight that are volatile (e.g., alcohols, ketones, acids, amines, and esters) tend to readily migrate through the capsule shell (Hom and Jimerson, 1990). Emulsions (oil-in-water or water-in-oil), although occasionally investigated as softgel fills (Bauer and Dortunc, 1984), are typically unsuitable for encapsulation because they eventually become destabilized, thus releasing water that migrates into the gelatin shell (Ebert, 1977). In general, strong acids or bases break non-covalent and covalent cross-links within the gelatin structure. Therefore, acidic liquids (pH < 2.5) encapsulated into softgels can hydrolyze gelatin and cause capsule leakage (Stanley, 1970). Highly alkaline liquids also can disrupt the shell structure, causing leakage. Salts of strong acids and bases (e.g., potassium, sodium, and choline chlorides) and ammonium salts (e.g., ammonium chloride) can also be destructive to the shell (Stanley, 1970; Ebert, 1977). Finally, compounds that are unstable in the presence of moisture, such as aspirin, are not suitable for encapsulation into softgels (Hom and Jimerson, 1990).

404

Gelatin, the product of partial hydrolysis of collagen, is the main component of the softgel shell. Its manufacture and characteristics are discussed in detail elsewhere in this book. It is estimated that about 7.6% of the total gelatin produced worldwide (19,000 out of 250,000 metric tons) in 1998 was used in softgel manufacturing (Pluvinet, 2000). In fact, the softgel business has been growing in recent years so that about 10% of the worldwide gelatin production is now allocated to softgels (Pluvinet, 2000). Both Type A and Type B gelatins (derived from acid and alkali hydrolysis of collagen, respectively), occasionally blended together by either the gelatin manufacturers or the softgel manufacturers, are used for preparing softgels. Gelatin type selection is influenced by both technical and economic considerations. Traditionally, bovine bones and skins (trimmings from the leather industry prior to tanning) have been used as collagenous raw materials for manufacturing Type A or Type B gelatin, while porcine skins have been used extensively for manufacturing Type A gelatin (Alleavitch et al., 1989; Johnston-Banks, 1990; GMIA, 1993). In recent years, porcine bones also have entered the stream of gelatin raw materials in Europe where they are either processed separately or co-processed with bovine bones to produce Type A or Type B gelatins. In the mid 1980s, fish gelatin became commercially available and has been marketed as an alternative to mammalian gelatins that present concerns for such religions as Judaism, Islam, and Hinduism. All fish are acceptable to most Islamic groups, while fish with removable scales are acceptable in Judaism with minimal restrictions (Choi and Regenstein, 2000). The skins of cod (a cold-water fish) were initially used for the commercial production of fish gelatin (Norland, 1987). However, due to its low content of hydroxyproline and proline amino acids, which contribute substantially to the gelatin gel structure through hydrogen bonding, cod skin gelatin is non-gelling at ambient temperatures (Norland, 1987; Leuenberger, 1991; Gudmundsson and Hafsteinsson, 1997; Gilsenan and Ross-Murphy, 2000) and, therefore, is not suitable for softgel manufacturing. Processes for extracting gelling gelatin having higher contents of hydroxyproline and proline amino acids from the skins of other fish species, such as tilapia (a warm-water fish), were later developed (Grossman and Bergman, 1992; Holzer, 1996). Although these gelatins have lower melting and gelling points than mammalian gelatins (Choi and Regenstein, 2000; Ross-Murphy, 2000), they are suitable for softgel manufacturing, and softgels

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made of fish gelatin are commercially available. Increasing the melting and gelling temperatures of fish gelatin is feasible through the addition of salts such as magnesium sulfate (Sarabia et al., 2000). The recovery of poultry gelatin has also been investigated (Kim et al., 2000) but not commercialized yet.

The gelatin used for softgel manufacturing meets compendial requirements such as those set forth in the United States Pharmacopeia/National Formulary (USP/NF) or the European Pharmacopeia (EP). For example, the NF monograph for gelatin (USP24/NF19, 1999) sets limits for residue on ignition ( 2.0%); sulfur dioxide ( 0.15%); arsenic ( 0.8 ppm); and heavy metals ( 0.005%). Tests for identification, odor, and water-insoluble substances, as well as microbial contamination (total bacterial count of 1000/g; absence of Salmonella species; and absence of Escherichia coli), also are included. Compendial requirements are general in scope and apply to gelatins used in all types of pharmaceutical dosage forms. Several other physical and chemical properties affect the encapsulating performance of gelatin and are monitored by softgel manufacturers. Gelatin batches used for softgels, or any other application, are prepared by blending a number of individual gelatin extracts or components (often up to 35) into a final product that meets customer specifications. The component selection (“recipe”) associated with this blending process can drastically affect the encapsulating performance of the gelatin batch. From a commercial standpoint, Bloom value (gel strength) and viscosity (related to average molecular weight), both determined on 6.67% w/w gelatin test solutions, are typically the two most important properties in assessing gelatin grade and quality (GMIA, 1993). Bloom strength is determined using texture analyzers, while viscosity is determined using U-tube viscometers at 60 C (GMIA, 1993). When comparing the Bloom strength and viscosity of various gelatins, it should be acknowledged that test measurements are affected by moisture content, ash content, and the pH of the gelatin (Stevens et al., 1995). In general, acid-processed gelatins have lower viscosity values (lower molecular weight) than lime-processed gelatins of the same Bloom strength. This is related to the ability of the acid treatment to cleave the acid-labile peptide bonds, in addition to disrupting the non-covalent bonds, in collagen (Rose, 1987). Gelatins having a Bloom strength from 150 to 250 g and a viscosity from 25 to 45 mP (at 60 C) may be used in the softgel industry (Stanley, 1986). However, in the interest of consistency, the softgel manufacturers’ specifications for various types of gelatins are much tighter, particularly in terms of viscosity (not more than 3 mP). In general, gelatin solutions show Newtonian flow at most temperatures, except those just above the gel setting point where the viscosity becomes time-dependent (Johnston-Banks, 1990; Wulansari et al., 1998). The

406 dissolution rate of softgel capsule shells made with Type B gelatin was shown to decrease linearly with increasing gelatin Bloom values (Hom et al., 1973). In addition to the “standard” viscosity of 6.67% (w/w) solutions, softgel manufacturers may also set specifications for the viscosity of more concentrated gelatin solutions (e.g., 12.5% w/w), because solution viscosity values are not directly proportional to gelatin concentration. Furthermore, viscosity drop (loss or breakdown) of a 12.5% (w/w) gelatin solution after 24 h at 60 C is used as a measure of the anticipated “pot-life” of a gelatin gel mass that is being heated. Usually, the greater the initial viscosity, the greater the viscosity drop in 24 h with the values for softgel grade gelatins ranging from 10 to 20%. In addition, microorganisms present in the test solutions accelerate the viscosity loss. As mentioned, the NF monograph for gelatin sets a limit of 1500 ppm for sulfur dioxide (SO2). However, SO2 can affect the stability of several synthetic dyes used with softgels. For example, FD&C Yellow #5 and FD&C Yellow #6 dyes are susceptible to fading upon interaction with SO2 (Downham and Collins, 2000). Therefore, stricter limits (e.g., 40 ppm) for the SO2 content of gelatin are used in the softgel industry. Iron, which may be introduced into the gelatin from raw materials or process water during manufacturing, can also affect the stability of synthetic dyes and can also participate in color-generating reactions with reducing substances that may be present in the capsule fill (Wilkinson and Hom, 1990). For example, ascorbic acid in fill formulations can react with iron, causing dark “spotting” on the capsule shell. Typically, the gelatin used for softgels has an iron content of 15 ppm (Stanley, 1986). Gelatin color and clarity, usually determined using spectrophotometric techniques, also are important, particularly for softgels that do not contain dyes. Generally, Type A pigskin gelatin has a lighter color and greater clarity than Type B bone or hide gelatin of the same Bloom strength. In terms of particle size, the gelatin used for softgel manufacturing ranges in mesh from 8 to 60 (2.36 mm to 0.25 mm). Fine particles in large amounts can cause dusting during handling, aggregates (“fish eyes”) in the gel reactors, and even foaming during gel manufacture. Coarse particles in large amounts can reduce the hydration rate of the gelatin in the gel reactors and also can cause handling problems with reactor feeding systems. From a microbiological standpoint, gelatin, although deficient in certain essential amino acids, may support microbial growth at higher moisture contents. Therefore, in addition to the mentioned NF microbiological testing (i.e., total microbial count, Salmonella, and E. coli), softgel gelatin specifications typically require absence of Pseudomonas aeruginosa and Staphylococcus aureus, and a total mold and yeast count of 100 organisms/g. The physical and chemical properties discussed above establish a basic set of quality attributes for softgel-grade gelatin. However, the overall performance of a gelatin blend during gel manufacturing, encapsulation, capsule drying, and long-term capsule storage is dictated by additional functional characteristics and can only be assessed through actual use tests. Examples of such

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characteristics include the viscosity of the gel mass and the loss of viscosity with heating; the Bloom strength of the gel mass and its decrease with heating; the stickiness of the gel mass and of the capsules; the strength of the capsule seams; the capsule hardness and brittleness; the drying rate of the capsules; and the physical stability of the capsules during storage.

Gelatin, plasticizer, and purified water are the main components of softgel shell formulations. In general, glycerin is widely used to plasticize various protein films and coatings (Gennadios et al., 1994). Glycerin of animal, vegetable, or synthetic origin is an effective plasticizer for softgels containing lipophilic fills. Gelatin and glycerin are highly compatible because strong intermolecular interactions are formed between the hydroxyl groups of glycerin and the hydrophilic groups of gelatin (Pouradier and Hodot, 1972). In fact, in high amounts, glycerin can be a solvent rather than a plasticizer for gelatin (Pouradier and Hodot, 1972). However, glycerin-plasticized softgels with hydrophilic, PEG-based fill compositions become brittle over time as both glycerin and water (which itself plasticizes gelatin) migrate from the shell into the hygroscopic fill (Brox, 1988a, b; Shah et al., 1992). To prevent this embrittlement, glycerin typically is used in combination with non-crystallizing sorbitol solutions to plasticize softgels having hydrophilic fill formulations (Brox, 1988a, b). Sorbitol, being less hydrophilic, having a higher molecular weight, and being less soluble in PEG, tends to not migrate into PEG-based fills as easily as glycerin does. Non-crystallizing sorbitol solutions are mixtures of sorbitol and sorbitol anhydrides or hydrogenated oligosaccharides (Reich, 1996). They are preferred over crystalline sorbitol, which tends to re-crystallize over time, forming white spots on the surfaces of the capsules (“blooming”). In addition to using non-crystallizing sorbitol solutions, the migration of glycerin and moisture from the shell into the hydrophilic fill can be further reduced through the incorporation of glycerin or propylene glycol into the fill to bring the two phases closer to equilibrium (Brox, 1988a, b; Shah et al., 1992). Other polyhydric alcohols, such as propylene glycol (Brox, 1999), PEG 200 (Rouffer, 2000), or maltitol syrup (Borkan et al., 1990; Chiprich et al., 1997), may also be used, often in combination with glycerin, as softgel plasticizers in specialty shell formulations. Cast gelatin ribbons containing propylene glycol as a plasticizer were reported to be tackier than ribbons plasticized with glycerin or sorbitol (Brox, 1999). The ratio (by weight) of dry plasticizer to dry gelatin is an important parameter that defines the “hardness” of the capsule shell and usually ranges from 0.4:1 to 0.8:1 (Stanley, 1986). This ratio is selected based mainly upon the fill formulation and the anticipated storage conditions of the finished product. Hy-

408 drophilic fills require greater plasticizer to gelatin ratios than lipophilic fills to compensate for any plasticizer migration into the fill over time. Softgels shipped to hot, humid areas require lower amounts of plasticizer than those shipped to cold, dry areas (Stanley, 1986). Capsule size may also influence the selection of the plasticizer to gelatin ratio. For the same fill composition, orally administered capsules larger than 10 minims typically have a higher content of plasticizer to increase the ease of swallowing. Low plasticizer concentrations are recommended for softgels containing oxygen-labile active compounds because the oxygen permeability of gelatin films increases with increasing amounts of glycerin (Hom et al., 1975). Finally, “overplasticization” of the capsule shell is a simple approach for preparing chewable softgels (although this is of limited effectiveness without additional shell/fill formulation adjustments).

Softgel shells need to be made opaque when the encapsulated active compounds are light-sensitive. Also, opacified shells are typically used with paste fill formulations for aesthetic purposes. Titanium dioxide (TiO2), a white pigment (insoluble in water) that has found universal regulatory approval, is the opacifier of choice for softgels. It also is used to prepare white-colored softgels, often in combination with minuscule amounts of FD&C Blue #1 dye that is used to neutralize the yellowish background color of gelatin. The natural mineral ilmenite (FeTiO2) is the source of the TiO2 used in various industrial applications. However, only synthetically produced TiO2 can be used as a color additive for food, drug, and cosmetic applications (Francis, 1999). In addition to its opacifying and whitening ability, TiO2 can also reduce the oxygen permeability of capsule shells, presumably by increasing the path tortuosity of diffusing oxygen molecules (Hom et al., 1975). Kellaway et al. (1978) reported that high amounts (2% w/w) of TiO2 had negligible effects on the mechanical properties (tensile strength, elongation at break, and Young’s modulus) of cast films from Type A or B gelatins. Similarly, Samura et al. (1993) determined that TiO2 did not notably affect the tensile strength of cast softgel shell formulations. In fact, TiO2 at high levels increased the strength of gelatin gels, presumably via strong TiO2 particle-gelatin interactions and a degree of hydrogen bonding (Johnston-Banks, 1990). Nevertheless, TiO2 usually is not added to softgel shell formulations in amounts greater than 1% (wet weight) (Hom et al., 1975). At greater amounts, this insoluble gel additive (filler) can negatively affect the encapsulating characteristics, particularly seam strength, of softgel shell formulations. In addition, over time, it may harm encapsulation dies and other processing equipment due to its abrasiveness. Zinc oxide (ZnO) is another opacifying/whitening inorganic pigment. How-

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ever, its hiding power is five times lower than that of TiO2 (Marmion, 1991). ZnO is widely used in the cosmetics industry, but is not an approved color additive for dietary supplements or ingestible drugs in the U.S. However, it is occasionally added to the shells of nutritional softgels as a nutrient (elemental zinc source). Another whitener that has a weak opacifying ability is calcium carbonate (CaCO3), itself a dietary supplement. It is used widely in the food industry in various roles, but is not currently listed by the FDA as an approved color additive for foods (Francis, 1999). However, it is an approved color additive for drugs in the U.S. CaCO3 can be problematic as an opacifier for softgels due to its weak hiding power and low tinctorial strength (coloring power). Furthermore, dissociated divalent calcium cations in the gel can interact with negatively charged sites on the gelatin, thus causing cross-linking (“bridging” together of adjacent protein chains). TiO2-coated mica pigment can be added to the shells of cosmetic softgels to introduce a pearl-like appearance (Benford, 1970). Mica is a white powder obtained from the naturally occurring mineral muscovite mica, and it consists primarily of potassium aluminum silicate (Marmion, 1991). Finally, small amounts of oily substances can introduce a degree of haziness into the shell, offering an opacified appearance. For example, silicone oil has been used as an opacifier in softgels (Davies et al., 1987). In addition, large amounts of vegetable or animal oils were incorporated into shell compositions to protect capsule contents from light and to increase shell hydrophobicity and lubricity (Yamada et al., 1988).

The psychological effects of color upon consumers of oral dosage forms have long been recognized (Woznicki and Schoneker, 1991). Therefore, the color of softgels is quite important from both a marketing and product identification/differentiation standpoint. Oil fills (e.g., vitamin E, marine oils, and lecithin) are often encapsulated into softgels that do not have dyes or pigments within the shell. Such transparent softgels have light yellow to light brown colors imparted to them by the combination of the oil color and the inherent amber color of gelatin (which varies slightly by gelatin type, grade, and manufacturer). It is possible to completely “neutralize” the inherent color of gelatin, thus obtaining clear, colorless softgels, by combining various dyes and pigments in appropriate amounts (Tanner and Orange, 1997). In addition, Ext. D&C Violet #2 (Alizarin Violet), which is approved in the U.S. only for externally applied drug and cosmetic products, may be used to “neutralize” the inherent yellowish color of gelatin. Softgels containing suspensions (pastes) and hydrophilic fills are typically colored. Because two gelatin ribbons are fed into the encapsulation machine to form softgels, each ribbon may have a different

410 color, thus resulting in bi-colored (two-toned) softgels. Generally, the shell color should not be lighter in hue than the encapsulated fill (Stanley, 1986). Also, darker colors are more appropriate for large, orally administered softgels (e.g., greater than 14 oblong) because they do not accentuate the capsule size (Stanley, 1986). The selection of colorants for softgels containing pharmaceuticals or dietary supplements requires careful consideration of the regulatory approvals and restrictions for the various colorants that apply to the targeted markets (Jones, 1993). Both chemically synthesized colorants and colorants derived from plant, animal, or mineral sources are used to color softgels. The latter are often referred to in the food and dietary supplement industries as natural colorants, although the term “natural colorant” is not appropriate from a regulatory standpoint in the U.S. unless the colorant is natural to that food (Boyd, 1998). In the U.S., the approved colorants for food, pharmaceutical, and cosmetic uses are listed in Title 21 of the Code of Federal Regulations (CFR), Parts 73 and 74. The CFR specifies two groups: colorants that are certified to comply with the established purity specifications of the FDA and color additives that are exempt from certification (Francis, 1999). In the U.S., the Nutritional Labeling and Education Act of 1990 mandates that certified color additives are specifically declared with their individual names on product labels (Francis, 1999). However, exempt colorants can still be declared generically as “artificial color” or with any other specific or generic name for the colorant (Francis, 1999). In the European Union (EU), 43 colorants (17 synthetic and 26 either naturally derived, synthesized to match naturally occurring counterparts, or inorganic pigments found in nature) are approved as food additives with each one assigned an “E number” (Downham and Collins, 2000).

In the U.S., seven synthetic FD&C (“food, drug, and cosmetic”) dyes are certified without restrictions for use in coloring foods and, therefore, dietary supplements (Francis, 1999). These are FD&C Blue #1 (Brilliant Blue), FD&C Blue #2 (Indigo Carmine or Indigotine), FD&C Green #3 (Fast Green FCF), FD&C Yellow #5 (Tartrazine), FD&C Yellow #6 (Sunset Yellow FCF), FD&C Red #3 (Erythrosine), and FD&C Red #40 (Allura Red AC). Due to their water-soluble nature, these FD&C dyes can be readily incorporated into a gel mass and, thus, be used to color softgels. However, FD&C Blue #2 has poor heat, light, and oxidative stability (Kuramoto et al., 1958; Kellaway et al., 1978; Downham and Collins, 2000) and, therefore, is typically avoided in the softgel industry. In addition, FD&C Blue #1 has only a fair light stability (Downham and Collins, 2000) and, therefore, may be problematic in softgels. In addition to the seven FD&C dyes, several other dyes designated as D&C (“drug and cosmetic;” considered safe in drugs and cosmetics when in contact

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with mucous membranes or when ingested) are available (Marmion, 1991) and may be used with pharmaceutical softgels. Among them, D&C Yellow #10 (Quinoline Yellow WS), D&C Red #27 (Tetrabromotetrachlorofluorescein), D&C #28 (Phloxine B), and D&C Red #33 (Acid Fuchsine) are the ones most commonly used with softgels. Synthetic dyes are available both in powder and granular form. However, powder forms, although less expensive than granular forms, can cause dusting (Dziezak, 1987). For this reason, and also for avoiding “fish eyes” upon incorporation into the gel masses, granular dyes are preferably used in the softgel industry. Although incorporating synthetic dyes (and other colorants) directly into shell formulations prior to capsule manufacturing is the norm in the softgel industry, surface dyeing of finished softgels using solutions of various water-soluble dyes in aqueous isopropyl alcohol has been studied (Smith, 1974). The FD&C and D&C dyes commonly used with softgels can be broadly assigned to one of six chemical classifications based on their chemical structure (Marmion, 1991). These are azo dyes (i.e., FD&C Yellow #5, FD&C Yellow #6, FD&C Red #40, and D&C Red #33), indigoids (i.e., FD&C Blue #2), xanthenes (i.e., FD&C Red #3 and D&C Red #28), quinolines (i.e., D&C Yellow #10), triphenylmethanes (i.e., FD&C Blue #1 and FD&C Green #3), and fluorans (i.e., D&C Red #27). Over the years, some of these dyes, particularly azo dyes, have faced consumer distrust due to their perceived association with largely unsubstantiated health concerns, such as hyperactivity and food intolerances (Downham and Collins, 2000). In particular, FD&C Yellow #5 was suspected of causing allergic-type reactions, including asthmatic symptoms, especially in individuals allergic to aspirin (Rumore et al., 1992). As a result, several companies marketing pharmaceutical or nutraceutical compounds in softgels presently avoid the use of azo dyes in shell formulations. Besides the water-soluble FD&C and D&C dyes, their corresponding water-insoluble lakes (still provisionally listed by the FDA with the exception of FD&C Red #40 lake, which has full approval and FD&C Red #3 lake, which is not permitted due to concerns over its iodine content) are occasionally used to color softgels (or gelatin-enrobed or gelatin-coated tablets/caplets). Lakes are the aluminum salts of water-soluble dyes that usually are adsorbed onto an alumina substrate (other substrates, such as clay, TiO2, and CaCO3, also are approved for D&C lakes) and impart color by dispersion (Francis, 1999). The dye content of lakes usually ranges from 10 to 40% (Marmion, 1991). Lakes have excellent light and heat stability (Francis, 1999). The main use of lakes in the softgel industry is with bi-colored capsules where they minimize color “bleeding” at the seams and color transfer marks between capsule halves. In addition to color, lakes also introduce a high degree of opacity to the softgel shells. Association of synthetic dyes and lakes with gelatin through ionic bonding and also hydrogen and hydrophobic interactions has been documented (Sheppard et al., 1942; Cooper et al., 1973; Gautam and Schott, 1994). Factors

412 that affect these dye-gelatin interactions include pH, temperature, and gelatin type (A or B). At the typical pH (around 5) of softgel gel mass preparations, the synthetic dyes, due to their anionic nature, will electrostatically associate to a greater extent with the positively charged Type A gelatin than the slightly negatively charged Type B gelatin (Cooper et al., 1973; Kellaway et al., 1978). However, because they are present in small amounts, synthetic dyes and lakes are not considered to appreciably affect the disintegration and dissolution characteristics of softgels. Nevertheless, Cooper et al. (1973) reported that FD&C Red #3 dye notably reduced the disintegration rate of type A gelatin. UV or visible irradiation under high relative humidity (RH) conditions notably reduced the disintegration and in vitro dissolution rates of hard gelatin capsules colored with various dyes such as FD&C Red #3 and FD&C Yellow #5 (Murthy et al., 1989a). Another study showed that FD&C Red #3, FD&C Blue #1, and FD&C Blue #2 at 0.1 or 1% (w/w) had minimal effects on the tensile strength and Young’s modulus of cast gelatin (Type A or B) films, but they reduced film contraction at a constant strain (Kellaway et al., 1978). Synthetic iron oxides also find use as softgel colorants. Yellow, red, and black iron oxides, typically produced from ferrous sulfate, are available commercially (Marmion, 1991). They are water insoluble (Francis, 1999), thus coloring gelatin gel masses by dispersion and also introducing a degree of opacity. Their stability to heat, light, pH, and oxidation is excellent (Downham and Collins, 2000). In the EU, synthetic iron oxides are approved color additives for both foods and pharmaceuticals, and are used to color softgels containing nutraceutical or pharmaceutical products. In the U.S., synthetic iron oxides are permitted colorants for ingestible drugs, but not for foods/dietary supplements. When used with softgels, synthetic iron oxides do not offer colors as vibrant and glossy as those obtained with synthetic dyes. However, the synthetic iron oxides are used often with pharmaceutical softgels because of their wide regulatory approval worldwide as colorants for drug products (Jones, 1993).

The list of color additives exempt from certification in the U.S. includes 26 colorants, but some of them are only approved for use with animal feed (Francis, 1999). Five of these 26 colorants (i.e., caramel, annatto extract, cochineal extract, turmeric, and turmeric oleoresin) are commonly used, often in combinations of two or more, to color softgels containing nutraceutical compounds. Three of them (caramel, annatto extract, and cochineal extract) also are permitted for use with pharmaceuticals in the U.S. However, their use in pharmaceutical softgels has been scarce, if at all. Caramel is a brown colorant manufactured by the controlled heat treatment of food-grade carbohydrates, typically corn syrup solids (Kamuf et al., 2000).

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Sulfite and/or ammonia compounds may be used as catalysts in the caramelization process (Downham and Collins, 2000). Due to its high tinctorial strength, good solubility in gelatin gel masses, good stability, and low cost, caramel is widely used to impart brown colorations to nutritional softgels. Annatto is a yellow carotenoid obtained from the seeds of the plant Bixa orellana (Francis, 2000a). The main pigments in annatto extract are the water-insoluble bixin and the water-soluble norbixin (Boyd, 1999). Bixin, the monomethyl ester of a dicarboxylic carotenoid, is the naturally occurring pigment form, while norbixin is the saponified form (Francis, 2000a). Aqueous preparations of norbixin at various concentrations are often added to the hydrophilic softgel shell to obtain orange/yellowish colorations. Oil-based bixin preparations also are used in softgels with the oil introducing a degree of haziness to the shells. Some fading upon exposure to light may occur with annatto-colored softgels. From a regulatory standpoint, there are no restrictions on the use of annatto as a color additive in the U.S. However, since 1994, the EU has significantly restricted the use of annatto colorant by categorizing it as a “color only permitted for certain uses” (Boyd, 1999). Saffron is another approved yellow color additive in the U.S. (no longer listed in the EU). Its carotenoid pigments (crocetin, crocin, and zeaxanthin) are chemically similar to those of annatto (Francis, 2000a). However, although suitable for coloring softgels, it is rarely used in the softgel industry due to its prohibitively high cost. Cochineal extract is the red, concentrated solution obtained after removing the ethanol from aqueous ethanol extracts of the dried bodies of the female insect Dactylopius coccus Costa (Francis, 1999). The main pigment in a cochineal extract is carminic acid, a hydroxyanthraquinone linked to a glysosyl group (Schul, 2000). Carmine is the lake of carminic acid (typically about 50%) (Marmion, 1991). The water-insoluble carmine pigment can be converted to a water-soluble dye by treatment with alkali (Schul, 2000). Cochineal extract, the carmine lake, and water-soluble carmine (often called carmine hydrosoluble) are stable to light and heat, resistant to oxidation, and not affected by sulfur dioxide (Francis, 1999). They function well as colorants for softgels where they impart pink/red/purplish colorations. However, their use is self-limiting due to cost considerations. Turmeric is a yellow colorant prepared from the dried roots of Curcuma longa, a tuberous rhizome (Pszczola, 1998). It owes its yellow color to three structurally related pigments (i.e., curcumin, demethoxy curcumin, and bis-demethoxy curcumin) (Chaterjee et al., 1998). Turmeric oleoresin is prepared by extracting turmeric powder with one or more organic solvents (Marmion, 1991). Both turmeric powder (ground rhizomes) and turmeric oleoresin (typically sold as a dispersion in propylene glycol or glycerin) are used as softgel colorants. Due to its water-insoluble nature (Francis, 1999), turmeric also introduces an opacifying effect to softgel shells, thus offering a degree of light protection to the capsule contents. Softgel shells colored with pure

414 curcumin (0.02 to 0.04% w/w) effectively increased the half-life of encapsulated photolabile test compounds (i.e., nifedipine, chloramphenicol, furosemide, or clonazepam) compared to non-colored softgel shells following irradiation with light of 240–600 nm (Tønnesen and Karlsen, 1987). The main drawback of turmeric as a colorant is its high susceptibility to light degradation (Boyd, 1998). Sodium copper chlorophyllin is a green colorant obtained from chlorophyll by replacing the methyl and phytyl ester groups with sodium and replacing the magnesium with copper (Marmion, 1991). Commercial sources of sodium copper chlorophyllin include alfalfa, mulberry tree leaves, and nettle. It finds wide use in the EU, Japan, and other parts of the world as a “natural,” green colorant in various applications, including softgels. Sodium copper chlorophyllin is susceptible to thermal degradation during heating (Milosevic et al., 2000). Although it may degrade to some extent during the heating of a gel mass in softgel manufacture, it yields quite stable color shades in stored softgels. In the U.S., sodium copper chlorophyllin is sold in powder and liquid forms as a dietary supplement. However, its use as a color additive is only permitted for dentifrices provided that it is derived from alfalfa (Francis, 1999). Several other “natural” colorants, such as anthocyanins, betalains, -carotene, riboflavin, and paprika oleoresin, are approved for use with foods/dietary supplements in the US and other countries. Commercial sources of anthocyanin pigments include grape skins, elderberries, chokeberries, black carrots, and red cabbage (Boyd, 2000; Downham and Collins, 2000). These pigments are presently receiving attention due to the health-related benefits ascribed to them (Boyd, 2000; Espín et al., 2000). However, anthocyanins have not typically been used for coloring softgels because of their pH sensitivity (lower color intensity at pH > 4) and susceptibility to degradation by light, heat, and oxygen (Francis, 2000b). Recently, commercialized acylated anthocyanins, such as those extracted from black carrots and red cabbage, were reported to have enhanced heat and light stability (Downham and Collins, 2000). Betalains, which are commercially extracted from beet roots, are largely unsuitable for coloring softgels because they have poor light and heat stability (Francis, 2000b). -Carotene and riboflavin are susceptible to light degradation (Marmion, 1991) and are used sparingly with softgels. Paprika oleoresin is produced by extracting dehydrated ground paprika with organic solvents (Locey and Guzinski, 2000). The pigments in paprika oleoresin are a mixture of carotenes and xanthophylls (Locey and Guzinski, 2000). Paprika oleoresin also is prone to degradation (Jarén-Galán et al., 1999) and is largely unsuitable for the softgel manufacturing process. Lycopene, the principle carotenoid pigment (red/orange) in tomato, is a listed color additive in the EU (Downham and Collins, 2000). However, it is not currently approved as a color additive in the U.S., although efforts are under way to obtain its approval (Boyd, 2000). The use of lycopene as a colorant for softgels in Europe has been hindered by its high cost and poor sta-

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bility, as it tends to isomerize and degrade over time (Anguelova and Warthesen, 2000). Arguably, softgels colored with “natural” colorants are more susceptible to variability in color shades from lot to lot compared to softgels colored with synthetic colorants. To improve lot to lot color consistency, commercial preparations of “natural” colorants often are “standardized” with excipients. Such excipients may include reducing sugars (e.g., maltodextrin and corn syrup solids); however, they can cross-link gelatin, and, therefore, their presence should be monitored when coloring softgels. The demand for “natural” colorants in the dietary supplement industry is quite strong and is expected to keep growing. This demand is generated by the increasing consumer pressure for “all-natural” products. In addition, several of the “natural” colorants are perceived to be functional ingredients themselves because they possess antioxidant and/or antimicrobial properties (Boyd, 2000). Undoubtedly, the stability concerns associated with various “natural” colorants present technical challenges regarding their use with softgels. Microencapsulation is currently receiving attention in the color industry as an approach for improving colorant stability (Downham and Collins, 2000).

Several minor additives may occasionally be added to softgel shell compositions. In the past, preservatives often were incorporated into the gelatin shell, particularly with pharmaceutical products. However, at present, the use of preservatives with softgels is uncommon. The large amount of humectants, such as glycerin and/or sorbitol, present in the softgels suppresses water activity (Hom and Jimerson, 1990). Due to the low water activity of dried, properly stored softgels, molds and yeasts had been the main concern in terms of microbial growth rather than bacteria. For this reason, the preferred preservatives for softgels were the alkyl esters of p-hydroxybenzoic acid (methyl, ethyl, propyl, butyl, or heptyl), which are known as parabens. Parabens, which are widely used antimicrobial agents in foods, pharmaceuticals, and cosmetics, have little effect on flavor, effectively inhibit molds and yeasts, and are relatively ineffective against bacteria, especially gram-negative bacteria (Lindsay, 1985). Two or more parabens often are used in combination because together they exhibit enhanced antimicrobial activity (Thompson et al., 1993). Typically, a mixture of methyl paraben and propyl paraben at a ratio of 4:1 w/w was added at 0.2% by weight of the wet gel mass (Stanley, 1986). Thompson et al. (1993) reported that parabens can interact with glycerin or sorbitol forming substances that could present problems with the assay validations of pharmaceutical formulations. Sorbates (Heizi et al., 1979) or propionates (Wittwer and Mayer, 1986) also were suggested as shell-added preservatives (Heizi et al., 1979). Flavorings, such as ethyl vanillin and essential oils, can be used to mask the

416 unpleasant odors and tastes of fill formulations (Stanley, 1986). Multivitamin/multimineral compositions are examples of softgels that often contain shell flavorings. However, aldehydes present in flavorings can be a concern because, as mentioned earlier, they cross-link gelatin. The use of flavorings is becoming less and less common. Buffering salts and acids (e.g., citric acid, tartaric acid, fumaric acid, and malic acid) have been used to adjust the pH of the gel mass (Mima et al., 1969; Henmi et al., 1987). Fumaric acid, in particular, incorporated at up to 1% (w/w) in a wet gel mass can prevent gelatin cross-linking by aldehydes (Stanley, 1986). Addition of fumaric acid or other acids at amounts greater than 1% is not recommended because the acids increase the viscosity degradation rate of gel mass preparations (Johnston-Banks, 1990), most likely due to acid hydrolysis of the protein chains. In general, the rate of viscosity loss of gelatin preparations is affected by the type of acid in the order (from least to most degradation) of citric, malic, tartaric, adipic, and fumaric acid (Johnston-Banks, 1990). In addition, Hom et al. (1973) reported that incorporating organic acids (e.g., fumaric, tartaric, or maleic acid) notably increased the dissolution rate of a typical softgel shell formulation. Besides fumaric acid, hydrolyzates of gelatin or other proteins as well as amino acids can be added to soft or hard gelatin capsules to “scavenge” aldehydes (or other reactive agents), thus preventing gelatin cross-linking (Tatematsu et al., 1991; Cade and Madit, 1997). Naturally, added hydrolyzates, which lack gelling strength, have a weakening effect on gelatin gels (Surówka, 1997). Antifoaming agents are used in the shell compositions of paintballs, which are typically manufactured with Type A pigskin gelatin (150 to 220 Bloom), to prevent excessive foaming during gel manufacturing. In contrast, the manufacture of softgels using foamed gelatin ribbons that were prepared by microdispersing a gas into the gel mass was described (Wittwer and Mayer, 1986). The purpose was to introduce opacity, to accelerate shell disintegration, and to reduce manufacturing costs through lower gel consumption and faster capsule drying (Wittwer and Mayer, 1986). In theory, incorporation of active compounds into capsule shells is feasible. However, this is not economical because a substantial proportion of the gelatin ribbons used in softgel manufacturing ends up as waste (Stanley, 1986). During the storage of softgels, migration of active compounds, particularly water-soluble compounds, from the fill into the shell can occur. For this reason, when the softgels are intended for oral use, the entire capsule needs to be assayed for the active compounds rather than just the fill (Stanley, 1970).

As previously mentioned, gelatin, plasticizer, and purified water are the main ingredients of the softgel shell. Typical gel formulations contain 40 to

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50% gelatin, 20 to 30% plasticizer, and 30 to 40% purified water (w/w) (Hom and Jimerson, 1990) with most of the water subsequently lost during capsule drying. The ingredients are combined to form a molten gel mass using either a cold melt or a hot melt process. The cold melt process involves mixing gelatin, plasticizer, and chilled water and then transferring the resulting light, “fluffy” mixture to a jacket-heated tank (melter) (Wilkinson and Hom, 1990). Alternatively, mixing and melting can be successively accomplished within a single jacket-heated, batch reactor equipped with a blade agitator (Figure 16.3). Typically, gelatin is first added to the slightly chilled plasticizer (14 to 18 C) followed by mixing for 10 to 30 min, a step referred to as “creaming.” The purpose of creaming is to “wet” the gelatin particles and facilitate their subsequent hydration without clumping. Chilled (7 to 12 C), purified water is then dispensed, and the mass is mixed without heating for another 15 to 30 min, a step referred to as “fluffing.” Water may be added to the creamed mass in one (total quantity) or more cycles (equal or unequal partial quantities). The fluffed mixture is melted (“cooked”) under a vacuum of 15 to 30 in. Hg (50.8 to 101.6 kPa) to a homogeneous, deaerated gel mass. Vacuum application may be continuous or intermittent during the melting process. The cooking temperature can range from 57 to 95 C, depending on the type of melter/reactor, and the process lasts 1.5 to 3 h. Shell additives, such as colorants/opacifiers, preservatives, and flavorings, can be added to the mass at any point during the gel manufacturing process, or they may be incorporated into the finished gel mass using a high-torque mixer. Colorants/opacifiers, in particular, often are added during the creaming stage. The hot melt process involves adding the gelatin, under mild agitation, to a preheated (60 to 80 C) mixture of plasticizer and water and stirring the blend until complete melting is achieved. Compared to the cold melt method, the hot melt method is faster but is often more susceptible to foaming and dusting. When the gel mass is intended for manufacturing non-ingestible capsules, such as paintballs, antifoaming agents often are added to the gel formulation. Once the gel masses are prepared, they are transferred to preheated, temperature-controlled, jacketed holding tanks (receivers) where they are kept (“aged”) at 50 to 60 C until encapsulation. Often, the gel mass is filtered prior to being transferred to the receivers so that any undissolved gelatin aggregates or impurities initially present in the raw materials are removed. In addition, a visual comparison of the gel mass with a color standard (e.g., gel swatch or finished capsule) commonly is made in the case of colored gels. Gel aging can last from two to 72 hours or ever longer depending on the gel formula. Besides serving logistical needs, the aging of the gel mass for certain time periods is often needed to “break down” the gel to a desired viscosity. The functional life of the molten gel mass in the receivers can be extended by cooling it to ambient temperature, thus gelling it, a process referred to as “caking.” They are then remelted (reheated to 50 to 60 C) and used. In general, the time-temperature

418 Figure 16.3 Process flow diagram for softgel manufacturing (courtesy of Banner Pharmacaps Inc., High Point, NC).

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history of gelatin gels affects their thermal stability, melting characteristics, and tensile properties (Castello and Goyan, 1964; Michon et al., 1997). Gelatin gels conditioned (matured) at higher temperatures can have higher melting points than gels of the same solution prepared by rapid chilling at a lower temperature (Te Nijenhuis, 1981; Michon et al., 1997; Choi and Regenstein, 2000). Aging “caked” (4 C), dilute (1.5 or 2.0% w/w) Type A or Type B gelatin gels for up to 90 h did not markedly affect the viscoelastic properties of the gels (Robinson et al., 1975). However, a decrease in gel structure was evident, and this was attributed to hydrolysis of amide links (Robinson et al., 1975). The rotary die encapsulation process inherently generates a notable amount of gel mass waste in the form of unused gel ribbons remaining after the formed capsules are cut out by the dies. Due to its appearance, this unused portion of the ribbon is referred to as “netting.” Additional, smaller amounts of gel mass waste are generated through leftover gel (overage) at the completion of an encapsulating run and through unused gel ribbons cast during the start-up of the encapsulation machines. The percentage of unused gel mass typically varies between 15 and 30%. This excess gel mass is for the most part discarded. However, it can be recovered and reused, a practice that is driven by economic factors and is mainly limited to non-colored, low-value, non-pharmaceutical products such as nutritional oils. In this case, the reused gel mass is combined with a new (“virgin”) gel mass and may account for up to 50% of the total mass. In the past, the reprocessing of netting involved chopping and washing with naphtha to remove the lubricant oil applied to the ribbon during the encapsulation process (Wilkinson and Hom, 1990). This residual oil needs to be removed because it causes gel “cloudiness.” The oil-free, chopped netting was then remelted and added to a new gel mass or was “caked” until needed (Wilkinson and Hom, 1990). Recently, as softgel manufacturers have been reducing their use of organic solvents, different approaches have been implemented for removing residual oil (and fill material) from the netting. The simplest approach is decanting, which involves remelting the netting (no chopping necessary), possibly with added purified water, in heated gel receivers. A phase separation occurs in the molten mass with a thin, oil-rich phase forming at the top. The clean gel (lower aqueous phase) is then recovered from the bottom of the tank. Although effective, the decanting method for netting recovery is time- and energy-consuming, because the remelted mass is kept heated for several hours to ensure a thorough oil-gel phase separation. Hot filtering and subsequent vacuum distillation of the recovered gel has been suggested to enhance purification (Schmidt et al., 1994). Furthermore, diafiltration, a filtration technique that uses ultrafiltration membranes and washing, may be employed to remove plasticizers and other water-soluble components from the recovered gel, thus yielding a recyclable gelatin solution (Schmidt, 1999). Alternatively, a gel netting recovery device was recently described that removed lubricant oil by feeding cut, rod-like gel pieces between adjacent, rotating belts of differing speeds

420 (Ohzeki and Ishikawa, 2000). The lubricant adhered to the belts and was removed from the belt surface by blades (Ohzeki and Ishikawa, 2000).

