Characterization of food: emerging methods [1 ed.] 044481499X, 9780444814999

Citation preview

CHARACTERIZATION OF FOOD Emerging Methods

This Page Intentionally Left Blank

zyx

CHARACTERIZATION OF FOOD Emerging Methods

edited by ANILKUMAR

zyxwv

G. GAONKAR

Technology Center Kraft Foods, Inc. Glenview, IL 60025 USA

1995 ELSEVIER Amsterdam - Lausanne - New York-

Oxford - Shannon - Tokyo

ELSEVIER SCIENCE B.V. Sara Burgerhartstraat 25 EO. Box 211, 1000AEAmsterdam The Netherlands

Library

oF Congress C a t a l o g i n g - I n - P u b l i c a t i o n

Data

Characterization oF Food : e m e r g i n g methods / e d i t e d by A n i l k u m a r O. Gaonkar. p. cm. Includes bibliographical references and i n d e x . ISBN 0 - 4 4 4 - 8 1 4 9 9 - X 1. F o o d - - A n a l y s i s . I . G a o n k a r , A n l l k u m a r G . , 1954. TX541.C42 1995 95-35144 664'.07--dc20 CIP

ISBN 0 444 81499 X 9 1995 Elsevier Science B.V. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior written permission of the publisher, Elsevier Science B.V., Copyright & Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. Special regulations for readers in the U.S.A. - This publication has been registered with the Copyright Clearance Center Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the U.S.A. All other copyright questions, including photocopying outside of the U.S.A,. should be referred to the copyright owner, Elsevier Science B.V., unless otherwise specified. No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. This book is printed on acid-free paper. Printed in The Netherlands

To my Family, Relatives, Teachers, Friends and Colleagues,

and to all Children and Senior Citizens of the World

This Page Intentionally Left Blank

vii

Preface

Rapid and continued developments in electronics, optics, computing, instrumentation, spectroscopy, and other branches of science and technology led to considerable improvements in various methodologies. Due to this revolution in methodology, we are now able to solve problems which were thought to be extremely difficult to solve a few years ago. The new methods enabled us to better characterize foods and enriched our understanding of foods. The aim of this book is to assemble, for a handy reference, various emerging, state-of-the-art methodologies used for characterizing foods. Although the emphasis is placed on real foods, model food systems are also considered. The book contains invited chapters contributed by scientists actively involved in research, most of whom have made notable contributions to the advancement of knowledge in their field of expertise. It is not possible to discuss all the methods available for characterizing foods critically and systematically in a single volume. Methods pertaining to interfaces (food emulsions, foams, and dispersions), fluorescence, ultrasonics, nuclear magnetic resonance, electron spin resonance, Fourier-transform infrared and near infrared spectroscopy, small-angle neutron scattering, dielectrics, microscopy, rheology, sensors, antibodies, flavor and aroma analysis are included. This book is an indispensable reference source for scientists/engineers/technologists in industries, universities, and government laboratories who are involved in food research and/or development, and also for faculty, advanced undergraduate, graduate and postgraduate students from Food Science, Food Engineering, and Biochemistry departments. In addition, it will serve as a valuable reference to analytical chemists, and surface and colloid scientists. I wish to thank all the contributing authors for their dedication, hard work and cooperation and the reviewers for valuable suggestions. Last, but not least, I would like to thank my family, friends, relatives, colleagues, and the management of Kraft Foods Research for their encouragement.

April 1995

Anilkumar G. Gaonkar Kraft Foods, Inc. Glenview, IL 60025

zy

This Page Intentionally Left Blank

zy zyxwvut ix

CONTRIBUTORS

Niels M. Barfod

Grindsted Products A/S, Edwin Rahrs Vej 38, DK-8220 Brabrand, Denmark.

Wendy E. Brown BBSRC Institute of Food Research, Whiteknights Road, Reading RG6 2EF, United Kingdom.

Charles R. Buffler Microwave Research Center, 126 Water Street, Marlborough, NH 03455, USA.

S. Chakrabarti Kraft Foods, Inc., 801 Waukegan Road, Glenview, IL 60025, USA.

S.G. Greg Cheng Kraft Foods, Inc., 801 Waukegan Road, Glenview, IL 60025, USA.

D.C. Clark Institute of Food Research, Norwich Laboratory, Norwich Research Park, Colney, Norwich NR4 7UA, United Kingdom.

Mika Fukuoka Food Science and Technology Department, Tokyo University of Fisheries, Konan 4, Minato, Tokyo 108, Japan.

R.G. Fulcher Department of Food Science and Nutrition, University of Minnesota, St. Paul MN 55108, USA.

R. Gray Food Science Division, Department of Agriculture for Northern Ireland, Newforge Lane, Belfast BT9 5PX, Northern Ireland.

M.C.M. Gribnau Unilever Research Laboratorium, Olivier van Noortlaan 120, 3133 AT Vlaardingen, The Netherlands.

Sumio Kawano

zyxwvutsr zyx

National Food Research Institute, 2-1-2 Kannondai, Tsukuba 305, Japan.

K. Koczo Department of Chemical Engineering, Illinois Institute of Technology, 10 West 33rd Street, Chicago, IL 60616-3793, USA.

D.J. McClements Department of Food Science, University of Massachusetts, Amherst, MA 01003, USA.

Zohar M. Merchant Kraft Foods, Inc., 801 Waukegan Road, Glenview, IL 60025, USA.

M.M.W. Mooren Unilever Research Laboratorium, Olivier van Noortlaan 120, 3133 AT Vlaardingen, The Netherlands.