The principles of the rotary die encapsulation process have been described (Stanley, 1986; Hom and Jimerson, 1990; Wilkinson and Hom, 1990; Stringer, 1994). A schematic diagram of the process is presented in Figure 16.4. The gel mass is fed either by gravity (Figure 16.3) or by positive displacement pumping to two heated (48 to 65 C) metering devices (spreader boxes), usually made of brass. The dynamic viscosity of the gel mass as it is fed to the encapsulation machine typically ranges from 5000 to 20,000 cP, depending on temperature, gel formulation, and raw gelatin viscosity. However, a range of 8000 to 12,000 cP often is preferred. The spreader boxes control the flow of gel through adjustable openings onto cooled, rotating casting drums (Figure 16.4). The temperature on the surface of the casting drums, which is controlled by either cool air or water, is maintained at 20 to 30 C. Ribbons are formed as the cast gel masses set (sol-gel transformation) on contact with the cool drum surface. Controlled ( 10%) wet ribbon thickness can vary between 0.50 and 1.14 mm. However, a range of 0.63 to 0.81 mm is more representative (Stanley, 1986). Besides gel mass temperature, gelatin concentration in the gel mass, and drum surface temperature, the blending profile (molecular weight distribution) of the gelatin greatly influences the setting

Figure 16.4 Schematic diagram for the rotary die process of softgel manufacturing (courtesy of Banner Pharmacaps Inc., High Point, NC). A gel hoses; B medicine (fill) hose; C (medicine) fill pump; D leads; E injection wedge; F guide rolls; G gel ribbons; H reciprocating dies; I gel casting drums; and J spreader boxes (gel metering devices).

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rate of the cast gel mass. Generally, the setting rate of gelatin solutions at a given cooling temperature is reduced by high molecular weight components (Johnston-Banks, 1990; Kobayashi et al., 1992). The formed ribbons are fed through a series of guide rolls and between the injection wedge and the counter-rotating, cylindrical capsule-forming dies (Figure 16.4). A food-grade lubricant oil is applied onto the ribbons to reduce their tackiness and facilitate their handling by the rollers. In addition, the lubricant prevents sticking among freshly formed capsules and also seals the ribbons along the injection wedge, thus preventing air from entering the capsules as they are formed (Stroud et al., 1998). The oil may be applied by passing the ribbons through an oil bath or, to better control the amount of applied lubricant, by having the ribbons directly contact application rolls. Mineral oil or medium chain triglycerides are the most commonly used lubricants, although other oils, such as soybean oil, may be used. Eliminating lubricant application has been proposed (Makino, 1996). Fill formulations, often referred to as medicines, are held in stainless-steel, jacketed tanks (receivers) and are fed to the encapsulation machine by gravity (Figure 16.3). Fill preparation may involve simple blending or, in the case of suspensions, successive mixing, milling or homogenizing, and deaeration steps (Stanley, 1986). Nitrogen blanketing of the medicine tank head space often is practiced with fills susceptible to oxidation. The fill material is precisely metered by a positive displacement pump through a series of leads and the injection wedge, and is pumped between the gelatin ribbons, which are passing through the cylindrical dies (Hom and Jimerson, 1990). Brass, aluminum, or stainless-steel may be used to manufacture the dies, which are typically six or 10 inches (15.2 to 25.4 cm) long. The maximum fill formulation viscosity that can be handled by the process is around 20,000 cP. However, materials that are highly tacky may be unsuitable for processing through the pump filling mechanism even if their viscosities are low (Stanley, 1986). Small orifices at the bottom of the wedge are lined up with the die pockets (Hom and Jimerson, 1990). The leading edge of the capsule is already sealed when the pumped fill material forces the ribbons into the die pockets where the capsules are then filled and shaped, their trailing edge sealed, and the filled softgels cut from the gelatin ribbon. Capsule sealing is achieved by the combination of heat from the wedge (33 to 43 C) and mechanical pressure (0.3 to 0.6 MPa) on the die rolls. Ribbon thickness, seam thickness, and fill weights are the most commonly monitored process parameters during the encapsulation operation and often are included in process validation programs (Berry, 1984). The seam strength of the finished softgels may also be assessed by determining their bursting strength. A device for measuring the bursting strength of softgels was described by Trenktrog et al. (1998). Alternatively, a compression test with a texture analyzer may be used. The theoretical output of an encapsulation machine depends on capsule size, die width, and the rotating speed of the casting drums/dies,

422 which typically varies between 3 and 5 rpm. Usual production rates range from 30,000 to 120,000 capsules per hour. Environmental conditions in the encapsulation areas are usually controlled within 18 to 22 C and 20 to 40% RH. For manufacturing pharmaceutical products, the encapsulation lines are usually segregated from each other into individual suites. In the past, the formed capsules were typically conveyed through a washing unit of naphtha or another organic solvent to remove residual lubricant and traces of fill material from the capsule surfaces (Stanley, 1986). Recently, as softgel manufacturers have focused on reducing their use of organic solvents, the capsule washing step has been viewed as not essential and often is skipped. Limiting lubricant application to a minimum helps negate the need for the washing step (Stroud et al., 1998). Subsequently, the capsules are transferred pneumatically or by a conveyor belt to a stainless-steel tumbler (dryer) consisting of successive rotating baskets (hollow drums) with perforated walls (Figure 16.3). The air coming in contact with the capsules in the tumbler typically is at the same temperature as the encapsulating room (18 to 22 C). Alternatively, heated (30 to 35 C) air may be pumped through the rotating baskets. Absorbent towels often are added to the tumblers to remove oil residues from the capsule surfaces. Immediately cooling the formed capsules to 2 to 7 C with low RH (20%) air prior to transferring them into the tumbler was suggested in a patent disclosure (Herman, 1992). The purpose was to prevent deformation of the warm capsules and to facilitate moisture removal from the capsule surfaces (Herman, 1992). However, the potentially negative influences of rapid cooling upon the shell structure should be considered. The tumbling action, which may last from 45 to 90 min, removes a large amount of moisture from the capsules. Alternatively, an infrared dryer may be used, which removes 60 to 70% of the total moisture eventually lost (Stanley, 1986).

After the capsules are “dumped” from the tumbler into a stainless-steel container (“back bin”), they are manually “scooped” and spread onto shallow fiberglass trays. The trays are then stacked and kept under controlled environmental conditions (18 to 24 C and 20 to 30% RH), often in drying tunnels with forced air circulation, to complete the drying process. Drying may last anywhere from 0.5 to 15 days depending on several factors such as environmental conditions, shell composition, shell thickness, fill formulation, number of capsules on the drying trays, and amount of residual oil on the capsule surfaces. The “dry” softgels have a final moisture content of 5 to 10%. Naturally, softgels with hydrophilic fills (e.g., PEG-based) require longer drying times. Determination of moisture content (usually by Karl Fischer titration) or hardness are the most commonly used criteria for releasing softgels from the drying room. It is possible to remove additional moisture from the dry capsules through

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heating at higher temperatures (e.g., 40 C), but such practice is not considered practical or necessary (Stanley, 1986). Also, a “stress-relieving” step where dry softgels are tempered at 32 to 43 C and 35 to 60% RH to remove dimples and other physical defects from the capsule shells has been suggested (Steele and Dietel, 1993). However, gelatin insolubility due to dehydrothermal cross-linking induced by heat treatments at elevated temperatures (55 to 105 C), especially at a reduced pressure, is well documented (Yannas and Tobolsky, 1967; Marks et al., 1968; Welz and Ofner, 1992; Vassileva et al., 1999). Therefore, subjecting dry softgels to elevated temperatures, even as low as 40 C, should be viewed cautiously as it may negatively affect shell solubility characteristics. In general, both the drying conditions and the drying rate of softgels are critical parameters. Accelerated (“aggressive”) drying, although economically appealing (faster throughput), can result in brittle capsules (Reich, 1995). On the other hand, incomplete drying results in soft, tacky capsules. Dry softgels typically are polished in revolving pans using absorbent towels carrying trace amounts of lecithin, isopropyl alcohol, or naphtha. Lecithin is not used for this purpose if the softgels are to be subsequently printed because it interferes with the printing inks. This polishing removes oil residues and other impurities from the capsule surfaces and maximizes the shiny, glossy appearance of the softgels. The polished softgels are passed through a mechanical sizer/sorter that removes any undersized/oversized capsules. From the sizer, the softgels are deposited onto a conveyor belt where they are visually inspected to detect and manually remove any defective capsules. Alternatively, the visual inspection may be accomplished prior to sizing while the capsules are still spread one layer thick on the shallow drying trays. Finally, the softgels are automatically filled by an electronic counter into bulk shipping cartons lined with polyethylene bags (moisture barrier) that are then labeled, sealed, palletized, and shipped. Subsequent retail packaging into plastic bottles, glass bottles, or blisters is accomplished with the same packaging equipment used for other solid dosage forms.

Pharmaceutical (and occasionally nutritional) softgels are printed (parallel or perpendicular to the long axis) for product identification. Both contact printing (i.e., rotogravure) and non-contact printing (i.e., ink jet or laser) may be used for this purpose. Rotogravure using Ackley, Hartnett, and Markem machines is the most common printing method for softgels and offers good quality printing. On-line printing of the cast gelatin ribbons prior to softgel formation using ink rollers also is feasible. The basic ingredients of ink formulations are pigments that impart color and opacity; resins that bind pigments and facilitate adherence onto printable surfaces; solvents that alter the physical characteris-

424 tics of the inks and control the drying rate; and special additives such as wetting agents, defoamers, and pH adjusters (Lykens, 1979). Similar to the selection of colorants, printing ink ingredients must be carefully monitored for regulatory compliance in the intended product markets. Traditionally, the softgel industry has been using ink formulations that combine organic solvents (e.g., SDA 3A alcohol, SD-45 alcohol, and N-butyl alcohol); shellac as a resin; and synthetic iron oxides, titanium dioxide, and/or lakes as pigments. However, water-based ink formulations, although generally more expensive than solvent-based formulations, have recently been gaining popularity as softgel manufacturers are undertaking efforts to reduce solvent use for environmental reasons. Water-based ink formulations use purified water with small amounts of isopropyl alcohol and/or methanol as the solvent system; hydroxypropyl methylcellulose (HPMC) as the resin; and synthetic dyes, synthetic iron oxides, and/or titanium dioxide as pigments. The removal of the oil residue from the softgel surface is particularly critical to good quality printing when using water-based inks. Solvent-based inks dry onto the softgels as the solvent evaporates, while water-based inks dry by adsoprtion onto the hydrophilic softgel surfaces (Spicer, 2000). Compared to solvent-based inks, water-based inks have a wider regulatory acceptance and are easier to clean up, but they offer a slightly less sharp image, dry slower, and are more expensive (Spicer, 2000). Ink-jet printing of softgels results in images of lower aesthetic quality than contact printing. In addition, only dyes, not iron oxides or lakes, can be used with jet printing, which presents stability concerns due to the light sensitivity of dyes (Spicer, 2000). Laser printing involves directing a high-intensity light beam onto the product to burn away a surface layer, thus imparting the printed image. As an obvious advantage, laser printing does not require pigments or solvents. However, it offers no color choices, is expensive, and the printed image tends to lose its sharpness over time.

Because softgel shells are hydrophilic, the environmental conditions during storage of bulk or retail packaged softgels can affect product quality. For optimum physical and chemical stability, softgels should be stored in the temperature range of 15 to 30 C and at RH less than 50% (Murthy and Ghebre-Sellassie, 1993). Packaging materials, such as high-density polyethylene (HDPE) liners in bulk cartons or retail HDPE bottles, protect softgels from moisture, but they are not an absolute moisture barrier. Therefore, upon prolonged exposure to elevated RH, packaged softgels can pick up moisture. The absorbed moisture softens, tackifies, and bloats the capsules (Stanley, 1986). Sticking (“bricking”) of capsules within the package can occur in such cases. In addition to the adverse physical effects, absorbed moisture can also negatively

Softgels with Modified Release Characteristics

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affect the chemical stability of hydrophilic fills as moisture migrates from the shell into the fill. For example, increased water content can reduce the solubility of poorly water-soluble drugs in PEG or other fill excipients, thereby causing drug crystallization (Serajuddin et al., 1986). Drug hydrolysis is another possible negative effect of absorbed moisture (Yoshioka et al., 1992). The disintegration characteristics of the capsule shells may also be affected by increased water content. However, Armstrong et al. (1983) reported that oil-filled softgels that absorbed moisture through exposure to elevated RH did not have significantly (P > 0.01) lower disintegration times. Increased storage temperatures, especially combined with increased RH (as is the case during stressed stability testing at 40 C and 75% RH), can have even more drastic effects on softgels. Capsule shell softening can occur, possibly causing the softgels to stick to each other or to even fuse together. Hakata et al. (1981, 1983) reported that softgels stored at 40 and 50 C had increased disintegration times. This was attributed to structural changes within the shell and, in particular, to denaturation of the collagen-like triple helix structure assumed by the gelatin in the dry shells (Hakata et al., 1981, 1983). The increase in disintegration time due to the storage of softgels at 40 C was prevented by the presence of desiccating agents (Hakata et al., 1992). Light is another environmental parameter that can cause physical changes to softgels, such as color fading or discoloration. This is particularly a concern with bath beads and other personal care softgel products that often are marketed in transparent retail packages. In contrast, pharmaceutical and nutritional softgels are generally marketed in light-resistant packages. A study by Hakata and Sato (1992) showed that exposure to fluorescent light did not affect the disintegration times of softgels. In a recent study, exposure to UV and visible illumination at 40 C and 75% RH for 8 days considerably retarded the dissolution rate of nimesulide softgels (Singh et al., 2000).

Orally administered softgels are intended to provide immediate release of their encapsulated contents upon exposure of the gelatin shell to the acidic environment and gastric enzymes in the stomach. Over the years, there has been an interest in developing softgels with modified release characteristics, such as delayed (sustained or continual) release or enteric release. A few fill formulation approaches to achieving a delayed release of active compounds from softgels have been proposed (Cohen et al., 1987, 1989; Hom and Ebert, 1989; Hom and Jimerson, 1990). The concept involves the use of insoluble natural or synthetic polymers (e.g., chicle, polyvinyl acetate, ethylcellulose, and calcium alginate) to prepare liquid to semi-solid cohesive matrices or microcapsules

426 containing the active compounds that are subsequently filled into softgels. The cohesive matrices or microcapsules allow for a delayed release of the actives by diffusion once the gelatin shell disintegrates. An Abbreviated New Drug Application (ANDA) for such a sustained release softgel (300 mg theophylline) was approved in the U.S. by the FDA (Stringer, 1994). Imparting delayed release characteristics to gelatin microcapsules or films by cross-linking the gelatin with aldehydes or reducing carbohydrates has been investigated (Po and Mhando, 1984; Bower et al., 1992; Akin and Hasirci, 1995; Cortesi et al., 1999). This approach also is applicable to softgels (Rolle, 1972; Hakata et al., 1994). In fact, softgels cross-linked with aldehydes or reducing sugars have found limited commercialization for products such as garlic oil (which causes reflux when released rapidly in the stomach). However, residual aldehydes are a cause for concern due to their toxicity. Other chemical cross-linkers may also be effective in imparting delayed release properties to softgels. For example, cross-linking of hard gelatin capsules by terephtaloyle chloride allowed for the in vitro sustained release of the asthma medication theophylline (Guyot et al., 1996). Recently, cast gelatin films were enzymatically cross-linked by microbial Ca2+-independent transglutaminase (Lim et al., 1999). This enzyme, which catalyzes the formation of -( -glutamyl)lysine cross-links in proteins (Nielsen, 1995), may offer the potential for preparing softgels with delayed release characteristics. Thiolated gelatin that has been cross-linked by disulfide bonds also holds a potential to create a delayed release system (Johnson, 1965; Barron and Tsuk, 1967; Okamoto et al., 1973). However, thiolated gelatin is not acceptable for edible applications. Finally, cross-linking gelatin using aluminum cations to prepare gelatin capsules with modified release characteristics was suggested (Shank, 1985). Overall, commercialization of delayed release softgels has been limited, mainly due to the difficulties in precisely controlling the release rate of active compounds.

Softgels having enteric release properties may be desirable for certain active compounds that degrade in the stomach (e.g., biologically active proteins and peptides); have the intestinal tract as their intended site of action; or cause gastric irritation or nausea. In general, solid pharmaceutical dosage forms can be coated with enteric polymers, which are resistant to the acidic stomach environment but disintegrate in the neutral or slightly alkaline intestinal environment. Enteric polymers include cellulose derivatives, such as hydroxypropyl methylcellulose phthalate and cellulose acetate phthalate (Sakellariou and Rowe, 1995); polyacrylates (Lehmann, 1989); and shellac (Specht et al., 1999). For example, application of enteric polymers onto hard gelatin capsules by pan or fluidized-bed coating, typically in combination with an aqueous subcoat (e.g., hydroxypropyl methylcellulose, or polyvinylpyrrolidone) to improve ad-

Softgels with Gelatin Extenders or Substitutes

427

hesion of the enteric coatings, has been discussed (Funakoshi et al., 1982; Tsuji, 1987; Murthy et al., 1988; Nielsen et al., 1999). However, applying enteric coatings to softgels, as a unit operation, is even more challenging due to their inherent flexibility and their potential susceptibility to mechanical deformation and sticking during the coating process (Felton et al., 1995). A few studies or patent disclosures focused on the enteric-coating of softgels with acrylates (Leiberich and Gabler, 1976; Davies et al., 1987; Pagay and Stetsko, 1994; Felton et al., 1995, 1996) or polyvinyl acetate phthalate (Matthews and Virgilio, 1989) have demonstrated the viability of the concept, although several parameters need to be controlled to obtain a successfully coated softgel. For example, adhesion of an enteric acrylate polymer onto the gelatin shell depended upon both the plasticizer incorporated into the coating formulation and the fill formulation of the softgel (Felton et al., 1996). Consideration should also be given to the aesthetic appearance of the coated softgels. Despite the ongoing interest in developing softgels with enteric properties, commercialization has been limited, if any. Besides the technical challenges, coating softgels with enteric polymers is a post-processing step that adds to the cost of the product. As an alternate approach to coating, enteric polymers may be directly incorporated into the gelatin-based film formulation prior to softgel manufacturing. For example, the preparation of enteric softgels by combining alginates with gelatin in the shell formulation was described in a Japanese patent (Takashi and Tetsuo, 1999). Incorporation of most enteric polymers into the gelatin shell is limited mainly by their likely incompatibility with gelatin.

Bicomponent gels comprised of gelatin and other biopolymers, such as maltodextrin (Alevisopoulos and Kasapis, 1999), starch (Ring and Stainsby, 1982; Khomutov et al., 1995), carrageenan (Michon et al., 1995; Kasapis et al., 1999), locust bean gum (Alves et al., 2000), pectin (Al-Ruqaie et al., 1997), microcrystalline cellulose (Kasapis, 1999), gellan gum (Chilvers and Morris, 1987), and whey proteins (Walkenström and Hermansson, 1997), have been studied. They can be mixed gels (independent polymer networks) or complex gels (coupled polymer networks) (Zasypkin et al., 1997). There has been interest in incorporating natural or synthetic polymers into softgel shell formulations to impart certain attributes (e.g., chewability), partially replace gelatin with a cheaper polymer, or substitute part of the gelatin with a smaller amount of a stronger gelling polymer. Enhancing the strength of the gelatin compositions used for capsule manufacturing through the addition of dialdehyde starch (0.5–5.0% w/w of gelatin) was disclosed in a patent by Helmstetter (1977). Dialdehyde starch, a polymeric aldehyde prepared by reacting starch with peri-

428 odic acid (Pfeifer et al., 1960), can cross-link proteins (Gennadios et al., 1998; Rhim et al., 1998) similar to the actions of low molecular weight aldehydes. However, it is not approved for edible applications in the U.S. Fischer et al. (1989) described softgels containing shell additives (at least 1% w/w), such as starches, starch derivatives, microcrystalline cellulose, and cellulose derivatives. In their pure states, these additives are capable of absorbing at least 10% (w/w) of their own weight in water. Such softgels were considered suitable for encapsulating fills containing water-miscible components of low volatility (e.g., glycerin and propylene glycol) at levels greater than the typical 10% (w/w of fill) limit. Softgels that had textured (frosted or satin) finishes and were resistant to sticking and to shape changes were described by Stroud (1996). These attributes were imparted to the capsules by partially substituting gelatin with starch of high amylose content (50–90%) so that the starch accounted for 3–60% of the dry shell weight. In addition, the textured surface allowed for better adhesion of subsequently applied enteric coatings (Stroud, 1996). Shell formulations that combined chemically modified gelatin, starch derivatives, and propylene glycol (plasticizer) with the intent to minimize gelatin-drug interactions over the capsule shelf life were described in a Japanese patent (Yamada and Makino, 1985). High amylose starch or starch derivatives were incorporated into the shells (at 5–20% of dry weight) of softgels having “break-off” features that were intended for delivering medicaments to an external body surface (Schurig et al., 1996). Such capsules had a drier feel, which enhanced their gripping and handling characteristics. Softgel shell formulations containing a non-hygroscopic plasticizer (e.g., maltitol, maltitol syrup, or hydrogenated starch hydrolyzate) and an elasticity-reducing extender (8–30% w/w of dry shell) selected from a large group of natural or synthetic polymers, such as modified starches and cellulosics, were described by Chiprich et al. (1997). Such softgels had an increased brittleness and could be broken with manual pressure to deliver their contents. Partial replacement of gelatin with gum acacia in softgel shell formulations also was proposed recently (Gennadios, 2001a). The incorporation of potato starch acetate into capsule shells at 3–12% of the gel mass weight was suggested for preparing chewable softgels (Hutchison et al., 1998). The starch acetate allowed greater amounts of plasticizer to be used, thus improving chewability. Furthermore, the starch acetate formed a gel that lacked substantial cross-bonding within the gelatin matrix. Upon chewing, the starch acetate and the gelatin swelled at different rates and became readily separable (Hutchison et al., 1998). Another approach to preparing chewable softgels involved the incorporation of insoluble masticatory substances (at 1 to 75% by wet weight), natural (e.g., chicle and natural rubber) or synthetic (e.g., petroleum wax), into the gel mass compositions (Ebert et al., 1984). Besides incorporating gelatin extenders into the capsule shell, applying

Softgels with Gelatin Extenders or Substitutes

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coatings onto softgels has been proposed. For example, a carnauba wax coating was used to improve shell strength and moisture resistance of softgels (Mizuno and Kayano, 1982). The use of chocolate coatings to improve palatability was discussed in a Japanese patent (Takemori et al., 1985). Coatings based on polysaccharides (e.g., gum arabic, pectin, and agar), possibly containing water-soluble nutrients (e.g., vitamin C), were applied onto softgels to improve taste, flavor, and stability (Ozawa et al., 1984). In a different type of invention, Sundararajan et al. (1996) prepared bilayer (or multilayer) softgels having one (or more) barrier layer(s) between the gelatin shell and the fill. The barrier layer prevented the migration of hydrophilic or hydrophobic ingredients from the fill into the shell, thus allowing for encapsulation of fill formulations having higher than typical contents of water and other low molecular weight substances (e.g., propylene glycol and ethanol). The barrier layer was a mixture of a polymer (e.g., ethylene acrylic acid co-polymer) and a microcrystalline wax. The capsules were manufactured using rotary die encapsulation machines that were modified so that the two capsule shell layers were formed simultaneously by co-extrusion on the casting drums using double-exit sheet dies (Sundararajan et al., 1996).

Due to its animal origin, gelatin is associated with religious and dietary concerns in certain segments of consumer populations. Over the years, such concerns have sustained interest in identifying viable, non-animal alternatives for gelatin in various edible and pharmaceutical applications, including hard and soft gelatin capsules. Injection-molded hard capsules made of potato starch (Kenyon et al., 1994) were commercialized in the early 1990s, but are not currently marketed. Also, the manufacture of gelatin-free hard capsules based on cellulose ethers has been discussed in patent disclosures over the years (Eli Lilly & Company, 1950; Sarkar, 1977; Chiba et al., 1990; Grosswald et al., 1998). In recent years, cellulose ether-based hard capsules have been commercialized but remain a niche product in terms of market share. The rotary die process for manufacturing softgels was developed to match the properties of gelatin, particularly its thermoreversibility. Adapting the same process to polysaccharides that form thermoreversible gels, such as gellan gum and carrageenan (Winston et al., 1994; Viaud, 1999; Gennadios, 2001b) has been attempted. Gellan gum is an extracellular, linear, anionic polysaccharide produced by the bacterium Pseudomonas elodea (Chilvers and Morris, 1987). Winston et al. (1994) used blends of gellan gum, -carrageenan, and mannan gums (e.g., locust bean gum) to prepare the gel mass for softgel manufacturing. The mannan gums were added to increase gel elasticity. The main limitation in processing such gels with the rotary die process is their notably higher melting/setting temperatures compared to gelatin gels. At similar

430 concentrations, -carrageenan gels have lower melting/setting temperatures than gellan gum gels (Nishinari et al., 1996). Recently, the formation of softgels from gel formulations based on either -carrageenan (Viaud, 1999) or -carrageenan (Gennadios, 2001b) was discussed. In the late 1980s, soft capsules made of glycerin-plasticized mixtures of sodium alginate and agar were commercialized for the encapsulation of dietary supplements. These spherical-shaped, seamless capsules were manufactured using the Globex process, which also is suitable for making gelatin capsules (Rakucewicz, 1988). With this process, two-layer droplets are ejected from a double orifice type nozzle. The inner layer is comprised of a lipophilic fill formulation, while the outer layer is a liquid shell formulation. The droplets are dropped into a liquid bath that hardens (solidifies) the outer layer (shell) forming the seamless capsules (Suzuki et al., 1995, 1999). For gelatin capsules, solidification is accomplished with a cooling bath (typically paraffin oil), whereas for sodium alginate-based capsules, solidification is accomplished with an aqueous solution of a calcium salt (Suzuki et al., 1995). Encapsulation of hydrophilic fills into seamless gelatin, alginate, or agar capsules also is possible with this manufacturing approach by ejecting three-layer droplets from concentrically arranged nozzles (Kikuchi and Kamaguchi, 1994). The innermost layer is a solution of the hydrophilic active substance, the outermost layer is the film-forming solution, and the middle layer is a viscous liquid scarcely miscible in water (Kikuchi and Kamaguchi, 1994). In general, the seamless capsules discussed above are limited in terms of their size and shape and have remained a niche product. An experimental technique for preparing soft capsule-like dosage forms from gliadin, the prolamin (alcohol-soluble) fraction of wheat proteins, was described by Stella et al. (1995). They prepared masses consisting of 18 to 36% (w/w) gliadin, 9 to 18% (w/w) plasticizer (sorbitol/glycerin, 2:1 w/w), and 46 to 73% (w/w) aqueous ethanol (50% v/v). The masses were cast and dried into films that were molded on stainless-steel mold pins, filled with the active compounds, and heat-sealed at 37 C and 70% RH. These gliadin-based, soft capsule-like systems showed promise for the controlled release of paracetamol (Stella et al., 1995). Recently, gelatin-free soft capsules made of polyvinyl alcohol (PVAL), a non-ingestible material, were developed, and bath oils encapsulated into such capsules have become commercially available (Anonymous, 1998). These capsules are formed by feeding preformed, dry PVAL films to a rotary die-like encapsulation machine (Brown, 1997). Instead of heat-sealing the two halves of the capsule, the capsule seal is formed by applying a solvent to the surface of at least one of the films prior to encapsulation (Brown, 1997). Besides PVAL, this method may be used with other natural or synthetic polymers, such as alginate, HPMC, and polyethylene oxide (Brown, 1997). In addition, the formation of gelatin-free soft capsules from multilayer films comprised of a sealing layer, a

Caplets/Tablets Enrobed/Coated with Gelatin

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barrier layer, and a binding layer between the sealing and the barrier layers was recently described by Brown et al. (2000). Preferably, the internal sealing layer is HPMC while the outer barrier layer is sodium alginate. The binding layer between the HPMC and the sodium alginate layers is propylene glycol alginate. Preformed, dried films are used for this process, and sealing is preferably accomplished by radio frequency signals (Brown et al., 2000). Softgels made of potato starch were commercially launched in July 2000 (VegaGelsJ, Swiss Caps, Kirchberg, Switzerland). Such capsules are formed from extruded starch-based ribbons (Anonymous, 2000). Because the extruded ribbons have a low moisture content, the capsule drying step is not necessary with this process (Anonymous, 2000). Manufacturing of flexible capsules using modified starches also was described in a recent European patent application (Laba et al., 2000). Finally, soft capsule shell compositions comprised of modified starch (preferably hydroxypropylated acid modified corn starch) and -carrageenan were described by Tanner et al. (2001) in a recent international patent application. They suggested a weight ratio of modified starch to -carrageenan in the range of 1.5:1 to 4:1.

Technologies have been developed for enrobing or coating solid cores (caplets or tablets) with gelatin. Such technologies may differ, more or less, from the softgel technology, but are briefly discussed here due to their relevance. Gelatin-coated caplets or tablets are typically referred to as gelcaps or geltabs, respectively. Within the past decade in the U.S., these solid oral dosage forms have captured a sizable share of the OTC drug market, and their use is expanding within the prescription drug and dietary supplement fields as well (largely driven by line extension opportunities). The main advantages of applying an outer gelatin layer onto caplets or tablets include, similar to softgels, ease of swallowing, taste and odor masking, protection from oxidation and light, color differentiation, printability, resistance to tampering, and increased core durability. The effect of the added gelatin layer upon the dissolution of the active is minimal due to the rapid solubilization of gelatin upon ingestion. In the early 1990s, Banner Pharmacaps Inc. (High Point, NC) developed and commercialized a method for enrobing solid cores with gelatin using modified rotary die softgel encapsulation equipment (Figure 16.5). The cores were aligned through a feeding mechanism and dispensed between two cast gelatin ribbons, which were supported on a pair of rotary dies (Sadek and Dietel, 1992, 1995). The gelatin films deformed around the cores and were heat-sealed together before the dies cut the enrobed cores from the ribbons. Both monocolored and bicolored enrobed cores can be manufactured by using gelatin ribbons of the same or differing color, respectively. Similar to softgels, the

432

Figure 16.5 Schematic diagram of the process used by Banner Pharmacaps (High Point, NC) to enrobe caplets and tablets with gelatin (Soflet gelcaps or geltabs, respectively) (courtesy of Banner Pharmacaps Inc., High Point, NC).

enrobed cores, referred to as Soflet gelcaps or Soflet geltabs, require drying (to a gelatin shell moisture content of 5 to 8%) prior to inspection, polishing, printing, and packaging. Cores used in this process are typically subcoated with a water-soluble polymer, such as HPMC or hydroxypropyl ethylcellulose, to give a weight gain of around 1%. The subcoat, which typically is clear (non-pigmented), prevents dusting and improves adhesion of the gelatin outer layer. The gelatin layer applied onto the cores is thinner (by 30 to 50%) than the gelatin shell of softgels and contains a lesser amount of plasticizer. For example, the wet gel formulation may contain 45% (w/w) gelatin and 9% (w/w) plasticizer (Sadek and Dietel, 1992, 1995). McNeil-PPC (Skillman, NJ) developed and commercialized a dipping process for coating caplets with gelatin (Berta, 1989a, b). This method involved the application of a gelatin coating onto one end of the caplet followed by application of a second gelatin coating onto the other end. The second coating was thicker than the first and could be applied using a gelatin bath having a higher viscosity than the bath used for the first coating. Because the second coating partially overlapped the first coating, which had a different color, the interlocking halves of a hard gelatin capsule were simulated (Berta, 1989a, b). The smoothness and uniformity of the outer gelatin coating was improved by using caplets that were subcoated with a mixture of a water-soluble polymer, such as HPMC, and a hydrophobic plasticizer, such as castor oil (Batista and Markley,

References

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1994; Parekh et al., 1998). This dipping process also was applied to tablets after the appropriate equipment modifications were made (Berta, 1993, 1995, 1997). Gelatin solutions also can be sprayed onto solid cores using various coating pan systems. For example, Becker (1992) spray-coated tablets and caplets (subcoated with HPMC or methylcellulose) with solutions of hydrolyzed (Bloom strength below 80 g) gelatin. The thin (25 to 100 m) gelatin coatings reduced the coefficient of friction of the cores, thus increasing their ease of swallowing. Gelatin coating compositions having improved drying characteristics and a reduced stickiness were discussed by Daher et al. (2000). Gelatin (preferably from fish) having a Bloom strength up to 200 g was combined with a surfactant (e.g., sodium stearoyl lactylate, calcium stearoyl lactylate, or glyceryl monostearate), a drying agent (e.g., sodium sulfate), and optionally a plasticizer (preferably propylene glycol monostearate). The surfactant was incorporated to reduce stickiness, while the drying agent accelerated the drying of the gelatin coating (Daher et al., 2000). These coating compositions could be applied to both subcoated and uncoated (“raw”) solid cores (1 to 5% weight gain) (Daher et al., 2000). Another approach to covering caplets with gelatin is by encapsulation into hard gelatin capsules. Barshay and Mayer (1990) described a method for adhesively bonding a caplet to the inner end surfaces of a hard gelatin capsule using an edible, water-based or organic solvent-based adhesive. An automatic device for filling caplets into hard gelatin capsules was designed by Boyd et al. (1992). In recent years, Warner-Lambert (Morris Plains, NJ) has developed and commercialized an improved process for encapsulating caplets that involves treating the two halves of the hard gelatin capsule with cold shrinking (Amey et al., 2000).

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Collagen Casings

C

ASINGS are used in sausage manufacture for forming sausage products into

a specific shape and/or for portion control. Natural casings, regenerated cellulose casings, and reconstituted or regenerated collagen casings are widely used in sausage manufacture (Wang, 1986). Collagen casings are important in the production of a variety of value-added or processed meat products. This chapter describes the fundamental manufacturing processes used in collagen casing manufacture, innovations and improvements in collagen casing manufacture, and the functionality of collagen casings for use in processed meat manufacturing. Additional emphasis is given to the use, performance, and problems associated with application of collagen casings during sausage manufacture (i.e., stuffing) and thermal processing (i.e., smoking and cooking). A glossary of terms commonly associated with the manufacture and use of collagen casings is also provided.

Casings from beef, pork, and lamb intestines stuffed with comminuted meat were the earliest forms of “natural” collagen casings. Current production of sausage products relies heavily on use of casings made from extruded, reconsti445

446 tuted collagen fibers, rather than natural casings. Sausage casings allow shaping and/or forming of comminuted meat batters in a specific shape while undergoing thermal processing to “heat-set” the meat proteins. Sausage casings, whether natural or manufactured, must be strong enough to hold the meat batter during thermal processing. Sausage casings must have the ability to shrink and stretch to accommodate contraction and expansion of meat batter during processing (stuffing, linking, cooking, chilling, packaging, and storing) (Schmidt, 1986). Tubular collagen casings are convenient edible packaging materials for processed sausage products (Hood, 1987). Fabricated collagen casings offer the advantages of uniformity, strength, and flexibility to withstand a variety of processing environments; a cleaner, more sanitary product; and greater consistency in net product weight compared to natural casings. They also provide sausage manufacturers with the ability to produce portion-controlled, value-added products from lower-value meat trimmings at high-speed automated production levels.

Hood (1987) provided an excellent historical perspective on collagen casing manufacture. The development of collagen casings arose due to the labor-intensive process in procurement and cleaning of natural gut casings. High bacterial loads, preservation requirements (salting or salting combined with refrigeration), lack of uniformity (thickness, color, and diameter), and breakage were also identified as inherent problems associated with use of natural gut casings for sausage production. The performance and supply of natural casings hindered the ability of the processed meat industry to move toward modern automated sausage manufacture at higher production capacities. Collagen casing technology initially originated through research associated with using gelatin from partially hydrolyzed collagen. The fibers of cotton bags or tubes dipped in a gelatin solution were described by Marshall (1864) and identified by Hood (1987) as the earliest individual sausage casings. Harvard and Harmony (1869) and Morris and Parker (1896) reported coating meat and other foods with gelatin. Hood (1987) suggested that the low peptide-chain molecular weight and lack of a collagen-like fibrillar structure caused inelasticity and brittleness of gelatin films, thus limiting the practicality of commercially producing gelatin-based casings. Gelatin coatings and films have limited resistance to water vapor transmission and require significant amounts of plasticizers (e.g., sorbitol and glycerin) to impart flexibility (Gennadios et al., 1997). Therefore, producing gelatin-based casings that are strong and flexible enough to withstand shirring and handling during processing (i.e., stuffing operations) would be challenging for casing manufacturers.

History of Collagen Casing Manufacture

447

The work of the German scientist Oscar Becker during the 1920s and 30s on the manufacture of synthetic collagen casings was cited (Hood, 1987) as the fundamental impetus for the manufacture of regenerated collagen casings, particularly for collagen casing technology based on the dry extrusion principle (Becker, 1936, 1938, 1939). The industrial manufacture of collagen casings is divided into two distinct processing systems, the “dry” and the “wet” manufacturing process. The “dry” manufacturing process (“dry spinning technology”) was developed in Europe during the 1930s, followed by the “wet” manufacturing process. The dry industrial manufacturing process includes the following processing steps (Hood, 1987): (1) Dehairing of cattle hides (2) Machine splitting of the hide to separate the outer layer (leather layer) from the corium (inner collagenous) layer (3) Alkaline treatment of the corium layer of bovine hides (primary collagen source, >90% collagen on a dry basis), followed by acidification to pH 3.0, which swells the collagen fibers (4) Shredding of the acid-swollen corium to preserve the fibrous structure of collagen (5) Forming of a viscous “collagen dough” (>12% solids) from the acid-swollen collagen fibers by mixing (6) Adding of plasticizing and cross-linking agents (7) High-pressure extrusion of the collagen dough to form tubular casings (8) Drying, conditioning, neutralizing, and/or providing additional cross-linking of the collagen tubes (9) Packaging of varying lengths of shirred, pleated, or folded casings. This dry extrusion process was not adopted by American casing manufacturers due to: (1) restricted knowledge of the process due to World War II; (2) unfamiliarity with the pumps and extruders necessary to extrude the collagen dough; (3) the inedible nature of the final product; and (4) the tough fibrous texture that was not acceptable by the sausage industry at the time (Hood, 1987). The second method described by Hood (1987) to manufacture collagen casings is termed the “wet” process (“wet spinning technology”). This was the dominant process for collagen casing manufacture during the early 1960s. For more detailed technical information on the early development of this process, the reader is referred to several patent disclosures (Hochstadt and Lieberman,

448 1960; Lieberman 1964a, 1964b, 1965; McKnight, 1964a, 1964b; Fagan, 1970). The process consists of using the hide corium from acid- or alkaline-dehaired cattle hides, which are decalcified, ground into small pieces, and then mixed with acid to produce a viscous suspension (4–5% solids) through high shear homogenization. Cellulose and carboxymethylcellulose may be added to improve mechanical properties of the casing. The acidified collagen slurry is then extruded to form a woven fiber structure, passes through a coagulation bath of brine, and is formed into tubular casing shape. The high salt concentration and acid neutralization shrinks the collagen fibers to form a strong casing. The casings are then washed to remove the salt and treated with plasticizing (glycerol or sorbitol) and cross-linking (gluteraldehyde) agents to improve casing strength and pliability. The plasticized collagen casings are then collapsed like an accordion (“shirred”) so that they can fit over various-sized sausage stuffing horns. The collagen casings are then dried under special temperature and humidity conditions to 13–18% moisture content before they are sealed in plastic bags and packed in boxes (Hood, 1987). The shelf life of these products depends upon maintaining favorable (cool and dry) storage conditions. Most collagen casings are ready to use “out of the box” (Ockerman and Hansen, 1999). A third process was referred to by Maser (1999) as “hybrid technology” that combines elements of the dry and wet processing technologies. Citing Fagan (1970), Maser (1999) described the process as follows. A collagen content of no more than 5% (similar to wet processing) is used. The collagen tube formed after extrusion is inflated with an ammonia-air mixture, preventing the collapse of the tube, and reacts with the acid present in the extrusion gel, resulting in coagulation of collagen fibrils. This results in increased tensile strength of the collagen tubing. The casing tube is horizontally extruded onto an endless conveyor belt (similar to dry extrusion processing). The conveyor belt passes the collagen tubing through an ammonia atmosphere that coagulates the collagen fibers. The casing tube is then flattened, washed, and plasticized in a series of baths (similar to wet extrusion processing), re-inflated, dried in its inflated state, shirred, and dried to its final moisture content. Maser (1999) further stated that the wet or hybrid manufacturing processes manufacture only small-diameter straight collagen casings. Neither of these processes is suitable for producing curved collagen casings. For production of natural curved casings and large diameter straight collagen casings ( 32 mm), the classical dry extrusion process is better suited.

The dry and wet extrusion processes both manufacture collagen casings suitable for sausage manufacture (Figure 17.1). Hood (1987) described differences between the two processes in the following summary. After

Figure 17.1 Flow chart of wet and dry technologies for manufacturing collagen sausage casings. (Adapted from Hood, 1987.)

449

450 decalcification, the dry processed cattle hides are swollen (pH 3) and then ground, whereas wet processed cattle hides are ground in the unswollen state (pH 4–5) and are mixed with water and acid (HCl) to swell. This results in a wet processed collagen containing 4–5% solids while dry processed collagen has about 12% solids. The dry processed collagen is mixed with water (resulting in 10% solids), and the collagen from both processes undergoes either a fluid two-stage (wet) or a shearing plate (dry) homogenization step. The resulting dry processed collagen dough consists of longer and larger collagen fibers compared to the wet processed collagen dough. Afterward, both processes require metering or pumping the collagen dough for extrusion. The primary purpose of extrusion is to “realign” the collagen dough fibers as the fibers exit the extruding horn or orifice. Due to their fibrous structure, collagen fibers tend to align parallel to the longitudinal axis of the extruding horn. To produce a woven, cross-hatched fibrous collagen casing structure, the collagen fibers are mixed prior to exiting the extruder horn or orifice. One difference between the two processes during the extrusion step is that no water is used during extrusion of collagen dough in the dry process. The wet process utilizes a disc extruder that mixes the collagen dough against the sides of the extruder case, whereas a counter-rotating nozzle extruder is used for the dry process. Twin counter-rotating serrated surfaces are used to mix and align the collagen dough fibers (Hood, 1987). The extruder rotating disc design and action impacts the collagen fiber alignment, thereby affecting the final strength and flexibility of manufactured collagen casings. Subsequent steps for wet processing include drying, conditioning, neutralizing, and/or providing additional cross-linking of the collagen tubes, and packaging of varying lengths of shirred, pleated, or folded casings. Following extrusion, dry processed collagen casings undergo the mentioned steps of brine coagulation, formation into tubular shape, washing, plasticizing, cross-linking, and finally shirring. The shorter collagen fibers produced by wet extrusion contribute to faster line speeds and larger production volumes compared to the slower (thicker, longer collagen fibers) and more expensive (high-pressure pumps and plumbing) dry extrusion technology (Hood, 1987).

Deacon and Kindleysides (1973) identified a method of co-extruding collagen with added methylcellulose to reduce the tendency of the extruded collagen coating of a collagen-cased sausage to disintegrate when the sausage was cooked in a humid environment. They reduced disintegration of the collagen casing by incorporating a thermogelling, non-ionic cellulose ether, such as methylcellulose. They further described the process of simultaneously extruding the collagen casing with the sausage emulsion. Smits (1985) also described a continuous process where collagen was co-extruded around the sausage bat-

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ter. This co-extrusion technology is an alternative to the established method of encasing sausage into preformed collagen casings and is normally associated with large production volumes. The process of simultaneous co-extrusion of the sausage batter and casing requires the collagen dough or suspension to be prepared in situ and pumped to an extrusion nozzle (Figure 17.2). The extrusion nozzle is comprised of a central tube for the sausage batter to flow through. The tube is enveloped by inner and outer cones through which the collagen dough is extruded to form a thin layer of oriented collagen fibers onto the surface of the sausage. Hood (1987) further described this process. The meat batter or emulsion is pumped through the inner orifice of a nozzle, and the collagen suspension is applied to the outside surface of the meat batter as it emerges through counter-rotating concentric cones that align the collagen fibers to form a woven structure. The encased sausage passes through a brine bath to dehydrate and “set” the collagen casing. The continuous sausage rope is then crimped into links of a selected size. Additional drying and smoking further aid interactions between extruded collagen

Figure 17.2 Diagram of nozzle used for co-extrusion of collagen onto a sausage batter. (Adapted from Smits, 1985).

452 fibers and meat proteins. Co-extruded sausage casings are more tender than preformed or reconstituted collagen casings, thus enhancing the sensory properties of the finished sausage product after cooking (Waldman, 1985). A recently developed extrusion attachment for sausage stuffing machines was developed (Kobussen et al., 1999) consisting of three connecting tubes forming three passageways: the inner tube; the space between the inner tube and the intermediate tube; and the space between the intermediate tube and the outer tube forming the third passageway. The meat emulsion passes through the inner tube under high pressure. Emulsion generated from the second passageway is thinly coated on the sausage strand extruded from the inner tube. A thin layer of collagen is extruded from the third passageway and deposited on the outside surface of the meat layer extruded through the second passageway. Morgan et al. (1998) developed a process for producing a linked, co-extruded, edible sausage by co-extruding a uniform layer of an acid collagen gel around extruded sausage. The collagen gel was then chemically coagulated with liquid smoke and a dehydrating agent (alkali or salt solution) to produce a collagen casing with sufficient mechanical strength to allow formation of sausage links via twist-linking.

Several patents have focused on improving the manufacture of collagen casings. Burke (1976) described a process for preparing collagen casings from limed bovine hide collagen. The limed hide collagen was soaked in a dilute edible acid (pH < 5.5) to reduce the pH of the hide center to 4.0 to 5.5. The neutralized hide was then washed, ground, and formed into collagen slurry that was then processed into edible collagen casings. Ziolko (1977, 1979) described a method of manufacturing tubular collagen casings by extruding collagen gel into two sets of collagen gel “ropes,” each individual set consisting of several ropes. The collagen fibers were oriented in the proper directions by development of an apparatus that consisted of an inner and outer extruder with orifices that extruded the collagen gel ropes with the surface fibers oriented in the direction of extrusion. The ropes in one set were directed outward in a radial and helical direction to form the first tubular layer. The second rope set was directed inward in a radial and helical direction opposite from the first layer to form a second tubular layer that was concentric to the first. A hardening step followed this process to unite the two layers. Wilson and Burke (1977) developed a process that deposited a humidifying fluid made of water and a mixture (0.8 to 3%) of partial fatty acid esters of glycerin and sorbitol to the inside wall of unshirred collagen casings. Application of this fluid enhanced the shirring process for preparation of humidified, shirred artificial collagen sausage casings.

Collagen Casing Properties

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Collagen casings must possess a variety of product attributes to aid in manufacturing of various sausage products with numerous types of sausage manufacturing equipment. Casing strength is important for withstanding the rigors of high-speed filling and linking operations. The strength of collagen casings is affected by solids content (collagen fiber content), drying conditions, and addition of cross-linking agents. Subjecting collagen casings to a solution of a collagen setting salt (sucrose or polyethylene glycol and sodium chloride) and then drying the casings to a final moisture content of 15–40% increased casing strength (Kidney, 1970). Treatment with a gluteraldehyde solution produced a strong, coherent film of collagen fibrils with great longitudinal and transverse strength whether the casing was dry or re-wet (Rose, 1968). Treating collagen casings with the sodium salt of carboxymethylcellulose to improve resistance of sausages stuffed in collagen casings to bursting (“burst strength”) during frying was described by McKnight (1964a). Exposing edible tubular collagen casing to ultraviolet irradiation (180–420 nm) increased casing strength (Miller and Marder, 1998). Enzymatic treatments were used to produce collagen casings with a more uniform diameter, greater wall thickness, and increased tenderness (Tsuzuki and Lieberman, 1972). A proteolytic enzyme obtained from Aspergillus niger var. macroporous was added to the collagen dough prior to extrusion to partially solubilize collagen. Boni (1988) prepared collagen casings by incorporating a lower alkyl diol. Such casings had improved mechanical properties at low temperatures due to reduced internal hydrogen bonding and increased molecular spacing. Miller (1983) described an enhancement of the wet process of collagen casing manufacture. Prior to extruding the collagen dough, a proteolytic enzyme (e.g., papain, bromelain, ficin, fungal protease, trypsin, chymotrypsin, or pepsin) was injected into the collagen. Access to water was restricted, thereby reducing enzymatic activity (immobilizing the enzyme). Upon stuffing the collagen casing with sausage and subsequent thermal processing, the enzyme was reactivated and continued to break down collagen, thereby tenderizing the casing.

Collagen casings sometimes contain food-grade colorants approved by the Food and Drug Administration (FDA). Casings with several colors are marketed including blush, brown, red, and smoke. Sausage manufacturers use colored collagen casings to enhance external color and appeal of various sausage products. Blush (yellow tinged) casings enhance the “lean appearance” of fresh sausage products. Talty (1969) described one of the first processes to counteract the parchment-like appearance observed when pork sausage links were

454 stuffed in collagen casings. Incorporation of small amounts (e.g., 5%) of edible vegetable or animal oil produced strong and translucent casings. Sausage stuffed in such casings showed the sausage meat in an attractive manner similar to natural casings. Crooks (1985) described a process that comprised treating buffered hide corium collagen with a solution containing glycerin (2–5%), hydrogen peroxide (0.5–3%), and ferrous iron (25–75 ppm). Collagen casings produced with this process exhibited improved clarity or translucency. A collagen casing containing 50–80% by weight of bovine collagen and 20–50% by weight of an acidified brine extracted pork skin had improved clarity (Fu Lu and Hood, 1986). Brown-colored collagen casings are used primarily for precooked sausages, such as pork breakfast link sausages. Steam cooking of these sausages results in a gray rather than a caramelized or brown exterior color. Brown-colored collagen casings enhance the “browning effect” of the exterior surface of these sausages, resulting in a cooked product with more appealing color. Bright red-colored collagen casings are used for hot, spicy sausages, such as the traditional Southern-style “hot link.” Smoke-colored collagen casings can be used to replace or enhance the natural or liquid smoke process. If used with a sausage product that undergoes either natural or liquid smoke application, the smoke-colored casing provides a richer, more uniform smoke-colored product surface. Coloring agents such as annatto, FD&C Red #3 dye, and caramel are normally used singularly or in combination to impart the desired collagen casing color (Devro-Teepak, 1996). A process for impregnating collagen casings with a caramel solution was described by Rahman and Coleman (1977). After the tubular collagen casing was impregnated with the caramel solution, the caramel was cross-linked and insolubilized in the casing to create a permanently colored casing. Stribling (1997) developed a collagen sausage casing by incorporating encapsulated liquid smoke into the casing that releases the smoke component during curing or smoking and prior to consumption. Another method was described that incorporated encapsulated liquid smoke directly into the collagen slurry, then extruded the slurry onto the surface of the sausage. Baldwin (1988) used a chocolate dye to develop a brown collagen casing. The chocolate was added during extrusion of the collagen gel or was applied to the extruded tubular casing prior to drying.

The following information is provided as a general guide to assist in determining the proper set-up of sausage stuffing/filling machines (Devro-Teepak, 1996). The major components and steps in sausage stuffing/filling are ad-

Techniques and Procedures for Stuffing Collagen Casings

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dressed with general processing guidelines, and troubleshooting tips are provided to aid in processing efficiency. The filling horn should be kept straight, clean, and free from damage, particularly at the open end. It is generally best to check if the horn is straight and running true with the filling machine empty but with the horn rotating. A straight horn will rotate within its own diameter. A bent horn tip will revolve in a wider arc. Holding a pencil close to the horn tip as it rotates can identify a bent horn. If the stuffing horn is bent, the pencil will mark it. The stuffing machine should be stopped and the horn straightened by pressing along the length of the horn toward the tip on the side identified by the pencil mark. This procedure should be repeated until the pencil can make a mark around the full circumference of the horn tip. A straight horn will rotate in the center of the chuck/brake-ring assembly on the automatic linking attachment. If not, then the manufacturer’s instructions should be followed to bring it in line with the straight horn. It is important to examine the linking attachment (chuck/brake-ring) for excess wear or damage. Problems that may arise during the sausage stuffing/filling operation are usually associated with casing breakage and can be traced to one or more causes as listed in Table 17.1. The availability of horn diameters can vary with respect to machine type. In general, the casing strand (slug) must be able to rotate freely on the largest horn available. Use of a horn diameter that is too small for the casing, or if the casing does not rotate freely on the horn (i.e., horn is too large), can cause casing breakage during filling. The table on which the stuffed sausage drops should be equal to at least one and a half sausage lengths below the filling horn (maximum drop height of 30 cm). If the drop is too small, the sausage will be supported by

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the table, which may result in inadequate “pull,” or resistance, to allow the rest of the sausage to twist. If the drop is too large, the weight of the “sausage chain” may cause sausage breakage, or the sausage casing may pass through the chuck/break-ring assembly too quickly, resulting in underfilled sausage. Table 17.2 provides general guidelines for filling horn diameter and corresponding collagen casing diameters. If a roller assembly is used at the outlet of the chuck/break-ring, the roller(s) must be attached so that the roller gently directs the filled sausage downward with minimum contact between the roller(s) and the filled sausage. If the resistance is too great, the casing will burst. A wide gap between the roller(s) will result in poor linking. Collagen casings require minimal preparation prior to use. They should be kept as dry as possible prior to stuffing. When operating semi-automatic filling machines, operators should keep their hands as dry as possible during loading. The filling horn should also be kept dry on the outside and wiped clean of any sausage batter or emulsion. Prior to loading the strand onto the horn, any emulsion that may have extruded from the top of the horn between each filling cycle should be removed. The conical end of the collagen strand (slug) should be fed onto the horn first, and care should be taken to avoid breaking the shirred strand. After threading the end of the casing through the chuck, the casing tension should be checked to ensure that it is not too tight by pulling a small length of casing through the chuck/brake-ring assembly. Then, the filling machine can be started. If the casing breaks during stuffing/filling, the machine should be stopped, the “fresh end” of dry casing should be rethreaded through the chuck/brake-ring assembly, and the machine should be restarted. The filled caliber (filled sausage casing diameter) should be checked at this stage and casing tension either loosened or tightened if the sausage is either overfilled or underfilled, respectively. The sausage should be considered as a “cylinder” of fixed dimensions with a

Techniques/Procedures for Smoking/Cooking Collagen Casings

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desired weight and length of each sausage portion. The desired weight and length of the sausage determines the sausage caliber. The density of sausage emulsions can vary due to recipe formulation and method of preparation. For dried or semi-dried sausage, where cold, coarse, chopped/minced meat mixes are used, it is normal to get filled calibers from 0.5–1.5 mm below the nominal caliber of the casing used. This may be due to absence of “free water” within the sausage mix that aids in casing expansion at filling coupled with a drop in filling pressure due to a more viscous meat mixture. The hanging configuration of sausage that is to be smoked and/or cooked is based upon sausage weight and length, size of the smokestick used, and the available hanging space on the smoketruck or trolley used.

The precise recipe formulations and smoke/cook programs can only be determined by the individual sausage processor who must consider the type of sausage produced and the available smoking/cooking equipment. The following information is summarized from product information provided by Alkar (1996) and Devro-Teepak (1996) and should be used as a general guide. The thermal processing programs (Tables 17.3, 17.4, 17.5, and 17.6) can be used as a starting point for first-time use. It is common that modifications must be made to account for the variability in smokehouse efficiency, sausage formulation, and the processor’s preference for final product color and internal temperature. The basic stages of the smoke/cook process are conditioning (precook), drying, smoking, color set, cooking, and finish (Alkar, 1996). Conditioning creates uniform surface conditions for the product load prior to smoking and cooking. This can be done simply by showering or by using warm (50 C) temperature and high relative humidity (95–100%) at this stage (Alkar, 1996). If the conditioning stage is too hot and dry, the sausage emulsion may not coagulate properly, and the collagen casing may be tough. High temperature can also contribute to poor meat-to-casing binding (meat adhesion). Poor meat adhesion may be attributed to poor emulsion stability and/or excessive added water (Devro-Teepak, 1996). The drying stage uniformly dries the product surface so that the product can uniformly absorb an application of smoke to produce a desirable smoked product color (Alkar, 1996). A longer drying stage generally lightens the smoke color on the product surface. Conversely, shorter drying times tend to produce a darker colored product surface. The smoking of frankfurter-type sausages filled in collagen casings is optimized in the presence of heat and humidity (Devro-Teepak, 1996). If sausages are kept slightly moist during this state, smoke uptake will be rapid, resulting in a uniform, desirable final product color. If the smoking stage is too long and too

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460

Glossary of Terms (Devro-Teepak, 1996)

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dry, poor product color will result. With good temperature and humidity control, collagen casing sausage products should develop a desirable smoked color in 30 to 40 min. The smoking procedure should not be carried out under conditions that are too dry or too warm (Alkar, 1996; Devro-Teepak, 1996). The purpose of the color setting step is to fix the smoke color before proceeding to the cooking stage. Hot, dry smokehouse or oven conditions promote development and stabilization of the desired smoke color (Alkar, 1996). The cooking stage ranges from 70 to 85 C dry bulb temperature and from 55 to 65 C wet bulb temperature. During the finishing step, the product is cooked to its final target core temperature. After cooking, it is best to shower the product with cold water to quickly reduce the temperature to below 30 C (Alkar, 1996; Devro-Teepak, 1996). Table 17.7 lists typical problems that occur during thermal processing (smoke/cook cycles) of sausages stuffed in collagen casings (Devro-Teepak, 1996).

Collagen casings are regenerated from the corium layer of cattle hides. Primarily either a wet (North American) or dry (European) extrusion process accomplishes collagen casing manufacture. Co-extrusion technology has allowed the coating of a thin layer of collagen as the sausage batter is extruded, resulting in a faster, higher-volume, continuous processing system. Innovations in collagen casing manufacture have improved casing strength and flexibility allowing improved processing efficiency of numerous sausage products manufactured by a variety of processing equipment within a wide range of processing environments. General techniques, guidelines, and troubleshooting information were provided to assist in the manufacture of products made with collagen casings.

The author acknowledges assistance in preparation of this book chapter from Mr. K. Stribling, Devro-Teepak Inc., Lisle, IL, USA; Dr. F. Maser, Naturin GmbH & Co., Weinheim, Germany; and Mr. R. Hanson, Alkar, Lodi, WI, USA.

• Air Pockets: Small voids in the meat mix in encased links. • Bite: The initial resistance of penetration by the teeth into the casing. • Buckling (Fissures): The wrinkled condition of collagen casing after processing. • Caddy Stick Length: The unrestrained shirred strand length in inches.

462 • Caliber: Diameter of sausage casing. • Casing Shirring (Oils): Oil is used during the shirring operation to reduce friction during sausage manufacture and to enhance the clarity of the casing. • Chew: The type of mouth-feel experienced during mastication of cooked sausage casings. Casings should integrate with the meat during mastication. The casing should not be rubbery, nor leave a coating or residue in the mouth after the meat is swallowed. • Clarity: Ability of casing that allows lean and fat color and particle definition of the product to be seen. • Closed Ends: A type of casing that has the lead end of the casing strand secured for use to run on automatic filling/stuffing machinery. • Collagen Gel: Suspension of acid-swollen collagen fibers. • Crocking: Transfer of dye from the casing surface to work surfaces, machinery, and hands of workers. • Dye Acceptance: The ability of the casing to accept an application of coloring during casing manufacture. • Dye Retention: The ability of the stuffed and processed product to retain the dyed surface color uniformly for a specified period of time. • Extrusion: A condition that occurs when encased sausage meat extends from the ends of the casing after cooking due to meat expansion, casing shrinkage, or both. • Fatting Out (Greasing Out): Collection of fat or oil between the surface of the product and the inside surface of the casing after cooking and/or smoking. • Fishmouthing: A characteristic found in processed meat products whereby the casing in the twist opens during cooking in the end use environment. A fishmouth is objectionable if the opening of the fishmouth exceeds 25% of the link diameter. • Frill: A pigtail that untwists itself shortly after linking. • Horn Fit: A term used to describe how well a strand slides onto and off of a specified horn (nozzle, stuffing tube). The inside diameter of a strand must be large enough to enable the strand to fit onto the outer diameter of a specified horn. When placed onto the stuffing horn, the strand should be loose enough to deshir and/or spin on the horn freely. • Hydrothermal Stability: Ability of a collagen casing to withstand cooking. • Machinability: Ability of a casing to withstand the rigors of stuffing, linking, and reeling (if applicable) within specified breakage limits. • Meat Adhesion: The formation of a protein bond between the meat emulsion and the casing wall during processing. • Nominal Deshirred Length: The minimum length of shirred casing in each strand.

Glossary of Terms (Devro-Teepak, 1996)

463

• Pencil Pointing/Tear Dropping: The effect created when the casing relaxes after stuffing and during cooking. The end of the link becomes pointed rather than maintaining a well-rounded (square shoulder) appearance. • Pigtail: The end of the casing that remains twisted shut after linking. • Porosity: Ability of a casing to allow steam and fat to render through the casing wall during consumer cooking, or to allow moisture to migrate through the casing wall during processing, or to dry at a proper rate while minimizing grease or oil migration from the sausage product. • Portion Control: The ability of a casing to be manipulated within a narrow range resulting in replication of the required weight of meat product per given length. • Reeler: A device used in the production of beefsticks. Stuffed (but not linked) casing is wrapped around bars on a rapidly rotating frame. The bars and casing are then transferred to smoke “trees” for transport into further processing areas. • Sheen: The appearance of a stuffed fresh sausage link created by the shirring oil used. • Shrinkage: This occurs when the casing shrinks longitudinally, or radially, leaving the meat exposed at one or both ends after cooking. • Size Uniformity: Ability of a stuffed and linked casing to maintain a consistent diameter throughout the length of the strand within the specifications of a given code. This consistency must be maintained from strand to strand. • Slip: (1) Table/Smoke Stick Slip: The ability of a stuffed strand to slide across a dry metal table and on a dry metal smoke stick with a competitive coefficient of friction. (2) Equipment/Machine Slip: The ability for a casing to maintain size control throughout stuffing without causing the following links to back-up. Also described as the ability of the links to slide properly on automatic collating and packaging machinery. (3) Packaging Slip: The ability of links to slide against each other and against the packaging materials during packing. • Smoke Acceptance: Ability of the casing to accept and retain smoke color and flavor during smoke application and throughout the shelf life of a product. • Snap: Bending of a sausage link until it breaks. After completion of cooking, this is evaluated by bending the links until they snap. • Splits: Casing splits during cooking. • Stuffing Equipment: Machinery used to fill sausage casings with meat. • Wet-Out (Linkability): Ability of casing to assume a state of non-elastic

464 pliability (during a specified time measured in seconds) prior to entering the linker. • Wet Strength: Ability of a collagen casing when wet to withstand the rigors of machining and subsequent handling without breaking or relaxing sufficiently to cause breakage or tapering of the stuffed link.

Alkar. 1996. Guidelines for the Development of Cooking and Smoking Processes for Smoked Meats. ed., R.E. Hanson, Lodi, WI, USA. Alkar. 1997. Cooking and Smoking Schedules for Alkar Batch Ovens. ed., R.E. Hanson, Lodi, WI, USA. Baldwin, W.L. March 1, 1988. U.S. patent 4,728,523. Becker, O.W. October 6, 1936. U.S. patent 2,056,595. Becker, O.W. April 26, 1938. U.S. patent 2,115,607. Becker, O.W. June 13, 1939. U.S. patent 2,161,908. Boni, K.A. December 27, 1988. U.S. patent 4,794,006. Burke, N.I. January 13, 1976. U.S. patent 3,932,677. Crooks, H.B. July 2, 1985. U.S. patent 4,526,580. Deacon, M.J. and Kindleysides, L. 1973. U.S. patent 3,767,821. In Sausage Casing Technology, Food Technology, Review No. 14, ed., E. Karmas, Park Ridge, NJ: Noyes Data Corp., pp. 88–89. Devro-Teepak, Inc. 1996. Product Information—Collagen. Internal document, Lisle, IL, USA. Fagan, P.V. October 20, 1970. U.S. patent 3,535,125. Fu Lu, M.Y. and Hood, L.L. October 7, 1986. U.S. patent 4,615,889. Gennadios, A., Hanna, M.A. and Kurth, L.B. 1997. “Application of Edible Coatings on Meats, Poultry and Seafoods: A Review,” Lebensm.-Wiss. U.-Technol., 30:337–350. Harvard, C. and Harmony, M.X. June 8, 1869. U.S. patent 90,944. Hochstadt, H.R. and Lieberman, E.R. January 5, 1960. U.S. patent 2,920,000. Hood, L.L. 1987. “Collagen in Sausage Casings” in Advances in Meat Research, Vol. 4, eds., A.M. Pearson, T.R. Dutson and A.J. Bailey, New York: Van Nostrand Reinhold, pp. 109–129. Kidney, A.J. 1970. U.S. patent 3,505,084. In Sausage Casing Technology, Food Technology Review No. 14, ed., E. Karmas, Park Ridge, NJ: Noyes Data Corp., pp. 122–123. Kobussen, J., Kobussen, J., Kobussen, M. and Basile II, V.L. March 30, 1999. U.S. patent 5,888,131. Lieberman, E.R. March 3, 1964a. U.S. patent 3,123,482. Lieberman, E.R. March 3, 1964b. U.S. patent 3,123,653. Lieberman, E.R. December 7, 1965. U.S. patent 3,221,372. Marshall, W.G. February 23, 1864. U.S. patent 41,712. Maser, F. 1999. Personal written communications. Naturin GmbH & Co., Weinheim, Germany. McKnight, J.T. March 3, 1964a. U.S. patent 3,123,483. McKnight, J.T. October 6, 1964b. U.S. patent 3,151,990. Miller, A.T. June 14, 1983. U.S. patent 4,388,331. Miller, A.T. and Marder, R.B. October 13, 1998. U.S. Patent 5,820,812. Morgan, T.F., Frame, G. and Kobussen, P.J. August 18, 1998. U.S. patent 5,795,605. Morris, A. and Parker, J.A. March 17, 1896. U.S. patent 556,471. Ockerman, H.W. and Hansen, C.L. 1999. Animal By-Product Processing & Utilization, Lancaster, PA: Technomic Publishing Company, Inc.

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Rahman, M. and Coleman, H.R. July 26, 1977. U.S. patent 4,038,438. Rose, H.J. 1968. U.S. patent 3,413,130. In Sausage Casing Technology, Food Technology Review No. 14, ed., E. Karmas, Park Ridge, NJ: Noyes Data Corp., pp. 102–103. Schmidt, G.R. 1986. “Processing and Fabrication” in Muscle as Food, ed., P.J. Bechtel, London: Academic Press, Inc., pp. 201–238. Smits, J.W. 1985. “The Sausage Coextrusion Process” in Trends in Modern Meat Technology, eds., B. Krol, P.S. van Roon and J.H. Houben, Wageningen, The Netherlands: Center for Agricultural Publishing and Documentation, pp. 60–62. Stribling, K.V. February 4, 1997. U.S. patent 5,599,570. Talty, R.D. 1969. U.S. patent 3,446,633. In Sausage Casing Technology, Food Technology Review No. 14, ed., E. Karmas, Park Ridge, NJ: Noyes Data Corp., p. 117. Tsuzuki, T. and Lieberman, E.R. August 11, 1972. U.S. patent 3,681,093. Waldman, R.C. 1985. “Co-extrusion-High Tech Innovation,” The National Provisioner, January 12 issue:13–16. Wang, P-Y. 1986. “Meat Processing I” in Encyclopedia of Food Engineering, 2nd edition, eds., C.W. Hall, A.W. Frall and A.L. Rippen, Westport, CT: AVI Publishing Co., Inc., pp. 545–550. Wilson, J.R. and Burke, N.I. December 13, 1977. U.S. patent 4,062,980. Ziolko, F.J. November 29, 1977. U.S. patent 4,060,361. Ziolko, F.J. February 6, 1979. U.S. patent 4,138,503.

Application of Edible Coatings on Muscle Foods

HROUGHOUT history, packaging materials have been specifically developed to prevent the deterioration of foods resulting from exposure to microbes, air, moisture, or pH changes associated with the food or the surrounding atmosphere. Both flexible and rigid packaging materials, alone or in combination with other preservation methods, have been developed to offer barriers, inactivation, and containment properties required for successful food packaging. Examples of flexible packaging associated with foods include controlled atmosphere, vacuum, modified atmosphere, and edible packaging (Cutter, 2001). During the 1950s, rubber and adhesive components were available, heat shrinkable polyvinyl chloride (PVC) was introduced, and nylon was integrated into packaging films. In the 1960s and 1970s, most of the major packaging accomplishments involved the development of jars, bottles, tubs, and films plastic derivatives (Cutter, 2001). Based on properties such as sealability, tensile strength, rigidity, temperature, and water and vapor barrier ability, the use of plastic products for food storage appears limitless. The use of vacuum packaging (VP) or modified atmosphere packaging (MAP) to improve stability and safety of fabricated or further processed muscle foods is well documented (Genigeorgis, 1985; Hintlian and Hotchkiss, 1986; Farber, 1991; Ooraikul and Stiles, 1991; Labuza et al., 1992; Church and Parsons, 1995; Garcia et al.,

T

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468 1995). In fact, more than 90% of meat and poultry currently produced in the US is marketed under VP or MAP (personal communication, American Meat Institute, 1997). The importance of VP or MAP is increasing as meat exportation becomes more prevalent (Siragusa et al., 1999). However, there are concerns that increased use of plastic-based packaging will be accompanied by significant packaging waste disposal problems in the future. Just as VP and MAP offer certain desirable properties, gels/films/coatings made from food-grade compounds are being formulated and used in the food industry and under research conditions to prevent moisture loss, drip, and rancidity due to lipid oxidation or to improve flavor attributes, handling properties, color retention, and microbial stability of meats, including pork, lamb, and beef (Gennadios et al., 1997). Other major benefits of using edible films are that they can be consumed with the food, provide additional nutrients, enhance organoleptic properties, or deliver quality-enhancing antimicrobials (Guilbert et al., 1996). Edible films made with natural biopolymers such as proteins, lipids, and polysaccharides are more suitable than plastic or synthetic polymers in that they are biodegradable, can be made from renewable raw materials, and ultimately reduce packaging waste (Guilbert et al., 1996). With consumers desiring more environmentally friendly packaging and leaning toward more natural products, edible films will continue to play an important role in the food industry by improving the quality of many products, including meats (Hernandez, 1994; Ben and Kurth, 1995).

Application of edible films to meat products may be accomplished by foaming, dipping, spraying, casting, brushing, individual wrapping, or rolling (Donhowe and Fennema, 1994; Grant and Burns, 1994). Emulsions for coatings may be applied with a foam applicator. In doing so, a foaming agent is added to the coating or compressed air is blown into the applicator tank with the agitated foam applied to the meat as it moves over rollers. The foam is then distributed over the product by flaps or brushes (Grant and Burns, 1994). When a product requires several applications of a coating to obtain uniformity on an irregular surface, or when cost is a factor, submerging the product into a tank of the emulsion may work best. After dipping and draining of excess coating, the film is allowed to set or solidify on the product (Donhowe and Fennema, 1994). To speed up the process, part of the process may entail the use of a drier to remove excess water or to dry the product under ambient conditions (Grant and Burns, 1994). When a thinner, more uniform film is required, spraying may be suitable for film application. In the U.S., early coating procedures involved spraying materials onto rollers or brushes, then allowing the tumbling action of the meat to evenly spread the coating (Grant and Burns, 1994). Spraying of meat products also is desirable when application is only needed on

Properties and Types of Edible Films

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one side or when a dual application must be used for cross-linking, as is practiced with alginate coatings (Donhowe and Fennema, 1994). Heat may be used after spraying to hasten drying and to improve uniform distribution of the film solution on the surfaces (Grant and Burns, 1994). Casting is a technique in which film-forming solutions are poured onto a smooth, flat, and level surface and allowed to dry, usually within a confined space. Casting produces free-standing films that exhibit a specified thickness, smoothness, and flatness (Donhowe and Fennema, 1994). Depending upon firmness and flexibility, cast films can then be used to wrap surfaces. Films also can be applied by brushing and/or rolling the compounds directly onto food surfaces and allowing them to set or dry in place.

For successful application to meat products, edible films must meet a range of functional requirements such as moisture barrier ability, water or lipid solubility, color, appearance, mechanical and rheological characteristics, and non-toxicity (Guilbert et al., 1996). These properties are influenced by addition of plasticizers, cross-linking agents, antimicrobials, antioxidants, or texture agents. Additionally, the effects of cohesion on flexibility, permeability, gas and solute migration, or porosity may be influenced by these properties (Guilbert et al., 1996). Films also should be resistant to breakage and abrasion.

Application of lipid coatings to meats to prevent shrinkage was first documented in the sixteenth century when animal fats were used to coat meat cuts (Kester and Fennema, 1986; Baker et al., 1994). This enrobing of meats with fat was called larding (Kamper and Fennema, 1984). Animal fats are still used today to coat a variety of foods, including frozen poultry (Hernandez, 1994). Waxes and oils also may be added to films to impart flexibility and to improve coating performance (Baldwin et al., 1995). Lipid-based release agents or lubricants are occasionally added to films to prevent coated foods from sticking (Baldwin et al., 1995). Lipid compounds include neutral lipids of glycerides, which are esters of glycerol and fatty acids, and the waxes, which are esters of long-chain monohydric alcohols and fatty acids (Hernandez, 1994). Lipid and resin coatings prepared from waxes (e.g., carnauba, beeswax, and paraffin), oils (vegetable, animal, and mineral), and surfactants are commonly used as edible coatings in the food industry. Fats as edible coatings maintain the quality of the coated product during extended storage by acting as excellent oxygen and moisture barriers. They have been used to coat shrimp, meat patties, and sausages (Kroger and Igoe, 1971; Hernandez, 1994). Lipid materials also offer several advantages for coating

470 foods. They have been used in edible films to impart hydrophobicity, cohesiveness, and flexibility (Kester and Fennema, 1986; Hernandez, 1994). Lipid films make good moisture barriers because the tightly packed crystalline structure of lipids restricts the passage of water vapor molecules. However, lipid coatings also can produce anaerobic conditions at higher storage temperatures (Baldwin et al., 1995), lack structural integrity, and do not adhere well to hydrophilic surfaces (Ben and Kurth, 1995). Additionally, some lipid films may have lower permeability to gases such as oxygen, carbon dioxide, and ethylene (Hernandez, 1994). Carnauba wax, derived from palm tree leaves, is considered GRAS (generally recognized as safe) in the U.S. and is permitted in coatings for fruits and vegetables, chewing gum, and confections. Beeswax, also a GRAS material, is harvested from honeycombs by separating the honey from the comb, melting the comb, and refining the wax (Hernandez, 1994). While very pliable at room temperature, beeswax becomes brittle at colder temperatures (Hernandez, 1994). Paraffin is derived from crude petroleum with synthetic versions allowed for food use in the U.S. Currently, paraffin waxes are used as protective coatings on fruits, vegetables, and cheese, in chewing gum, and for microencapsulation of spice flavoring substances (Hernandez, 1994). Coating frozen meats with waxes eliminated down-grading of fresh meat caused by discoloration and reduced packaging labor (McGrath, 1955). The wax coating offered a transparent film capable of withstanding rugged supermarket handling (McGrath, 1955). Long-chain fatty acids, such as stearic and palmitic acids, and corresponding alcohols are used as additives in edible coatings due to their high melting point and hydrophobic characteristics (Hernandez, 1994). Letney (1958) proposed the use of molten fats by allowing them to solidify over fresh meat surfaces to provide a film. Coating meats in this manner lengthened the storage life of the meat at refrigerated conditions, lessened surface dehydration, and maintained the meat “bloom” (Letney, 1958). Ayers (1959) demonstrated that an acetylated glyceride coating containing chlortetracycline reduced off-odors and retained moisture, yet resulted in an unappealing meat color. Anderson (1960, 1961a, b) was issued patents for a lipid-based film used to reduce moisture loss, maintain color integrity, and lessen freezer burn of frozen meats or fish. In the process, cetyl alcohol was mixed with 40% mono-, 40% di-, and 20% triglyceride, heated to 50 C, and the emulsion was either sprayed directly on the meat or the meat was dipped before the temperature dropped. Initial studies with fresh lamb carcasses were not successful, yet application of the emulsion to fresh meat prior to freezing improved color and texture of the meat surface (Anderson, 1960). Brissey and Hill (1961) proposed the application of an acetylated mono- and diglyceride coating to whole cuts of meat, processed with artificial casings, smoked and heated, as a means of keeping the meat and casing separate. A mixture of lard and tallow containing a small amount of lactic


471

acid-fatty acid triglyceride, with and without an antioxidant, was used to coat freeze-dried and fresh meat, including beef steaks, pork chops, and beef cubes (Sleeth and Furgal, 1965). After long-term storage, thiobarbituric acid levels were lower in the antioxidant-treated meat than in the control samples (Sleeth and Furgal, 1965). Schneide (1972) treated the surfaces of beef, veal, and pork steaks as well as fish fillets with different compositions of mono-, di-, and triglycerides of paraffinic and/or carboxylic acids. Formation of a coherent film over the meat surface improved meat storageability and quality with acceptable organoleptic properties and no moisture loss (Schneide, 1972). Griffin et al. (1987) and Leu et al. (1987) treated vacuum-packaged strip loins and top rounds with an acetylated monoglyceride and stored treated product at 2 C for up to 49 days. There were no differences in the storage life (microbial growth, color, or odor) of vacuum-packaged steaks or roasts after four and seven weeks of storage at 2 C, respectively. Stemmler and Stemmler (1976) described coating formulations containing cellulose propionate and fully acetylated monoglyceride obtained from lard. The coatings prolonged the freshness, color, aroma, tenderness, and microbiological stability of fresh beef and pork cuts (Stemmler and Stemmler, 1976). Heine et al. (1979) reported that a completely acetylated mixture of fatty acid mono-, di-, and triglycerides applied to fresh beef and pork pieces and stored at 2 C for 14 days retained desirable color and did not exhibit appreciable weight loss. Water-in-oil emulsions composed of animal fats or vegetable oils have been used as flavor carriers and protective containments for meat and to form continuous coatings that are largely impervious to water vapor (Bauer et al., 1968; Hernandez, 1994; Baldwin et al., 1997). In using these types of emulsions, frozen chicken pieces and pork chops were protected from dehydration, meat yield increased, moisture loss decreased, and flavor and tenderness improved (Zabik and Dawson, 1963; Kroger and Igoe, 1971; Baldwin et al., 1997). Bauer and Neuser (1969) also demonstrated that an edible meat coating comprised of methylcellulose in a water-in-oil emulsion remained stable and intact at low temperatures and prevented excessive loss during cooking on various cuts of pork, beef, chicken, comminuted meat, sausages, fish, and seafood. In another instance, acetostearin films exhibited oxidative stability on meats, especially if derived from hydrogenated vegetable oils, while acetoolein films were less resistant to oxidation (Feuge, 1955; Baldwin, 1994). Waxes, including carnauba, beeswax, and candelilla, also have been successfully used to coat frozen meat pieces and extend their storage life without substantial dehydration (Daniels, 1973). Despite the previously mentioned successes with lipid coatings, highly saturated, acetylated monoglycerides flake and crack during cold storage (Hirasa, 1991), pick up off-flavors (Zabik and Dawson, 1963), and have bitter aftertastes (Morgan, 1971). Unsaturated glycerides and acetylated glycerides appear more susceptible to oxidation (Gennadios et al., 1997).

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Water-soluble or water-dispersible polysaccharides are long-chain polymers that are used in the food industry to provide hardness, crispness, compactness, thickening, adhesiveness, and gel-forming ability (Glicksman, 1983; Whistler and Daniel, 1990; Nisperos-Carriedo, 1994). Generally, polysaccharide films are made from starch, alginate, cellulose ethers, chitosan, carrageenan, or pectins and exhibit good gas barrier properties (Baldwin et al., 1995; Ben and Kurth, 1995). The gas permeability properties of such films result in desirable modified atmospheres, thereby increasing product shelf life without creating anaerobic conditions (Baldwin et al., 1995). However, due to their hydrophilic nature, polysaccharide films are poor water vapor barriers (Ben and Kurth, 1995). An ideal edible polysaccharide-based coating will mimic controlled or modified atmosphere storage associated with plastic-type films while providing an environmentally friendly or biodegradable coating (Baldwin et al., 1995). The aesthetically appealing, non-greasy, and low-calorie polysaccharide films and coatings can be used to extend the shelf life of several foods, including meat products, by preventing dehydration, oxidative rancidity, and surface browning (Nisperos-Carriedo, 1994). Polysaccharide films are used in Japan for packing processed meats, including ham and poultry (Labell, 1991). Films are wrapped around the meat product, which is then smoked and steamed. Not only does the smoke penetrate the film, but the film also is dissolved during the steaming process. Overall, the coated meats exhibit improved yield, structure, and texture and reduced moisture loss (Labell, 1991; Stollman et al., 1994).

Starch is mainly derived from cereal grains, potatoes, tapioca, and arrowroot (Nisperos-Carriedo, 1994). Starch-based films are odorless, tasteless, colorless, non-toxic, biologically absorbable, semi-permeable to carbon dioxide, resistant to passage of oxygen, and exhibit physical characteristics similar to plastic films (Rankin et al., 1958; Nisperos-Carriedo, 1994). Amylose and amylopectin are the two polymers that comprise starch, a polysaccharide occurring widely in nature (Baldwin et al., 1995). Amylose forms coherent, relatively strong, free-standing films (Zobel, 1988). In the late 1960s, hydroxypropylated, high amylose starch films, known as Ediflex, were extruded and used to wrap frozen meat, poultry, and fish (Anonymous, 1967; Kroger and Igoe, 1971; Morgan, 1971; Sacharow, 1972). These flexible, oxygen impermeable, oil-resistant, heat-sealable, and water-soluble films protected meat products during frozen storage and also dissolved during thawing and cooking (Kroger and Igoe, 1971; Morgan, 1971; Sacharow, 1972).