A.D. Nikolov Department of Chemical Engineering, Illinois Institute of Technology, 10 West 33rd Street, Chicago, IL 60616-3793, USA. D.G. Pechak Kraft Foods, Inc., 801 Waukegan Road, Glenview, IL 60025, USA. Peter Schieberle Bergische Universitat/GH, Food Chemistry/FB 9, Gauf~straB e 20, D-42097 Wuppertal, Germany.

M.G. Smart Kraft Foods, Inc., 801 Waukegan Road, Glenview, IL 60025, USA.

Philip H. Stothart 33, Betchworth Avenue, Earley, Reading, Berkshire RG6 2RH, United Kingdom.

K. Toko Department of Electronics, Faculty of Engineering, Kyushu University 36, 6-10-1 Hakozaki, Higashi-Ku, Fukuoka 812, Japan.

zyxwvuts zyxwvu xi

M.A. Voorbach Unilever Research Laboratorium, Olivier van Noortlaan 120, 3133 AT Vlaardin gen, The Netherlands.

D.T. Wasan Department of Chemical Engineering, Illinois Institute of Technology, 10 West 33rd Street, Chicago, IL 60616-3793, USA.

Hisahiko Watanabe Food Science and Technology Department, Tokyo University of Fisheries, Konan 4, Minato, Tokyo 108, Japan. Tokuko Watanabe Food Science and Technology Department, Tokyo University of Fisheries, Konan 4, Minato, Tokyo 108, Japan.

This Page Intentionally Left Blank

xiii

zy

Contents

Page No. Preface Contributors Interfacial Characterization of Food Systems D. T. Wasan, K. Koczo and A. D. Nikolov .

zyxwv o ~

Vll

Application of State-of-the-Art Fluorescence and Interferometric Techniques to Study Coalescence in Food Dispersions

ix

23

D. C. Clark

3. Methods for Characterization of Structure in Whippable Dairy-based Emulsions

59

Niels M. Barfod

4. Ultrasonic Characterization of Foods

93

D.J. McClements

5. Recent Advances in Characterization of foods using Nuclear Magnetic Resonance (NMR)

117

Hisahiko Watanabe, Mika Fukuoka and Tokuko Watanabe

6. Determination of Droplet Size Distributions in Emulsions by Pulsed Field Gradient NMR

151

M.M. W. Mooren, M. C.M. Gribnau and M.A. Voorbach ,

The Application of EPR Spectroscopy to the Detection of Irradiated Food

163

R. Gray

Progress in Application of NIR and FT-IR in Food Characterization Sumio Kawano

185

xiv

zyxwvut

Developments in the Application of Small-Angle Neutron Scattering to Food Systems Philip H. Stothart

10. Advances in Dielectric Measurement of Foods

201

213

Charles R. Buffier

11. Recent Developments in the Microstructural Characterization of Foods

233

M.G. Smart, R.G. Fulcher and D.G. Pechak

12. Some Recent Advances in Food Rheology

277

S. Chakrabarti

13. The Use of Mastication Analysis to Examine the Dynamics of Oral Breakdown of Food Contributing to Perceived Texture

309

Wendy E. Brown

14. Biosensors in Food Analysis

329

S.G. Greg Cheng and Zohar M. Merchant

15. Developments in Characterization of Foods Using Antibodies

347

Zohar M. Merchant and S.G. Greg Cheng

16. Taste Sensor

377

K. Toko

17. New Developments in Methods for Analysis of Volatile Flavor Compounds and their Precursors

403

Peter Schieberle

Index

433

Characterization of Food: Emerging Methods A.G. Gaonkar (Editor) 9 1995 Elsevier Science B.V. All rights reserved.

zyxwv

Chapter 1 Interfacial characterization of food systems D. T. W a s a n , K. K o c z o and A. D. Nikolov D e p a r t m e n t of C h e m i c a l Engineering, Illinois Institute of T e c h n o l o g y , Chicago, I L 60616, U S A

1. INTRODUCTION Many food products (salad dressings, whipped toppings, ice cream etc.) are dispersed colloid systems, such as emulsions, suspensions or foams. Texture, structure and stability of these dispersions have fundamental importance for the food manufacturer. Our chapter presents new methods, most of them developed in our laboratory, and mechanisms which can be very helpful for the food researcher or developer. The interfacial area of fine dispersions is very high and thus these interfaces strongly influence the behavior of the dispersions. The rheological characteristics of interfaces can be investigated by a new, versatile, but very simple technique, the controlled drop tensiometer, as will be described. The stability of foams and emulsions strongly depends on the structure and stability of the liquid films which form between approaching bubbles, emulsion drops or a bubble and an oil drop, respectively. Another application of the controlled drop tensiometer as well as optical interferometric techniques, will be discussed which allow the study of liquid films. A new mechanism of film stability involving thin film microlayering by small particles (sub-micron particles, surfactant micelles, macromolecules or protein aggregates) is also presented. The texture and s~cture of foods is very delicate, therefore experimental methods which cause no or very little structural damage has to be applied in their investigations. Such techniques, the surface force balance and back-light scattering methods and dielectrometry will be also discussed in the chapter. 2. INTERFACIAL RHEOLOGY Interfacial rheology deals with the flow behavior in the interfacial region between two immiscible fluid phases (gas-liquid as in foams, and liquid-liquid as in emulsions). The flow is considerably modified by surface active agents present in the system. Surface active agents (surfactants) are molecules with an affinity for the interface and accumulate there forming a packed structure. This results in a variation in physical and chemical properties in a thin interfacial region with a thickness of the order of a few molecular diameters. These

surfactants alter the hydrodynamic resistance to interfacial flow. Therefore, study of this variation, especially of the rheological properties, is important since many properties of dispersions, such as foam and emulsion stability, emulsification (making emulsions) and demulsification (breaking up emulsions) processes, are controlled by the interfacial flow behavior. To study the flow behavior in the interfacial region we use interfacial rheometers. In these instruments, a stress is imposed on an interface containing surfactants and the response is studied by measuring the velocity profile or the capillary pressure change. We can distinguish between two types of stresses on an interface: a shear stress and a dilatational stress. In a shear stress experiment, the interfacial area is kept constant and a shear is imposed on the interface. The resistance is characterized by a shear viscosity, similar to the Newtonian viscosity of fluids. In a dilatational stress experiment, an interface is expanded (dilated) without shear. This resistance is characterized by a dilatational viscosity. In an actual dynamic situation, the total stress is a sum of these stresses, and both these viscosities represent the total flow resistance afforded by the interface to an applied stress. There are a number of instruments to study interfacial rheology and most of them are described in Ref. [1]. The most recent instrumentation is the controlled drop tensiometer. The controlled drop tensiometer is a simple and very flexible method for measuring interfacial tension (IFT) in equilibrium as well as in various dynamic conditions. In this technique (Fig. 1), the capillary pressure, Pc, of a drop, which is formed at the tip of a capillary and immersed into another immiscible phase (liquid or gas), is measured by a sensitive pressure transducer. The capillary pressure is related to the IFT and drop radius, R, through the Young-Laplace equation [2,3]:

zyxwvutsrq zyxwvuts

2o Pc = Pa-Pb = R

(1)

where o is the instantaneous IFT, and Pa and Pb are the pressures inside and outside the drop, respectively. Deformation of the drop by gravity can be avoided by using capillaries with sufficiently small radius (Re). The size of the drop is varied by using a computer controlled microsyringe attached to the capillary and the output of the transducer is also fed into a computer. The volume and radius of the drop at any instant are determined by the position and speed of the microsyringe plunger. For the measurement of equilibrium IFT, a drop is formed at the capillary tip and maintained at that size. After sufficient time, chemical equilibrium is achieved and the equilibrium thermodynamic IFT can be calculated from the measured, steady state capillary pressure and drop radius by Eq. 1. The adsorption and desorption kinetics of surfactants, such as food emulsifiers, can be measured by the stress relaxation method [4]. In this, a "clean" interface, devoid of surfactants, is first formed by rapidly expanding a new drop to the desired size and, then, this size is maintained and the capillary pressure is monitored. Figure 2 shows experimental relaxation data for a dodecane/aq. Brij 58 surfactant solution interface, at a concentration below the CMC. An initial rapid relaxation process is followed by a slower relaxation prior to achieving the equilibrium IFT. Initially, the IFT is h i g h , - close to the IFT between the pure solvents. Then, the tension decreases because surfactants diffuse to the interface and adsorb, eventually reaching the equilibrium value. The data provide key information about the diffusion and adsorption kinetics of the surfactants, such as emulsifiers or proteins.

zyxwvu

Figure 1. Schematics of controlled drop tensiometer

Desorption kinetics can also be studied by contracting a drop from a known state to a new state. The sudden reduction in interfacial area causes desorption of surfactants, which is deduced from the IFT change over time. Dynamic interfacial tension of dilating or contracting liquid-liquid interfaces can be measured by monitoring the capillary pressure for an expanding (or contracting) liquid drop. The IFT, as a function of time, is computed from the capillary pressure change and radius change with time. To study the effect of dilation of the interface under controlled conditions, first, a drop is formed and an initial equilibrium state is established by maintaining this drop size for sufficient time. Then, in expansion experiments, the drop volume is increased with constant flow rate and the capillary pressure is monitored over time. In pure systems, or in systems where surfactant adsorption is fast (high surfactant concentration) the IFF does not change during drop expansion and the capillary pressure decreases, as shown by Eq. 1. In surfactant systems, if the surfactant adsorption is not fast, the dynamic IFT can be significantly higher than the equilibrium values and the capillary pressure can increase during interface expansion. Figure 3 shows the dynamic IFT of soybean oil/water interfaces under expansion with constant flow rate as a function of the relative change of the interfacial area, with various surfactants in the oil and aqueous phases, respectively. The IFT is lowest if both phases contain surface active additives, and it hardly changes due to the presence of the fast adsorbing, low molecular emulsifier SPAN 80 in the oil phase. The increase of the dynamic IFT with the interface expansion is most pronounced with 0.01% BSA in the aqueous phase due to the slow adsorption of the protein.

zyx

Figure 2. Stress relaxation of the dodecane/aqueous Brij 58 interface at c=10 -6 mol/dm 3, Rc---O.141 mm, 25 ~

A similar technique can be used to study the rheological properties of liquid films. Figure 4 shows the formation of a W/O/W emulsion film with two, identical aqueous phases (such as in water-in-oil emulsions) at the tip of the capillary. A pre-requisite of the experiment is that the surface of the capillary must be well wetted by the film phase, i.e., it should be hydrophobic in this case. First, an aqueous drop is formed inside the oil (film liquid) and the aqueous phase is in the bottom of the cuvette. Then, the level of the aqueous phase is slowly increased. As the oil/water interface passes the drop, a cap shaped oil film, bordered by a circular meniscus, covers the drop. This film can be studied in equilibrium and in dynamic conditions, similar to the single interfaces (See above). The technique can be used to study films from oil or aqueous phase which can be sandwiched between identical or different liquid or gas phases. For relatively thick films (higher than about 30 nm), the pressure drop at the film is the sum of the capillary pressures at the two film interfaces. In this case, the Young-Laplace equation for the film can be written as

_2f I where the film tension (f) is given by:

zyxwvu t2)

Figure 3. Dynamic interfacial tension of soybean oil/water systems. expanding in water, pH=7, flow rate: 3.10 .4 mm3/s, Rc=0.141 mm.