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Alginates are salts of alginic acid, a linear co-polymer of D-mannuronic and L-guluronic acids, derived from seaweed, which possess good film-forming properties that make them particularly useful for food applications (Kester and Fennema, 1986; Nisperos-Carriedo, 1994). Divalent cations are used as gelling agents in alginate film formation because they bridge alginate chains together via ionic interactions followed by hydrogen bonding (Kester and Fennema, 1986; Gennadios et al., 1997). Calcium is more effective in gelling alginates than magnesium, manganese, aluminum, ferrous, or ferric ions (Kester and Fennema, 1986). However, calcium chloride imparts a bitter flavor to foods when used at concentrations greater than 5M, in which case, other calcium salts may be substituted or lower levels used (Baker et al., 1994). Allen et al. (1963) demonstrated that stronger alginate coatings were formed with calcium chloride than with calcium gluconate, nitrate, or propionate. Alginate film strength also can be improved by adding modified starches, oligosaccharides, or simple sugars (Gennadios et al., 1997). Berlin (1957) coated beef cuts with a combination of sodium alginate, calcium chloride, and glycerin (plasticizer) to reduce dehydration. Allen et al. (1963) prepared alginate coatings by dipping beef steaks, pork chops, and chicken drumsticks in alginate or cornstarch solutions, followed by dipping in calcium chloride solutions. Following refrigerated storage for up to seven days, the coatings reduced product shrinkage and improved product texture, juiciness, color, and odor. However, the bitterness imparted by the calcium chloride required for gelling yielded a coated product with unacceptable sensory properties (Allen et al., 1963). In addition, free calcium and other metal cations also may increase proteolytic enzyme activity on meat surfaces by acting as enzyme activators (Gennadios et al., 1997). The use of calcium propionate in alginate coatings instead of calcium chloride resulted in more acceptable flavor; however, due to the weak ionizing properties of calcium propionate, prolonged gelling was necessary to achieve coatings similar in strength to those formed with calcium chloride (Hartal, 1966). Alginate coatings gelled by calcium chloride were used by Lazarus et al. (1976) to reduce weight loss of lamb carcasses. While the alginate coatings resulted in greater weight loss than plastic films, bacterial counts were reduced in alginate-coated samples. Other studies reported that alginate coatings did not affect cooking loss, flavor, odor, or overall acceptability of coated beef (Mountney and Winter, 1961), pork (Nisperos-Carriedo, 1994), and poultry products (Williams et al., 1978). When alginate coatings were applied to beef cuts, weight loss, off-odor, and drip decreased after 96 hours of storage at 5 C compared to non-coated samples (Williams et al., 1978). The coated product also had acceptable flavor, tender-

474 ness, appearance, and cooking loss (Williams et al., 1978). After 144 hours of storage at 5 C, alginate-coated steaks exhibited moisture loss due to drying of the coating. The calcium alginate-based films had antimicrobial activity against natural microflora and coliform bacteria, which was attributed to the presence of calcium chloride (Williams et al., 1978; Cuq et al., 1995). When applied to lamb carcasses, alginate coatings resulted in slower microbial growth and faster chill rate than plastic films (Lazarus et al., 1976). Alginate coatings have been used to retard oxidative off-flavors in re-heated pork patties (Wanstedt et al., 1981). Compared to untreated patties, alginate-treated patties exhibited improved flavor and juiciness, with no lipid oxidation (Wanstedt et al., 1981). Other researchers have extended the shelf life of shrimp, fish, and sausage with alginate coatings (Earle and Snyder, 1966; Daniels, 1973). Earle (1968) and Earle and McKee (1976) extended the shelf life of meat and seafood products by coating them with an aqueous dispersion of algin and dextrose, followed by application of calcium chloride. Alginate coatings also improved adhesion between batter and product, thus reducing the loss of batter from the surface of meat and fish (Fischer and Wong, 1972). Pork chops treated with alginate-starch coatings containing the natural antioxidant tocopherol were juicier and less susceptible to lipid oxidation, but they also developed off-flavors (Hargens-Madsen, 1995). Finally, sodium alginate coatings were successfully used to extend the shelf life of salted and dried mackerel (Shetty et al., 1996).

Carrageenan is extracted from seaweed with water and a small amount of alkali, filtered, and recovered by alcohol precipitation (Nisperos-Carriedo, 1994). It is a complex mixture of several polysaccharides and is used for stabilizing suspensions and emulsions, gelation, thickening, and improving viscosity of food systems (Nisperos-Carriedo, 1994; Baldwin et al., 1995). Food applications using carrageenan-based coatings are made from kappa-, iota-, and lambda-polymers and have been used to prolong the shelf life of poultry (Pearce and Lavers, 1949; Meyer et al., 1959) and fish (Stoloff et al., 1948). To improve product shelf life, antioxidants, such as gallic or ascorbic acids or lecithin, were added to the coatings (Stoloff et al., 1948; Allingham, 1949), as well as salt (Pearce and Lavers, 1949). While earlier research by Meyer et al. (1959) described the addition of antibiotics into carrageenan coatings, such coatings would likely not be approved for food use today.

Another seaweed-derived polysaccharide is agar. Used extensively in mi-


475

crobiological media to provide firmness, agar exhibits characteristics that make it useful for coating meats. It is a galactose polymer that forms strong gels characterized by melting points far above the initial gelation temperature (Sanderson, 1981; Whistler and Daniel, 1985; Gennadios et al., 1997). While combining antibiotics with agar coatings extended the shelf life of coated poultry (Meyer et al., 1959) and beef (Ayres, 1959), the coatings did not reduce moisture loss. Recently, addition of the bacteriocin nisin to agar coatings in combination with food-grade chelators (EDTA, citric acid, or polyoxyethylene sorbitan monolaureate) effectively reduced the levels of Salmonella typhimurium on poultry surfaces (Natrajan and Sheldon, 1995).

Cellulose ethers can be used to produce edible films that are water-soluble, resistant to fats and oils, tough, and flexible (Krumel and Lindsay, 1976; Baldwin et al., 1995). They may be combined with an oil as a plasticizer to coat both fresh and frozen meats (Daniels, 1973). Meat coatings made with methylcellulose (MC) or hydroxypropyl methylcellulose (HPMC) can minimize run-off during cooking and reduce moisture loss when applied as glazes for poultry and seafood (Baker et al., 1994). Bauer and Neuser (1969) described a process that prevented the loss of coating composition during cooking and increased absorption of the coating on pork and poultry pieces by MC, seasoning, and preservative agents. During deep-fat frying of meats, use of HPMC coatings reduced oil uptake and moisture loss (Meyers, 1990). Since the 1980s, considerable research has been directed at developing cellulose ether-based films that exhibit excellent mechanical, oxygen barrier, and oil barrier properties for foods such as pizza and ice cream cones (Kamper and Fennema, 1985; Kester and Fennema, 1989; Rico-Pena and Torres, 1990). However, such films have not been evaluated on meats. Ethylcellulose (EC) has been combined with lipid components and applied to frozen meat products. In addition to being transparent and readily peelable, the resulting coatings prevented desiccation and extended the shelf life of beefsteaks (Ayres, 1959).

Pectins are a group of plant-derived polysaccharides. They differ in methyl ester content and degree of esterification that may affect solubility and gelation properties (Baldwin et al., 1995). Pectin-based films appear to work well with low-moisture foods. However, they are not adequate moisture barriers (Baldwin et al., 1995). Stubbs and Cornforth (1980) demonstrated that a calcium pectinate gel coating reduced shrinkage and bacterial growth on beef plates.

476

Chitin is the major organic skeletal substance of invertebrates and the cell wall of fungi and green algae (Nisperos-Carriedo, 1994). It is found in association with calcium carbonate, protein, or organic compounds. Chitosan, produced by treating chitin with alkali, inhibits fungi and phytopathogens and can produce semi-permeable coatings (Nisperos-Carriedo, 1994). Simpson et al. (1997) and Chen et al. (1998) demonstrated that chitosan coatings reduced microbial contamination on shrimp and oysters, respectively.

Protein films can be prepared from casein, whey protein, gelatin/collagen, fibrinogen, soy protein, wheat gluten, corn zein, and egg albumen (Ben and Kurth, 1995). Such films adhere well to hydrophilic surfaces and provide good barriers for oxygen and carbon dioxide, but do not resist water diffusion (Gennadios and Weller, 1990; Baldwin et al., 1995). Despite the advantages of using proteins in film formation, there are additional concerns. Specifically, enzymes associated with muscle foods may degrade the protein in the films or protein films may present health problems, especially for individuals with food allergies associated with milk, egg, peanut, soybean, or rice proteins (Gennadios et al., 1997).

Gelatin-derived films have been discussed in patent disclosures as early as 1869 (Gennadios et al., 1997). Warm gelatin solutions are applied to products, and coatings are formed during the cooling process. However, gelatin lacks strength and requires a drying step to form more durable films (Daniels, 1973). Formulations of gelatin with polyhydric alcohols (e.g., propylene glycol, ethylene glycol, glycerol, or sorbitol) produce quick-setting, flexible films that exhibit good barrier properties (Whitman and Rosenthal, 1971). Gelatin films have been used as carriers for applying antioxidants to poultry or have been applied directly to poultry meat surfaces or processed meats to prevent microbial growth, salt rust, grease bleeding, handling abuse, water transfer, moisture loss, and oil adsorption during frying (Klose et al., 1952; Childs, 1957; Gennadios et al., 1997). The use of collagen-based films has been proposed for processed meats, including hams, netted roasts, roast beef, fish fillets, and meat pastes (Gennadios et al., 1997). One commercially available collagen film has been purported to reduce shrink loss, increase juiciness, allow for easy removal of nets after cooking or smoking, and absorb fluid exudates for a variety of cooked meat products


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including hams and sausages (Gennadios et al., 1997). Collagen coatings also have been used to reduce transport of gas and moisture in meats (Baker et al., 1994).

Limited information exists on the use of cereal and oilseed proteins as edible films for meats. Corn zein coatings reduced lipid oxidation in precooked pork chops, but did not reduce moisture loss (Hargens-Madsen, 1995). Addition of an antioxidant, emulsifier, and plasticizer to corn zein films reduced rancidity, yet produced off-flavors in cooked turkey breast slices (Herald et al., 1996). In another study, coatings based on 7S soy protein fraction were not as effective as phosphate in preventing development of warmed-over flavor in precooked chicken breast (Kunte, 1996). Wu et al. (2000) coated precooked beef patties with wheat gluten or soy protein coatings. Both coatings were as effective as polyvinyl chloride film in reducing moisture loss after 3 days at 4 C. Both protein coatings controlled lipid oxidation as evidenced by reduced thiobarbituric acid and hexanal values for coated samples compared to non-coated controls.

Due to the advantages and disadvantages of various components, edible film formulations often used combinations of the aforementioned materials (Baldwin et al., 1995). By using composite coatings, two or more materials are combined to improve gas exchange, adherence to coated products, and moisture vapor permeability properties (Baldwin et al., 1995). For example, composite films consisting of lipids and hydrocolloids (proteins or polysaccharides) combine the advantages of each type of material. They may be applied as emulsions or bilayer films. Plasticizers can be used to modify film mechanical properties (Ben and Kurth, 1995). For example, adding glycerol, polyethylene glycol, or sorbitol can reduce film brittleness. Cross-linking proteins and hydrocolloids with enzymes (e.g., transglutaminase) or polyvalent ions (e.g., calcium cations) may also be used for preparing composite films (Ben and Kurth, 1995). A composite coating containing vegetable oils, glycerin, citric acid, and antioxidants prevented rancidity by acting as a moisture barrier, restricting oxygen transport, and serving as a carrier for antioxidants (Baldwin et al., 1995). In a study by Ben and Kurth (1995), sodium caseinate films were cast on trays and dried at ambient temperature, and their barrier properties were determined at 4 C and 90% relative humidity. Additional films were made by adding lipid and/or enzyme (transglutaminase) to sodium caseinate. They reported that lipid addition notably improved moisture barrier properties, but the films appeared slightly cloudy. Enzymatic cross-linking produced water-insoluble films that

478 could gel at 4 C. Enzymatically cross-linked films containing lipids had the best moisture barrier properties. However, when these particular films were applied to meats, the meat surface was masked by the white gel and had an unappealing appearance. Subsequent addition of three enzymes to sodium caseinate produced a cross-linked protein film with limited resistance to water vapor and oxygen. However, this film remained clear, did not alter the color of the meat, and also prevented drip loss (Ben and Kurth, 1995). Further research demonstrated that sealing meat with a three-enzyme cross-linked protein gel produced a juicier product due to reduced drip loss, reduced packaging waste and handling by negating the use of absorbent pads, and presented a more attractive product (Ben and Kurth, 1995). Wong et al. (1992) demonstrated that a chitosan-lauric acid film was less permeable to water vapor, but more permeable to gases than chitosan alone. The microstructure of this film consisted of a sheet-like arrangement stacked in layers, thereby producing an efficient barrier to moisture (Wong et al., 1992).

The benefits of using edible films as packaging materials to minimize microbial growth are threefold. Because water activity (aw) is critical for microbial, chemical, and enzymatic activities, edible films may resist the migration of outer moisture into the food during storage (Wong et al., 1994). Edible films also may serve as gas and solute barriers and compliment other types of packaging by improving the quality and shelf life of foods (Wong et al., 1994). For example, edible films may be formulated to reduce drip loss of meat products while also retarding microbial growth by lowering the aw within a package (Wong et al., 1994). Finally, edible films are excellent vehicles for incorporating a wide variety of additives, such as antioxidants, antifungal agents, antimicrobials, colors, and nutrients (Baldwin, 1994; Wong et al., 1994; Gennadios et al., 1997). Application of antimicrobial compounds in combination with edible packaging materials is receiving considerable attention as a potential intervention for a variety of muscle foods (Cutter, 1998). Several researchers have demonstrated that antimicrobial compounds can be added to edible films to reduce microbial loads on various meat products. Antifungal compounds, organic acids, potassium sorbate, or the bacteriocin nisin were more effective for reducing the levels of foodborne microorganisms when immobilized or incorporated into edible gels, such as starch, carrageenan, waxes, cellulose ethers, or alginate, and then applied to meat surfaces than when these compounds were applied alone. Meyer et al. (1959) were among the first researchers to demonstrate that antibiotics and antifungal compounds could be added to a carrageenan film to re-

Antimicrobial Films

479

duce bacteria by 2 log10 on poultry. Antimycotic agents have been incorporated into edible coatings prepared from waxes and cellulose ethers (Hotchkiss, 1995). Siragusa and Dickson (1992, 1993) demonstrated that organic acids were more efficacious for reducing levels of Listeria monocytogenes, Salmonella typhimurium, and E. coli O157:H7 when immobilized in calcium alginate and applied to beef carcass tissue than when these compounds were applied alone. Baron (1993) demonstrated that potassium sorbate and lactic acid could be incorporated into an edible cornstarch film to inhibit S. typhimurium and E. coli O157:H7 on poultry. Cutter and Siragusa (1996, 1997) immobilized the bacteriocin nisin in calcium alginate gels. This resulted in greater reductions of bacterial populations on lean and adipose beef surfaces and also in greater and sustained bacteriocin activity when the tissues were ground and stored under refrigerated conditions for up to seven days, as compared to control samples treated with only nisin (Cutter and Siragusa, 1996, 1997). When nisin and lysozyme were incorporated into edible heat-set and cast films made from corn zein or soy protein, activity was observed against Escherichia coli and Lactobacillus plantarum (Dawson et al., 1996; Padgett et al., 1998). According to Labuza and Breene (1989), Union Carbide also marketed a sausage casing containing sorbate and glycol that formed a preservative-bearing surface. Natrajan and Sheldon (2000) also incorporated nisin and chelators into protein- and polysaccharide-based films to inhibit Salmonella on poultry skin. Recently, Miller and Cutter (2000) suggested that nisin can be incorporated into collagen-based films and retain activity against a pathogenic and spoilage bacterium following heat treatments and long-term refrigerated storage. A method known as Fibrimex (F.N.A. Foods, Calgary, Alberta, Canada) has been developed for preparing restructured meat products (Wijngaards and Paardekooper, 1988; Paardekooper and Wijngaards, 1989). This cold meat-binding system results from an enzymatic reaction between the blood proteins, fibrinogen and thrombin, and the meat components, collagen and fibronectin (Wijngaards and Paardekooper, 1988). Fibrimex is approved in the U.S. for use in the meat industry as a means of adhering whole muscle or pieces of fresh meat, poultry, fish, or seafood together into one piece, but also forms a gel if applied to a meat surface (Cutter and Siragusa, 1998). Cutter and Siragusa (1998) demonstrated that addition of the bacteriocin nisin to Fibrimex may provide an added antimicrobial advantage to restructured raw meat products that incorporate surface tissues into the product interior or as a delivery system for antimicrobials to meat surfaces. These results highlight the potential for incorporating antimicrobial compounds with a wider and different range of inhibitory activity directly into edible packaging materials. Ultimately, these materials can be used for controlling spoilage, enhancing shelf life, and improving the microbial safety of muscle foods.

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Allen, L., Nelson, A.I., Steinberg, M.P. and McGill, J.N. 1963. “Edible Corn-Carbohydrate Food Coatings. II. Evaluation on Fresh Meat Products,” Food Technol., 17:1442–1446. Allingham, W.J. May 17, 1949. U.S. patent 2,470,281. Anderson, T.R. August 9, 1960. U.S. patent 2,948,623. Anderson, T.R. June 20, 1961a. U.S. patent 2,989,401. Anderson, T.R. June 20, 1961b. U.S. patent 2,989,402. Anonymous. 1967. “Edible Packaging Offers Pluses for Frozen Meat, Poultry,” Quick Frozen Foods, 29:165–167, 213–214. Ayers, J.C. 1959. “Use of Coating Materials or Film Impregnated with Chlortetracycline to Enhance Color and Storage Life of Fresh Beef,” Food Technol., 13:512–515. Baker, R.A., Baldwin, E.A. and Nisperos-Carriedo, N.O. 1994. “Edible Coatings and Films for Processed Foods” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 89–104. Baldwin, E.A. 1994. “Edible Coatings for Fresh Fruits and Vegetables: Past, Present and Future” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 25–64. Baldwin, E.A., Nisperos, N.O. and Baker, R.A. 1995. “Use of Edible Coatings to Preserve Quality of Lightly (and Slightly) Processed Products,” Crit. Rev. Food Sci. Nutr., 35:509–524. Baldwin, E.A., Nisperos, M.O., Hagenmaier, R.D. and Baker, R.A. 1997. “Use of Lipids in Coatings for Food Products,” Food Technol., 51(6):56–62, 64. Baron, J.K. 1993. Inhibition of Salmonella typhimurium and Escherichia coli O157:H7 by an Antimicrobial Containing Edible Film. M.S. thesis, University of Nebraska, Lincoln, NE, USA. Bauer, C.D. and Neuser, G.L. December 9, 1969. U.S. patent 3,483,004. Bauer, C.D., Neuser, G.L. and Pinkalla, H.A. October 15, 1968. U.S. patent 3,406,081. Ben, A. and Kurth, L.B. 1995. Edible film coatings for meat cuts and primal. Meat ’95, The Australian Meat Industry Research Conference, CSIRO, September 10–12, 1995. Berlin, A. 1957. “Calcium Alginate Films and Their Application for Meats Used for Freezing,” Chem. Abstracts, 51:17007. Brissey, G.E. and Hill, R.C. May 2, 1961. U.S. patent 2,982,660. Chen, C.S., Liau, W.Y. and Tsai, G.J. 1998. “Antibacterial Effects of N-Sulfonated and N-Sulfobenzoyl Chitosan and Application to Oyster Preservation,” J. Food Prot., 61:1124–1128. Childs, W.H. October 29, 1957. U.S. patent 2,811,453. Church, I.J. and Parsons, A.L. 1995. “Modified Atmosphere Packaging Technology: A Review,” J. Sci. Food Agric., 67:143–152. Cuq, B., Gontard, N. and Guilbert, S. 1995. “Edible Films and Coatings as Active Layers” in Active Food Packaging, ed., M.L. Rooney, New York: Blackie Academic and Professional, pp. 111-142. Cutter, C.N. 1998. “New Intervention Technologies” in Proceedings of the 51st Annual Reciprocal Meat Conference, Kansas City, MO: American Meat Science Association, pp. 133–140. Cutter, C.N. 2001. “Microbial Control by Packaging: A Review,” Crit. Rev. Food Sci. Nutr. (in press). Cutter, C.N. and Siragusa, G.R. 1996. “Reduction of Brochothrix thermosphacta on Beef Surfaces Following Immobilization of Nisin in Calcium Alginate Gels,” Lett. Appl. Microbiol., 23:9–12. Cutter, C.N. and Siragusa, G.R. 1997. “Growth of Brochothrix thermosphacta in Ground Beef Following Treatments with Nisin in Calcium Alginate Gels,” Food Microbiol., 14:425-430.

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482 Hotchkiss, J.H. 1995. “Safety Considerations in Active Packaging” in Active Food Packaging, ed. M.L. Rooney, New York: Blackie Academic & Professional, pp. 238-255. Kamper, S.L. and Fennema, O.R. 1984. “Water Vapor Permeability of an Edible Fatty Acid, Bilayer Film,” J. Food Sci., 49:1482–1485. Kamper, S.L. and Fennema, O.R. 1985. “Use of an Edible Film to Maintain Water Vapor Gradients in Foods,” J. Food Sci., 50:382–384. Kester, J.J. and Fennema, O.R. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Kester, J.J. and Fennema, O.R. 1989. “An Edible Film of Lipids and Cellulose Ethers: Performance in a Model Frozen-Food System,” J. Food Sci., 54:1390–1392, 1406. Klose, A.A., Mecchi, E.P. and Hanson, H.L. 1952. “Use of Antioxidants in the Frozen Storage of Turkeys,” Food Technol., 6:308–311. Kroger, M. and Igoe, R.S. 1971. “Edible Containers,” Food Prod. Dev., 5:74, 76, 78–79, 82. Krumel, K.L. and Lindsay, T.A. 1976. “Nonionic Cellulose Ethers,” Food Technol., 30(4):36–38, 40, 43. Kunte, L.A. 1996. Effectiveness of 7S Soy Protein and Edible Film in Controlling Lipid Oxidation in Chicken. M.S. thesis, University of Nebraska, Lincoln, NE, USA. Labell, F. 1991. “Edible Packaging,” Food Process., 52:24. Labuza, T.P. and Breene, W.M. 1989. “Applications of ‘Active Packaging’for Improvement of Shelf Life and Nutritional Quality of Fresh and Extended Shelf-Life Foods,” J. Food Proc. Pres., 13:1–69. Labuza, T.P., Fu, B. and Taoukis, P.S. 1992. “Prediction for Shelf Life and Safety of Minimally Processed CAP/MAP Chilled Foods: A Review,” J. Food Prot., 55:741–750. Lazarus, C.R., West, R.L., Oblinger, J.L. and Palmer, A.Z. 1976. “Evaluation of a Calcium Alginate Coating and a Protective Plastic Wrapping for the Control of Lamb Carcass Shrinkage,” J. Food Sci., 41:639–641. Letney, L.R. January 24, 1958. U.S. patent 2,819,975. Leu, R., Keeton, J.T., Griffin, D.B., Savell, J.W. and Vanderzant, C. 1987. “Microflora of Vacuum Packaged Beef Steaks and Roasts Treated with an Edible Acetylated Monoglyceride,” J. Food Prot., 50: 554–556. McGrath, E.P. 1955. “Packaging Costs Cut, Quality Protected by Wax-Coating Frozen Meats,” Food Engr., 27(8):50–51, 77. Meyer, R.C., Winter, A.R. and Weister, H.H. 1959. “Edible Protective Coatings for Extending the Shelf Life of Poultry,” Food Technol., 13:146–148. Meyers, M.A. 1990. “Functionality of Hydrocolloids in Batter Coating Systems” in Batters and Breadings in Food Processing, eds. K. Kulp and R. Loewe, St. Paul, MN: American Association of Cereal Chemists, pp. 117–141. Miller, B.J. and Cutter, C.N. 2000. “Incorporation of Nisin into a Collagen Film Retains Antimicrobial Activity Against Listeria monocytogenes and Brochothrix thermosphacta Associated with a Ready-to-Eat Meat Product.” Presented at the Fall Meeting of the Allegheny Branch of the American Society for Microbiology, State College, PA, USA, October 27, 2000. Morgan, B.H. 1971. “Edible Packaging Update,” Food Prod. Dev., 5(6):75–77, 108. Mountney, G.J. and Winter, A.R. 1961. “The Use of Calcium Alginate Film for Coating Cut-up Poultry,” Poultry Sci., 40:28–34. Natrajan, N. and Sheldon, B.W. 1995. “Evaluation of Bacteriocin-based Packaging and Edible Film Delivery Systems to Reduce Salmonella in Fresh Poultry,” Poultry Sci., 74(Suppl. 1):31. Natrajan, N. and Sheldon, B.W. 2000. “Inhibition of Salmonella on Poultry Skin Using Protein- and Polysaccharide-based Films Containing a Nisin Formulation,” J. Food Prot., 63:1268–1272. Nisperos-Carriedo, M.O. 1994. “Edible Coatings and Films Based on Polysaccharides” in Edible Coatings and Films to Improve Food Quality, eds. J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, pp. 305–335.

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Protein-based Edible Films and Coatings Carrying Antimicrobial Agents

CTIVE packaging technologies involve interactions between the food, the packaging (or coating) material, and the internal gaseous atmosphere to extend the shelf life of foods while maintaining their quality and safety (Labuza and Breene, 1988). In general, active food packaging can provide several functions that do not exist in conventional packaging systems. Antimicrobial, edible films and coatings have two characteristic functions compared to the conventional barrier and protective functions of food packaging systems. These functions are antimicrobial activity and edibility/biodegradability. In addition to antimicrobial activity, possible functions include scavenging of oxygen, moisture, or ethanol, and emission of ethanol (Floros et al., 1997; Han, 2000). Edible films and coatings are protective barriers while also being an edible portion of the packaged or coated food. Because of the barrier function of edible films and coatings, the coated foods may not require high-barrier packaging systems. Therefore, the entire packaging structure can be simplified by satisfying the requirement of barrier properties. The protective functions include oxygen, moisture, and aroma barriers, as well as physical damage prevention. In addition to enhanced barrier properties, edible films and coatings control adhesion, cohesion, and durability, and improve the appearance of coated foods (Krochta, 1997).

A

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486 Active ingredients and seasonings can be incorporated into edible films and coatings. They may carry antioxidants, antimicrobial agents, colorants, flavors, fortified nutrients, and/or spices. The active ingredient carrier function of edible films and coatings needs further studying due to its variety of potential applications and benefits. These functions are promising and will receive more attention in the near future. Food packaging materials protect food from physical, chemical, and biological hazards (e.g., moisture, light, gases, aromas, microorganisms, rodents, and insects), thereby maintaining its quality and prolonging shelf life. Food packaging also provides important information to consumers, such as expiry dating, and facilitates convenience during distribution. Contrary to the importance of packaging functions, packaging materials generate approximately 30% by weight of municipal solid waste and account for two-thirds of the volume in trash cans due to their bulk (Hunt et al., 1990). Among the 30% packaging waste, 13% is due to plastic materials, which do not biodegrade although they are convenient, safe, strong, and economical (Krochta and De Mulder-Johnston, 1997). Because of the environmental aspect of packaging materials, edible films and coatings, as well as biodegradable materials, offer alternative packaging systems without environmental costs, unlike the use of non-degradable synthetic materials (Krochta and De Mulder-Johnston, 1997). Edible and/or biodegradable materials may totally replace some synthetic packaging films or reduce the use of synthetic materials by partially replacing them. Because they will be consumed with foods or will be decomposed before they reach the landfill sites, the overall amount of solid waste may be reduced.

Food spoilage has been studied throughout history in a search for efficient preservation methods. Traditionally, several methods have been used to preserve foods including changes in temperature (heating or cooling), water activity reduction, pH control, modified/controlled atmospheres, curing, salting, addition of antimicrobials, irradiation, and packaging (Wagner and Moberg, 1989). Antimicrobial agents generally control the growth of spoilage and pathogenic microorganisms. Inhibition of spoilage microorganisms has been studied to develop effective preservation methods. Currently, food preservation by controlling spoilage and pathogenic microorganisms is generally practiced well in modernized manufacturing systems. However, compared to spoilage microorganisms, contamination of foods by pathogenic microorganisms causes much more serious problems and endangers public health. Because edible films and coatings are a part of the food packaging system, they possess an important function of safety when antimicrobial agents are incorporated into the film or coating materials. A variety of antimicrobial agents have tradition-

Antimicrobial Edible Films and Coatings

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ally been used for food preservation and may be added to the film or coating materials to provide antimicrobial functions to edible films and coatings. Antimicrobial edible films and coatings, as well as antimicrobial synthetic plastic films, have emerged recently. However, antimicrobial edible film and coating studies go far back into history in the development of sausage casings that contain chemical preservatives and organic acids. Natural casings are mainly edible protein tubes that are filled with sausage batter. Salting, curing, smoking, and acid treatments on natural casings provided effective preservation and have been applied for a long time with homemade sausages before modern scientific studies were initiated. In addition to the sausage casing applications, meat loaf and beef carcasses were coated with gels carrying antimicrobial agents to inhibit growth of spoilage and pathogenic bacteria. However, most of the used gel-based coatings were non-protein gums, such as calcium alginate. Antimicrobial agents incorporated in calcium alginate coatings were nisin (Fang and Lin, 1995; Cutter and Siragusa, 1997) and lactic/acetic acids (Siragusa and Dickson, 1992, 1993). Only a few research reports deal with coating of meat products with protein (e.g., gelatin) coatings carrying antimicrobial agents. Gill (2000) applied gelatin-based gel coatings containing lysozyme, nisin, and EDTA on cooked ham and bologna sausage to control the growth of pathogens. The antimicrobial gels had a bactericidal effect on gram-positive pathogens (i.e., Brochothrix thermosphacta, Lactobacillus sakei, Leuconostoc mesenteroides, and Listeria monocytogenes) and Salmonella typhimurium (Gill, 2000). Besides the sausage casing studies, research on edible films and coatings containing antimicrobial agents started in the mid-1980s, mostly using sorbic acid or potassium sorbate. However, proteins were not the selected coating materials. Instead, carbohydrates and lipids were commonly used as coatings containing sorbates. Such coating materials included methyl cellulose (Rico-Pena and Torres, 1991), hydroxypropylmethyl cellulose (Vojdani and Torres, 1990), starch (Baron and Sumner, 1993), fatty acids (Vojdani and Torres, 1990; Rico-Pena and Torres, 1991), and carnauba wax (Guilbert, 1988). Since the mid-1990s, various antimicrobial films were prepared by combining chitosan and organic acids (Chen et al., 1996; Ouattara et al., 2000). Some studies on protein-based edible films and coatings used as microbial hurdle technology also started in the mid-1980s. Torres et al. (1985) studied the diffusion and permeation of sorbic acid in a zein coating applied onto an intermediate moisture food. They reported that surface microbial stability increased after the food was coated with sorbic acid and zein. Torres and Karel (1985) also demonstrated the effectiveness of sorbic acid incorporated into zein coatings against Staphylococcus aureus. Guilbert (1988) determined the effectiveness of sorbic acid incorporated in casein or gelatin films and showed improved microbial stability. Since the late 1990s, natural antimicrobials have been in-

488 corporated into edible protein films. Nisin-containing zein films (Padgett et al., 1995) and lysozyme- or nisin-containing soy protein films/zein films (Padgett et al., 1998) were tested for their antimicrobial activity against Lactobacillus plantarum. Potassium sorbate was incorporated into whey protein isolate films and the antimicrobial activity of the films, against Saccharomyces cerevisiae, Aspergillus niger, and Penicillium roqueforti was determined (Ozdermir, 1999). Cooksey et al. (2000) incorporated nisin in zein pouches for packaging shredded cheese. However, they reported that the nisin in the zein films had no effect on aerobic bacteria and the zein films had inadequate barrier properties at low temperature. Rodrigues and Han (2000) demonstrated the growth inhibition effects of whey protein films containing lysozyme, nisin, EDTA, or propyl-p-benzoic acid against Listeria monocytogenes, E. coli O157:H7, Brochothorix thermosphacta, Staphylococcus aureus, and Salmonella typhimirium. Films carrying lysozyme or nisin substantially inhibited B. thermosphacta and adequately inhibited S. aureus. EDTA-containing films showed good inhibition against L. monocytogenes and S. typhimirium, and had a marginal effect on E. coli O157:H7. Propyl-p-benzoic acid did not significantly affect the tested bacteria. A case-by-case optimization of the antimicrobial edible protein films based on food characteristics, physical properties of protein layers, storage environments, combinations of protein layers and antimicrobials, and potential microflora is suggested for maximizing the effectiveness of the films (Cooksey et al., 2000; Rodrigues and Han, 2000).

Non-edible packaging systems may contain any type of food-grade preservative in their packaging materials to create an antimicrobial activity. They may include organic acids and their salts, fungicides, bacteriocins, antibiotics, enzymes, alcohols, thiols, antioxidants, or sanitizing gases (Han, 2000). However, in the case of edible films and coatings, selection of the incorporated antimicrobial agents is limited to edible compounds. Because they have to be consumed along with the edible film/coating layers and the foods, their edibility and safety are essential.

Food-grade chemical antimicrobial agents used for food preservation may be incorporated into film and coating materials that contain edible proteins. Such agents include organic acids and their salts, such as benzoic acid, sodium benzoate, sorbic acid, potassium sorbate, propionic acid, lactic acid, and acetic acid (Cuppett, 1994; Han, 2000). Most chemical antimicrobial agents naturally exist in plants or fermented products. However, they mainly are chemically

Antimicrobial Substances

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synthesized and categorized as chemical preservatives. Table 19.1 shows chemical, food-grade antimicrobial agents that may be incorporated into edible films and coatings as functional additives with or without regulatory restrictions (Lueck and Jager, 1997).

As shown in Table 19.1, many compounds found in nature or naturally in foods have antimicrobial activity. Natural antimicrobials consist of enzymes, proteins, organic acids, fatty acids, pigments, flavors, and spice oils. Due to their natural characteristics, it is expected that they have great potential for use as antimicrobial additives in foods or edible films and coatings without the ad-

490 verse effects associated with chemical preservatives. However, the inhibitory levels, antimicrobial spectrum, and toxicity of natural preservatives must be studied more thoroughly to ensure their antimicrobial effectiveness in edible films and coatings and in foods.

Bacteriocins are protein-containing macromolecules that have bactericidal effects on susceptible bacteria. They are produced by various bacteria and possess different antibacterial spectra, modes of action, and chemical properties (Daeschel, 1989). Common bacteriocins are colicins (E. coli bacteriocins), lactic acid bacteriocins, pediocins (Pediococcus bacteriocins), nisin, diolococcin, propionibacterium bacteriocins, and reuterin (Daeschel, 1989). Lysozyme is a widely studied antimicrobial enzyme. It possesses antimicrobial activity due to its ability to hydrolyze the peptidoglycan that composes the cell wall of gram-positive and gram-negative microorganisms. Though lysozymes can be isolated from a wide range of sources, including mammalian tears, milk, serum, and avian eggs, hen’s egg lysozyme is mainly studied and utilized for food applications. Although lysozyme is effective against many gram-positive bacteria, the resistance of gram-negative bacteria, which have a lipid-based outer membrane over the peptidoglycan cell wall, limits its antimicrobial activity.

Figure 19.1 shows the structure of edible protein film and coating systems. In the case of edible film applications, protein films function as packaging materials. Packaged food may be contaminated by microorganisms before packaging or by post-process contamination after the sealed package is opened. Therefore, surface contamination is the most probable and needs to be prevented. Contaminating microorganisms will locate themselves on the food surface, i.e., in the area between the package and the food. In the case of edible coating systems, protein coatings completely cover the food. However, the food surface may potentially get contaminated before the coating process. Microorganisms positioned between the coating layer and the food product may not become active and grow due to lack of oxygen and to direct contact with the antimicrobial agents. Therefore, coated foods are most likely to become contaminated on the external coated surface where microorganisms will position themselves and start to grow. In both film and coating systems, the food layers that do not contain antimicrobial agents initially have very large volume com-

Factors Considered in Designing Antimicrobial Edible Film

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Figure 19.1 Structure of edible protein film and coating systems and migration of antimicrobial agents in food. In the case of incorporating volatile antimicrobial agents (not illustrated), the antimicrobial agent would escape from the film and coating layer to the surrounding environment.

pared to the volume of the thin films or coatings. Because of the almost infinite volume of the food layer compared to the film/coating layer and the migration of antimicrobial agents from the films/coatings into the food, the concentration of the antimicrobials in the protein film and coating layers will be reduced. Eventually the antimicrobials will be depleted from the protein films/coatings. Therefore, the release rate must be controlled to prevent the early depletion of antimicrobial agents due to fast migration. Therefore, Figure 19.1 suggests that the two systems (the film system and the coating system) should have different protective functional designs. In the protein film system, the incorporated antimicrobials should migrate slowly from the film layer into the food, thereby acting against contaminating microorganisms. In contrast, in the protein coating system, the antimicrobial agents must remain in the coating layer to protect the food product from invasion of contaminating microorganisms. Controlled release of the antimicrobials with an intermediate diffusion rate is expected to achieve effective antimicrobial activity at the food surface in a protein film system. In a coating system, preserved high concentration of the antimicrobials is required with a very slow diffusion rate to maintain the efficiency of the antimicrobial functions against spoilage and pathogenic microorganisms.

492 A mass transfer model of the migration phenomena can be used to describe the concentration profile in the film/coating layer and food over time. Figure 19.1 shows that a two-layer diffusion model can represent both the protein film and coating systems. When volatile antimicrobial agents are incorporated, they can evaporate out from the system to the surrounding environment. Most non-volatile antimicrobial agents would penetrate into the food layer during storage and distribution. Though various models are available, Han (2000) summarized traditional mass transfer models and his own proposed models that may be used to describe the migration of antimicrobial agents through food packaging systems consisting of single, double, or triple layers. Because a mass transfer model describes the relationship between concentration and time, it allows for calculating the storage period that maintains the antimicrobial concentration above the critical inhibitory concentration and permits the estimation of the microbiologically safe shelf life.

Antimicrobial agents have their own growth inhibition mechanisms and kinetics against a target microorganism. Various mechanisms may exist against a single microorganism and may vary depending on the target microorganisms. Growth inhibition kinetic studies deal with the inhibition rate of antimicrobial agents against a target microorganism at certain conditions (e.g., storage temperature, carbon source, nitrogen source, partial pressure of oxygen, pH, moisture, and other environmental factors). Most environmental factors are controlled by the nature of the food product and by the storage conditions. Therefore, food components significantly impact the effectiveness of contained antimicrobials and the sensitivity of the microorganisms. For example, the acidic pH of a food increases the antimicrobial activity of most organic acids and their salts due to the association of the acids. Moisture content of the food relates to the water activity of the food and affects the microbial growth rate. Also, each food has its own characteristic microflora. Therefore, selection criteria of antimicrobial agents should include food composition and the contaminating microorganisms that are likely to be present. Most proteins have isoelectric points at acidic pH values. Therefore, most protein films and coatings have a negative net charge at neutral pH, an environment that is more susceptible to microbial growth. In this neutral pH region, lysozyme and nisin, which also are proteins, are positively charged because their isoelectric points are in the highly alkaline pH region. Therefore, at the neutral pH of edible protein films and coatings, the film-forming proteins and lysozyme/nisin form strong ionic interactions. The movement of lysozyme and nisin in the film/coating protein structure is not free, thereby decreasing the effective concentration of the antimicrobial agents per microorganism at the food


493

surface due to the limitations in the migration of lysozyme and nisin. This phenomenon has to be considered when designing protein edible film/coating systems because the migration at the specific release rate of antimicrobials from the protein layer into the food is important to the effectiveness of the antimicrobial packaging system. Both attractive and repulsive forces between the migrating antimicrobials and the protein film/coating layers are factors that affect the diffusion coefficient. This knowledge of molecular interaction and repulsion can provide the clue to altering the diffusion coefficient, so that controlled release of antimicrobials is achieved by designing a modified structure of the edible film and coating system.

Storage temperature can affect antimicrobial activity, migration rate of antimicrobials, and microflora. High temperature treatments may cause loss of the antimicrobial activity because most chemicals are not stable at high temperatures. If the packaged or coated food needs a heat treatment, the residual activity of the incorporated antimicrobials and the heat effect on the activity should be determined. High temperatures also increase the volatility of incorporated antimicrobials. If highly or partially volatile antimicrobials are used, the residual antimicrobial activity may decrease due to evaporation of the active chemicals. Generally, the mass transfer rate follows an Arrhenius relationship to the storage temperature. Increased storage temperature can accelerate the migration of the antimicrobials in the film/coating layers, while refrigeration slows down the migration rate. Temperature also is a critical factor in microbial growth. Every microorganism has a characteristic temperature response and an optimal growth temperature. Controlling the storage temperature has been an effective way to prevent microbial contamination of food products. However, changes in storage temperature also affect other essential factors in designing antimicrobial edible film and coating systems, such as migration rate, chemical stability, and cost.

Edible films are typically prepared by a wet casting method, which uses a film-forming protein/plasticizer solution. This method will likely have to be replaced by extrusion methods for future manufacturing production. Dry extrusion methods (e.g., for thermoplastic films) use high temperature and pressure combinations, while wet extrusion methods (e.g., for cellophane films) require relatively lower temperature and pressure. A heat-press method was used to prepare soy protein films containing nisin (Padgett et al., 1998). Using a heat-press mold is an important transition toward developing an extrusion method, because it also uses high temperature and pressure treatments. To ex-

494 trude films, the process conditions have to be optimized to minimize degradation of the antimicrobial agents during the extrusion process. Generally, proteins have lower melting and gelation temperatures than synthetic plastic materials. Therefore, the extrusion temperatures for protein films would be lower than the processing temperatures for conventional plastic films. However, protein extrusion is still a high-temperature process. High temperature and pressure processes used in film formation reduce antimicrobial activity. For example, potassium sorbate significantly lost its antimicrobial activity by heat treatment (Han and Floros, 1999). Most casting processes of protein films and all coating processes are designed to use protein film-forming solutions. To produce antimicrobial protein films/coatings, the antimicrobial agents must be dissolved in the protein solutions before film casting or food coating. It is recommended that the active antimicrobials have a high solubility in protein solutions, do not form coagulates with the film/coating protein in the solutions, and have high stability that does not allow the antimicrobials to evaporate or degrade during the drying process after casting or coating.

Addition of antimicrobial agents into edible film/coating solutions generally causes the film properties to deteriorate due to increased heterogeneity in the edible film/coating structures. The material properties of protein films and coatings include mechanical properties (e.g., tensile strength, elongation, stiffness, and tearing resistance); physical properties (e.g., oxygen permeability, water vapor permeability, aroma/volatile component permeability, wettability, water absorptiveness, and grease resistance); optical properties (e.g., brightness, haze, gloss, transparency, and color); and process/machinability properties (e.g., stickiness, static, seal strength, and ink absorptiveness). The performance of edible films or coatings must be maintained when antimicrobial agents are added. Film-forming and coating materials have large molecular size and are present in high amounts compared to the small molecular size and the small amounts of the antimicrobial. Addition of antimicrobials into plastic films generally does not affect the physical strength of the films. For example, addition of potassium sorbate to polyethylene films did not affect their mechanical strength (Han and Floros, 1997). This may be explained by the location of the very small molecule (i.e., sorbate) in the large amorphous region of the plastic polymer. However, the transparency of the plastic films decreased with addition of the antimicrobial. This could have been caused by the poor compatibility of the water-soluble antimicrobial with the hydrophobic plastic structure. Edible film/coating structures that contain water-soluble chemicals or peptide antimicrobials can possibly produce clear films without noticeable


495

changes in mechanical strength. For example, when lysozyme was incorporated (10% w/w) into whey protein isolate (WPI) films, where both the antimicrobial agent (i.e., lysozyme) and the film-forming protein (i.e., WPI) were compatible and highly water-soluble, the transparency and tensile strength of the WPI films was not affected (Han, 2000). Also, the physical properties of WPI films did not change by adding nisin, EDTA, or propyl-p-benzoic acid (Rodrigues and Han, 2000). However, films from corn zein, an alcohol-soluble protein, have a disadvantage in that addition of water-soluble antimicrobials decreases their physical strength. For example, adverse effects of added nisin on corn zein film tensile strength were reported by Cooksey et al. (2000). This loss of film strength may be caused by the poor compatibility between water-soluble nisin and water-insoluble zein. Although more extensive studies on zein films containing heterogeneous additives are required, it is expected that addition of alcohol-soluble antimicrobials will not affect the mechanical properties of zein films.