Soybean oil drop

Figure 4. Method to form and study an oil film between aqueous phases at the tip of capillary

f-

4 0 to o Ol+O o

zyxwvutsrqp zyxwvutsrqp (3)

where Rf is the film radius, t~i and % are the interfacial tensions at inner and outer film interfaces, respectively. For emulsion (or foam) films: ~i=% and f=2t~. Figure 5 shows dynamic film tension of soybean oil films stabilized by 0.5 wt % SPAN 80 emulsifier between aqueous phases under expansion by various flow rates. The increase in film tension from equilibrium is higher at higher rates of interface expansion because the flux of surfactant that can adsorb during expansion is lower at higher rates.

Figure 5. Dynamic film tension of soybean oil film containing 0.5 wt% SPAN 80 between water phases, expanding with various flow rates as a function of the relative film area. Rc=0.32 mm, h=0.03 mm, pH=7, at 25 ~

3. INTERFEROMETRY 3.1. Common interferometry - mechanisms of liquid film stability

zyxw

Liquid films which form between approaching drops or bubbles are important structural elements of dispersed systems. The stability of these films controls the dispersion stability because the drops or bubbles cannot coalesce until the intervening film ruptures. The drainage and stability of thin liquid films attracted the attention of scientists already centuries ago [5,6]. The thinning process of plan-parallel liquid films have been generally observed using reflected light interferometry [7-9]. The experimental setup to form and study such film contains a small, vertically oriented cylindrical tube of hydrophilic inner walls with a horizontal capillary side arm, as shown in Figure 9 of Chapter 2. The film liquid is filled into the vertical tube and, then, a horizontal liquid film encircled by a biconcave meniscus is formed by slowly sucking out the liquid through the side arm. The driving force for film thinning is the capillary pressure which, for small film contact angles and complete wetting of the capillary wall, is given for this film configuration by:

PC

zyxwvutsrq (4)

Rc2

2

where R c is the inner radius of the capillary, Rf is the film radius and c is the interfacial tension. Thus, an increase in Pc leads to an increase in Rf and the film area. The film is observed by a microscope using reflected light. The film holder and the objective are immersed in air in the case of foam (i.e., air/liquid/air) film and in the oil phase, in the case of an O/W/O emulsion film, respectively. The film thickness can be determined by measuring the intensity of the light reflected from the film surfaces [9]. Further details of the technique will be discussed in Chapter 2. We have used film interferometry to reveal a new mechanism for the stabilization of foams and emulsions due to layering inside the thinning films, as will be discussed below. When two emulsion drops or foam bubbles approach each other, they hydrodynamically interact which generally results in the formation of a dimple [10,11]. After the dimple moves out, a thick lamella with parallel interfaces forms. If the continuous phase (i.e., the film phase) contains only surface active components at relatively low concentrations (not more than a few times their critical micellar concentration), the thick lamella thins on continually (see Fig. 6, left side). During continuous thinning, the film generally reaches a critical thickness where it either ruptures or black spots appear in it and then, by the expansion of these black spots, it transforms into a very thin film, which is either a common black (10-30 nm) or a Newton black film (5-10 nm). The thickness of the common black film depends on the capillary pressure and salt concentration [8]. This film drainage mechanism has been studied by several researchers [8,10-12] and it has been found that the classical DLVO theory of dispersion stability [13,14] can be qualitatively applied to it by taking into account the electrostatic, van der Waals and steric interactions between the film interfaces [8]. The hydrodynamic stability of such films is controlled by the capillary pressure, film

Figure 6. Mechanisms of liquid film stability

area, the interfacial rheological properties (such as surface shear viscosity etc.) and the surface tension gradients (Gibbs-Marangoni effect) of the surfactant adsorption layer at the film interfaces [8,15-21]. The properties of these films, in relation to food systems, are discussed in Chapter 2. Film studies in the past decade have revealed the existence of another film stability mechanism: If the continuous phase contains not only a small amount of surface active substances but also a "sufficient amount" of "small particles", these particles can form layers inside the draining film (see Fig. 6, right)[9,22-32]. As a result, such films thin step-wise, by several step-transitions (also called stratification) when at a step transition a layer of small particles leaves the film. Sodium caseinate is commonly used in foods as emulsion and foam stabilizer. The photomicrographs of Figure 7 show the phenomenon of film step-transitions for a foam film which was formed from 2 wt% sodium caseinate solution at 40 ~ Shortly after lamella formation, a dimple forms (See Fig. 7, picture a). After the dimple leaves the lamella the film drains continually. After about 100 nm film thickness, however, the film thinning becomes step-wise. First, most of the film turns uniformly bright. Then, uniform, light grey (i.e., thinner) areas with sharp borders appear and start to cover the bright areas (See Fig. 7, pict. b), i.e., the first step-transition occurs. Shortly later dark grey spots form near the border of the film, inside the light grey region (See Fig. 7, pict. c, upper left section). The dark grey spots expand, unify and occupy the film area (second transition). During this process, a black, thinner spot forms inside the dark grey film (See Fig. 7, pict. d), followed by the formation and expansion of several other black spots (See Fig. 7, pict. e). Finally, the black spots unify (See Fig. 7, pict. 3') and the film turns uniformly black (third steptransition). No more step-transitions could be observed and the color, i.e., the thickness, of the foam film did not change any more. These observations show that microlayering takes place in the foam film containing 2 wt% caseinate. During a step-transition a layer leaves the film, until no layer is left (black film). Thus, the bright film contained three layers, the light grey two, the dark grey one and the black film contained zero layers. The average thickness differences between films containing zero, one, or two layers, respectively (i.e., the heights of the step-transitions), were measured by interferometry and it was found that they are approximately equal and about 20 nm. It was found by several researchers [33-36] that casein molecules form aggregates in aqueous solutions the so-called casein sub-micelles with approximately the same size as these step-transitions. (The casein miceUes are much larger particles and they form from the sub-micelles by calcium [33,34]. In the sodium caseinate there is practically no calcium and thus, the caseinate solution contains a significant amount of sub-micelles.) It can be concluded that the foam film containing caseinate solution thins by step-transitions because the caseinate sub-micelles form layers in the film. The step-transition phenomenon resembles the common black film/Newton black film transition (Fig. 6, left), however, there are basic differences between the two processes. The step transitions, due to microlayering, can occur at very high thicknesses (depending on the size and concentration of the small particles, as high as several hundred nanometers [27]) and the number of the step-transitions can be much higher than one [27]. The investigations in our laboratory showed that the film microlayering mechanism is a universal phenomenon which fundamentally differs from the classical film thinning mechanism by common black film/Newton black film transition as summarized in Fig. 6. It has been found that the "small particles" can be virtually any kind of isotropic structures with about 10-100 nm size,