Antimicrobial edible protein films and coatings also are edible portions of the packaged or coated foods. Therefore, the components of the protein films and coatings should not interfere with the organoleptic characteristics of the food product. Exceptionally, some flavor compounds may be intentionally incorporated into the edible films/coatings to create unique tastes or mask undesirable tastes of the foods. Generally, tasteless protein films and coatings are desirable to minimize taste interference. Because the antimicrobial edible protein layers mainly consist of film-forming proteins, plasticizers, and antimicrobial agents, all of these three major components should be tasteless. Though WPI has a creamy milk flavor, dried, denatured WPI films are tasteless. Soy protein isolate films and peanut protein films have a slight beany flavor, which is generally acceptable. Collagen and gelatin films are known to be tasteless. Casein films have an acceptable milk flavor. Corn zein produces yellowish, tough films with a corn meal flavor that can be objectionable (Guilbert, 1986). Generally, aroma compounds of protein sources affect the residual flavor of protein film layers more significantly than taste compounds. Plasticizers are the second major component of the antimicrobial edible protein layers. Common edible plasticizers are glycerol, sorbitol, and polyethylene glycol. Glycerol and sorbitol taste sweet. However, the sweetness of glycerol in whey protein films is negligible, while the sweetness of sorbitol is noticeable. Polyethylene glycol is tasteless. Most antimicrobial agents have their own characteristic flavor. However, because the antimicrobial concentration in protein films is generally very low, their taste effects may be negligible. When natural antimicrobials are added to

496 protein film layers in high concentration levels, the protein layers may possess the strong flavor of the incorporated antimicrobials. This phenomenon is more significant when plant and herb essential oils/extracts or phenolic flavors are added to edible protein layers. Several studies have focused on edible protein film and coating technology. However, few studies have dealt with sensory examination of edible film and coating layers. More research on the organoleptic characteristics of antimicrobial edible protein films, as well as conventional edible films, is required.

Edible protein films and coatings contain natural food proteins and are biodegradable. The use of natural ingredients instead of synthetic chemicals is a worldwide trend affecting most consumer products. Due to this consumer preference, more research and development will focus on the use of natural antimicrobials. Their use as crude extracts or in partially purified forms may provide additional functionalities beyond antimicrobial activity, such as nutraceutical activity and different flavors.

Food safety is one of the primary concerns of food processors, and consumer demand for safer food increases. Research and development on antimicrobial films and coatings was initiated to satisfy this food safety requirement. In relevant research studies, only a single antimicrobial compound has been used in the edible protein layer. Because of the characteristic inhibitory mechanism and the specific activity of each antimicrobial against a specific target microorganism, the antimicrobial edible films/coatings had a limited antimicrobial spectrum. Combining two or more antimicrobials can widen this limited spectrum. Bactericidal agents, such as lysozyme, that destroy peptidoglycan in cell walls can effectively eliminate most gram-positive pathogens. However, it is difficult to inhibit the growth of gram-negative bacteria by cell wall destruction. For example, to simultaneously inhibit Listeria monocytogenes and E. coli O157:H7 using a single antimicrobial film/coating, the antimicrobial film must carry a mixture of antimicrobial agents (e.g., lysozyme and EDTA). Studies on gram-negative inhibition are required to develop films and coatings with a wider antimicrobial spectrum.

Research and Development Trends for Antimicrobial Edible Protein Films 497

The use of mixed antimicrobials also increases the sensitivity of pathogens and provides a synergistic inhibitory effect. Padgett et al. (1998) reported a synergistic antimicrobial effect of lysozyme or nisin with EDTA in soy protein and zein films against E. coli. Gill (2000) reported a significant synergistic effect among lysozyme, nisin, and EDTA in gelatin casings against B. thermosphacta, Leuconostoc mesentroides, Lactobacillus curvatus, L. monocytogenes, S. typhimurium, and E. coli O157:H7. In porous foods (e.g., breads and hamburger patties), in-package headspaces (e.g., powdered milk and peanut pouch), or in food with air gaps between the protein layer and the food, the irregular surface and the air space prevent migration of the antimicrobials into the food structure. In these cases, only volatile antimicrobials can be utilized and vaporized into the air spaces. They will migrate via the air gaps into the food. Use of volatile antimicrobials provides a greater benefit to the preservation of heterogeneous foods and food products for which coatings are not available. Volatile antimicrobial systems require a headspace concentration controlling system, which maintains the effective gaseous concentration of the antimicrobial vapors.

Antimicrobial protein films and coatings are a type of food packaging. Protein layers are expected to possess barrier functions to satisfy the requirement of the food packaging materials. If the protein film and coating layers have adequate physical and mechanical strength and have the additive antimicrobial activity to protect the food from physical, chemical, and microbial hazards, this single protein film or coating layer can eliminate or reduce the use of non-edible synthetic packaging materials. Compositional modifications of film-forming materials can alter their barrier properties. Most edible protein film research has focused on increasing film barrier ability against moisture and oxygen and in improving mechanical properties, and this will continue in the future. Plant and herb essential oils/extracts and phenolic flavors possessing antimicrobial activity can be used as additional nutraceutical ingredients, antioxidants, colorants, or flavors in protein films and coatings. The multi-functional edible films/coatings can then be utilized for value-added confections, medicinal and therapeutic foods, pharmaceutical products, and other nutraceuticals as well as conventional perishable foods. Several edible proteins are potential allergens. For example, film-forming proteins derived from nuts, peas, beans, and milk, as well as protein-based antimicrobials, may cause allergy problems. Studies on the potential allergenicity of film-forming proteins and protein-based antimicrobial agents should be pursued intensively in order for them to be used as an antimicrobial

498 edible protein film/coating system for wider and safer food preservation methods.

Recently, edible films and coatings have been receiving interest in the food industry due to their distinct possible advantages over non-edible packaging systems. Their inherent edibility and biodegradability are strong advantages of these edible films and coatings. In addition to their environmentally friendly nature, increased food safety can be achieved by incorporating antimicrobial agents into the films/coatings. Edibility, biodegradability, and microbial safety are three major benefits of antimicrobial protein films and coatings.

Baron, J.K. and Sumner, S.S. 1993. “Antimicrobial Containing Edible Films as an Inhibitory System to Control Microbial Growth on Meat Products,” J. Food Protect., 56:916. Beuchat, L.R. and Golden, D.A. 1989. “Antimicrobials Occurring Naturally in Foods,” Food Technol., 43(1):134–142. Chen, M.-C., Yeh, G.H. and Chiang, B.-H. 1996. “Antimicrobial and Physicochemical Properties of Methylcellulose and Chitosan Films Containing a Preservative,” J. Food Process. Preserv., 20:379–390. Cooksey, D.K., Gremmer, A. and Grower, J. 2000. “Characteristics of Nisin-Containing Corn Zein Pouches for Reduction of Microbial Growth in Refrigerated Shredded Cheddar Cheese” in 2000 IFT Annual Meeting Book of Abstracts, Chicago, IL: Institute of Food Technologists, p. 188. Cuppett, S.L. 1994. “Edible Coatings as Carriers of Food Additives, Fungicides and Natural Antagonists” in Edible Films and Coatings to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Co., Inc., pp. 121–137. Cutter, C.N. and Siragusa, G.R. 1997. “Growth of Brochothrix thermosphacta in Ground Beef Following Treatments with Nisin in Calcium Alginate Gels,” Food Microbiol., 14:425–430. Daeschel, M.A. 1989. “Antimicrobial Substances from Lactic Acid Bacteria for Use as Food Preservatives,” Food Technol., 43(1):164–167. Fang, T.J. and Lin, C. 1995. “Inhibition of Listeria monocytogenes on Pork Tissue by Immobilized Nisin,” J. Food Drug Anal., 3:269–274. Floros, J.D., Dock, L.L. and Han, J.H. 1997. “Active Packaging Technologies and Applications,” Food Cosmet. Drug Pkg., 20(1):10–17. Gill, A.O. 2000. Application of Lysozyme and Nisin to Control Bacterial Growth on Cured Meat Products. M.S. thesis, The University of Manitoba, Winnipeg, MB, Canada. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation: Theory and Practice, ed., M. Mathlouthi, London, UK: Elsevier Applied Science Publishers, pp. 371–394. Guilbert, S. 1988. “Use of Superficial Edible Layer to Protect Intermediate Moisture Foods: Application to the Protection of Tropical Fruit Dehydrated by Osmosis” in Food Preservation by Moisture Control, ed., C.C. Seow, London, UK: Elsevier Applied Science Publishers, pp. 119–219. Han, J.H. 2000. “Antimicrobial Food Packaging,” Food Technol., 54(3):56–65.

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Han, J.H. and Floros, J.D. 1997. “Casting Antimicrobial Packaging Films and Measuring Their Physical Properties and Antimicrobial Activity,” J. Plastic Film & Sheeting, 13:287–298. Han, J.H. and Floros, J.D. 1999. “Modeling Antimicrobial Activity Loss of Potassium Sorbate against Baker’s Yeast after Heat Process to Develop Antimicrobial Food Packaging Materials,” Food Sci. Biotechnol., 8:11–14. Hunt, R.G., Sellers, V.R., Franklin, W.E., Nelson, J.M., Rathje, W.L., Hughes, W.W. and Wilson, D.C. 1990. Estimates of the Volume of MSW and Selected Components in Trash Cans and Landfills. Prairie Village, KS: Franlin Associates, Ltd., and Tucson, AZ: The Garbage Project. Krochta, J.M. 1997. “Edible Protein Films and Coatings” in Food Proteins and Their Applications in Foods, eds., S. Damodaran and A. Paraf, New York: Marcel Dekker, pp. 529–549. Krochta, J.M. and De Mulder-Johnston C. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Labuza, T.P. and Breene, W.M. 1988. “Applications of Active Packaging for Improvement of Shelf-life and Nutritional Quality of Fresh and Extended Shelf-life Foods,” J. Food Process. Preserv., 13:1–69. Lueck, E. and Jager, M. 1997. Antimicrobial Food Additives: Characteristics, Uses, Effects. Translated by S.F. Laichena, New York: Springer. Naidu, A.S. 2000. Natural Food Antimicrobial Systems. Boca Raton, FL: CRC Press. Ouattara, B., Simard, S., Piette, G.J.P., Holley, R.A. and Begin, A. 2000. “Diffusion of Acetic and Propionic Acids from Chitosan-based Antimicrobial Packaging Films,” J. Food Sci., 65:768–773. Ozdermir, M. 1999. Antimicrobial Releasing Edible Whey Protein Films and Coatings. Ph.D. dissertation, Purdue University, West Lafayette, IN. Padgett, T.M., Han, I.Y. and Dawson, P.L. 1995. “Incorporation of Lysozyme into Biodegradable Packaging Films,” Poultry Sci., 74(Suppl. 1):175. Padgett, T., Han, I.Y. and Dawson, P.L. 1998. “Incorporation of Food-Grade Antimicrobial Compounds into Biodegradable Packaging Films,” J. Food Protect., 61:1330–1335. Rico-Pena, D.C. and Torres, J.A. 1991. “Sorbic Acid and Potassium Sorbate Permeability of an Edible Methylcellulose-Palmitic Acid Film: Water Activity and pH Effects,” J. Food Sci., 56:497–499. Rodrigues, E.T. and Han, J.H. 2000. “Antimicrobial Whey Protein Films against Spoilage and Pathogenic Bacteria” in 2000 IFT Annual Meeting Book of Abstracts, Chicago, IL: Institute of Food Technologists, p. 191. Siragusa, G.R. and Dickson, J.S. 1992. “Inhibition of Listeria Monocytogenes on Beef Tissue by Application of Organic Acids Immobilized in a Calcium Alginate Gel,” J. Food Sci., 57:293–296. Siragusa, G.R. and Dickson, J.S. 1993. “Inhibition of Listeria Monocytogenes, Salmonella Typhimurium and Escherichia coli O157:H7 on Beef Muscle Tissue by Lactic or Acetic Acid Contained in Calcium Alginate Gels,” J. Food Safety, 13:147–158. Torres, J.A. and Karel, M. 1985. “Microbial Stability of Intermediate Moisture Food Surfaces. III. Effects of Surface Preservative Concentration and Surface pH Control on Microbial Stability of an Intermediate Moisture Cheese Analog,” J. Food Process. Preserv., 9:107–119. Torres, J.A., Motoki, M. and Karel, M. 1985. “Microbial Stabilization of Intermediate Moisture Food Surfaces. I. Control of Surface Preservative Concentration,” J. Food Process. Preserv., 9:75–92. Vojdani, F. and Torres, J.A. 1990. “Potassium Sorbate Permeability of Methylcellulose and Hydroxypropyl Methylcellulose Coatings: Effect of Fatty Acids,” J. Food Sci., 55:841–846. Wagner, M.K. and Moberg, L.J. 1989. “Present and Future Use of Traditional Antimicrobials,” Food Technol., 43(1):143–147, 155.

Use of Proteins in Edible Coatings for Whole and Minimally Processed Fruits and Vegetables1

DIBLE coatings have been used on fruits and vegetables since the twelfth century in China (Hardenburg, 1967). Traditionally, coatings have been used to reduce water loss and improve appearance of whole produce such as apple and citrus. Certain coatings have been shown to improve the appearance of products by imparting gloss (Hagenmaier and Baker, 1994), intensifying peel color (Perkins-Veazie and Collins, 1991), and reducing pigment degradation (Chen et al., 1996) or undesirable pigment formation (Baldwin et al., 1996). Coatings are also being used on fruit that continues to ripen after harvest (climacteric fruit) to delay ripening and thereby extend product postharvest life or “shelf life.” Prolonged shelf life expands shipping distances from harvest sites and extends strorage and marketing periods. Delayed ripening can also mean shipment of produce by sea instead of air, which reduces transport costs and, thus, prices for consumers. Coating permeability, which is important to coating performance, is affected by the materials used in coating formulations. Protein materials have

E

1Mention of a trademark or proprietary product is for identification only and does not imply a guarantee or warranty of the product by the U.S. Department of Agriculture. Mention of brand or firm names does not constitute an endorsement by the U.S. Department of Agriculture over others of a similar nature not mentioned.

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502 been used as main film-formers and as supplemental ingredients that alter film structure and therefore permeability, appearance, and general performance of the final coating. Protein coatings have the added benefit of offering nutritional value, giving a more wholesome appeal.

To understand the development and function of edible coatings for fresh fruit and vegetable products, it is necessary to understand fresh fruit and vegetable postharvest physiology. These products continue to undergo metabolic reactions after harvest, resulting in ripening and/or senescence. Fresh produce requires oxygen (O2) for respiratory metabolism and releases carbon dioxide (CO2) and water (H2O) as by-products (glucose + O2 CO2 + H2O). Fruits and vegetables also synthesize the gaseous hormone ethylene, which is required by climacteric fruits to initiate ripening and eventually causes senescence and shortens shelf life (Wills et al., 1981; Baldwin, 1994). Synthesis of ethylene requires oxygen and is inhibited by relatively high levels of CO2. Both processes are affected by the modified atmosphere created by a semi-permeable barrier as occurs when produce is packaged or coated. In such situations, O2 is not freely replaced, nor can the CO2 produced freely escape. This results in a “modified atmosphere” (MA) within the package or coating of relatively high CO2 and low O2. Protein coatings, for example, present an effective semi-permeable barrier to these respiratory gases and can lead to establishment of MA when applied to fresh produce. This MA slows down metabolism, including respiration and ethylene production, and subsequently retards ripening and senescence (Kader, 1986). The end result is an extension of produce shelf life. However, if O2 levels fall too low, the product may convert to anaerobic respiration, which results in ethanol production, subsequent off-flavor, and accelerated deterioration (Kader, 1986). In addition, if CO2 levels become too high, CO2-injury can occur, which is manifested in various postharvest disorders (e.g., impaired ripening, discoloration, and deterioration) (Shewfelt, 1992). Fruits and vegetables have a high water content, and the relative humidity (RH) in the intercellular spaces of most fresh produce is generally 100%. Therefore, fruits and vegetables lose water to the storage atmosphere in the form of water vapor. This results in weight loss, shriveling, shrinking, and/or wilting of fresh produce (Franklin, 1961; Woods, 1990). The outer coating of fruits and vegetables, called the cuticle, is comprised of wax-like material. The purpose of this waxy outer layer is to protect the fruit surface and to retard water loss. This natural coating is often partially removed during washing of fruits after harvest, which necessitates the application of a replacement or supplemental coating. Hydrophobic coating materials, such as waxes or oils, generally present a more effective barrier to water vapor than do hydrophilic compounds such as proteins (Kester and Fennema, 1986; Gennadios and Weller, 1990).

Types of Coatings Used on Fruits and Vegetables

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Lightly or minimally processed fruits and vegetables (also called “fresh-cuts”) include shredded, diced, sliced, peeled, cored, trimmed, or otherwise altered fruit and vegetable products that have not been frozen or cooked and are, thus, still in their fresh state (Rolle and Chism, 1987; Shewfelt, 1987). This subcategory of fresh produce is one of the fastest growing in the industry and presents special problems for storage and marketing. The additional processing removes some, if not all, of the natural protective outer coating and peel, which results in not only enhanced water loss problems, but also opens an avenue for microbial contamination. Removal of the outer coating or peel also allows entrance of O2 and subsequent oxidation reactions. These reactions often produce undesirable color changes, such as discoloration of the cut surface of apples and carrots (Tatsumi et al., 1991; Brecht et al., 1993). Furthermore, minimal processing “wounds” the fruit tissue resulting in plant defense responses such as increased production of ethylene; increased production of enzymes that alter fruit cell wall structure and subsequently texture; and de novo synthesis of secondary metabolites of various types, some of which can cause off-flavors and odors and undesirable color changes (Bolin and Huxsoll, 1991; Wong et al., 1994). Application of coatings to these products is tricky because they often present a wet surface of broken cellular material. If the surface is not too moist, then hydrophilic coatings, such as protein or protein-carbohydrate formulations, adhere better to the cut surface than do hydrophobic materials (Baldwin et al., 1995a, b). Respiration is also affected by storage temperature, in that rates increase with increasing temperature (Zagory and Kader, 1988). Temperature abuse of fresh produce can occur during transit (e.g., refrigeration breakdown and standing on loading docks) or marketing (e.g., display at super markets). Coatings or packaging materials designed to allow appropriate permeability for normal produce respiration in chilled storage may cause products to go anaerobic at warmer temperatures (Cameron et al., 1995).

Fruit coatings have been formulated from several major chemical classes including lipids, resins, carbohydrates, and proteins. The different classes of materials have advantages and disadvantages when used as film-formers or as ingredients of coatings. For this reason, most commercial fruit coatings contain a mixture of components and, thus, are called composite coatings (Kester and Fennema, 1986). Understanding the characteristics of the different available materials for use in coatings is needed to combine compatible and complimentary components for optimal coating performance. Interestingly, fruit and vegetable coatings are generically referred to as “waxes” even though some do not contain any wax components. Generally, lipid-based coatings offer resistance to water vapor and, there-

504 fore, reduce water loss from coated fresh produce. This, in turn, reduces weight loss, shriveling, and shrinkage of fruit and vegetable products, which, if left unchecked, render these commodities unmarketable. These coatings, however, are relatively permeable to gases and, therefore, result in less modification of the fruit internal atmosphere. As a result, they have less of an effect on ripening and senescence. On the other hand, lipid coatings are less likely to result in anaerobic conditions and the accompanying off-flavors (Davis and Hofmann, 1973; Hagenmaier and Baker, 1993). Lipid-based formulations based on carnauba, candelilla, or polyethylene waxes can form fairly high-gloss coatings and are used on apples and citrus fruits where high shine boosts sales (Baldwin et al., 1997). Resin-based coatings include the materials shellac and wood rosin (Hernandez, 1994). The latter is only allowed for use on citrus products where the peel is generally not ingested. These coatings impart a very glossy, high shine appearance to coated products and are used mainly on apples and citrus for this reason. Resin coatings are fairly effective at reducing water loss, but are the least permeable to gases among the available coating film-formers (Hagenmaier and Shaw, 1992). This means that fruit can easily undergo anaerobic respiration and flavor changes that are usually undesirable. Some climacteric fruit do not tolerate resin coatings at all due to impaired ripening from the severe MA created by these materials. Protein and carbohydrate coatings are intermediate in gas barrier properties between lipid and resin coatings. They generally offer limited resistance to water vapor transfer; however, due to their hydrophilic nature, they are not effective in reducing water loss (Kester and Fennema, 1986; Gennadios and Weller, 1990; Park and Chinnan, 1990; Gennadios et al., 1994). Certain protein materials containing higher levels of hydrophobic amino acids present more effective moisture barriers, especially in combination with lipids (soy protein, casein). Protein and carbohydrate coatings are more effective for ripening control via creation of a gentle MA. Protein films can impart a nice sheen to coated products. But of all the food-grade derivatives available, only the corn protein, zein, can result in a high-gloss appearance (Gennadios and Weller, 1990) that equals or even surpasses that of resin-based coatings.

Proteins used in coatings include zein, soy protein, wheat gluten, whey protein, casein, gelatin, and collagen. Reconstituted collagen has long been used for extruded casings on meat products, and zein has been used commercially on confectionery and nut products (Turner, 1993; Gennadios et al., 1994). Proteins reported for use in coatings for fresh produce include zein, soy, wheat gluten, and gelatin. Zein, a protein from corn endosperm, is a prolamin (aqueous alco-

Factors that Affect Coating Permeability

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hol-soluble protein). It consists of monomers and disulfide-linked oligomers of varying molecular weight. Zein is only soluble in water at very high or low pH and is insoluble in anhydrous alcohols due to its amino acid composition (mostly non-polar leucine, proline, and alanine as well as glutamine) (Gennadios and Weller, 1990; Gennadios et al., 1994). Soybeans have a relatively high protein content. Soy proteins are classified by relative sedimentation rates. The main fractions are 11S and 7S globulins (saline-soluble) (Okamoto, 1978) with 2S and 15S fractions making up the other globulin fractions (Gennadios and Weller, 1991). Soy protein is available as soy concentrate (70% protein) or soy isolate (>90% protein). The amino acids are quite evenly represented in soy protein with glutamic and aspartic acids being somewhat more prominent (Gennadios et al., 1994). When soy-milk is heated, a surface film forms that is a mixture of soy protein and oil (Okamoto, 1978). Gluten is derived from wheat kernels and falls mainly into the prolamin and glutelin classifications called gliadin and glutenin, respectively. This protein is very elastic, hence its usefulness in making dough (Gennadios et al., 1994). It is characterized by a high content of glutamic acid, mainly in the form of glutamine. It also contains cystine, which is partly responsible for its film-forming ability via disulfide bonds (Gennadios and Weller, 1990). These films are good barriers to O2 and CO2 but are highly permeable to water vapor. Gelatin is hydrolyzed collagen, which is derived from animal skin, tendon, and connective tissues. Collagen and gelatin contain relatively high contents of glycine, proline, and hydroxyproline (Gennadios et al., 1994). This protein lacks internal order and exhibits a random configuration of polypeptide chains in aqueous solutions. Gelatin forms strong gels, which has led to its use as an encapsulating medium by the pharmaceutical and dietary supplement industries. Gelatin films have low permeability to O2 and offer a moderate moisture barrier for a protein (Anonymous, 1992). Milk proteins are excellent emulsifiers (McHugh and Krochta, 1994a) and are therefore good candidates for composite protein-lipid emulsion coatings. Milk contains two main classes of proteins: caseins (80%) and whey proteins (20%). Caseins are mostly phosphoproteins characterized by low cysteine levels and, therefore, a paucity of disulfide bonds. Whey proteins are mainly lactalbumin and lactoglobulin and are commercially available as whey protein isolates (>80% protein) and concentrates (25–80% protein) (Gennadios et al., 1994).

Factors affecting coating permeability that are relevant to fruit storage are ingredients, temperature, humidity, and interaction with the fruit cuticle and stomates. Fresh produce is often stored at low temperature (anywhere from just

506 above freezing to 15 C) prior to marketing. Marketing temperatures can vary from 12 C to ambient. Temperature is probably the least important factor affecting coating permeability, but, nevertheless, increased temperature increases mobility of polymer chains, thus increasing permeability (McHugh and Krochta, 1994b). Plasticizers are added to coating formulations to increase elasticity of filmformers. This helps to reduce coating brittleness and breakage, which is observed as flaking or peeling of coatings from the coated surface (Kester and Fennema, 1986). Plasticizers are generally small molecules such as polyols (e.g., sorbitol, glycerol, and propylene glycol) that intersperse and intercalate among and between polymer chains, disrupting hydrogen bonding and spreading the chains apart, which not only increases flexibility, but also permeability (Kester and Fennema, 1986; McHugh and Krochta, 1994b). Water vapor permeability is more affected by plasticizers than gas permeability (Park and Chinnan, 1990; McHugh and Krochta 1994b, c). Water vapor plasticizes coatings and increases coating permeability. This is especially true of hydrophilic coatings such as those made from carbohydrates and proteins. For example, the effect of RH on permeability properties of soy and wheat gluten films showed that transmission rates for O2 and CO2 increased sharply from 60 to >90% RH (Guilbert et al., 1996). Fruits and vegetables are often stored under conditions of high RH (85–99%) to reduce, as much as possible, the water vapor gradient compared to inside the fruit (100% RH), thereby reducing water loss from fresh produce via transpiration. This high storage RH can generally increase fruit coating permeability (Donhowe and Fennema, 1992; Lerdthanangkul and Krochta, 1996), thus reducing the ability of coatings to create MA within the fruit or vegetable. Permeability to gases for films made with zein or wheat protein were also determined to be a function of film thickness (Park and Chinnan, 1990), whereas this is not always true for polysaccharide films (Park et al., 1993). Film-forming polymer structure affects barrier properties of edible films. Highly polar polymers, including many proteins and polysaccharides, are capable of hydrogen bonding, thus forming low gas permeability films at low RH (McHugh and Krochta, 1994b). Establishment of cross-links between film-forming polymers can also affect coating permeability (McHugh and Krochta, 1994b). This has been shown with protein-carbohydrate coatings along with manipulation of coating pH (Baldwin et al., 1996). Increasing or decreasing the pH away from the protein isoelectric point resulted in denaturation and unfolding of proteins, which, theoretically, would allow more hydrogen bonding between protein and film-former polymer functional groups. The increased bonding between proteins and film-former chains decreased coating permeability to water vapor. On the other hand, adjustment of pH to 4.6 (isoelectric point of casein) with calcium ascorbate buffer in combination with calcium ion cross-linking improved water barrier characteristics of calcium caseinate films (Avena-Bustillos and Krochta, 1993).

Protein-Based Coatings for Fresh Fruits and Vegetables

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Combining hydrophilic materials, such as proteins, with lipids to prepare composite films can enhance water barrier properties. For example, acetylated monoglycerides or beeswax were combined with casein to form emulsion coatings (Krochta et al., 1990a, b; Avena-Bustillos and Krochta, 1993; Avena-Bustillos et al., 1993). Addition of these lipid materials reduced casein film permeability to water vapor.

Peeled carrots undergo a discoloration or whitening of the peeled surface, referred to as “white blush,” which is determined using a colorimeter and a whiteness index. This is thought to be associated with dehydration (Tatsumi et al., 1991) and possibly lignin formation (Bolin and Huxsoll, 1991) as a wound response to the peeling process. Casein-monoglyceride films applied to peeled carrots increased the water vapor resistance of that product, thereby reducing white blush (Avena-Bustillos and Krochta, 1993). The whitish index of carrots was estimated (Judd, 1963) from L, a, and b measurements on the CIE color scale by a Minolta Chromameter. A sodium caseinate/stearic acid emulsion coating reduced the whitish index from 48 for uncoated carrots to 42 after 14 days at 10 C (Figure 20.1) and the respiration rate from 4.75 mL CO2/kg h for uncoated to 3.74 mL CO2/kg h for coated carrots after 30 days at 2.5 C (Avena-Bustillos et al., 1993, 1994a). Similar coatings increased water vapor resistance (decreased water loss) of coated compared to uncoated peeled carrots (Figure 20.2). Calcium caseinate-acetylated monoglyceride aqueous emulsions (2.5–7.0% total solids) increased water vapor resistance of coated zucchini (Avena-Bustillos et al., 1994b). Coating bell peppers with whey protein isolate, sodium caseinate, or sodium caseinate/beeswax emulsion did not result in MA due to the high humidity storage conditions (80–85% RH), although these proteins were good gas barriers at low RH (Lerdthanangkul and Krochta, 1996). This demonstrates the necessity of testing coatings under simulated commercial storage conditions for a product. A coating of gum acacia and gelatin (Sealgum, Colloides Naturels Inc., Bridgewater, NJ) inhibited darkening of potatoes after cooking (Mazza and Qi, 1991), while a carboxymethyl cellulose/soy protein coating carrying antioxidants and acidulants reduced browning of fresh cut potato (Baldwin et al., 1996). Zein coatings delayed color change, softening, and weight loss of tomato fruit, extending their shelf life by 6 days (Park et al., 1994). No adverse off-flavors were noted and no ethanol was produced, indicating the fruit maintained aerobic respiration.

Figure 20.1 Whitish index during storage of coated peeled carrots packaged in LDPE bags ( 38 m thick) at 10 C. Edible coating formulations: T1 calcium caseinate/beeswax; T2 calcium caseinate/stearic acid; T3 calcium caseinate/acetylated monoglyceride; T4 sodium caseinate/beeswax; T5 sodium caseinate/stearic acid; and T6 sodium caseinate/acetylated monoglyceride. Reprinted from Postharvest Biology & Technology, Volume 4, Avena-Bustillos et al., “Optimization of Edible Coatings on Minimally Processed Carrots Using Response Surface Methodology,” p. 326, Copyright 1994, with permission from Elsevier Science.

Figure 20.2 Water vapor resistance of coated and uncoated peeled carrots (2.5 C, 70% RH, and 20 m/min). Letters indicate significant (P < 0.05) difference. Bars are standard deviations. Treatment coatings: 1 calcium caseinate/beeswax; 2 calcium caseinate/stearic acid; 3 calcium caseinate/acetylated monoglyceride; 4 sodium caseinate/beeswax; 5 sodium caseinate/stearic acid; 6 sodium caseinate/acetylated monoglyceride; and 7 uncoated control. Reprinted from Postharvest Biology & Technology, Volume 4, Avena-Bustillos et al., “Optimization of Edible Coatings on Minimally Processed Carrots Using Response Surface Methodology,” p. 325, Copyright 1994, with permission from Elsevier Science.

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Protein coatings have been applied to raisins where water loss is a problem. Casein coatings did not reduce water loss (Watters and Brekke, 1961), but egg albumen and soy protein reduced moisture loss from raisins stored in bran flake cereal (Bolin, 1976). Dried fruit has been successfully coated with an amylose ester of a fatty acid/protein bilayer coating that included zein to reduce stickiness (Gunnerson and Bruno, 1990). Zein-based coatings (14 to 20% zein) were developed for whole apple fruit as an alternative to shellac for a high-gloss fruit coating (Alleyne et al., 1999). The zein coating resulted in fruit firmness and weight loss values similar to those of commercial shellac- and carnauba-coated apples (Table 20.1). Gloss values fell below those of shellac after 14 days at marketing conditions, yet remained higher than carnauba wax-coated apples. The internal atmosphere was less modified for zein than for shellac-coated apples and was not significantly different in the zein-coated fruit from uncoated controls. Internal ethylene levels were significantly higher in shellac-coated fruit and in fruit coated with experimental coatings compared to uncoated controls. Zein- and carnauba wax-coated fruit internal ethylene levels, however, were not significantly different from control (uncoated) fruit. The problem with both shellac and especially zein coatings is that they tend to whiten (“blush”) when in contact with water. This occurs when apples come out of cold storage and water vapor in the

510 air condenses on the cold apple surface. Other formulations made with less zein (5–10%) did not blush but exhibited lower gloss values than those with higher zein content, although they were still comparable to a commercial shellac product (Alleyne et al., 1999). Casein-lipid (acetylated monoglyceride) emulsion films resulted in improved film strength, integrity, and ability to adhere to fruit surfaces. These coatings retarded water loss and browning in cut apple (Krochta et al., 1990a). The casein/lipid idea was further explored using acetylated monoglyceride, stearate, or beeswax. Beeswax was the most effective lipid for reducing water vapor permeability, perhaps due to its crystalline structure. Calcium ion cross-linking also reduced water vapor permeability of caseinate-based films, probably as the result of reduced protein polymer segmental mobility and protein solubility in water (Avena-Bustillos and Krochta, 1993). Various carboxymethyl cellulose-soy protein concentrate formulations reduced browning of cut apple, microbial counts, and water loss (Baldwin et al., 1996).

Addition of protein to carbohydrate coatings can affect permeability to water vapor depending on protein type. For example, protein isolate of Pistacia terebinthus L. (pistachio relative), a highly hydrophilic protein, increased solubility, but decreased moisture barrier ability, of hydroxypropyl methylcellulose films (Ayranci and Cetin, 1995). Thickness of such films also increased as a result of water retention within the structure by protein hydrophilic groups (Ayranci and Cetin, 1995). In contrast, when soy protein concentrate was added to a carboxymethyl cellulose film, the water vapor permeability decreased along with water loss from coated apple discs. Soy peptone (a peptide) was even more effective (Baldwin et al., 1996). Adjustment of coating pH away from the isoelectric point of the protein resulted in even lower weight loss of apple discs (Table 20.2). A coating (Nature Seal , EcoScience Corp., Orlando, FL) was designed and tested on cut apples where it reduced browning of the cut surface (Figure 20.3). There was a problem of protein solubility in the cellulose matrix, which was improved by addition of xanthan gum. Reportedly, xanthan gum improved solubility and emulsification properties of soy protein isolate (Xie and Hettiarachchy, 1997). However, the gum also increased viscosity, resulting in substantially thick coatings. Different proteins were added to a candelilla formulation to prepare wax microemulsion coatings (Hagenmaier and Baker, 1996) with higher gloss characteristics. Soy protein is occasionally added to commercial coatings to

Protein as an Ingredient of Fruit and Vegetable Coatings

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improve shine. In the study by Hagenmaier and Baker (1996), gelatin, soy protein isolate, and zein were added, but only soy isolate and gelatin improved gloss. A composite coating of alginic acid (a polysaccharide polymer of mannosyluronic and gulosyluronic acids, which contains carboxyl groups), casein, and a lipid (acetylated monoglyceride) that cross-links with the addition of calcium into a three-dimensional network was developed by the USDA Agricultural Research Service. Casein is contained in the cross-linked polysaccharide matrix with the lipid dispersed in the interstices. This type of coating may also cross-link with the carboxyl groups on the endemic pectin on the cut surface of fruit, such as apple, if the cut fruit is first dipped in a calcium solution. On cut apple stored at 23 C and 50% RH, this coating increased water vapor resistance by 92%, decreased respiration by 70%, and decreased ethylene evolution by 90% (Wong et al., 1994).

Figure 20.3 Effect of water (H2O, ✚), antioxidants (0.5% ascorbic acid and 0.02% TBHQ), preservatives (0.1% sodium benzoate), and acidulants (citric and other acids) in aqueous solution (AQ.SOL, ❍) or as additives to the CMC-based Nature Seal® coating (NS 1020, ●) on brown color development of apple (A) or potato (B) cores during two weeks of storage at 4 C. Data are means of five samples cut from the same fruit/tuber, each read three times, and expressed as ‘a*’Chromameter values and % inhibition of browning compared to water-treated controls. Visual rating scale: 1, no browning, 5, severe browning. Reprinted from Postharvest Biology & Technology, Volume 9, Baldwin et al., “Improving Storage Life of Cut Apple and Potato with Edible Coating,” p. 157, Copyright 1996, with permission from Elsevier Science.

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References

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Coated intact fruit and vegetables should be labeled on their marketing display identifying the coating as a vegetable-, animal-, resin-, petroleum-, or beeswax-based coating (Lecos, 1982; Anonymous, 1993). This is important because a small portion of the population has intolerances and/or allergic reactions to certain proteins. For example, gluten intolerance (celiac disease) causes poor absorption of nutrients due to toxic effects of gluten (Gennadios et al., 1994). Milk proteins contained in whey and casein fractions can cause allergic reactions. Furthermore, whey protein concentrates and caseinates may contain elevated levels of lactose. Certain individuals are lactose intolerant due to low levels of the enzyme lactase (Gennadios et al., 1994). Finally, vegetarians would be concerned about consumption of animal-derived protein coatings, such as those that contain gelatin and possibly milk protein. Therefore, proper labeling of products enrobed in protein coatings is necessary.

Alleyne, V., Baldwin, E.A. and Hagenmaier, R.D. 1999. Unpublished data. Citrus & Subtropical Products Laboratory, Agricultural Research Service, U.S. Department of Agriculture, Winter Haven, FL, USA. Anonymous. 1992. “Ingredient Capsules Beg Processing Solutions,” Prepared Foods, 161(12):81. Anonymous. 1993. “Waxes: A Common Sense Rule,” Citrograph (Feb.). Avena-Bustillos, R.J. and Krochta, J.M.. 1993. “Water Vapor Permeability of Caseinate-Based Edible Films as Affected by pH, Calcium Crosslinking and Lipid Content,” J. Food Sci., 58:904–907. Avena-Bustillos, R.J., Cisneros-Zevallos, L.A., Krochta, J.M. and Saltveit, M.E., Jr. 1993. “Optimization of Edible Coatings on Minimally Processed Carrots Using Response Surface Methodology,” Trans. ASAE, 36:801–805. Avena-Bustillos, R.J., Cisneros-Zevallos, L.A., Krochta, J.M. and Saltveit, M.E., Jr. 1994a. “Application of Casein-Lipid Edible Film Emulsions to Reduce White Blush on Minimally Processed Carrots,” Postharvest Biol. Technol., 4:319–329. Avena-Bustillos, R.J., Krochta, J.M., Saltveit, M.E., Jr., Rojas-Villegas, R.J. and Sauceda-Perez, J.A. 1994b. “Optimization of Edible Coating Formulation on Zucchini to Reduce Water Loss,” J. Food Engr., 21:197–214. Ayranci, E. and Cetin, E. 1995. “The Effect of Protein Isolate of Pistacia terebinthus L. on Moisture Transfer Properties of Cellulose-based Edible Films,” Lebens. Wiss Technol., 28:241–244. Baldwin, E.A. 1994. “Edible Coatings for Fresh Fruits and Vegetables: Past, Present, and Future” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster PA: Technomic Publishing Co., Inc., pp. 25–64. Baldwin, E.A., Nisperos-Carriedo, M.O. and Baker, R.A. 1995a. “Edible Coatings for Lightly Processed Fruits and Vegetables,” HortSci., 30:35–38. Baldwin, E.A., Nisperos-Carriedo, M.O. and Baker, R.A. 1995b. “Use of Edible Coatings to Preserve Quality of Lightly (and Slightly) Processed Products,” Critical Rev. Food Sci. Nutr., 35:509–524. Baldwin, E.A., Nisperos, M.O., Chen, X. and Hagenmaier, R.D. 1996. “Improving Storage Life of Cut Apple and Potato with Edible Coating,” Postharvest Biol. Technol., 9:151–163.