zy

10

Figure 7. Photomicrographs on the various drainage stages of a foam film containing 2 wt% sodium caseinate, at 40 ~ Film diameter: 0.35 mm.

zyxwvutsr zyx 11

including micelles of ionic or non-ionic surfactants [9,22-24] - fine solid particles, such as silica or latex particles [9,27] - macromolecules, such as globular protein molecules or random coil shaped polysaccharide molecules protein aggregates, such as caseinate sub-micelles, as was shown above [29] for the occurrence of film microlayering. Note that all of these substances are commonly used in foods. The reason for film microlayering is that the restrictive geometrical conditions, i.e., the presence of the "walls", the film surfaces, force the (sub)-micelles or Brownian particles inside the film to be layered and organized [30]. A pre-requisite of the film microlayering phenomenon is that the effective volume fraction of the small particles should be sufficiently high, at least about 5-1.0 vol% [27]. It is important to emphasize that the effective volume fraction of such sub-micron sized particles, i.e., the volume that the particles really occupy in the solution, is much higher than their geometrical volume fraction. Thus, the above volume fraction range can be reached with about 0.1-1 wt% of small molecule surfactants or with less than 0.1 wt% macromolecules [29]. The number of layers increases with the effective volume fraction of the small particles [127]. It is of great importance to the film microlayering phenomenon that these concentration ranges are typical in practical applications such as in food emulsions and foams. (It can be mentioned that at very high concentrations, from about 10 wt%, ordering of the small particles, such as surfactant micelles, takes place not only inside the films but also in the bulk phase [37]. These concentrations are, however, impractical and therefore, this phenomenon will not be discussed here.) Lower polydispersity enhances the film microlayering process, thus the film stability [27]. Liquid films containing layers cannot be described by the DLVO theory because their disjoining pressure is controlled by the repulsive particle/particle and particle/interface interactions and not by the interface/interface interactions because the interfaces are too far apart when layers are inside the film [25]. Due to these interactions, the disjoining pressure isotherm of a film containing layers is oscillatory (Fig 6.), which explains that the film thinning has several steps [30]. Because of this, the occurrence of layering and the height of the step-transitions do not depend on the nature of the interface: the same steps (by number and height) can be observed in a foam and in an O/W/O emulsion film, respectively, if the two types of films contain the same amount of small particles, such as sodium caseinate [29,32]. A very important feature of the film microlayering phenomenon is that the occurrence of a step-transition also depends on the area (diameter) of the film. If the film area is smaller than a critical value, the step-transition is inhibited and a layer or layers of fine particles stay inside the film for an unlimited time [27,29,31,32]. It is interesting to note that the capillary pressure of drops or bubbles, which is the driving force of film drainage, increases with decreasing drop or bubble size, i.e., film area (See Eq. 1). However, the presence of strong structural forces in the film overrides the effect of capillary pressure in this case. The phenomenon can be explained by the vacancy mechanism of the step-transitions [28]. The existence of critical film size has great practical importance: when layer or layers of small particles stay trapped in the liquid films between small drops (or bubbles) the stability of these films is extremely high. -

-

12

zyxwvutsrq

3.2. Differential interferometry- characterization of the pseudoemulsion film When an oil drop in an aqueous phase rises to the surface of the solution or an oil drop approaches a bubble inside a foam an asymmetrical, oil/water/oil film, the so-called pseudoemulsion film forms between the oil and air phases (Figure 8.) The importance of