514 Baldwin, E.A., Nisperos, M.O., Hagenmaier, R.D. and Baker, R.A. 1997. “Use of Lipids in Edible Coatings for Food Products,” Food Technol., 51(6):56–62. Bolin, H.R. 1976. “Texture and Crystalization Control in Raisins,” J. Food Sci., 41:1316–1319. Bolin, H.R. and Huxsoll, C.C. 1991. “Control of Minimally Processed Carrot (Daucus carota) Surface Discoloration Caused by Abrasion Peeling,” J. Food Sci., 56:416–418. Brecht, J.K., Sabaa-Srur, A.U.O., Sargent, S.A. and Bender, R.J. 1993. “Hypochlorite Inhibition of Enzymic Browning of Cut Vegetables and Fruit,” Acta Hort., 343:341–344. Cameron, A.C., Talasila, P.C. and Joles, D.W. 1995. “Predicting Film Permeability Needs for Modified Atmosphere Packaging of Lightly Processed Fruits and Vegetables,” HortSci., 30:25–34. Chen, X., Campbell, C.A., Grant, L.A., Li, P. and Barth, M. 1996. “Effect of Nature Seal on Maintaining Carotene in Fresh-Cut Carrots, Proc. Fla. State Hort. Soc., 109:258–259. Davis, P.L. and Hofmann, R.C. 1973. “Effects of Coating on Weight Loss and Ethanol Buildup in Juice of Oranges,” J. Agric. Food Chem., 21:455–458. Donhowe, I.G. and Fennema, O. 1992. “The Effect of Relative Humidity Gradient on Water Vapor Permeance of Lipid and Lipid-Hydrocolloid Bilayer Films,” JAOCS, 69:1081–1087. Franklin, E.W. 1961. “The Waxing of Turnips for the Retail Market,” Canada Dept. of Agric., Publication No. 1120, 3 pp. Gennadios, A. and Weller, C.L. 1990. “Edible Films and Coatings from Wheat and Corn Proteins,” Food Technol., 44(10):63-69. Gennadios, A. and Weller, C.L. 1991. “Edible Films and Coatings from Soymilk and Soy Protein,” Cereal Foods World, 36:1004-1009. Gennadios, A., McHugh, T.H., Weller, C.L. and Krochta, J.M. 1994. “Edible Coatings and Films Based on Proteins” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster PA: Technomic Publishing Co., Inc., pp. 201–277. Guilbert, S., Gontard, N. and Gorris, L.G.M. 1996. “Prolongation of the Shelf-Life of Perishable Food Products Using Biodegradable Films and Coatings,” Lebens. Wiss. Technol., 29:10–17. Gunnerson, R.E. and Bruno, R.C. August 7, 1990. U.S. patent 4,946,694. Hagenmaier, R.D. and Baker, R.A. 1993. “Reduction in Gas Exchange of Citrus Fruit by Wax Coatings,” J. Agric. Food Chem., 41:283–287. Hagenmaier, R.D. and Baker, R.A. 1994. “Internal Gases, Ethanol Content and Gloss of Citrus Fruit Coated with Polyethylene Wax, Carnauba Wax, Shellac or Resin at Different Application Levels,” Proc. Fla. State Hort. Soc., 107:261–265. Hagenmaier, R.D. and Baker, R.A. 1996. “Edible Coatings from Candelilla Wax Microemulsions,” J. Food Sci., 61:562–565. Hagenmaier, R.D. and Shaw, P.E. 1992. “Gas Permeability of Fruit Coating Waxes,” J. Amer. Soc. Hort. Sci., 117:105–109. Hardenburg, R.E. 1967. “Wax and Related Coatings for Horticultural Products. Bibliography,” Agr. Res. Bull., 51-15, U.S. Dept. of Agric., Washington, DC. Hernandez, E. 1994. “Edible Coatings from Lipids and Resins” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin, and M.O. Nisperos-Carriedo, Lancaster PA: Technomic Publishing Co., Inc., pp. 279–303. Judd, D.B. 1963. Color in Business, Science and Industry. New York: John Wiley, pp. 299-300. Kader, A.A. 1986. “Biochemical and Physiological Basis for Effects of Controlled and Modified Atmospheres on Fruits and Vegetables,” Food Technol., 40(5):99–104. Kester, J.J. and Fennema, O.R. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Krochta, J.M., Hudson, J.S. and Avena-Bustillos, R.J. 1990a. “Casein-Acetylated Monoglyceride Coatings for Sliced Apple Products,” Presented at the 1990 Annual Meeting of the Institute of Food Technologists, June 16–20, Anaheim, CA. Krochta, J.M., Pavlath, A.E. and Goodman, N. 1990b. “Edible Films from Casein-Lipid Emulsions for Lightly-Processed Fruits and Vegetables: Preservation, Process and Related Techniques” in

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Engineering and Food, Vol. 2, ed., W.E.L. Spiess, London: Elsevier Applied Science, pp. 329–340. Lecos, C. 1982. “How to Shine an Apple,” FDA Consumer (February):8–11. Lerdthanangkul, S. and Krochta, J.M. 1996. “Edible Coating Effects on Postharvest Quality of Green Bell Peppers,” J. Food Sci., 61:176–179. Mazza, G. and Qi, H. 1991. “Control of After-Cooking Darkening in Potatoes with Edible Film-Forming Products and Calcium Chloride,” J. Agric. Food Chem., 39:2163–2166. McHugh, T.H. and Krochta, J.M. 1994a. “Milk-Protein-Based Edible Films and Coatings,” Food Technol., 48(1):97–103. McHugh, T.H. and Krochta, J.M. 1994b. “Permeability Properties of Edible Films” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster PA: Technomic Publishing Co., Inc., pp. 139–187. McHugh, T.H. and Krochta, J.M. 1994c. “Sorbitol- vs. Glycerol-Plasticized Whey Protein Edible Films: Integrated Oxygen Permeability and Tensile Property Evaluation,” J. Agric. Food Chem., 42:841–845. Okamoto, S. 1978. “Factors Affecting Protein Film Formation,” Cereal Foods World, 25:256–262. Park, H.J. and Chinnan, M.S. 1990. “Properties of Edible Coatings for Fruits and Vegetables,” ASAE Paper No. 90-6510, 19 pp. Park, H.J., Weller, C.L., Vergano, P.J. and Testin, R.F. 1993. “Permeability and Mechanical Properties of Cellulose-Based Edible Films,” J. Food Sci., 58:1361–1364, 1370. Park, H.J., Chinnan, M.S. and Shewfelt, R.L. 1994. “Edible Coating Effects on Storage Life and Quality of Tomatoes,” J. Food Sci., 59:568–570. Perkins-Veazie, P.M. and Collins, J.K. 1991. “Color Changes In Waxed Turnips During Storage,” J. Food Qual., 14:313–319. Rolle, R.S. and Chism, G.W., III. 1987. “Physiological Consequences of Minimally Processed Fruits and Vegetables,” J. Food Qual., 10:157–177. Shewfelt, R.L. 1987. “Quality of Minimally Processed Fruits and Vegetables,” J. Food Qual., 10:143–156. Shewfelt, R.L. 1992. “Stress Physiology: A Cellular approach to Quality” in Postharvest Handling, a Systems Approach, eds., R.L. Shewfelt and S.E. Prussia, New York: Academic Press, Inc., pp. 258–276. Tatsumi, Y., Watada, A.E. and Wergin, W.P. 1991. “Scanning Electron Microscopy of Carrot Stick Surface to Determine Cause of White Translucent Appearance,” J. Food Sci., 56:1357–1359. Turner, E. 1993. “Edible Packaging,” Sustain Notes, 5(1):1–4. Watters, G.G. and Brekke, J.E. 1961. “Stabilized Raisins for Dry Cereal Products,” Food Technol., 15(5):236–238. Wills, R.H.H., Lee, T.H., Graham, D., McGlasson, W.B. and Hall, E.G. 1981. Postharvest: An Introduction to the Physiology and Handling of Fruit and Vegetables. Westport, CT: AVI Publishing Co., Inc., pp. 1–150. Wong, D.W.S., Camirand, W.M. and Pavlath, A.E. 1994. “Development of Edible Coatings for Minimally Processed Fruits and Vegetables” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin, and M.O. Nisperos-Carriedo, Lancaster PA: Technomic Publishing Co., Inc., pp. 65–88. Woods, J.L. 1990. “Moisture Loss from Fruits and Vegetables,” Postharvest News and Information, 1:195–199. Xie, Y.R. and Hettiarachchy, N.S. 1997. “Xanthan Gum Effects on Solubility and Emulsification Properties of Soy Protein Isolate,” J. Food Sci., 62:1101–1104. Zagory, D. and Kader, A.A. 1988. “Modified Atmosphere Packaging for Fresh Produce,” Food Technol., 42(9):70–77.

Protein-based Edible Coatings for Fried Foods

T

HE film-forming properties of several proteins have been utilized in devel-

oping edible, protective films and coatings (Gennadios et al., 1994). Currently, edible coatings formed by proteins are being investigated for applications on a broad range of food products (e.g., processed meats, seafood, nuts, confections, fruits, vegetables, and eggs). Due to their film-forming ability, cohesiveness, adhesiveness, and lipid barrier ability, proteins form coating layers that can preserve and improve the quality of food products. Potential advantages of using protein-based edible films and coatings on deep-fat fried foods are discussed in this chapter. Deep-fat frying is a widely used method for preparing foods with an attractive and tasty surface. The soft, moist interior and the porous, crispy crust increase food palatability. In most industrialized countries, deep-fat frying is one of the most rapidly growing culinary techniques (Varela and Ruiz-Roso, 1992). In the U.S., the fried food business is flourishing. More than 500,000 institutional and commercial restaurants are involved in deep-fat frying operations. Fried chicken is a $6 billion, fast-growing industry. Despite their popularity, deep-fat fried foods contribute greatly to dietary fat intake as they contain large amounts of absorbed fat (Saguy and Pinthus, 1995). Fats and oils are important sources of certain nutrients and food energy, while their smooth mouthfeel and flavor improve food palatability (Khalil, 1999). 517

518 However, reducing dietary fat intake to less than 30% of calories has been recommended by health organizations during the past several years (Saguy and Pinthus, 1995). This has generated interest in reducing the amount of absorbed oil during deep-fat frying without negative effects on other interrelated quality attributes such as taste, texture, and surface color (Hansen, 1998). Prior to deep-fat frying, products often are battered and/or breaded. Cunningham and Suderman (1981) defined a batter as “liquid mixture composed of water, flour, starch, and seasonings into which food products are dipped prior to cooking.” Also, they defined a breading as “dry mixture of flour, starch, and seasonings of coarse composition, and applied to moistened or battered food products prior to cooking” (Cunningham and Suderman, 1981). Proteins (e.g., egg albumen, gelatin, and soy protein) and polysaccharides (e.g., xanthan gum, alginates, and guar gum) have been incorporated into traditional batter and breading formulations or used as dry predusts to promote adhesion of the outer crust onto deep-fat fried products (Baker et al., 1972; Toloday and Andres, 1975; Suderman et al., 1981; Hsia et al., 1992). In addition, edible coatings based on proteins and other hydrocolloids can play a role in reducing oil absorption during frying (Anonymous, 1997).

Corn zein, wheat gluten, soy protein, gelatin, whey protein, and casein are among the proteins that have been studied for their film-forming properties (Gennadios et al., 1994). In general, protein films contain plasticizers, such as glycerin or sorbitol, which reduce internal hydrogen bonding, thereby imparting flexibility to the films (McHugh and Krochta, 1994a). Corn grains have a protein content ranging from 7 to 11% (Wall and Paulis, 1978). Zein is the prolamin (aqueous alcohol-soluble) fraction of corn proteins. It is insoluble in water due to its high content of non-polar amino acids (Pomes, 1971). There has been a long-standing interest in zein due to its film-forming ability. Zein films are characterized as tough, glossy, scuff-resistant, and grease-resistant (Pomes, 1971). Wheat gluten is comprised of the prolamin and glutelin (alkali- and acid-soluble) fractions of the wheat proteins (Kasarda et al., 1976). Films can be prepared by casting and drying aqueous alcoholic solutions (in acidic or basic conditions) of wheat gluten (Cuq et al., 1998). Such factors as pH, gluten concentration, and ethanol concentration of film-forming solutions have strong interactive effects on film opacity, water solubility, and water vapor permeability (Gontard et al., 1992). Soybeans have a high content of proteins (38 to 44%), which are mainly classified as globulins (saline-soluble) (Snyder and Kwon, 1987). Films can be prepared on the surface of heated soymilk or by casting aqueous solutions of

Film Properties

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soy protein isolates (Gennadios and Weller, 1991). Gelatin is obtained from partial hydrolysis of collagen using porkskins and bovine hides and bones as principal raw materials (Alleavitch et al., 1989). It forms clear, flexible, and strong films when cast from aqueous solutions (Gennadios et al., 1994). Whey proteins produce transparent, flavorless, and flexible edible films that are water-insoluble due to the presence of covalent disulfide bonds (McHugh and Krochta, 1994a). Typically, such films are cast from heated (75 to 100ºC) aqueous whey protein solutions (8 to 12% w/w) (McHugh and Krochta, 1994a). Caseinates, when cast from aqueous solutions, readily form transparent, water-soluble films through extensive hydrogen bonding and electrostatic interactions (McHugh and Krochta, 1994a; Chen, 1995).

Protein-based films are good barriers to the transport of gases, though moderate barriers to the transport of moisture (Krochta, 1992). The limited moisture barrier ability of protein-based films is mainly due to the hydrophilic nature of proteins. The barrier and mechanical properties of protein films are affected by water absorption (Krochta and De Mulder-Johnston, 1997). Therefore, film conditioning is an important factor when characterizing protein films as relative humidity (RH), temperature, and time period of conditioning can substantially affect film properties. At low RH conditions (0 to 50%), protein films are good oxygen barriers (Gennadios et al., 1993; McHugh and Krochta, 1994b; Miller and Krochta, 1997). However, at higher RH, the oxygen barrier ability decreases due to the plasticization effect of absorbed water (McHugh and Krochta, 1994b; Miller and Krochta, 1997). Multicomponent films with improved water vapor barrier properties can be prepared by combining proteins or polysaccharides with lipids (Krochta and De Mulder-Johnston, 1997). For example, considerable research has been conducted on multicomponent films prepared from lipids and cellulose ethers (Kester and Fennema, 1986), whey protein (McHugh and Krochta, 1994c), casein (Avena-Bustillos and Krochta, 1993), wheat gluten (Gontard et al., 1994), or soy protein (Gennadios et al., 1998). Due to their hydrophilic nature, protein films can function as lipid barriers (Krochta and De Mulder-Johnston, 1997). This attribute is widely utilized in the encapsulation of lipophilic compounds within protein (mainly gelatin) capsules (Gennadios et al., 1994). Also, zein-coated paper had grease resistance comparable to that of polyethylene-coated paper (Trezza and Vergano, 1994). This lipid barrier ability of protein-based films can be of value in reducing fat adsorption by fried foods. Mallikarjunan et al. (1998) developed a method to estimate the lipid conductivity through edible coatings and films at frying temperatures. They used film-forming solutions prepared from corn zein (CZ) or hydroxypropyl

520

Figure 21.1 Effect of frying temperature and solids concentration on the lipid conductivity of corn zein (CZ) and hydroxypropyl methylcellulose (HPMC) edible films (data from Mallikarjunan et al., 1998).

methylcellulose (HPMC) at two different solids concentrations. For both CZ and HPMC films, increasing the solids concentration decreased lipid conductivity by reducing the rate of oil adsorption (Figure 21.1). With an increase in oil temperature, lipid conductivity also increased. HPMC films were better barriers to lipid transfer than CZ films (Mallikarjunan et al., 1998). The developed lipid conductivity method can be used to select a suitable edible film providing better lipid barrier properties during deep-fat frying. However, further investigations on estimation of lipid barrier properties of various protein-based coatings at frying temperatures are needed.

Deep-fat fried foods are immersed in oil and heated to frying temperatures between 155 and 200 C. A sudden increase in the surface temperature of the food causes rapid transfer of heat, moisture, and oil between the food and the surrounding oil. Heat is transferred to the product surface by convection and to the geometric center by conduction. During deep-fat frying, a crust layer forms. In addition to moisture transfer, frying oil is transferred from the medium to the product by diffusion, and fat from the product migrates to the medium due to capillary forces.

Application of Coatings on Fried Foods

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Figure 21.2 shows a schematic diagram representing the moisture and lipid transfer during frying of a product coated with an edible film. The edible film acts as a selective barrier for moisture removal and oil penetration. During frying, edible coatings form a thermally induced barrier between food and frying medium. In addition, the edible coatings provide stability and shape to the product and reduce loss of solids from the product surface during frying.

The use of hydrocolloid coatings to reduce oil absorption during frying and to improve textural characteristics of fried foods has received attention (Saguy and Pinthus, 1995). For example, treating fried potato strips with coatings based on starch, modified starches, or dextrins has been extensively discussed (Arenson, 1972; El-Hag and Shanbhag, 1982; El-Hag et al., 1985; Sloan et al., 1991; Calder et al., 1994; Busacker et al., 1995; Higgins et al., 1999). Recently, Khalil (1999) reported that pectin or sodium alginate coatings that were “fixed” with calcium cations effectively reduced oil uptake by french fried potatoes. The oil content of the french fries was even lower when a second coating of carboxymethyl cellulose was applied on top of the first coating (Khalil, 1999). Cellulose ethers and gellan gum are additional examples of film-forming polysaccharides that decreased oil uptake by coated potatoes and/or other foods during frying (Meyers and Conklin, 1990; Anonymous, 1997;

Figure 21.2 Schematic representation of reduction in oil uptake and of moisture retention during deep-fat frying of a food coated with an edible film. Reprinted from Balasubramaniam et al., 1995, with permission from ASAE.

522 Balasubramaniam et al., 1997; Williams and Mittal, 1999a, b). Coatings based on synthetic polymeric materials, such as polyvinylpyrrolidone (Prosise, 1990) and colloidal silica (El-Nokaly and Hiler, 1992), also were suggested for use on French fried potatoes. Protein-based coatings also have been investigated for their potential to reduce oil absorption by coated foods during frying and, secondarily, to retain natural juices and flavors, enhance texture and appearance, and reduce moisture loss. Spraying aqueous gelatin solutions on battered and breaded meats reduced oil absorption upon frying due to the grease resistance of gelatin films formed around meat products (Olson and Zoss, 1985). Feeney et al. (1992, 1993) applied prolamin (preferably zein) coatings to coated potatoes and other foods prior to deep-fat frying. The coatings were prepared in the form of prolamin latices substantially free of organic solvents according to a method described by Stark and Gross (1991). Reportedly, such prolamin coatings reduced the absorbed oil content of finished fried foods by 20 to 40% compared to non-coated products (Feeney et al., 1992, 1993). Polansky (1993) dipped potato slices in dilute collagen solutions (1 to 2% w/w) prior to frying. The protective collagen coatings reduced the amount of absorbed oil by more than 40% compared to non-coated potato slices (Polansky, 1993). Rayner et al. (2000) recommended a solution of soy protein isolate (10% w/w) and gellan gum (0.5% w/w) for coating foods to reduce oil uptake during deep-fat frying. The coating solutions were heat-treated at 80 C for 20 min and cooled to ambient temperature prior to application. Coated and fried discs of doughnut mix had a notably reduced fat content (by about 55%) compared to non-coated samples. Also, a preference evaluation by an untrained sensory panel indicated no significant difference between coated and non-coated French fries (Rayner et al., 2000). Mallikarjunan et al. (1997) reported that coating mashed potato balls with zein prior to frying reduced oil uptake by 59% compared to non-coated control samples. Also, moisture loss from zein-coated products was lower by 15% compared to non-coated samples. Most likely, zein coagulated upon contact with moisture on the product surface and formed a thin and water-insoluble film at the coating/potato interface. However, HPMC and methylcellulose coatings were even more effective than zein coatings in decreasing oil uptake and retaining moisture (Mallikarjunan et al., 1997). The greater moisture loss from zein-coated samples compared to cellulose ether-coated samples can partially be attributed to the presence of a highly hydrophilic plasticizer (glycerin) in the zein coatings. Huse et al. (1998) applied zein coatings on partially fried akara that was subsequently frozen, thawed, and finish-fried. Akara is a protein-rich, deep-fat fried, West African traditional food prepared from whipped cowpea paste. Zein coatings applied by either spraying or dipping were equally effective in reducing (by about 40%) the fat content and increasing the moisture content (by about 5%) of the final product. However, the increased moisture re-

References

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tention resulted in a soggier product. Similar results were obtained in the same study with HPMC, methylcellulose, and amylose coatings.

Deep-fat frying is a widely used cooking process. The appeal of fried food is universal and enjoyed by many cultures around the world. Recently, there has been substantial interest in the use of edible films and coatings to perform different functions in food processing and preservation. However, limited research has focused on the use of protein-based edible coatings on fried foods. Investigations determined such advantages of applying protein-based edible films and coatings as improved breading adhesion, increased retention of natural juices and flavors, enhanced texture and appearance, and reduced moisture migration and fat uptake during frying. By suitable selection of edible films, it is possible to control moisture and fat transfer between the frying medium and the food. Consumer demand for low-fat fried foods with good organoleptic attributes stimulate interest in new applications of edible films in the food industry.

Alleavitch, J., Turner, W.A. and Finch, C.A. 1989. “Gelatin” in Ullmann’s Encyclopedia of Industrial Chemistry, Vol. A12, 5th edition, eds., B. Elvers, S. Hawkins, M. Ravenscroft, J.F. Rounsaville and G. Schultz, Weinheim, Germany: VCH Verlagsgesellschaft, pp. 307–316. Anonymous. 1997. “Edible Films Solve Problems,” Food Technol., 51(2):60. Arenson, S.W. 1972. “Novel Process Controls Quality Variables for French Fried Potatoes—Extends Shelflife of Potato Chips,” Food Prod. Develop., 6(3):42–44, 46. Avena-Bustillos, R.J. and Krochta, J.M. 1993. “Water Vapor Permeability of Caseinate-Based Edible Films as Affected by pH, Calcium Crosslinking, and Lipid Content,” J. Food Sci., 58:904–907. Baker, R.C., Darfler, J.M. and Vadehra, D.V. 1972. “Prebrowned Fried Chicken. 2. Evaluation of Predust Materials,” Poultry Sci., 51:1220–1222. Balasubramaniam, V.M., Chinnan, M.S. and Mallikarjunan, P. 1995. “Deep-fat Frying of Edible Film Coated Products: Experimentation and Modeling” in Proceedings of the FPAC IV Conference, St. Joseph, MI: American Association of Agricultural Engineers, pp. 486–494. Balasubramaniam, V.M., Chinnan, M.S., Mallikarjunan, P. and Phillips, R.D. 1997. “The Effect of Edible Film on Oil Uptake and Moisture Retention of a Deep-fat Fried Poultry Product,” J. Food Process. Engr., 20:17–29. Busacker, W.F., Calder, J.W., Erickson, L.J. and Pinegar, R.K. February 28, 1995. U.S. patent 5,393,552. Calder, J.W., Erickson, L.J. and Pinegar, R.K. April 12, 1994. U.S. patent 5,302,410. Chen, H. 1995. “Functional Properties and Applications of Edible Films Made of Milk Proteins,” J. Dairy Sci., 78:2563–2583. Cunningham, F.E. and Suderman, D.R. 1981. “Use of Batters and Breadings on Food Products—A Review” in Quality of Poultry Meat, eds., R.W.A.W. Mulder, C.W. Scheele and C.H. Veerkamp, Beekbergen, The Netherlands: Spelderholt Institute for Poultry Research, pp. 314–323.

524 Cuq, B., Gontard, N. and Guilbert, S. 1998. “Proteins as Agricultural Polymers for Packaging Production,” Cereal Chem., 75:1–9. El-Hag, N.A. and Shanbhag, S.P. March 2, 1982. U.S. patent 4,317,842. El-Hag, N.A., Cochran, S.A., Schwabe, K.R., Carricato, L. and Shanbhag, S.P. November 5, 1985. U.S. patent 4,551,340. El-Nokaly, M. and Hiler, G.D. March 31, 1992. U.S. patent 5,100,684. Feeney, R.D., Haralampu, S.G. and Gross, A. June 30, 1992. U.S. patent 5,126,152. Feeney, R.D., Haralampu, S.G. and Gross, A. June 8, 1993. U.S. patent 5,217,736. Gennadios, A. and Weller, C.L. 1991. “Edible Films and Coatings from Soymilk and Soy Protein,” Cereal Foods World, 36:1004–1009. Gennadios, A., Weller, C.L. and Testin, R.F. 1993. “Temperature Effect on Oxygen Permeability of Edible Protein-based Films,” J. Food Sci., 58:212–214, 219. Gennadios, A., McHugh, T.H., Weller, C.L. and Krochta, J.M. 1994. “Edible Coatings and Films Based on Proteins” in Edible Coatings and Films to Improve Food Quality, eds., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 201–277. Gennadios, A., Cezeirat, C., Weller, C.L. and Hanna, M.A. 1998. “Emulsified Soy Protein-Lipid Films” in Paradigm for Successful Utilization of Renewable Resources, eds., D.J. Sessa and J.L. Willett, Champaign, IL: AOCS Press, pp. 213–226. Gontard, N., Guilbert, S. and Cuq, J-L. 1992. “Edible Wheat Gluten Films: Influence of the Main Process Variables on Film Properties Using Response Surface Methodology,” J. Food Sci., 57:190–195, 199. Gontard, N., Duchez, C., Cuq, J-L. and Guilbert, S. 1994. “Edible Composite Films of Wheat Gluten and Lipids: Water Vapor Permeability and Other Physical Properties,” Int. J. Food Sci. Technol., 29:39–50. Hansen, S.L. 1998. “Effects of Prefry Drying Methods on the Quality of Deep Fat Fried Onion Slices,” J. Food Qual., 21:433–443. Higgins, C., Qian, J. and Williams, K. November 2, 1999. U.S. patent 5,976,607. Hsia, H.Y., Smith, D.M. and Steffe, J.F. 1992. “Rheological Properties and Adhesion Characteristics of Flour-Based Batters for Chicken Nuggets as Affected by Three Hydrocolloids,” J. Food Sci., 57:16–18, 24. Huse, H.L., Mallikarjunan, P., Chinnan, M.S., Hung, Y.-C. and Phillips, R.D. 1998. “Edible Coatings for Reducing Oil Uptake in Production of Akara (Deep-fat Frying of Cowpea Paste),” J. Food Process. Preserv., 22:155–165. Kasarda, D.D., Bernardin, J.E. and Nimmo, C.C. 1976. “Wheat Proteins” in Advances in Cereal Science and Technology, Vol. 1, ed., Y. Pomeranz, St. Paul, MN: American Association of Cereal Chemists, pp. 158–236. Kester, J.J. and Fennema, O. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Khalil, A.H. 1999. “Quality of French Fried Potatoes as Influenced by Coating with Hydrocolloids,” Food Chem., 66:201–208. Krochta, J.M. 1992. “Control of Mass Transfer in Foods with Edible Coatings and Films” in Advances in Food Engineering, eds., R.P. Singh and M.A. Wirakartakasumah, Boca Raton, FL: CRC Press, pp. 517–536. Krochta, J.M. and De Mulder-Johnston, C. 1997. “Edible and Biodegradable Polymer Films: Challenges and Opportunities,” Food Technol., 51(2):61–74. Mallikarjunan, P., Chinnan, M.S., Balasubramaniam, V.M. and Phillips, R.D. 1997. “Edible Coatings for Deep-fat Frying of Starchy Products,” Lebensm. Wiss. Technol., 30:709–714. Mallikarjunan, P., Chinnan, M.S. and Ngadi, M.O. 1998. “Determination of Lipid Conductivity through Edible Films” in 1998 IFT Annual Meeting Book of Abstracts, Chicago, IL: Institute of Food Technologists, p. 10.

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McHugh, T.H. and Krochta, J.M. 1994a. “Milk Protein-based Edible Films and Coatings,” Food Technol., 48(1):97–103. McHugh, T.H. and Krochta, J.M. 1994b. “Sorbitol- vs Glycerol-Placticized Whey Protein Edible Films: Integrated Oxygen Permeability and Tensile Property Evaluation,” J. Agric. Food Chem., 42:841–845. McHugh, T.H. and Krochta, J.M. 1994c. “Water Vapor Permeability Properties of Edible Whey Protein-Lipid Emulsion Films,” JAOCS, 71:307–312. Meyers, M.A. and Conklin, J.R. February 13, 1990. U.S. patent 4,900,573. Miller, K.S. and Krochta, J.M. 1997. “Oxygen and Aroma Barrier Properties of Edible Films: A Review,” Trends Food Sci. Technol., 8:228–237. Olson, S. and Zoss, R. April 16, 1985. U.S. patent 4,511,583. Polansky, S. August 3, 1993. U.S. patent 5,232,721. Pomes, A.F. 1971. “Zein” in Encyclopedia of Polymer Science and Technology: Plastics, Resins, Rubbers, Fibers, Vol. 15, eds., H.F. Mark, N.G. Gaylord and N.M. Bikales, New York: Interscience Publishers, pp. 125–132. Prosise, W.E. April 17, 1990. U.S. patent 4,917,908. Rao, V.N.M. and Delaney, R.A.M. 1995. “An Engineering Perspective on Deep-Fat Frying of Breaded Chicken Pieces,” Food Technol., 49(4):138–141. Rayner, M., Ciolfi, V., Maves, B., Stedman, P. and Mittal, G.S. 2000. “Development and Application of Soy-Protein Films to Reduce Fat Intake in Deep-Fried Foods,” J. Sci. Food Agric., 80:777–782. Saguy, I.S. and Pinthus, E.J. 1995. “Oil Uptake During Deep-Fat Frying: Factors and Mechanism,” Food Technol., 49(4):142–145, 152. Sloan, J.L., Middaugh, K.F. and Jacobsen, G.B. October 22, 1991. U.S. patent 5,059,435. Snyder, H.E. and Kwon, T.W. 1987. Soybean Utilization. New York: Van Nostrand Reinhold Company. Stark, L.E. and Gross, A.T. June 4, 1991. U.S. patent 5,021,248. Suderman, D.R., Wiker, J. and Cunningham, F.E. 1981. “Factors Affecting Adhesion of Coating to Poultry Skin: Effects of Various Protein and Gum Sources in the Coating Composition,” J. Food Sci., 46:1010–1011. Toloday, D. and Andres, C. 1975. “Vegetable Gum Dry Mix Improves Shrimp Breading,” Food Process., 36(5):32. Trezza, T.A. and Vergano, P.J. 1994. “Grease Resistance of Corn Zein Coated Paper,” J. Food Sci., 59:912–915. Varela, G. and Ruiz-Roso, B. 1992. “Some Effects of Deep Frying on Dietary Fat Uptake,” Nutr. Rev., 50:256–262. Wall, J.S. and Paulis, J.W. 1978. “Corn and Sorghum Grain Proteins” in Advances in Cereal Science and Technology, Vol. 2, ed., Y. Pomeranz, St. Paul, MN: American Association of Cereal Chemists, pp. 135–219. Williams, R. and Mittal, G.S. 1999a. “Low-Fat Fried Foods with Edible Coatings: Modeling and Simulation,” J. Food Sci., 64:317–322. Williams, R. and Mittal, G.S. 1999b. “Water and Fat Transfer Properties of Polysaccharide Films on Fried Pastry Mix,” Lebensm. Wiss. Technol., 32:440–445.

Application of Edible Protein Coatings to Nuts and Nut-Containing Food Products

OTANICALLY, nuts are fruits consisting of a kernel inside a hard, tough shell. Peanuts are botanically legumes; however, they are often called nuts because of their usage and characteristics. For the purposes of this chapter, nuts are considered any food product that is commonly thought to be a nut or eaten like a nut. This loose definition includes tree nuts (e.g., almonds, walnuts, and Brazil nuts), some oilseeds (e.g., sunflower kernels), and peanuts.

B

Nuts, peanuts, and oilseeds are versatile foods that can be used in a variety of forms and products. For example, in 1990–1991 in the U.S., pecans were utilized in bakery products (20.3%), for direct retail consumption (17.9%), as confection ingredients (9.7%), as inclusions in ice cream (5.7%), in gift packs (8.1%), for food service (3.2%), and a number of other uses (NPSA, 1997). Nuts are also integral components in products such as snack and trail mixes, granola and breakfast bars, breakfast cereals, and certain sauces. They often add a distinct flavor and a desirable crunch to many foods. Annual per capita consumption of peanuts in the U.S. is approximately 2.6 kg. These peanuts are con527

528 sumed as snack nuts (0.5 kg/year/person), in-shell nuts (0.2 kg/year/person), peanut butter (1.2 kg/year/person), and confection ingredients (0.6 kg/year/ person) with the balance (~ 0.1 kg/year/person) in other forms (Haumann, 1998).

Nuts and peanuts comprise a significant financial portion of the U.S. agricultural and food processing industries. In 1996, snack nuts accounted for $1.2 billion, or 7.8% of total value in the snack food industry (SFA, 1997). The export value to the U.S. economy in 1998 for almonds, hazelnuts, peanuts, and walnuts was more than $190 million (USDC, 1998a). In 1997, nut and peanut ingredients accounted for around 3% of the total value of U.S. confectionery products (USDC, 1998b). Sales of candy bars containing peanut butter rose 9% over the 52 weeks ending February 15, 1998 (Anonymous, 1998). Also, sales of chocolate-panned confections, many of which contain nuts, increased in value more than 9% from 1996 to 1997 (USDC, 1998b).

Nutritional and medical research has shown that diets rich in plant foods and with a high ratio of polyunsaturated to saturated fats can reduce the risk of humans developing certain diseases such as cardiovascular disease and stroke (Bernert and Browner, 1995; Haumann, 1998). Many nuts, oilseeds, and peanuts are rich in polyunsaturated fatty acids (PUFAs) (Table 22.1). Walnuts, which contain exceptionally high levels of PUFAs, favorably modified human

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serum lipid profiles and reduced cholesterol of men to recommended levels (Sabaté et al., 1993). Women also have reduced risks of coronary heart disease when consuming nuts (including peanuts) as part of a healthy diet (Hu et al., 1998). Peanuts contain high levels of protein, monounsaturated fatty acids, and other agents that are also gaining recognition by nutritionists as having significant roles in healthy diets (Haumann, 1998). Other nuts that are relatively high in PUFAs, such as almonds, pecans, sunflower seeds, and Brazil nuts, can likewise contribute to healthy diets.

Nuts are susceptible to a variety of modes of deterioration. Light, moisture, oxygen, high temperature, microbes, and other factors can significantly contribute to loss of quality and compromise the potential safety of nuts and nut-containing foods. The most common modes of spoilage of nuts are lipid oxidation (causing rancidity and staleness), sogginess due to moisture uptake, and loss of flavor from a combination of oxygen and moisture. Microbial spoilage can also occur.

Nuts are typically low-moisture, high-fat foods. The high polyunsaturated lipid content of many nuts makes them especially susceptible to oxidative rancidity. Rancidity is a general term used to describe the rank off-flavor development that occurs during long-term or high-temperature food storage. In high-fat foods, this rancidity is caused by oxidation of fatty acids through reactions with atmospheric oxygen (Coultate, 1989). Lipid oxidation is the main method of deterioration of nuts (Nawar, 1996; Agbo et al., 1992). In the presence of appropriate initiators, unsaturated lipids (LH) form alkyl radicals (L•) and peroxyl radicals (COO•) (Coultate, 1989; Frankel, 1991). These radicals propagate in the presence of oxygen to form hydroperoxides (LOO•) (Frankel, 1991). These lipid hydroperoxides readily decompose to form a variety of aldehydes, ketones, hydrocarbons, alcohols, and other compounds that contribute to the characteristic “rancid” flavor of spoiled nuts (Gray, 1978; Frankel, 1991). The most abundantly formed oxidation products from linoleic acid are hexanal, octenals, and heptenals; while, linolenic acid oxidation forms propanal and heptadienals (Belitz and Grosch, 1987). Rancid peanuts and nuts are typically described as having a grassy/green character, as painty or solvent, as reminiscent of cardboard, and as tasting metallic (Johnson et al., 1988; Civille and Dus, 1992 ). The major factors influencing the rate of lipid oxidation are light, temperature, oxygen concentration, moisture, degree of lipid unsaturation, length of storage of

530 the produce, pro-oxidant presence (such as metals and other compounds), and presence of antioxidant compounds.

The onset of detectable degrees of lipid oxidation varies depending on nut type, storage conditions, and composition of multi-phase foods (for nut-containing products). The induction period is the initial time when free radicals are being produced and oxidation products cannot be detected in the food. After the induction period, the oxidation rate is measured as the increase of flavor-altering compounds as a function of time. The fatty acid profile has a large influence on the induction period and oxidation rate of nut lipids (Table 22.2). The greater the content of PUFAs in the product, the greater the oxidation rate. Thus, nuts and oilseeds such as walnuts, sunflower seeds, pecans, and peanuts become rancid relatively quickly at ambient conditions. As a result, some raw and many roasted nuts without other treatments are considered to have short shelf lives, ranging from a few weeks to a few months (Table 22.3), compared to other dry food products.

Nuts and oilseeds can lose their characteristic flavors over prolonged storage. The term flavor fade is typically used to describe the loss of flavor of peanuts during storage (Dimick, 1994). The flavor of roasted peanuts is attributed to a large number of volatile compounds including pyrazines, methyl butanol, methyl propanal, sulfur compounds (such as methanethiol, carbon disulfide, and dimethyl sulfide), ketones, alcohols, and hydrocarbons (Walradt et al., 1971; Crippen et al., 1992). Flavor fade in peanuts was due primarily to masking of pyrazines and other flavor compounds by low-molecular weight aldehydes produced during lipid oxidation (Warner et al., 1996). Sensory scores of peanut flavor have paralleled changes in pyrazine content (Braddock et al., 1995). It is likely that flavor fade of peanuts can be limited by reducing the development of oxidative rancidity (Dimick, 1994; Warner et al., 1996). Other nuts also have loss of flavor associated with oxidative rancidity. For example, roasted filbert (hazelnut) flavor is attributed to low levels of 2-methylpropanal, 2-methyl- and 3-methyl butanol, and perhaps dimethyl sulfide (Sheldon et al., 1972). It is presumed that the loss of flavor of filberts, as well as other nuts, is due to factors similar to those found for peanuts. This has been confirmed qualitatively for hazelnuts (Keme et al., 1983a, b). Flavor fade can limit the shelf life of nuts and nut-containing products. Because the flavor of nuts generally “fades” as lipids oxidize, the nuts not only lose the desirable nut flavor, but they also gain the undesirable rancid flavor.

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Changing moisture content is a major mode of quality deterioration for nuts. Generally, a small range of optimal moisture content exists for nuts (Table 22.4). Peanuts, for example, are best stored at a moisture content of about 7% (Woodroof, 1966). Above 8% moisture, peanuts are much more susceptible to mold growth. Below 5% moisture, peanuts become brittle and are more likely to incur breakage during handling (Woodroof, 1966). Moisture uptake can result in stale and soggy nuts. Cosler (1958 a, b) and Alikonis and Cosler (1961) found that besides the development of rancidity, moisture was important in determining the quality of a variety of nuts. Increases in moisture can also increase the rate of oxidative rancidity. In general, the oxidation rate of lipids is greatest when the water activity is around 0.60 to 0.75 (Hall, 1991). These water activity values equate to moisture contents from about 4.0 to 5.5%, depending on the type of nut (Beuchat, 1978). Lastly, increasing moisture content increases susceptibility of nuts to microbial growth. Moisture changes in nuts have implications in many areas of the food industry. Nuts typically have water activities around 0.2 to 0.4 (1 to 4% moisture con-

532

tent, depending on the type of nut) (Beuchat, 1978). Nuts are often imbedded into confections containing nougats, caramels, cremes, and other higher water activity (0.5–0.7) components. Moisture migration from the center phase to the nut phase often results in loss of crunch and increased flavor fade of the peanuts. For bread and bakery items, the same scenario can occur. Nuts are also commonly used as inclusions in ice cream and other frozen desserts. It is not uncommon to find soggy nuts in a container of certain varieties of ice cream.

Transfer of oil from nuts to other components in a food is a primary concern in the confectionery industry. The shelf life of products such as chocolate-coated walnuts and Brazil nuts is severely compromised due to oil migration. The net result of oil migration is a softening of the chocolate, hardening of the nut center, and formation of a grayish white, hazy film on the product (Minifie, 1980; Talbot, 1996; Couzens and Wille, 1997). This film, called “bloom,” is actually a layer of fat that has migrated to the surface of the product and recrystallized (Lohman and Hartel, 1994). Fat bloom on chocolate is sometimes confused for mold growth, although it poses no health or nutritional hazards (Kleinert, 1969).

The quality and shelf life of nuts can be drastically altered with storage conditions, addition of antioxidants, and by packaging. With careful selection of

Traditional Strategies for Maintaining the Quality of Nuts

533

these three factors, the shelf life of nuts, peanuts, and oilseeds can be dramatically increased, and the quality of the nuts can be maintained to a high degree (Table 22.5).

Refrigeration or freezing of nuts is the most common and most reliable method to preserve them. Nuts, such as pecans and walnuts, can be stored for two years or more when frozen. In research on nut and oil rancidity, it is often assumed that freezing halts the reactions involved in oxidative rancidity development and quality loss. This has been confirmed in several studies and is commercially practiced and recommended for long-term storage of many nuts (Woodroof, 1966; Ahmed and Storey, 1975; NPSA, 1997). Although generally giving a reduced shelf life relative to freezing, refrigeration is also a recommended practice for the storage of many nuts, including pecans, peanuts, and walnuts (Woodroof, 1953; Payne, 1985; NPSA, 1997).

Although refrigerated and frozen storage minimize the deterioration of nuts, these methods of storage are relatively expensive and create distribution challenges. In a typical grocery store in the U.S., nuts are stored and displayed at

534 ambient temperatures. When nuts are put into other products, such as bread, that have different optimum storage conditions from those of nuts, the quality and shelf life of nuts can be severely compromised. Antioxidants are edible substances that delay the onset of rancidity of lipids and lipid-containing foods. Antioxidants quench free radicals that lead to peroxide development and generation of the off-flavor compounds found in nuts (Coultate, 1989). The antioxidants do not completely prevent lipid oxidation; however, many lengthen the induction period of lipid oxidation through resonance-stabilized free-radical reactions and scavenging of the singlet oxygen responsible for initiating the oxidation of unsaturated fatty acids (Nawar, 1996). There are a number of antioxidant compounds approved for food use around the world that fall into the two general categories of synthetic and natural. Synthetic antioxidants are prepared from petroleum products and are sometimes called chemical antioxidants (Botma, 1978). The most commonly used synthetic antioxidants are the phenolic compounds butylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT), and tertiary butylhydroquinone (TBHQ). Gallate esters (such as propyl- and octyl-gallate) are also used as antioxidants for foods. Combinations of BHT and BHA or TBHQ are often added to walnuts, chocolate-coated peanut butter/nut confections, and other nut-containing products. TBHQ effectively reduced the rancidity of roasted peanuts (Hoover and Nathan, 1980, 1981). Natural antioxidants, which are extracted from plants, include vitamins such as ascorbic acid (vitamin C), tocopherols (vitamin E), carotenoids (e.g., vitamin A and lycopene), and a wide variety of other naturally occurring compounds. Many “natural” products, such as vitamins, can also be synthesized commercially. Extracts from spices have been known for some years to have antioxidant potential (Chipault et al., 1952). In the past few years, a number of spice extracts, especially from rosemary, have been commercialized as antioxidant ingredients (Hudson, 1992; Nawar, 1996; Sandusky et al., 1999). Synthetic and natural antioxidants have similar antioxidant activities. Both types of antioxidants have greater antioxidant power in animal fat versus vegetable oil when subject to Rancimat testing (Table 22.6). Richheimer et al. (1996) reported that carnosic acid, the major antioxidant compound in rosemary, had antioxidant activity seven times that of BHT and BHA but only half the antioxidant activity of TBHQ. Recent studies have shown that natural antioxidants did not limit oxidative rancidity in restructured meat products as effectively as TBHQ (Lai et al., 1991; Stoick et al., 1991). Synergists, such as ascorbyl palmitate, citric and phosphoric acids, and lecithin, increased antioxidant effectiveness at levels as low as 0.1% (Belitz and Grosch, 1987; Nawar, 1996). The synthetic antioxidants BHT, BHA, and TBHQ are limited by the U.S. Code of Federal Regulations to maximum levels of 200 ppm. Natural antioxidants do not have the same limitations at this time and are generally GRAS ingredients allowed up to levels that will achieve the desired effect.