Figure 8. Formation of a pseudoemulsion film drop between an oil drop and air this film is that the effect of oil drops in foam stability is controlled by the stability of the pseudoemulsion film [31,32]. If this film is unstable, that is, it ruptures, the oil drop enters the air/water surface and spreads on it, generally resulting in antifoam action [38]. If, however, the pseudoemulsion film is stable, the oil drops cannot spread and instead of breaking it, the oil drops stabilize the foam [39]. Due to its asymmetrical nature, the pseudoemulsion film is always curved. Similarly, foams or emulsions which form between bubbles (drops) of differing size are not plane parallel but curved (cap) shaped [4]. When two fluid interfaces have a high radius of curvature, such as in the pseudoemulsion film, the distance between the interference patterns is too small to be measured by common reflected light interferometry. In this case, differential interferometry can be used for imaging the interface profile 140-45]. (Another technique for studying curved films is the controlled drop tensiometer, as was shown in section 2.) The basic principle of differential interferometry consists of splitting the original image into two images. An Aus Jena Epival Interphako microscope was used in our laboratory for film studies with common and differential interferometry. This microscope is capable of viewing objects in transmitted light as well as in reflected light and also equipped with a Max Zhender interferometer. The interferometer splits the original beam of the image into two beams of different optical paths which, when recombined, give a sheafing type differential interference pattern - this can be used to measure curvature of surfaces [43-45] (See Figure 9). The two images are shifted at a distance d at which the beams reflected by the interfaces Z(x,y) and Z'(x,y), respectively, interfere. As a result, a characteristic interference pattern forms, which contains streaks, tings and mustaches [41,42] (Fig. 9). The optical path length between the two beams is

zyxw

zyxwvu

A = 2(Z-Z')n/

(5)

where n r is the refractive index in the phase between the surface and the objective. When A =iL/2, (where i=0,1,2.., is the order of interference and ~, is the wavelength of

13 the monochromatic light used) dark fringes form for odd and bright ones for even i. By measuring the distance between the parallel bright and dark fringes the curvature of the film, Rf, can be calculated [44]. Lobo and Wasan [41] determined the exact profile of pseudoemulsion films, their meniscus and the film contact angles by using common interferometry (Newton tings) and differential interferometry in conjunction with the Laplace equation for the film menisci [42]. The differential interference image of a pseudoemulsion film between an octane drop and air inside a 4 wt% micellar solution of the non-ionic surfactant C1215AE30 (ethoxylated alcohol with C~z-C~5alkyl chain and 30 ethoxy groups) is shown in Figure 10. The drainage characteristics of the pseudoemulsion film were also observed, in reflected light by common interferometry, by submerging the oil (octane) drop and allowing it to rise in the solution. As the rising oil drop reached the gas-aqueous interface, a thick, nonuniform film (with a dimple) was initially formed as seen in Fig. 1 la. Fig. 1 lb and c show the film which was undergoing similar thickness transitions as the foam film in Figure 7. The step-wise thinning phenomenon observed here for the pseudoemulsion film is the result of microlayering of non-ionic surfactant micelles, as described in the previous section for foam or emulsion films. In Figure 11 it is also seen that the thin pseudoemulsion film appears bright, as opposed to the thin foam and emulsion films, which are black (see Fig. 7). The reason for this is the optical path difference between the reflected rays from the two film surfaces. Light rays which are incident on the film reflect from the two surfaces

Figure 9. Principle of differential interferometry

14

Figure 10. Photomicrograph of the differential interference pattern of a pseudoemulsion film (octane drop in 4 wt% C1215AE30 solution).

Figure 11. Thinning pseudoemulsion film. a) Thick film with dimple, b) Film undergoing stratification - two thickness transitions - and, c) Enhanced image of film undergoing stratification. The film has three discrete thicknesses resulting from the first two transitions.

zyxwvu zyxwvuts 15

of the film and these reflected light rays interfere. When light rays of wavelength k, are incident on a thin (zero thickness)foam or emulsion film, they encounter, alternately, optically dense and optically rare medium (the order depends on the type of emulsion). As a result, the two reflected rays from each of the film surfaces differ by a path length of ~./2. This is the condition for the destructive interference of light, which is why for film with thicknesses less than 100 nm the foam and emulsion films appear black in reflected light. On the other hand, light rays incident from the air side of an aqueous pseudoemulsion film encounter an optically denser medium at both the film surfaces (air to water and water to oil). Thus, the reflected rays from film surfaces are shifted by ~./2 and the path difference is ~,. This is the condition for constructive interference, which is why the pseudoemulsion film appears bright [42].

3.3. Capillary force balance The texture and stability of food foams or emulsions strongly depend on the interaction between the fat or other dispersed particles in the system. Aggregation of paricles, drops by polysaccharide macromolecules has been observed in food systems [46-48]. Aggregation phenomena and interparticle interactions can be directly observed by using transmitted light interference microscopy in conjunction with the capillary force balance technique recently developed in our laboratory. First, the emulsion or dispersion is filled in the film holder, then, the formed conical interfaces are pushed together by sucking out the liquid through the capillary side arm (Fig. 12). A thick film (lamella) several micrometers thick forms as a result of this increase in the capillary pressure. The film structure, its response to external stress, which can be manipulated by the capillary pressure and the stability of the lamella can be directly observed using transmitted light microscopy. Draining of the lamella can proceed until a thin liquid film is formed. A great advantage of the method is that the observed emulsion layer is "free", without having any connection to other surfaces (such as a glass slide, etc.). Figure 13 demonstrates the particle aggregation phenomenon induced by gums as observed using the surface force balance method. The figure shows photomicrographs of thick foam lamellae from O/W food emulsions containing 20 vol% fat, in the presence and in the absence of gums, respectively. The elementary particle size of these emulsions, as determined by light scattering after strong dilution, was mainly in the sub-micron range. It is seen, however, that in the undiluted emulsion containing gums (Fig. 13 a) the fat particles appear as 5,101am aggregates. It could be also observed that the particle aggregates move together as the lamella thickness is changed. In the emulsion without gums (Fig. 13 b) only a few larger particles can be seen, the rest of the particles are very small and cannot be seen under the low magnification used.

4. BACK-LIGHT SCATTERING - KOSSEL DIFFRACTION Another optical technique, called the back-light scattering (Kossel-diffraction) method can also be used to investigate structure in food emulsions and foams. In this method, the emulsion (or foam) in a transparent vessel is illuminated by a collimated laser beam (See

16

Figure 12. Principle of capillary force balance

Figure 13. Photomicrographs of lamellae formed from O/W food emulsions containing 20 wt% emulsified fat, illuminated by transmitted light. a) Particles aggregate in the presence of gums. b) Negligible aggregation without gums.

17

zyxwvut

Fig. 14). A portion of the light rays are scattered from the emulsion particles through the wall of the vessel and form a concentric interference pattern. The back-scattering phenomenon is analogous to the operation of diffraction gratings [49]. The measurement can be used to characterize the structure of the emulsion, because the shape of the light intensity profile depends on the particle size (2a), the average distance between the particles (d), the wavelength of the laser light used (~.), the angle of observation (O) and the regularity of the spacial arrangement of the particles (S) (See Fig. 14). The interference image can be recorded and the intensity profile along a vertical line, going through the center of the image, can be measured by an image analyzer. The optical conditions (laser beam diameter, magnification etc.) and the wall of the sample holder influence the intensity profile, thus, these parameters must be kept constant in the measurements. Figure 15 shows a typical light intensity profile of a food emulsion as a function of distance in arbitrary units. The curves are symmetrical with a large, primary maximum in the center, which is surrounded by minima and secondary maxima at both sides. When the parameters of the emulsion (d, a and S) change, it is generally reflected on i.) the width of the shoulder of the primary maximum; ii.) the depth and position of the minima and iii.) the height and position of the secondary maxima of the intensity profile. The shape of the intensity profile reflects the shape of the radial distribution function of the particles. The radial distribution of a highly ordered structure, such as a crystal, is periodic, i.e., the concentration of particles changes periodically as a function of the radial distance from a given point. If the order, the regularity of the structure, is lower, such as in a liquid, the radial distribution function is less periodic: the difference between the maxima and minima

Figure 14. Principle of back-light scattering measurement.

18

Figure 15. Intensity profile of light back-scattered from a food emulsion containing 20 wt% emulsified fat, at 5 ~ using green light (543 nm).

zyxwvutsr

are much smaller than in the ordered structure. If there is no order, such as in a gas, the periodicity of the radial distribution function vanishes and the intensity decreases monotonously as a function of distance. The effect of particle aggregation on the intensity profile is illustrated in Fig. 16 showing the profiles of a food emulsion without gums, in which no aggregation takes place, and the same emulsion in the presence of gums, where the particles aggregate, respectively. Aggregation changes several emulsion properties at the same time: it increases the particle size, polydispersity and the distance between the particles. As a result~ the shoulder of the primary maximum becomes wider (See Fig. 16). Moreover, aggregation generally decreases the order of the particles too, which results in a decrease of the secondary maximum. It is seen that the emulsion with the aggregates has, indeed, a very small secondary maximum.

5. D I E L E C T R O M E T R Y The dielectric properties of water have been extensively used to determine moisture content in food systems. However, only very recently have we used the complex dielectric properties of emulsions in the microwave frequency region to characterize both emulsion type and water content [50-52]. We have developed both a cavity resonance dielectrometer capable of operating at 8-11 GHz and an interference dielectrometer operating at 23.45 GHz.

19

Figure 16. Effect of particle aggregation on the back-scattered light intensity profile of food emulsion, containing 20 wt% emulsified fat, at 5 ~ (green light). We have employed these dielectric techniques to study the hydration characteristics of hydrocolloids widely used in food systems 1531. The rotational relaxation of water molecules is influenced by its immediate environment. The microwave dielectric characteristics of associated or bound water molecules are markedly different from those of free water molecules. During the hydration of hydrocolloids, water molecules go from a bound state to an unbound state and the change is detected dielectrically. In food systems, hydrocotloids are added to impart increased product viscosity as well as to stabilize the emulsions. The stabilizing action arises from the formation of complex structures between the water molecules and gums. The water molecules are held in a bound state through avariety of bonding mechanisms. The existence of continuous phase structures prevents the emulsion drops from approaching each other and therefore prevents coalescence. In addition to enhancing emulsion stability, the bound water is not available for microbial growth and this is clearly an important feature in food emulsion systems. The extent of hydration of hydrocolloid is important in determining the efficacy of inhibiting the coalescence of emulsion drops. The microwave dielectric measurements exclusively measure the rotational relaxation of the water molecule.

20 Figure 17 shows the change in permittivity as a function of time of hydration for 0.5 wt% ~c-carrageenan dissolved in double deionized water. The measurements were made by monitoring the changes in dielectric response of the hydrocolloid sample solution held in the cavity of resonance dielectrometer operating at 9.505 GHz. The measurements indicate ,hat the hydration process is complete after a period of 6 hours. During the early stages of hydration, a high dielectric permittivity value was measured corresponding to the large amount of free water present in the system. As the water molecules attach themselves to the numerous hydratable groups present in the hydrocolloid molecule, the permittivity values decline. When all the water molecules are held in a bound state, the hydration process is complete and no change in permittivity was observed.

Figure 17. Experimentally measured variation in permittivity with extent of hydration of 0.5 wt% K:-carrageenan hydrocolloid at 23.45 GHz.

6.

CONCLUDING REMARKS

New experimental techniques and several of their applications were presented which help in the understanding of structure, texture and stability of food systems. For future research, the mechanism of film stability by the microlayering of colloid particles seems to be the most promising - especially in food emulsions and foams. Work is in progress in our laboratory to calculate the oscillatory disjoining pressure inside liquid films containing microlayers [30]. The structure and stability of foamed emulsions, such as whipped cream, ice cream or whipped toppings, strongly depend on the interparticle interactions and on the orientation of drops/particles at the foam films. Further development of the surface force balance and

zyxwvuts 21

back-light scattering techniques will aid in the understanding of the stability mechanisms in food dispersions.