Edible Protein Coatings for Maintaining the Quality of Nuts

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Packaging selection can also greatly influence the quality and shelf life of nuts. Glass and metal containers provide a total barrier to oxygen and give product that is generally considered upscale or elegant. High-oxygen-barrier plastic films, such as polyvinylidene chloride (PVDC) and ethylene vinyl alcohol (EVOH), and metalized films are one step down in barrier protection from glass and metal. Other packaging strategies include the use of vacuum packaging and nitrogen flushing. If a gas-flushed package is used, a high-barrier material may be necessary to limit the amount of oxygen diffusing into, or the amount of nitrogen diffusing out of, the package during storage. Reducing product exposure to light by package design can also help protect against light-induced oxidative reactions.

As discussed elsewhere in this book, protein films possess excellent oxygen barrier properties—often better than the best conventional synthetic polymer films, PVDC and EVOH. If protein coatings are kept at low relative humidity, and if the coating can be evenly applied to nuts, then excellent oxygen protection of nuts can be maintained. Biopolymer films, especially those from pro-

536 teins, are also excellent flavor and aroma barriers (Miller and Krochta, 1998). Gelatin has been extensively used as an encapsulating agent to preserve the flavor and aroma of essential oils and other flavor compounds (Reineccius, 1994).

Protein coatings can also reduce the amount of packaging materials and processing steps required for nuts. For products packaged in high-barrier PVDC, EVOH, or other materials, the existence of an edible coating can potentially eliminate the need for the high-cost barrier polymer. This can lead to package cost savings. Also, PVDC and EVOH laminates are difficult to recycle because the barrier layers cannot be easily separated from other polymers in the film. There is interest in PVDC alternatives due to its recycling problems (Gavitt, 1996). Use of an edible coating potentially increases the recyclability of a package because the separation of polymers in the package is no longer required (Maté and Krochta, 1997).

Gloss is an important quality factor for many food products (Potter and Hotchkiss, 1995). Numerous studies have shown that the gloss of chocolate dramatically influences flavor judgment (Musser, 1973). Foods such as apples, citrus fruits, vegetables, and confectionery products are coated with waxes and glazes to provide a high gloss (Lecos, 1982; Hernandez, 1994). Trezza and Krochta (2000a) reported that whey protein coatings had a high gloss equal to that of shellac coatings. Shellac is commonly used to add gloss to confectionery products and fresh fruits and vegetables. Also, the color of nuts can be improved by the use of protein coatings. Wheat gluten and corn zein proteins also form high-gloss coatings (Cook and Shulman, 1998; Trezza and Krochta, 2000a). Taking advantage of the browning tendencies of protein-reducing sugar systems can induce controlled color development. For example, soy and whey protein coatings have the potential to create controlled color development (Gennadios et al., 1996; Miller et al., 1997; Trezza and Krochta, 2000b).

Protein films and coatings are considered excellent barriers to oil migration. Zein coatings were effective grease barriers as coatings on paper (Trezza and Vergano, 1994). De Mulder-Johnston (1999) found that whey protein coatings prevented peanut oil migration for more than 19 weeks. These coatings have high potential as oil barrier coatings on nuts or nut-containing centers that are coated or enrobed with chocolate.

Edible Protein Coatings for Maintaining the Quality of Nuts

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Within the past 15 years, more than 200 research articles on the physical properties and potential uses of edible films and coatings have been published. Of these, more than 120 papers were focused on edible protein films and coatings. Despite the extensive research on the subject, relatively few commercial applications for edible films and coatings actually exist. However, as food processing technology has advanced, functionality of food ingredients has improved. Researchers have discovered that exploitation of specific properties of food ingredients provide new opportunities for food coating applications. Use of protein coatings is highly promising due to the wide array of physical properties exhibited by proteins (e.g., film forming and emulsification). Pertinent reviews have been published (Koelsch, 1994; Krochta et al., 1994; Gennadios et al., 1997; Debeaufort et al., 1998). A summary of protein coating research on nuts is presented here (Table 22.7).

Reports from the late 1950s and early 1960s indicated that commercial corn zein coatings reduced the sensory-detectable rancidity of roasted Spanish and Virginia peanuts, pecans, almonds, and walnuts (Cosler, 1958a; Alikonis and Cosler, 1961). Zein-coated peanut bars and chocolate bars containing zein-coated nuts had increased shelf life compared to products with uncoated nuts (Cosler, 1958b). The coating also provided a superior gloss, moisture resistance, and an oil barrier (Cosler, 1957). Because these coatings were a commercial blend of zein, oil, BHT, and other antioxidants, the influence of the protein alone could not be determined. In other early work, Harris et al. (1972) coated unroasted and roasted diced almonds with a zein solution similar to that used by Cosler. Overall, the almonds did not have increased resistance to oxidative rancidity. In the same study, coating diced roasted almonds did not offer additional protection against oxidative rancidity when the nuts were incorporated into a chocolate confection (Harris et al., 1972). Hebert and Holloway (1992) patented a process and formulation for precoating nuts with a protein. The protein coating was a base coating to improve uniformity and adhesion of a second seasoning layer. Sodium caseinate and gelatin coatings prevented stickiness and oxidative deterioration of fig bars, brownies, and other bakery items (Durst, 1967).

In recent work, whey proteins showed potential for reducing the rancidity and extending the shelf life of roasted peanuts. Besides their oxygen and flavor barrier properties, additional benefits of whey protein coatings include the abil-

538

Application of Edible Coatings on Nuts

539

ity to form water-insoluble films even though they are cast from a water solvent, high gloss coatings, and excellent emulsifying capabilities. This emulsifying ability allows for incorporation of lipids to improve moisture barrier properties of the coatings (Krochta and McHugh, 1996; Shellhammer and Krochta, 1997; Trezza and Krochta, 2000a). Peanuts coated with whey protein isolate (WPI)/glycerol (GLY) coatings had hexanal levels reduced by more than 75% as compared to uncoated nuts after 70 days of storage at 37 C and 21% RH (Maté et al., 1996). Peroxide values followed the same trends. The same WPI/GLY coatings on English walnuts did not have as dramatic effect as on peanuts. After 70 days at 37 C and 53% RH, coated walnuts had hexanal values less than 50% of those of uncoated walnuts (Maté and Krochta, 1997). Addition of tocopherol from 0.11 to 1.1% lowered the rate of hexanal formation. The poorer performance of the WPI coatings on walnuts (versus peanuts) was attributed to difficulty in obtaining a uniform and complete coating on the irregularly shaped walnuts. To compensate, Maté and Krochta (1997) produced coatings of 12-20% (w/w) by dipping nuts into chilled protein solution. Shrinkage of the thick coatings produced oil expression and then leakage due to coating cracking, which led to greater than predicted rates of rancidity development (Maté and Krochta, 1998). WPI/acetylated monoglyceride coatings have also been applied as moisture barriers on chopped almonds (Adams et al., 1995). The nuts, which were then incorporated into ice cream, did not become soggy after six weeks of frozen storage. Uncoated, chopped almonds in ice cream became soggy after one week.

Besides proteins, various other materials were effective as quality-maintaining, shelf life-extending coatings for nuts, peanuts, and oilseeds. In their walnut coating study, Maté and Krochta (1997) found that acetylated monoglyceride coatings dramatically reduced hexanal production in walnuts stored for 70 days at 37 C. With ascorbyl palmitate, tocopherol, or a combination of the two, hexanal generation was at parity with walnuts treated with BHT. Coating with acetylated monoglycerides had a small effect on the shelf life of pecans (Senter and Forbus, 1979). Various other coatings increased shelf life and reduced the rate of rancidity of nuts including cellulosic coatings on pecans (Anonymous, 1999), pectin coatings on almonds (Swenson et al., 1953), and alginate coatings on sunflower seeds (Hettiarachchy et al., 1989).

The actual application of edible coatings presents several challenges to the food processor. Food composition, size and shape, coating conditions, and

540 other factors all influence the coating process and the final coated product quality. Irregularly shaped nuts, such as walnuts, presented difficulties in obtaining complete, uniform coatings (Maté and Krochta, 1997). Pieces with flat surfaces (e.g., split peanuts and sliced almonds) often “twin” or stick together during the coating process.

The current commercial processes used for pan coating of confectionery items are well suited for applying edible coatings to nuts, peanuts, and oilseeds. Typically, nuts that are to be coated or enrobed with chocolate are precoated with hydrocolloids consisting of gum arabic, starches, and other polysaccharides (Boutin, 1997; Copping 1997). These gum coatings provide an oil barrier to reduce lipid migration from the nut phase to the chocolate phase and a surface to promote chocolate adhesion and absorb moisture from the center (Copping, 1996, 1997). Pan coaters for chocolate confectionery are very well suited for coating nuts with fairly uniform shapes, such as whole peanuts, hazelnuts, almonds, and sunflower seeds. Sharma (1985) used conventional pan coating to apply flavored sugar slurry coatings to roasted peanuts. Hebert and Holloway (1992) also used conventional panning techniques for adding protein coatings to peanuts. For irregularly shaped nuts, such as walnuts and pecans, drum coaters, which are often used for products such as expanded snacks, may be more suitable (Dreier, 1991). Walnuts can be particularly difficult to coat with laboratory scale pan coaters. Trezza (1999b) found that only 80% of shelled walnut pieces were uniformly coated using traditional methods. Loss of coating liquid to the walls of the pan due to poor wetting of walnuts was also noticed. Vented pans, which provide airflow, temperature, and air humidity control in panning, have possible applications for edible coating applications to nuts. Coatings, which would be applied using a spray technique involving pneumatic spraying technology, could benefit from increased coating uniformity (with or without added colors and other ingredients), increased process control, and increased product throughput and efficiency (Strub, 1987). This has already been demonstrated to be an effective method for coating peanuts (Lee et al., 1999). Fluidized bed systems can also be used for coating nuts. These systems can precisely control coating conditions and offer gentle handling during the coating process (Adams et al., 1995). Trezza (1999b) has successfully coated sunflower seeds, peanuts, and hazelnuts in a laboratory scale, Wurster columntype, fluidized bed coater.

Application of Edible Coatings on Nuts

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As mentioned, it is difficult to achieve uniform coatings using conventional panning on irregularly shaped nuts, such as pecans and walnuts. The same problems will likely occur when using vented pans. However, porous, irregularly shaped products such as nut clusters are successfully enrobed with chocolate and similar coatings (Webb, 1996). Dipping gave 100% uniform coverage of whey protein and acetylated monoglyceride coatings on walnuts (Krochta, 1996; Maté and Krochta, 1997).

Many factors influence the performance of protein films and coatings. Film composition has the most dramatic effect on final film properties. This section will identify the major factors that affect application of edible coatings to nuts, oilseeds, and peanuts.

Many protein coatings need considerable plasticization to form useable coatings. Protein coatings without plasticizers are brittle and fragile. Most protein coatings need 15 to 50% plasticizer (on weight of the base polymer) to form coatings that will not crack (Krochta et al., 1994). In contrast, shellac coatings, which are commonly used as gloss coatings on chocolate-coated nuts and fruit, require only 5% plasticizer (Isganitis, 1988; Boutin 1997). The plasticizers currently used for edible coatings include propylene glycol, glycerol, polyethylene glycol, triacetin, triethyl citrate, and other hydrophilic materials. If too much plasticizer is added, the coatings will become unacceptably tacky and lose gas barrier properties (Porter, 1979). Increasing the plasticizer level in the coatings, for example, dramatically decreases the oxygen barrier properties of many protein coatings. This can limit effectiveness against oxidation of nuts and subsequent development of rancid off-flavors. It is important to use plasticizers that offer the right balance of desired mechanical and barrier properties for the desired application.

Because nuts have hydrophobic surfaces and water-soluble protein coatings are cast from water-based solvents, the wetting and adhesion of these coatings to nuts is of special concern. Due to their low oxygen permeability, WPI coatings effectively reduced oxygen uptake, slowed the onset of rancidity, and in-

542 creased the shelf life of peanuts (Maté et al., 1996). However, reduction in the development of rancidity by WPI coatings was not as significant as predicted from a model that used the oxygen permeability of the WPI coating (Maté and Krochta, 1998). Several factors likely contributed to this result. During the coating of peanuts and walnuts, de-wetting of the solution from nuts dipped in the coating solution occurred. De-wetting means that the coating liquid did not spread across the nut surface. Rather, the liquid retracted and did not have intimate contact with the entire nut surface. By increasing viscosity, de-wetting could be partially alleviated. However, shrinking and cracking of the coating during drying caused expression of oil to the coating-air surface, which also limited coating effectiveness in reducing nut rancidity. Also, after drying, flaking and de-adherence of the coating occurred (Maté et al., 1996). Therefore, to maximize the rancidity reducing potential of protein coatings on peanuts and other nuts, knowledge of how to maintain coating adhesion to the nut must be acquired. Added surfactants (e.g., lecithin, polysorbates, mono- and di-glycerides) can improve the wetting and adhesion of protein coatings to nuts. However, excess surfactant can reduce coating adhesion (Trezza, 1999a).

Various coating ingredients can also reduce coating gloss. Surfactants can decrease coating gloss and increase coating haze by migrating to the air-coating surface (Trezza, 1999a). Lipids added to impart moisture resistance can reduce gloss and increase haze if their particle size is not well controlled. Absorption of moisture can cause protein coatings to become tacky, hazy, and sometimes de-adhere from the product surface. In general, the fewer additives in a coating, the greater the gloss potential and the stability of that gloss over prolonged storage (Trezza, 1999b). Barrier and mechanical properties of the coating can also be compromised as lipids, surfactants, and other additives are incorporated into the coating.

Besides formulation issues, there are many process factors that can influence the efficacy and feasibility of commercial application of protein coatings to nuts. Drying temperature, drying time, airflow, and RH all have severe consequences on the quality and performance of edible coatings on nuts. Drying temperatures have an optimum range where they will effectively dry yet not overheat the nut. Temperature abuse can accelerate the oxidative rancidity process on most nuts (Woodroof, 1966; Perren and Escher, 1997). If roasting time

Research Needs

543

and temperatures are used as a guide to drying temperature profiles during coating, then nuts can have a wide variety of optimal drying temperature-time combinations. For example, a typical range of suitable roasting conditions for hazelnuts is 120 to 145 C for 5 to 12 min (Perren and Escher, 1997; Özugur, 1999). Each nut product can have a unique drying requirement based upon the characteristics of the nut and the thermal history (including roasting) of that product. Lee et al. (1999) reported that low heat (30 C) could be used to produce excellent coatings on peanuts, providing airflow and tumbling conditions were adequate. Adams et al. (1995) recommended coating temperature ranges for chopped almonds of 30 to 50 C, with an optimum at 35 to 45 C. It also is possible to apply coatings at low temperatures for certain applications.

The most commonly used edible coatings are added to impart gloss and moisture protection to confectionery products. These coatings consist primarily of shellac or corn zein applied from ethanol-based solutions (Isganitis, 1988; Boutin, 1997). A significant problem with the use of shellac and zein coatings is that their solvent systems are composed mostly of volatile organic compounds (VOCs). VOCs have become strictly regulated because of their impact on air and water quality (Sawyer, 1997). The U.S. Federal Government has mandated that confectionery manufacturers must gradually lower the amount of VOCs they emit into the atmosphere to an ultimate goal of zero discharge (Glesecke, 1998). The regulation of VOC emissions has forced confectionery manufacturers to seek alternatives to shellac and other VOC-containing coatings (Anonymous, 1998). In the pharmaceutical industry, water-based hydroxypropyl methylcellulose (HPMC) coatings have been successfully used as coatings for solid dosage forms (tablets) (Banker and Peck, 1981; Anonymous, 1988). Although water-based protein coatings have the potential to replace shellac and corn zein as coatings on some panned confectionery products, several issues must be considered. Solvent-based coatings work well because they are fast-drying. As solvent evaporates, the coating quickly passes through a tack phase where product can stick together in a coating pan or other coating equipment. Coatings using water dry at a much slower rate than coatings using ethanol. Typical ethanol-based coatings can become surface-dry in 20 to 30 min at ambient temperatures (Groves, 1992). Water-based coatings can take 16 to 24 h to dry at ambient conditions. The slow drying time of water-based coatings, even when heat is applied, can have severe consequences in terms of texture, susceptibility to spoilage, and product integrity.

To date, much of the work on the use of edible coatings for nuts has focused

544 on physical properties of free-standing films. Limited application research has been performed because of the unique coating requirements of each system. The development of water-based systems with short drying times at ambient temperatures presents challenges. For protein-coatings to be successfully applied, current commercial applications (such as panning and fluidized-bed coating) must be widely explored on a variety of nut systems. Although some application work is in progress, there is much to be discovered about how one can actually get the desired coating onto a food product. The major hurdle is that many nuts have processing limitations that influence the use of protein coatings. For example, shellac coatings cast from ethanol-based solutions dry very rapidly. That is partially the reason they are used for chocolate-coated confections. When coating pharmaceutical tablets and nutritional supplements, heat is often applied to shorten the drying times. For nuts, a clear understanding of the heat that nuts can withstand without compromising quality must be achieved. Applications must be optimized in terms of coating temperatures, application rates, and drying conditions (e.g., RH and airflow rate). Each product must be optimized individually, based on its inherent properties and the desired properties of the final product. Technologies, such as electrostatic coating used for snack food seasoning application (Pannell, 1980; Clark, 1995), may be useful for applying protein coatings to nuts with good uniformity and adhesion. Other expanding technology such as Atomic Force Microscopy (AFM) might be useful for studying the film-forming kinetics of the coatings and the result of particles on the gloss and adhesion of coatings (Goudy et al., 1995; Gilcinski, 1997). Coating methods from the pharmaceutical industry may be utilized to realize new potential in food coatings.

There are possibilities for using protein-based coatings to increase the quality of nuts and nut-containing products during prolonged storage. A solid knowledge base about the physical properties of a large number of protein coatings exists. A number of researchers have demonstrated that protein coatings can be used successfully to add value to nuts and nut-containing products. However, the results are mixed. Application methods and product properties and requirements must be linked to maximize the opportunities for successful applications of edible protein coatings on nuts.

Adams, R.M., Melachouris, N., Tonner, G. and Vadehra, D.V. March 28, 1995. U.S. patent 5,401,518. Agbo, O.F., Anderson, J.C. and Singh, B. 1992. “Lipid Oxidation of Edible Peanut Pastes During Storage with Variation of Environmental and Processing Factors,” Peanut Sci., 19:101–105. Ahmed, H.S. and Storey, J.B. 1975. “Effect of Temperature on the Quality of Vacuum Packed Pecans,” Libyan J. Agric., 4:87–89.

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Protein-based Films and Coatings for Military Packaging Applications1

T

HERE is strong interest in understanding and exploiting the unique capabili-

ties of edible films and coatings within government research, particularly for military applications. It is important to realize that military operational rations must meet stricter storage requirements than those for commercial food products. It is difficult for an edible coating or film to meet these requirements, even though in some cases the edible coating extends shelf stability of the coated product to meet such requirements. The U.S. Army has conducted several edible coating studies over the years to solve problems such as the need to reduce the weight and volume of shelf stable food rations. Needs that foster military requirements and shape the focus of the research and development for military operational rations will be discussed. Supplementary Navy requirements, which drive many of the current studies, will also be discussed. Navy needs differ in that personnel are confined to ships and are charged with restrictions from dumping refuse overboard. This chapter covers several past and present studies conducted by the military dealing with edible coatings and films. Early military studies were con-

1This chapter was prepared by its author as an employee of the U.S. Government as part of his/her of-

ficial duties. There is no copyright on material prepared by a U.S. Government employee as part of his/her official duties.

551

552 ducted to determine the best-suited coatings for specific purposes. These studies evaluated several coatings and eliminated many because of non-suitability. Recent or current studies are reviewed along with possible reasons for either their success or failure. Carbohydrate- and fat-based coatings and films which also were studied will not be discussed here. The objectives of future studies will also be discussed.

When choosing a coating for an application to improve food quality or solve a specific food development problem, one must take into account the physical and functional properties of the coating or film material as well as the composition of the coated food. Physical and functional properties are specific for each film or coating material and are significant when assessing potential applications. Edible films and coatings can be used for a wide variety of purposes; for instance, to replace conventional packaging as a means of protecting food from the environment and to improve food quality. The permeability of coatings to moisture and gases can determine their ability to protect food from atmospheric damage. When atmospheric water or oxygen come into contact with a food product, they can promote degradation reactions. If an edible coating or film can retard moisture and/or oxygen transport, it will act as an additional parameter for improving shelf stability and overall quality of the food product. An edible coating can also improve structural integrity of products, such as dry bars, by strengthening the outer surface to prevent crumbling. Edible coatings that can inhibit fat leaching from a high fat food product can also improve food quality. A list of functions of edible coatings or films to fulfill particular applications is compiled in Table 23.1. Physical and functional properties of a coating material can be manipulated to a certain degree. The classic example is the limitations found when modifying film properties by using additives. The amount of additive needed to achieve a certain goal may surpass that which results in undesirable side effects, and therefore a balance between amount of additives and their effects must be maintained. For example, to manipulate film physical properties, such as flexibility and elasticity, a plasticizing agent is often added. Intermolecular forces responsible for film brittleness can be reduced by appropriate plasticizers, thereby increasing the mobility of polymer chains. However, this also consequently increases film permeability to gas and water vapor (Guilbert et al., 1995). When too much plasticizer is needed to accomplish a desired property change, it will also impact on odor and flavor. These property changes can affect film cohesiveness, continuity, and mechanical handling properties. The method of coating a food product can also affect film barrier properties. Often, the coating method is determined by the characteristics of the specific coating material. For example, certain coating materials are applied in their liq-

Unique Military Needs

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uid form directly to the outer surface of a food product, whereas films that are independent, self-supporting structures can be physically molded to contain a food product. Liquid coating materials can be applied to a food product by spraying, enrobing, dipping, spreading, or dripping into a revolving drum or pan coater (Dreier, 1991). Liquid coatings can also be used in encapsulation, agglomeration, and atomization processing (Sand, 1995). In contrast, other coating materials are extruded or cast into self-supporting films (Guilbert et al., 1995) that can be folded and heat-sealed to forms that resemble traditional packages. Producing coating and film materials generally requires a solvent carrier, such as water and/or ethanol. The drying process used to remove the solvent from the material can affect the uniformity of a coating or film. Ambient air and drying with mild heat are the primary film drying methods. Cracks, pinholes, peeling, non-continuous films, and non-cohesive films can result from rapid evaporation of solvent caused by applying excessive heat in the drying process. To be effective barriers, it is necessary for coatings and films to be continuous, have suitable cohesion to adhere to foods forming a second skin, and be flexible enough, which allows for bending or abuse from handling or to form a package or wrapping in the case of self-supported films (Guilbert, 1986; Cuq et al., 1995). In general, protein-based coatings and films are more permeable to water vapor than lipids and tend to be more capable of forming self-supporting films. The protein in the film acts as the support matrix and is the component that acts as the barrier to water vapor (Koelsch, 1994).

Military rations are designed with the specific needs of the soldier in mind, such as special nutritional requirements and ration configurations, which are dictated by the field conditions and location, and factors such as temperature, water availability, activity, and duration of maneuvers. The combat feeding system’s operational rations are designed for different situations and include

554 general operational rations, special operational subsistence rations, and survival subsistence rations. The shelf life requirement for military rations is a minimum of three years at 27 C (80 F) and six months at 38 C (100 F) (Anonymous, 1992a). It is critical that military rations be suitable for practical utilization or consumption, as well as holding up in extreme scenarios including fluctuating temperatures and rough handling (Wright and Taub, 1998). Modern battlefield requirements demand service support systems that adequately provide for the needs of military personnel in extremely intense and highly mobile combat situations. The military’s science and technology objectives for the Combat Feeding Program include designing a ration that is compact and consumable on the move. Ration development requirements to meet these needs continually involve maximizing convenience while minimizing weight and volume. Rations are therefore designed to be lightweight, portable, easy to eat under a variety of conditions, and of the highest quality within the given requirements. Edible coatings can help meet these objectives by reducing the amount of packaging needed and consequently reducing weight and volume. Edible-coated ration items may be eaten directly without removing the coating, making them consumable on the move as well. Additionally, developmental efforts must not only consider shelf stability but also quality and acceptability. Ration quality is continually improved with advancing processing technology. Increased quality usually means increased ration acceptance. As mentioned, incorporation of edible coatings into rations can also improve the quality of ration items. Furthermore, edible coatings can supply additional nutrients and serve as distinct packaging for small particle foods, such as powders and grains. Generally, they also are low cost and often use renewable resources. A list of these and other potential benefits provided by edible coatings and films, as well as advantages over conventional packaging, are shown in Table 23.2. Despite the mentioned benefits, the Army uses very few edible films in their rations due to performance and functional limitations of edible films in comparison to traditional polymer films. Currently, the military uses microencapsulated products, such as potassium sorbate preservatives, to prevent mold growth in the Meal-Ready-to-Eat (MRE) bread. Potassium sorbate is microencapsulated so that the yeast would not interfere with its activity. A microencapsulated flavor enhancer, glucono delta lactone, is used in intermediate moisture meat sticks. Shellac-based coatings are still used for macroencapsulation of components found in the raisin-nut mix to prevent oxidation of the high fat nuts and moisture transfer between components of different water activities. Other than chocolate coatings, this may be the extent of edible coating usage within the military ration system. However, increased environmental awareness has led to new policies restricting waste disposal. As a result, growing interest in development of biodegradable packaging and edible films has emerged.

Past Studies

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During the 1960s, the U.S. Army was committed to reducing the weight and volume (increase compactness) of military rations. These factors directly affected soldiers’ performance and ability to maneuver on the battlefield. The U.S. Army Natick Soldier Systems Center in Natick, MA, was instrumental in developing processing techniques such as freeze drying and in developing items such as freeze-dried compressed bars to decrease weight and volume of ration items (Blake, 1965; Lampi et al., 1965).

An encapsulation study conducted by Schuetze et al. (1962) investigated barrier materials meeting Food and Drug Administration (FDA) standards that may be suitable for keeping foods confined, moist, and sterile, while maintaining the essential properties of capsule contents under ambient conditions and also at high temperatures and in a high vacuum environment. Food capsules must remain uncontaminated and highly stable for six months without refrigeration. Several foods were coated using FDA-acceptable materials. For example, caramels were spray-coated with zein glazes and acetylated monoglycerides; cooked beef pieces were coated in a laminate consisting of methylcellulose and gelatin films lined with beeswax; and carrots, green beans, and pineapple were encapsulated with methylcellulose/gelatin laminates. No material was singly capable of satisfying all of the requirements. Many of the evaluated samples were not resistant to the unrealistically high temperatures, and were unpalatable and not calorically dense. The temperature requirement for this study was originally set at 71 5 C (160 5 F) and then lowered to 43

556 5 C (110 5 F). When the temperature requirement was lowered, some of the samples were suitable. For example, gelatin was a good film former and could support and serve as the capsule shell. There were severe limitations to scaling-up to manufacturing quantities while ensuring sterility.

Durst (1965) developed food films with various condiment flavorings such as catsup, maple syrup, pickle relish, barbecue sauce, onion gravy, fruit preserves, peanut butter, and soy sauce to accompany and be eaten along with dry, compressed food bars. This study was designed to develop film-configured condiments meant to improve acceptability of dry, compressed food bars without increasing weight and volume. Thirty different prototype condiment sheets were developed; each sheet flavor closely simulated that of its respective condiment. Storage stability tests were conducted on the condiment sheets by placing samples in 4, 23, and 38 C (40, 73, and 100 F) temperature boxes. When two or more film sheets were packaged together and stored for two, four, eight, or 13 weeks, there was no appreciable transfer of flavors from one sheet to another with the exception of flavors with strong volatiles, such as horseradish and pickle relish. Gelatin and sodium caseinate were the primary protein-based materials used as dispersions and self-support matrices. For several condiment film formulas, gelatin and/or sodium caseinate were reportedly the best materials for forming sheets.

The military investigated edible coatings as protective barriers for freeze-dried and dehydrated foods against the effects of atmospheric oxygen and moisture, microbial attack, and mechanical damage (e.g., fragmentation, breakage, abrasion, and erosion) incurred during handling and transport. Coatings were required to not unduly interfere with rehydration and to constitute not more than 16% of the total food bar in weight or volume. Also, they had to be composed only of materials approved by the FDA for human consumption and to, ideally, stand up to a wide range of temperatures (Cole, 1966a). Cole (1966a) conducted an intensive study screening readily available FDA-approved food ingredients as possible materials for edible films. Materials were initially evaluated for structural and barrier characteristics, such as tensile strength, elongation, tear test, water vapor transmission rate (WVTR), oxygen transmission rate (OTR), and equilibrium to relative humidity (RH). Materials that had suitable characteristics were combined into laminates. Those laminates that showed the best barrier properties were applied to dehydrated foods and tested under various conditions. Fragmentation, moisture uptake, oxidation, and resistance to microbial growth were some of the tested

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properties for the coated, dehydrated, and freeze-dried foods. Several protein-based edible coatings were tested, including gelatin, soy proteinate (soy protein isolate), wheat gluten, egg albumen, sodium caseinate, and zein, as well as several non protein-based films. Films prepared by casting, hot press, and coated fiberglass were used for WVTR and OTR analysis (Cole, 1966a). A number of the developed coating compositions exhibited suitable characteristics. Because this study was very extensive, it would be impossible to report all of the results here. However, WVTR values, observations of film properties at varying temperatures, and solubility in water of selected films are discussed here and are listed in Tables 23.3 and 23.4. Soy flour films had a low WVTR of 620 g/m2 day. They also showed low solubility in water at 82 C (180 F), but instead softened and swelled. Wheat gluten films with 20% glycerol had WVTR of 1224 g/m2 day. Films remained flexible at both 23 C ( 10 F) and 49 C (120 F), but softened and became elastic in hot water. Egg albumen films had very high WVTR (1659 g/m2 day), became very brittle at 49 C (120 F), and softened in water but remained insoluble. Sodium caseinate films cast from 50% aqueous ethanol showed a rather high WVTR of 1426 g/m2 day, remained flexible at 23 C ( 10 F) and 49 C (120 F), and were quite soluble in water. WVTR of sodium caseinate films decreased to 620 g/m2 day by substituting glycerol with polyglycerol ester. Cast zein films plasticized with propylene glycol or lactic acid had the lowest WVTR of all the tested protein-based films (589 and 372 g/m2 day, respectively). Zein films were insoluble in water, but they did soften. Gelatin (250 Bloom type A) films of various compositions had WVTR between 859 and 2077 g/m2 day. Higher WVTR values were seen for unplasticized gelatin and gelatin with glycerol. Gelatin films were soluble in water, flexible at freezer temperatures, and dried out and became brittle at 49 C (120 F). Films of unplasticized gelatin ruptured at higher temperatures and lower RH than gelatin films plasticized with glycerol. Laminates were more effective barriers than films composed of homogenous dispersions of two or more components. A laminated film of acetoglyceride and soy proteinate was effective only at high temperature and high RH. A three-layered laminate of amylose laurate, soy proteinate, and acetoglyceride performed well, except under conditions of low temperature and low RH. A laminate film composed of gelatin and monoglyceride showed excellent oxygen barrier properties between 27 and 32 C (80–90 F), but not between 6.5 and 1 C (20–30 F). Proteins showed the best oxygen barrier characteristics among tested film-formers. Cole (1966b) also reported that incorporated glycerol retarded moisture evaporation from films, thereby maintaining film continuity. At levels below 2.5%, glycerol also reduced OTR. Various dehydrated and freeze-dried food products were most effectively coated by edible coatings containing soy proteinate and gelatin.

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During the 1970s, the U.S. Army was exploring the restructuring process of flaked and formed meats. Various edible coatings were tested for their abilities to prevent moisture loss and deleterious oxidative changes occurring in frozen meats during lengthy storage periods required by the military. Most of the studied coatings were either fat- or carbohydrate-based. However, protein-based coatings such as gelatin/propylene glycol (Gelcote , Atlantic Gelatin, Woburn, MA) and soy protein isolate also were tested. Meat portions were dipped into the coating medium, allowed to air dry, packed between sheets of parchment paper, and blast frozen at 29 C ( 20 F) before being stored at 3 C (26 F). After 24 hours, the coated meat portions were visually inspected, grilled, and evaluated for off-odors and flavors. Materials that produced even, translucent coatings without off-odors or flavors were stored at 3 C (26 F) for eight weeks and at 12 C (10 F) for five months. After storage, coated meat samples were again inspected for visual appearance, odor, and flavor. Also, fat oxidation was measured by thiobarbituate TBA method (Shaw and Secrist, 1977). Both of the tested protein-based coatings failed and were not included in the storage phase of this study. The soy protein isolate coatings reportedly cracked at freezer temperatures. A glycerol/gelatin combination was reported to have an acrid odor upon cooking. Also, the gelatin/propylene glycol coating had a bitter taste, which was most likely attributable to the high concentration of propylene glycol.

Over the years, foods that accompanied astronauts into space have undergone several transitions. The Mercury (first manned space flights) flight food systems consisted of semi-solid, sterile, tubed foods of fruits and meat combinations packaged in collapsible aluminum tubes. This menu was supplemented with dry, bite-sized food cubes. These cubes were originally designed to withstand storage temperatures of 27 C (80 F). However, the cubes softened, or even melted, during some Mercury flights. The purpose of early coating studies conducted for the National Aeronautics and Space Administration (NASA) was to prevent free-floating crumbs from fouling the spacecraft’s instrumentation during flight. In addition, coatings had to withstand short-term exposure to temperatures exceeding 55 C (131 F). Therefore, the first coatings used to combat these problems included a fat with a high melting point of 58 C (137 F). However, these coatings were unpalatable and poorly absorbed in the digestive tract, often resulting in steatorrhea (Klicka and Smith, 1982). Further constraints of these studies included special consideration to reduce weight and volume, unavailability of refrigeration, lack of cooking facilities, need for

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ready-to-eat foods or simple food preparation, and food consumption in a weightless environment. Food requirements for the Gemini Project, similar to those for the Mercury Project, were for all food items to withstand temperatures of 43 C (110 F), 100% RH, 100% oxygen atmosphere, cabin pressures of 5.3 psi, launch acceleration, vibration, and acoustic noise without formation of crumbs, dust, or fragmentation. Some problems discovered during the Mercury Project resulted in routine coating of bite-sized food cubes to prevent crumb formation and to reduce cube greasiness or stickiness. Many of the bite-sized food cube formulas for the Gemini Project were also altered to control crumbs. Several coating formulas were developed to meet the needs of various dry, bite-sized cube categories such as entrees, sandwiches, desserts, and cereals. A mixture of water, vegetable oil, sodium caseinate, glycerol, and gelatin was used primarily to coat bite-sized entree cubes such as beef, barbecued beef, beef stew, creamed chicken and turkey, in addition to bite-sized beef, cheese, and chicken sandwich cubes. Aqueous gelatin coatings were applied to dessert cubes (e.g., date and pineapple fruitcake, graham cracker, and ice cream) and apricot, orange/lemon, and strawberry cereal cubes. Some of the food cubes were soaked in gelatin solutions, freeze dried, and then coated along the edges with acetylated monoglyceride. The fruitcake cubes were coated in gelatin and then wrapped in an edible starch paper to prevent stickiness. Combinations of zein, acetylated monoglycerides, citric acid, butylated hydroxyanisole (BHA), and butylated hydroxytoluene (BHT) were used to coat chocolate, coconut, apricot, pineapple, strawberry, and peanut dessert cubes. Special coating combinations of encapsulated protein, fat, and carbohydrate not only prevented crumbling and stickiness, but also preserved freshness. These coatings, which were developed by Pillsbury (Minneapolis, MN), were also used for other dessert items such as brownies and gingerbread (Hollender, 1965). The categories of bite-sized food cubes mentioned above, and others not mentioned, were coated with these formulas until coatings were phased out of the space food feeding system sometime after the Apollo Project (Hollender, 1965). When freezers, refrigerators, and food heaters were installed aboard Skylab and other spacecraft, the need for dry, bite-sized food cubes diminished. Additionally, irradiation technology, shelf stable breads, and improved rehydratable items significantly expanded the variety of available items and further reduced the dependence on dry, bite-sized food cubes (Bourland, 1996).

The military’s interest in edible coatings focuses on the need to reduce packaging waste in the field, which can be accomplished by replacing conventional packaging with edible or biodegradable coatings and films. Other benefits from using edible coatings important to the military include the decrease of troop sig-

562 nature (physical or visual evidence of soldier presence), the decrease in the amount of waste material to be buried or hauled off the field, and the reduction of ration weight and volume. The added convenience of consuming an edible coated ration not only eliminates the need to remove packaging, but also can increase ration intake (and therefore caloric intake) while on the move during extreme environmental and tactical situations. This, in turn, may reduce food waste and increase cost effectiveness of ration items. Increased ration intake may also result from enhanced quality and/or extension of shelf stability of a food item, which is also possible with edible coatings. A project entitled “Multi-Functional Edible Barrier Coatings” which involved the study of edible barrier coatings or films for potential use with military rations was conducted by the U.S. Army Natick Soldier Systems Center. One of the project objectives was to develop sensory guidelines by which commercial and new unique edible materials are evaluated. Sensory characteristics must be factored in when considering the adequacy of a coating or film for a particular application. Edible coatings and films that enhance acceptability, extend shelf stability, and improve food quality would allow the inclusion of more novel ration items into the military system. The primary project objective was to identify edible coatings and films that would serve a useful purpose. The military food ration system has undergone radical changes. Specifically, the military now uses a flexible food pouch for individual shelf stable ration items; there are many ethnic food items; and there are more ration items meant to be eaten on the move such as pocket sandwiches, fruit bars, granola bars, and performance-enhancing items such as high energy bars. The military mission in regard to the ration system now also includes use of rations for emergencies such as disaster relief, refugee situations, and other peacekeeping efforts in different parts of the world. All edible film and coating investigations take into account changes in the rations as well as the military’s changed mission, which resulted from the end of the Cold War. Many aspects encountered with use of edible films and coatings for military rations were studied or considered. Those edible films and coatings that possessed properties to best withstand the abusive storage and handling conditions frequently encountered during military operations (e.g., high temperatures, high RH, prolonged refrigerated storage, and rough handling) were considered for further study. Coatings or films that can enhance acceptability and quality of a ration item were tested for shelf stability. Part of the evaluation involved trials to determine how well film materials are formed and yet another phase to determine method of application. Film forming and casting methods best suited for each application were investigated for military foods including dehydrated and intermediate moisture foods. Other project objectives included reducing the need for conventional packaging or eliminating the primary packaging (the layer closest to the item) while still maintaining safety and stability. The safety of edible coatings and films in

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regard to microbial growth and surface contamination has been questioned. Edible coatings are generally low in moisture. However, if they inadvertently get wet, they can provide nutrition for microbial growth. Because military rations require long-term storage and also are used in atypical and extreme situations, this may give microbes adequate opportunity to grow if the coating is used as the primary package. Although several studies involving several edible coatings and films were conducted, only those based on proteins, such as collagen, gelatin, and corn zein, will be discussed.

The military’s focus is to provide nutritional benefits to soldiers. They must consume the rations, and the odds of consumption are increased if the rations, as well as the edible coating or film, are of high quality and acceptability. Consumers typically have very high standards for food, and these standards are important when considering acceptability. An acceptable edible film or coating should have good eating characteristics or be soluble upon preparation (heating, cooking, or dissolving when mixed with water). Coatings must be compatible with the food and have acceptable color, odor, flavor, and texture or be undetectable. Continuity, adhesion, and flexibility may affect overall appearance and texture. Typically, it is desirable that when dry, coatings and films are not tacky and do not stick to each other and to other packages (Conca and Yang, 1993). Many characteristics such as solubility, moisture and oxygen transmission rate, continuity, adhesion, flexibility, color, and texture can be measured and, therefore, are considered objective measurements. However, they can also be included in subjective sensory evaluations.

Sensory analysis is a subjective method used to evaluate the sensory quality of food products, which, in this instance, are edible coatings and films and/or coated foods. When initial acceptability has been established, military standards dictate that shelf stable products must remain acceptable for a minimum of three years storage at 27 C (80 F). Storage studies are conducted to determine ration acceptability, shelf stability, and quality under these conditions (Ross et al., 1985). Sensory evaluation of quality attributes (i.e., appearance, odor, and texture before eating; and flavor, texture, and mouth-feel during eating) is conducted by a trained panel. Quality attributes are rated using a nine-point hedonic scale with 1 extremely poor and 9 excellent. The elements listed under each of the subjective sensory attributes in Table 23.5 are

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used to characterize and determine acceptability of an edible material. This list also uses several elements that can be measured for appearance and texture to determine acceptability. Sensory characteristics of edible coatings or films must include acceptability, unless the coating or film will be removed prior to food consumption (in such case, only appearance and odor evaluation is needed).

The Navy’s needs are more immediate because of the Marine Pollution Treaty (MARPOL). MARPOL was designed to protect the marine environment from various types of garbage and restricted all ships from dumping plastics and other waste in the oceans. Prior to 1999, the Navy was allowed to discharge plastics outside of certain areas, which included the North Sea, Baltic Sea, and Antarctic Ocean, at least 50 nautical miles from shore if they were out at sea for longer than 20 days and storage space was not available. In 1999, it became illegal for Navy ships to discharge plastics overboard in any location. For submarines, the total worldwide ban for dumping plastics has been extended to 2009.

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The military considers the waste disposal issue as serious. The U.S. Navy is always searching for ways to implement source reduction or to reduce the amount of plastic that comes onboard ships. They have given priority to a comprehensive program that consists of source reduction through material and item substitution, development of reusable containers from biodegradable materials (includes edible films and coatings), and development and installation of onboard solid waste processing equipment (Borgemeister, 1994). The U.S. Army also is exploring source reduction (Derr and Donberg, 1995), biodegradable polymer packaging (Kaplan et al., 1993), and edible packaging (Kester and Fennema, 1986; Conca and Yang, 1993). The Navy has been successful in eliminating plastic waste discharges from ships. A program entitled “Plastics Removal in the Marine Environment (PRIME)” aims to reduce the amount (both by weight and by number of pieces) of plastic in ships’ trash. All plastic waste is separated and stored onboard Navy ships. Plastic waste processors were recently introduced onboard ships to melt plastic waste into 20-inch discs, which are stored until reaching port. Plastic odor-barrier bags are used to trap odors because most discs have some food contamination and storage becomes a health, sanitation, and odor problem.

Use of edible coatings for frozen meats, which allows for direct cooking without coating removal, is a potential solution to the problem of dumping or storing food-contaminated plastic waste. An edible collagen film was studied as a substitute for plastic bags used to ship frozen meats. Collagen is a complex, fibrous, structural animal protein found in skin, connective tissue, and tendons. Extruded collagen films are generally transparent with slight opacity and texture and adhere well to damp food products, such as fresh meat. Such films also are flexible when non-hydrated and become increasingly flexible with increased hydration. Collagen shrinks and can be sealed upon heating. It also is nearly sterile because it is extruded under high heat and pressure before passing through an acid solution. Also, collagen films are quite tasteless and even seem to acquire the flavor of the coated food. Collagen films could be described as ideal based on the sensory characteristics considered in Table 23.5. A commercial collagen film (Profile , Devro Co., Somerville, NJ) composed of 70% protein, 20% humectant, and 10% water was tested as a substitute for the heavy-duty plastic bags currently used in the military system as packing material for distribution and storage of frozen meats. The benefits of collagen-coated beef steaks are that collagen can withstand frozen storage, will prevent leakage during thawing, and can be cooked directly without being removed. Beef cubes (1.6 3.8 6.4 cm) were wrapped in collagen film with the edges heat-sealed. They were then individually blast frozen. The non-coated

566 beef samples were placed in re-closable, heavy-duty plastic freezer bags and then boxed with freezer paper on top and bottom, while coated samples were just boxed with freezer paper. Both were stored at 23 C (10 F) for five months. Samples were rated for sensory quality after 0, 1.5, 3, 4, and 5 months. Non-coated and collagen-coated meat samples were pan-fried from the frozen state without removing the collagen coating. Mean sensory ratings for all samples were within acceptable ranges. In general, higher ratings were given to the coated meat for odor, flavor, texture, and overall quality. Mean ratings for overall quality at each withdrawal period are shown in Figure 23.1. The film tended to hold in meat juices during cooking, thereby making the coated samples more flavorful, tender, and juicy. Texture was measured using a texture analyzer. The force measured during flat plate compression was used to calculate elasticity. Specifically, the ratio of the recoverable work (area under decompression curve) to the total work (area under the compression curve) was the percent recoverable work and was equivalent to the elasticity. There was little or no difference in elasticity between non-coated and collagen-coated beef samples during storage, although differences were observed among withdrawal periods for other cuts of meat (Conca, 1995). These and other storage study results have shown that collagen packaging maintains meat quality at frozen temperatures comparable to that of plastic bags currently used. OTRs of collagen films were measured using a MOCON Ox-Tran oxygen analyzer (Modern Controls, Inc., Minneapolis, MN). Testing conditions were set at 21 C (70 F) and 0% RH (ASTM, 1988; McHugh and Krochta, 1994), and the films had OTR of 0.495–0.504 cm3/m2 day. These OTR values were quite low and notably important because fat contained in beef is highly susceptible to oxidation. Film WVTR was measured using the standard ASTM desiccant method E96-80, and RH was maintained using standard ASTM E104-85 (ASTM, 1980, 1985; McHugh and Krochta, 1994). The WVTR at 23 C (73.5 F) and 50% RH ranged from 169 to 201 g/m2 day. The calculated WVTRs of three samples for four consecutive days are shown in Figure 23.2. These moderately high WVTR values are acceptable for this particular application. Although the moisture content of the meat is relatively high, most moisture will form ice and be fairly immobile during frozen storage. Very little variation in moisture content of beef cubes was noted between coated and non-coated samples and among withdrawal periods. Measured moisture contents for all samples ranged from 64.0 to 71.7% (wet basis). Thermomechanical analysis (TMA) measures linear or volumetric changes in the dimensions of a sample as a function of time, temperature, and force. The data can provide valuable information on coefficients of thermal expansion and softening. Thermomechanical analysis conducted using penetration mode showed film flexibility and resilience for collagen film samples at 40 C

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Figure 23.1 Overall sensory ratings for collagen-coated and non-coated beef cubes stored at 23 C ( 10 F).

( 40 F) while the force ranged from 0 to 1 N (Figure 23.3). The thickness of collagen samples measured ranged within 0.030–0.095 mm. Moisture contents of collagen films were measured using a thermogravimetric analyzer (TGA). The weight change was calculated at 100 C (212 F), where a definite peak observed in the derivative weight curve indicated water loss. Measured values obtained with TGA were 4.25–6.22% for dry films and 77.51–86.99% for hydrated films. These measurements were somewhat lower than measurements obtained using the conventional oven drying method.

Figure 23.2 Water vapor transmission rate of collagen film. Dry cup method with conditions set at 23 C (73.5 F) and 50% RH.

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Figure 23.3 TMA scan of collagen film (0.074 mm thick) with dimensional change seen at 40 C ( 40 F) while force ranged from 0 to 1 N.

The glass transition temperature and how it changes is important in predicting food shelf life and quality (Reid, 1996; Roos et al., 1996). Temperature, water, and time-dependent changes that occur in foods during storage can be reduced by not exceeding their determined critical glass transition temperatures. Reformulation of a product can provide sufficiently higher or lower critical glass transition temperatures depending on the product. A distinct glass transition was seen using a differential scanning calorimeter (DSC) at 98.7 C (210 F) upon heating dry collagen film. A glass transition is the conversion of an amorphous material from a solid glassy state to a supercooled rubbery state. Water, which is essentially a plasticizer, depresses the glass transition temperature (Roos et al., 1996). Because collagen-coated meat is a high moisture system, this principle will apply. After collagen samples were hydrated with water, the glass transition peak shifted to 4 C (25 F). These two DSC scans are shown in Figures 23.4 and 23.5.

Gelatin is the product of partially hydrolyzed collagen. It forms thermo-reversible gels when heated in water and, therefore, is moderately susceptible to moisture. The WVTR ranged within 32–56 g/m2 day for gelatin films with varying thicknesses of 0.326-0.575 mm. However, gelatin is an effective fat and oxygen barrier (Anonymous, 1992b). OTR ranged within 0.0268–0.1172 cm3/m2 day for the same gelatin film samples. Gelatin has been used extensively in liquid, semi-liquid, and film form.

Figure 23.4 DSC scan for two dry collagen film samples. Glass transition peak (Tg) seen at 98.7 C (210 F).

Figure 23.5 DSC scan for two hydrated collagen film samples. Glass transition peak (Tg) seen at 4 C (25 F).

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570 Soft gelatin capsules can hold a variety of condiment-type food products. These gelatin fillings must be low in moisture (typically high fat or sugar), flowable, and shelf stable. The capability to formulate and control its solvation rate and tensile strength has given gelatin a flexible advantage (Anonymous, 1990). The military was interested in evaluating gelatin macroencapsulated condiments such as peanut butter, jellies, butter, margarine, syrups, and sauces to accompany and enhance existing military rations. The U.S. Army Natick Soldier Systems Center, with assistance from Banner Pharmacaps Inc. (High Point, NC), considered various soft gelatin capsule shapes (e.g., pillows, spheres, bears, and capsules with break-off tabs). These individually macroencapsulated condiments could be co-packed with other food items appropriate for each condiment. The initial proposals consisted of using encapsulated condiments with shelf stable waffles and other baked items. Margarine and/or syrup encapsulated in gelatin capsules could be co-packed with waffles. The capsules would then melt and release their contents upon heating at 60 C (140 F) for about 20 min (Anonymous, 1992b). An accelerated storage study for shelf stable ration items requires four weeks at 52 C (126 F). A storage study was conducted on co-packed soft gelatin capsules with peanut butter and shelf stable MRE bread in the military trilaminate pouch under accelerated storage conditions. The capsule shells consisted of gelatin, water, and glycerol. The initial water activity (aw) of gelatin-encapsulated peanut butter was quite low (0.275–0.306), and the aw of the bread was 0.916. There was significant moisture transfer from the bread to the gelatin. The aw of the gelatin shell increased to an average of 0.755 after only two weeks of storage and to 0.774 after 4 weeks of storage. The average aw values of six samples at each withdrawal period are shown in Figure 23.6.

Figure 23.6 Water activity of gelatin-encapsulated peanut butter and bread when co-packed and stored at 52 C (126 F).

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Gelatin-encapsulated peanut butter was separately tested beforehand at accelerated storage temperatures and found not to melt in a closed container. However, after two weeks storage at 52 C (126 F), the gelatin capsule had completely collapsed and melted away from the peanut butter when co-packed with the MRE bread. The moisture transfer from the bread increased the aw of the gelatin, most likely lowering the gelatin melting point below 52 C (126 F). Use of secondary coating systems to improve capsule resistance to high heat and RH also were investigated. Specifically, soft gelatin capsules were coated with both the lipid-based ingredient Capol (Centerchem, Inc., Stamford, CT), which has moisture and oxygen barrier properties, and with a thin layer of beeswax (moisture barrier). These coated capsules also failed under the same test conditions. Gelatin’s susceptibility to moisture hinders co-packing gelatin with higher moisture products at relatively high storage temperatures. The WVTR of gelatin films is largely a function of RH due to the hygroscopic nature of gelatin (Glicksman, 1982). TMA was conducted on the original gelatin capsule shells. As expected, little flexibility was detected at the lower temperatures of 40 C ( 40 F) and at 25 C (77 F) with dimensional changes of only 3.8 and 12%, respectively. However, considerable mobility was detected at the higher temperatures of 52 C (126 F) and 200 C (392 F) with dimensional changes of 47 and 97%, respectively. The thickness of tested gelatin shells was 0.3–0.6 mm. TMA scans at 40, 25, 52, and 200 C are shown in Figures 23.7, 23.8, 23.9, and 23.10, respectively. These four figures showed increased mobility or softening of the gelatin as indicated by increasing dimensional changes with higher temperatures.

Zein, derived from corn gluten meal, belongs to a class of hydrophobic proteins known as prolamins. Because zein is insoluble in water, zein-based coating formulations often use organic solvents such as ethanol, acetone, and acetonylacetone (Gennadios and Weller, 1990). Zein’s low solubility in water is attributed to its high content of non-polar amino acids. Typically, zein is suspended in alcohol or acetone solutions and is applied onto items by spraying, tumbling, or dipping (Dreier, 1991). Because of its natural sheen, zein is commonly used to coat confections (Andres, 1984). Initially, two zein products supplied by Zumbro, Inc. (Hayfield, MN) and by Enzytech Food Ingredients, Inc. (Cambridge, MA; currently Opta Food Ingredients, Inc.) were tested for their film-forming properties. Films were formed by pouring or spray casting. The sample from Zumbro (Cozeen ) was a combination of alcohol, zein, vegetable oils, glycerin, citric acid, BHA, and BHT. Zein interacts with and dissolves in alcohol when in the liquid state. Then, the alcohol evaporates, and zein hydrophobically bonds with vegetable oil to pro-

Figure 23.7 TMA scan of gelatin film (0.343-mm thick) with dimensional change seen at 40 C ( 40 F) while force ranged from 0 to 1 N.

Figure 23.8 TMA scan of gelatin film (0.363-mm thick) with dimensional change seen at 25 C (77 F) while force ranged from 0 to 1 N.

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Figure 23.9 TMA scan of gelatin film (0.367-mm thick) dimensional change seen at 52 C (126 F) while force ranged from 0 to 1 N.

Figure 23.10 TMA scan of gelatin film (0.633-mm thick) dimensional change seen at 200 C (392 F) while force ranged from 0 to 1 N.

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574 duce a film (Andres, 1984). This product is intended for panning and did not produce a smooth nor continuous film. When tested on nuts, it did not significantly reduce lipid oxidation. The film was yellow, slightly greasy, and had a strong odor. The zein sample from Enzytech (Lita ) was functionally modified using a proprietary technology to produce water-soluble zein. Cast films from this material were also non-continuous. They were white and flaked apart easily. Varying amounts of polyethylene glycol (PEG) 400 were tested to increase film flexibility and continuity. Low levels of PEG (e.g., 2%) with and without ethanol formed fragile and brittle films, which were tough and did not break down upon chewing. Higher levels of PEG (e.g., 4%) with added ethanol formed films that were more flexible, but that were still very fragile and not resistant to tearing. Zein formulations supplied by Opta Food Ingredients, Inc. (Cambridge, MA) containing 13% zein solids, approximately 50% less than in previous formulations, which allowed atomized spray coating, also were tested. Zein in these formulations was purified enzymatically to remove residual starch, and alkali was used to remove residual fatty acids. It also was decolorized and deodorized. There was an alcohol and a water-based latex version of this formulation. The alcohol-based zein (Optazein ) was chosen for its quicker drying properties. It contained 13% solids, 80% ethanol, and 5% glycerol (added to aid in spreading and to impart flexibility). Cast films became tacky within 3 min and dried completely within 5 min. Sample films were tested for WVTR using the method described above for collagen films. The films were 0.18-mm thick, and the testing conditions were 50% RH and 24 C (75 F) with an effective film test area of 5 cm2. WVTRs fluctuated within 102–134 g/m2 day. This formulation was tested as a coating to improve quality of nuts in the “raisin nut mix,” an item found in the military Cold Weather Ration. This product is packaged in the military trilaminate pouch. The aw was measured after the packaged nut and raisin assembly had equilibrated. The aw values of the coated “nuts only” and for the “whole mix” were 0.483 and 0.476, respectively.

A latex coating material from wheat gluten was tested alongside the zein coating mentioned above in the “raisin nut mix” storage study. The wheat gluten coating (Optaglaze ), also supplied by Opta Food Ingredients, was a water-based colloidal suspension of a water insoluble protein that was formulated mainly for baked goods. It contained about 14% solids, of which 90% was protein solids. Cast films became tacky after 5 min and were dry after 15 min. Sample films were tested for WVTR using the method described for collagen films. The tested films were 0.09-mm thick, and the rates measured were within 130–159 g/m2 day.

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Roasted peanuts were pan-coated with zein- and wheat-based coatings (2% solids uptake after drying). The “raisin nut mix” was assembled and packaged in military pouches using the coated nuts. After the product had equilibrated, the aw was measured. Nuts coated with wheat gluten had aw of 0.545, and the whole mix had aw of 0.540. Samples were then stored at 27 C (80 F) or 38 C (100 F) for a three-year storage study. Product was withdrawn from storage, and sensory analysis was conducted initially and after six, 12, 18, 24, and 36 months of storage. The study is still in progress, but available results showed that zein-coated samples had higher sensory ratings and held up better than wheat gluten-coated samples. Wheat gluten-coated and zein-coated samples stored at 38 C (100 F) were dropped from the study after 12 and 18 months of storage, respectively, due to low sensory ratings. Zein-coated samples received a mean rating of 6.2 for appearance, odor, and texture, and of 5.5 for flavor and overall quality after 24 months at 27 C (80 F).

Future study of edible films and coatings by the military will focus on use of inter-component films and coatings to act as barriers between components of multi-component ration items. For example inter-component films will be placed between food layers during preparation and processing to improve product shelf stability and to enhance the impact on appearance and acceptability by preventing component ingredients from commingling and mixing (e.g., peanut butter, jelly, and bread; filling with sauce and bread). Inter-component films and coatings can also prevent cross-contamination of flavors, fat oxidation of a high fat component by entrapped oxygen within another component, and moisture and fat transfer among components. A decrease in moisture transfer can reduce the risk of creating an environment favorable for bacterial growth in minimally processed food items. Certain films will be chosen or designed to melt at appropriate temperature ranges providing selective barriers to primarily moisture until the final stages of processing. Other films will be chosen or designed to reform after processing or not to melt at all. Incorporation of preservatives in films will reduce the amount of preservatives or antioxidants added directly to a food product. The type of rations considered for this project are the new shelf stable bread items such as layered pocket sandwiches, burritos, and pizzas. These items are highly desirable, and inter-component films can enhance their quality and increase their acceptability as well as their shelf life.

Andres, C. 1984. “Natural Edible Coating Has Excellent Moisture and Grease Barrier Properties,” Food Process., 45(13):48–49.

576 Anonymous, 1990. “Exciting New Uses for Gelatin Encapsulation,” Good Packaging Magazine, Issue No. 12. Anonymous, 1992a. “Operational Rations of the Department of Defense,” U.S. Army SBCCOM, Natick Soldier Center, Natick Pamphlet 30-25/3rd Edition. Anonymous, 1992b. “Ingredient Capsules Beg Processing Solutions,” Prepared Foods, 161(12):81. ASTM 1980. “Standard Test Method for Water Vapor Transmission of Materials” in Annual Book of ASTM Standards, Vol. 4.06, Method E96-80, Philadelphia, PA: American Society for Testing and Materials, pp. 745–754. ASTM 1985. “Standard Practice for Maintaining Constant Relative Humidity by Means of Aqueous Solutions” in Annual Book of ASTM Standards, Vol. 4.06, Method E104-85, Philadelphia, PA: American Society for Testing and Materials, pp. 695–696. ASTM 1988. “Standard Test Method for Determining Gas Permeability Characteristics of Plastic Film and Sheeting” in Annual Book of ASTM Standards, Vol. 15.09, Method D1434-82, Philadelphia, PA: American Society for Testing and Materials, pp. 255–266. Blake, J.H. 1965. “Study of Carrier Gas Freeze-Drying at Low Pressures,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/FD-30. Borgemeister, USN Lt. L. 1994. “U.S. Navy Update,” R&D Associates, Activities Report, Boston, MA 46(1):117–121. Bourland, C. 1996. “Food for Space Flight,” NASA Facts, NP-1996-07-007JSC. Cole, M.S. 1966a. “Edible Coatings for Dried and Compacted Foods: Part I,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/66-38-FD. Cole, M.S. 1966b. “Edible Coatings for Dried and Compacted Foods: Part II,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/66-37-FD. Conca, K.R. 1995. “Evaluation of Collagen-based Film as an Edible Packaging Material for Frozen Meats,” Paper 5–6 presented at the Annual Meeting of the Institute of Food Technologists, Anaheim, CA, June 3–7. Conca, K.R. and Yang, T.C.S. 1993. “Edible Food Barrier Coatings” in Biodegradable Polymers and Packaging, eds., C. Ching, D.L. Kaplan and E.L. Thomas, Lancaster PA: Technomic Publishing Company, Inc., pp. 357–369. Cuq, B., Gontard, N. and Guilbert, S. 1995. “Edible Films and Coatings as Active Layers” in Active Food Packaging, ed., M.L. Rooney, London: Chapman & Hall, pp.111–142. Derr, J. and Donberg, D. 1995. “Spreading the Word on Source Reduction,” Converting Magazine, 3:56–61. Dreier, W. 1991. “The Nuts and Bolts of Coating and Enrobing,” Prepared Foods, 160(7):47-48. Durst, J.R. 1965. “Food Adjuncts Stabilized as Thin Sheets or Laminates,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/FD-29. Gennadios, A. and Weller, C.L. 1990. “Edible Films and Coatings from Wheat and Corn Proteins,” Food Technol., 44(10):63–69. Glicksman, M. 1982. Food Hydrocolloids, Vol. 1, Boca Raton, FL: CRC Press, pp. 47-100. Guilbert, S. 1986. “Technology and Application of Edible Protective Films” in Food Packaging and Preservation: Theory and Practice, ed., M. Mathlouthi, New York: Elsevier Applied Science Publishers, Ltd., pp. 371–394. Guilbert, S., Gontard, N. and Raoult-Wack, A.L. 1995. “Superficial Edible Films and Osmotic Dehydration: Application of Hurdle Technology without Affecting the Food Integrity” in Food Preservation by Moisture Control: Fundamentals and Applications, eds., G.V. BarbosaCanovas and J. Welti-Chanes, Lancaster, PA: Technomic Publishing Company, Inc., pp. 305–323. Hollender, H.A. 1965. “Technology of Space Foods,” R&D Associates, Activities Report, 17:19–31. Kaplan, D.L., Mayer, J.M., Ball, D., McCassie, J., Allen, A.L. and Stenhouse, P. 1993. “Fundamentals of Biodegradable Polymers” in Biodegradable Polymers and Packaging, eds., C. Ching, D.L. Kaplan and E.L. Thomas, Lancaster, PA: Technomic Publishing Company, Inc., pp. 1–42.

References

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Kester, J.J. and Fennema, O.R. 1986. “Edible Films and Coatings: A Review,” Food Technol., 40(12):47–59. Klicka, M.V. and Smith, M.C., Jr. 1982. “Food for U.S. Manned Space Flight,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/TR-82/019. Koelsch, C. 1994. “Edible Water Vapor Barriers: Properties and Promise,” Trends Food Sci. Technol., 5:76–81. Lampi, R.A., Takahashi, H., Lennon, J., Battey, R.F. and Sierra, S. 1965. “Ultra-High Compression of Dried Foods,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report/FD-35. McHugh, T.H. and Krochta, J.M. 1994. “Permeability Properties of Edible Films” in Edible Coatings and Films to Improve Food Quality, ed., J.M. Krochta, E.A. Baldwin and M.O. Nisperos-Carriedo, Lancaster, PA: Technomic Publishing Company, Inc., pp. 139–187. Reid, D.S. 1996. “Effect of Glass Transitions in Foods,” Food Technol., 50(11):20. Roos, Y.H., Karel, M. and Kokini, J.L. 1996. “Glass Transitions in Low Moisture and Frozen Foods: Effects on Shelf Life and Quality,” Food Technol., 50(11):95–108. Ross, E.W., Klicka, M.V., Kalick, J. and Branagan, M.E. 1985. “Acceptance of a Military Ration after 24-Month Storage,” J. Food Sci., 50:178–181, 208. Sand, C.K. 1995. “Technology and Packaging Applications of Edible Barrier Research,” Presented at IOPP Technology of Packaging Conference, Chicago, IL, November 3–4. Schuetze, C.E., McMahon, W.E., Adams, L.M. and Barnes, W.M. 1962. “Encapsulation of Foods,” Southwest Research Institute, Biomedical Laboratory, Aerospace Medical Division, Technical Documentary Report MRL-TDR-62-53. Shaw, C.P. and Secrist, J.L. 1977. “Edible Coatings for Individual Frozen Meat Portions,” U.S. Army SBCCOM, Natick Soldier Center, Technical Report / TR-77/028. Wright, B.B. and Taub, I.A. 1998. “Stored Product Quality: Open Dating and Temperature Monitoring” in Food Storage Stability, eds., I.A. Taub and R.P. Singh, Boca Raton, FL: CRC Press, pp. 353–367.

Edible Films and Coatings as Aroma Barriers

F

OODS must be preserved to maintain their initial qualities and properties by

protecting them from the surrounding environment and by limiting loss of some of their components. Food satisfies the biological need for a source of nutrition, but it is its flavor and aroma that provide the impetus to the consumer. Packaging reduces the transfer of gases, vapors, liquids, and solids between the food and its environment, thus extending food shelf life. Migration of moisture, oxygen, or various solutes, including aroma compounds, can occur, inducing deleterious changes in sensory, nutritional, microbiological, and/or physicochemical characteristics of food. Loss of volatile compounds diminishes flavor intensity, thus changing the aromatic note of the food product. Moreover, absorbed off-flavors permeating through packaging from the environment can modify sensory characteristics of the food. Finally, solvents, monomers, or residues originating from package (mainly plastics) manufacturing may come in contact with the packaged food. For these reasons, food processors reduce or control interactions and transfers of small volatile molecules by using appropriate multi-layered packaging films (Mohney et al., 1988). Such films are usually highly efficient, but they also are expensive and often non-recyclable. Therefore, the potential use of biodegradable or edible films and coatings to improve the aroma barrier ability of cheaper plastics or to retain aromas within foods has been receiving interest. 579

580

Aroma is the combined effect of the volatile compounds in food able to provide an olfactory perception to the consumer. These compounds are organic volatile molecules perceived by the human nose at extremely low concentrations. Therefore, the loss or absorption of even minuscule amounts of such volatiles can possibly be detectable by consumers. Consequently, it is fundamental that food remains stable in terms of aroma concentration and composition during the shelf life of the product. Compounds that comprise an aroma are molecules similar to most organic solvents (e.g., alcohols, esters, ketones, and aldehydes); have low molecular weight (g/mol); have a vapor pressure high enough to be partially in the vapor state at normal storage conditions; and, especially, are generally hydrophobic. This explains the strong affinity typically exhibited by aroma compounds for non-polar and hydrophobic polymers, such as most plastics used in packaging (Leufven and Stöllman, 1992). Because of their size and nature, volatile compounds interact with plastic polymers by adsorption (“scalping”) and by permeation, which can occur from the surrounding environment to the food (migration in), or the opposite (migration out). Adsorption and permeation can modify polymer film properties through plasticization, partial solubilization, or decreased crystallinity. Such phenomena can increase permeability, change appearance (opacification), reduce adhesion (multi-layered films), or form cracks. Aroma retention in food can be compromised by loss of package integrity (Shimoda et al., 1987; Sadler and Braddock, 1990). Damaged packages can alter food sensory quality and decrease food shelf life due to microbiological contamination, color and texture changes, biochemical and enzymatic reactions (mainly due to oxygen penetration), and, especially, the loss of aroma or the generation of off-tastes and off-flavors.

The significant loss of volatile compounds from fruit juice stored in packages based on polyethylene (PE), ethylene/vinyl alcohol copolymer (EVOH), or polyethylene terephthalate (PET) has been documented (Dürr et al., 1981; Charara et al., 1992; Konczal et al., 1992). For example, flavor losses from orange juice stored at 4 C for 20 days included 1 to 9% of alcohols, 0.3 to 64% of aldehydes, and 2 to 85% of esters. Such loss of volatiles is more substantial with packages from polyolefins, such as polyethylene (PE) or polypropylene (PP),

Control of Flavor Interactions and Transfers by Edible Barriers

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than with packages from EVOH or polyvinylidene chloride (PVDC). EVOH and PVDC are usually recommended as aroma or oxygen barriers because they have aroma and oxygen permeability values that are lower by 1000 to 100,000 times than those of PE (Lebosse et al., 1997). However, EVOH and PVDC are not heat sealable. Moreover, EVOH is a moisture-sensitive polymer. For these reasons, EVOH and PVDC are usually used as internal layers in composite films, with the sealable layer in contact with the outer environment or with the food product often being PE. These composite films are usually very potent barriers against aroma transfers because of their low permeation. However, they also trap (scalp) flavor compounds, which typically have high affinity for the PE layer. Therefore, even such composite packages do not fully preserve the sensory characteristics of packaged foods (Fayoux et al., 1997a, b). For example, sensory evaluation scores for orange juice decreased by half after one week of storage in a package composed of PE-paperboard-PE-aluminum-PE, a common “brick” package used for fruit juice. After six weeks, the hedonic response for the orange juice represented only 20% of the initial value, and 25% of the ascorbic acid was lost (Moshonas and Shaw, 1989). Besides flavor loss from the product, sensory characteristics can also change through the release of volatiles from the package. Stöllman (1995) reported that recycled bottles of PET that initially contained cola or citrus soda drinks gave off-tastes and off-flavors to packaged water and soft-drinks, even after thorough washing of the bottles with hot water and detergents before reuse. The above discussion indicates some of the challenges facing the food industry in regards to packaging of flavored foods.

Even though most published research focusing on interactions between packaging and volatile compounds is related to beverages, similar problems occur for viscous liquids, gels, and solid foods. A suggestion for decreasing interactions between aroma and plastics is to retain flavor molecules inside the food product by using an additional barrier. An edible film or coating that has low permeability to aroma transfers and can be eaten along with the protected food can serve as the additional barrier (Callegarin et al., 1997; Miller and Krochta, 1997; Debeaufort et al., 1998). Essentially, this is a concept of macroencapsulating the food product (Figure 24.1). The main envisioned application is using the edible barrier to retain aroma within the food and also to protect the food from volatiles originating in the packaging or surrounding environment. However, the edible film or coating may also be used to carry or support flavors at the product surface. These flavors will then be promptly released upon consumption of the product. In fact, a few products that use this flavoring concept have already been commercialized. For instance, there are

582

Figure 24.1 The concept of combining traditional packaging and edible barriers to improve aroma retention in stored food.

roasted peanuts with a curry-flavored coating that instantaneously dissolves in the mouth providing the perception of the Indian spice. Another example involves application of multiple sugar coatings on sweets with each coating containing different tastes and flavors. Gum arabic or another hydrocolloid layer separates the coatings to prevent aroma migration. For such an application, diffusivity of volatiles should be very low, and the volatiles should have high affinity for the highly soluble coating. The mentioned edible coatings are mainly composed of one or more film-forming materials, which provide matrix cohesiveness (continuous network) and a barrier against aroma migration. Various polysaccharides, proteins, and lipids are used alone or in mixtures as film-formers and/or barriers. A single substance can simultaneously be a good film former and a potent barrier, such as wheat gluten, which forms films with satisfactory mechanical resistance and very low oxygen permeability (Gontard et al., 1994). Also, whey protein-based films reportedly had lower permeability of d-limonene (one of the main flavor compounds in citrus) than EVOH or PVDC films (Fayoux et al., 1997a, b; Miller and Krochta, 1997). The functionality of edible films and coatings strongly depends on their composition. Protein- and polysaccharide-based films and coatings usually have good organoleptic and mechanical properties. Also, they are effective barriers against non-condensable gases (e.g., oxygen, carbon dioxide, and nitrogen), aroma compounds, and oils, but are highly permeable to moisture. In contrast, lipid-based coatings are good moisture barriers, but are poor oxygen and aroma barriers and have poor organoleptic and mechanical properties (Debeaufort et al., 1998; Quezada-Gallo et al., 2000). The efficiency of an edible coating to retain or limit transfers of aroma compounds is the highest if it has low affinity for volatile compounds and low diffusivity. Therefore, solubility, diffusivity, and permeability values of the film have to be low. By controlling either solubility or diffusivity of aroma in biopolymer networks, aroma losses can be significantly reduced. Research on model solutions mimicking a traditional Italian liquor showed that edible films

Aroma Barrier Properties of Edible Films and Coatings

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composed of whey proteins, wheat gluten, and oils retained 60 to 99.9% of the aroma compounds usually lost in control samples after eight days of storage at ambient temperature (Sensidoni et al., 2001). The results of the study on the model system were also validated by applying the edible films on liquor-stuffed chocolate pralines (Sensidoni et al., 2001).

The increasing need for barrier materials requires extensive measurements of permeability values, but also solubility and diffusivity values. Although moisture and oxygen permeability testing methods are well known and standardized, limited information is available on determining aroma permeabilities of both edible and plastics films. Various laboratories have developed their own apparatuses for measuring flavor permeability, and three systems have been commercialized. Mainly static or dynamic isobaric methods have been used. Isobaric methods use permeation cells with two compartments that are separated by the tested film specimen. The overall pressure is the same in both compartments, but one compartment contains a known concentration or vapor pressure of the volatile compound. In the other compartment, the concentration of the volatile compound transferred through the film is determined over time. In these methods, qualitative and quantitative analyses of the vapor phase are generally performed by gas chromatography using flame ionizing or catharometric detectors. With the static method, one compartment of the permeation cell contains the pure aroma compound in the liquid state or diluted in a solvent of low volatility, such as water (Figure 24.2). The other compartment is tightly closed, and the vapor phase is periodically sampled with a syringe or an automated sample valve and is analyzed by gas-liquid chromatography (GLC) (Apostolopoulos and Winters, 1991; Durand, 1996). The main advantage of this technique when

Figure 24.2 Schematic representation of the static isobaric method for aroma permeability determination.

584 using a dilute solution of flavor is that it better simulates what really occurs among the food product, its package, and the surrounding environment. Inversely, because of the tightness of the receiving compartment, the aroma concentration gradient between the two sides of the tested film decreases with time, and thus the transfer rate decreases too. Therefore, in these conditions, the permeability and the transfer rate cannot be determined at steady state and have to be extrapolated. For this reason, the dynamic method is generally replacing the static method. The dynamic method uses the same principle as the static method. However, the two cell compartments are continuously swept by a carrier gas (e.g., helium, nitrogen, hydrogen, or air) containing one or several flavor substances in the upper side of the cell (Gavara and Hernandez, 1993). At the other side of the film, the carrier gas is free of aromas, but can be wet or dry. The aroma-carrying gas is usually generated by stripping through the pure compound and diluting with an aroma-free carrier gas flow, or by stripping through a dilute aroma solution. The main disadvantage of stripping through a dilute aroma solution is the change of aroma concentration over time as the carrier gas continues to extract flavors from the solution. Gas stripping through pure compounds allows for mixing several carrier gas flow streams and for blending several aroma compounds and also oxygen or water vapor. Debeaufort and Voilley (1994) developed an apparatus that simultaneously measures the permeability of four aroma compounds, oxygen, and moisture (Figure 24.3). The main advantage of the dynamic method is maintaining the aroma concentration gradient constant during the experiment, thus allowing permeability to be determined at steady state. However, it is critical to eliminate stagnant layers in the cell that can lead to underestimation of permeability values, especially when testing highly potent aroma barriers such as EVOH or protein-based films. An aroma permeability measurement system (MAS2000 ) that uses the dynamic method was developed by MAS Technologies (Zumbrota, MN, USA). It uses only an FID detector at the cell exit, which gives a signal proportional to the aroma transfer rate of the tested film sample. However, it only works at saturated atmospheres (very high aroma concentrations). Moreover, only a combined permeability value is obtained when a mixture of volatile compounds transfers through the film, because there is no chromatographic column to separate each compound. The Lyssy GPM 500 (LYSSY, Zolikon, Switzerland) instrument is based on the same system shown in Figure 24.3, but it can only measure at moisture-free environments. Several other modified apparatuses have been proposed. Benet et al. (1992) placed a cryofocuser between the permeation cell and the gas chromatograph, which allowed entrapment and concentration of permeated aroma vapors before analysis. They obtained a very low detection threshold, and thus they could measure very low permeability values in the case of plastic bottle corks. This system (Aromatran ) is commercially available from Modern Controls Inc. (Minneapolis, MN, USA). The dynamic

Aroma Barrier Properties of Edible Films and Coatings

585

Figure 24.3 System developed by Debeaufort and Voilley (1994) and improved by Quezada-Gallo et al. (1999) for simultaneous measurement of film permeability to flavor compounds, gases, and water vapor.

technique has most commonly been used to quantify aroma transfers through synthetic polymers (Gilbert et al., 1983; Delassus et al., 1988; Franz, 1993; Thompson et al., 1994; Doyon et al., 1996) and also edible films (Debeaufort and Voilley, 1994, 1995, 1997; Debeaufort et al., 1995; Miller and Krochta, 1997; Miller et al., 1998; Quezada-Gallo et al., 1999). Regardless of the technique used to measure aroma permeability of edible films, particular attention must be given to controlling temperature and relative humidity (RH) conditions. Aroma permeability, solubility, and diffusivity vary according to the Arrhenius law: P

P0 exp( EaP / RT )

D

D0 exp( EaD / RT )

S

S0 exp(

HS / RT )

where EaP and EaD are the apparent activation energies of permeation and diffusion (kJ/mol); HS is the enthalpy of sorption (kJ/mol); P0, D0, and S0 are the reference values of permeability, diffusivity, and solubility, respectively; R is the ideal gas constant (8.31 kJ/mol·K); and T is the absolute temperature (K). From the definition of permeability, which is the product of solubility and diffusivity coefficients, the following relationship exists: EaP

EaD

HS

EaD is always positive, and D increases with temperature (Rogers, 1985). Conversely, HS depends on the sorption mechanism of the volatile compound in the edible film and can be either positive or negative. When sorption is

586 mainly due to hydrophobic interactions between flavor and film components, S increases with temperature. When sorption is principally driven by hydrogen bonding, S decreases with increasing temperature (Moisan, 1980). Consequently, the effect of temperature on aroma permeability cannot be easily predicted (Quezada-Gallo, 1999). In most cases, moisture tends to increase the aroma permeability of edible films and coatings (Debeaufort and Voilley, 1994; Miller et al., 1998). Water acts as a plasticizer, increasing the molecular mobility of biopolymer chains within the edible film structure, which facilitates aroma diffusion and transfer through the film. However, Quezada-Gallo (1999) observed that for highly hydrophobic compounds, such as 2-nonanone and ethyl octanoate, permeability decreased when the moisture content of the carrier gas increased. This was explained by a lower solubility of these flavor compounds when films were wet. Therefore, the aroma barrier ability of edible films can be notably affected by changes in both temperature and RH.

d-Limonene is the most studied flavor compound. It is considered the typical aroma probe of citrus juice and soda beverage. Figure 24.4 compares d-limonene permeability values (at 25 C and 0% RH) of selected edible and plastic films. Methylcellulose films had similar permeability to heat-sealable polymers, such as polyolefins. Protein-based films had d-limonene

Figure 24.4 d-Limonene permeability values of various plastic and edible films at 25 C and 0% RH. EVOH ethylene vinyl alcohol copolymer; PET polyethylene terephtalate; BOPP Biaxially oriented polypropylene; Co-VDC vinylidene copolymer; HDPE high-density polyethylene; LDPE low-density polyethylene; and EVA ethylene vinyl alcohol (Franz, 1993; Kobayashi et al., 1995; Quezada-Gallo, 1999; Paik and Writer, 1995; Miller and Krochta, 1997).

587

588 permeabilities as low as those of the most effective plastic aroma barriers (Figure 24.4). For example, a d-limonene permeability of 10 18 g/m s Pa was reported for wheat gluten-based films and of 10 14 to 10 16 g/m s Pa for whey protein-based films (Miller et al., 1998; Quezada-Gallo, 1999). Although d-limonene is probably the most studied flavor compound in the field of polymer permeability, it is not often used for flavoring solid food products. Therefore, Debeaufort and Voilley (1995) and Quezada-Gallo et al. (1999) studied mass transfers through edible films of several other aromatic molecules, commonly found in dairy products and fruits. As shown in Table 24.1, permeability (at 25 C and 0% RH) of methylketones and ethyl esters was much lower (by two to five orders of magnitude) through protein-based films than through carbohydrate-based films. These results suggest that protein-based films merit consideration as protective layers against scalping and permeation of aroma vapors through polyolefins such as PE.

Barrier properties of edible films and coatings, as often observed for synthetic polymers, are affected by several parameters, such as external conditions, physicochemical characteristics of the permeant, and biopolymer nature and structure.

The main external factors affecting aroma transfer through film packaging are aroma concentration gradients, temperature, and RH. With non-condensable gases permeating through synthetic polymers, the transfer rate increases linearly with the concentration gradient. However, for flavor compounds and biopolymer films, this behavior is scarcely observed. An exception was the permeation of 2-octanone and 2-nonanone through methylcellulose films plasticized with 25% polyethylene glycol 400 (Figure 24.5) as reported by Quezada-Gallo et al. (1999, 2000). In contrast, 2-pentanone, 2-heptanone, ethyl esters, and 1-octen-3-ol gave non-linear curves of aroma transfer rate versus concentration gradient. This indicated that permeability values were concentration-dependent (Quezada-Gallo et al., 1999, 2000). Factors and phenomena that could explain these results are plasticization, antiplasticization, glass transition, and changes in polymer crystallinity. Miller et al. (1998) measured d-limonene permeability of whey protein films at different concentration gradients from 62 to 226 ppm (mol/mol) as a function of temperature. They reported that concentration gradient did not affect permeability values, i.e., the transfer rates increased linearly with the gradient, but only in the temperature range of 40 to 50 C. At higher

Factors Affecting Aroma Transfers Through Edible Barriers

589

temperatures, d-limonene permeability increased with concentration gradient at 40% RH. At 60% RH, it increased up to a concentration of 150 ppm, and then decreased at higher concentration gradients. The authors did not offer any explanation for this behavior. Therefore, it appears that the effect of concentration gradient on aroma transfers also depends on temperature and moisture. Similar to certain plastic films, such as EVOH, aroma flux through whey protein films increased exponentially with temperature according to the Arrhenius model (Miller et al., 1998). Activation energy values were 63 kJ/mol at 50% RH and 117 kJ/mol at 80% RH (Miller et al., 1998). In the case of methylcellulose film and 1-octen-3-ol, the activation energy was about 80 kJ/mol at dry conditions (RH

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