REFERENCES

.

.

4.

5. 6. .

8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

25. 26.

D.A. Edwards, H. Brenner and D.T. Wasan, lnterfacial Transport Processes and Rheology, Butterworth-Heineman Publishers, Stoneham, MA, 1991. A. Passerone, L. Liggieri, N. Rando, F. Ravera and E. Ricci, J. Colloid Interface Sci., 146 (1991) 152. R. Nagarajan and D.T. Wasan, J. Colloid Interface Sci., 159 (1993) 164. R. Nagarajan, K. Koczo, E. Erdos and D.T. Wasan, AIChE J., (1994) in press. I. Newton, Optiks; Book II, Part I, Observation 17, Smith and Watford, London, 1704. K.J. Mysels, S. Shinoda and S. Frankel, Soap Films - A Study of Their Thinning and a Bibliography, Pergamon, New York, 1959. B.V. Derjaguin and A.S. Titievskaya, Kolloid Zh., 15 (1953) 416.. A. Scheludko, Adv. Colloid Interface Sci., 1 (1967) 391. A.D. Nikolov and D.T. Wasan, J. Colloid Interface Sci., 133 (1989) 1. I.B. lvanov (Ed.), Thin Liquid Films, Surfactant Sci. Ser., Marcel Dekker, New York, 1988. Clark, D.C., Coke, M., Wilde, P.J. and Wilson, D.R., in Food Polymers, Gels, and Colloids, E. Dickinson (ed.), Royal Soc. Chem. Spec. Publ. No. 82, p. 272, 1991. D. Exerowa, Khr. Khristov and I. Penev, in Foams, Proc. Symp. at Brunel Univ., R.J. Akers (ed.), pp. 109-126, 1975. B. Derjaguin and L. Landau, Acta Physicochim., 14, (1941) 633. E. Verwey and J.Th.G. Overbeek, Theory of The Stability of Lyophobic Colloids, Elsevier, Amsterdam, 1948. B.V. Derjaguin and M.M. Kusakov, Izv. Acad. Sci., U.S.S.R., 1 (1936) 256. D.T. Wasan and A.K. Malhotra, AIChE Symp. Ser., 82 (1986) 5. Z. Zapryanov, A.K. Malhotra, N. Aderangi, and D.T. Wasan, lnt'l. J. Multiphase Flow, 9 (1983) 105. I.B. Ivanov and D.J. Dimitrov, Coll. Polym. Science, 252 (1974) 982. B.P. Radoev, D.S. Dimitrov and I.B. Ivanov, Coll. Polym. Sci., 252 (1974) 50. A.D. Barber and S. Hartland, Can, J. Chem. Eng., 54 (1976) 279. A.K. Malhotra and D.T. Wasan, in Thin Liquid Films, l.B.lvanov (ed.), Marcel Dekker, p. 95, 1990. E. Manev, S.V. Sazdanova and D.T. Wasan, Dispersion Sci. Tech., 3 (1982) 435. A.D. Nikolov, D.T. Wasan, N.D. Denkov, P.A. Kralchevsky and I.B. Ivanov, Progress in Colloid and Polymer Sci., 82 (1990) 87. A.D. Nikolov, D.T. Wasan, P.A. Kralchevsky and I.B. Ivanov, in Ordering and Organization in Ionic Solutions, N. Ike and I. Sogami (eds.), World Scientific Publications, Singapore, 1988. A.D. Nikolov, P.A. Kralchevsky, I.B. Ivanov and D.T. Wasan, J. Colloid Interface Sci., 133 (1989) 13. E.S. Basheva, A.D. Nikolov, P.A. Kralchevsky, I.B. lvanov and D.T. Wasan, in Surfactants in Solution, vol. 11, K. Mittal (ed.), p. 467, 1990.

22 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53.

A.D. Nikolov and D.T. Wasan, Langmuir, 8 (1992) 2985. P.A. Kralchevsky, A.D. Nikolov, D.T. Wasan, and I.B. Ivanov, Langmuir, 6 (1990) 1180. K. Koczo, A.D. Nikolov, D.T. Wasan, R.P. Borwankar and A. Gonsalves, paper submitted to J. Colloid Interface Sci. (1994). X. L. Chu, A.D. Nikolov and D.T. Wasan, Langmuir, 10 (1994) 4403. D.T. Wasan, A.D. Nikolov, L.A. Lobo, K. Koczo and D.A. Edwards, Progr. Surf. Sci., 39 (1992) 119. D.T. Wasan, K. Koczo and A.D. Nikolov, in Foams: Fundamentals and Applications, L.L. Schramm (ed.), ACS, Chapter 2, 1994. D.G. Schmidt and T.A.J. Payens, Surface and Colloid Science, E. Matijevic (ed.), pp. 165-229, Wiley-Interscience, New York, 1976. P. Walstra and R. Jenness, Diary Chemistry and Physics. Wiley, New York, 1976. D.G. Schmidt and J.A.J. Payens, J. Colloid Interface Sci., 39 (1972) 655. T.F. Kumosinski, H. Pessen, H.M. Farrell and H. Bumberger, Arch. Biochem. Biophys., 266 (1988) 548. S. Friberg, S.E. Linden and H. Saito, Nature, 251 (1974) 495. K. Koczo, J.K. Koczone and D.T.Wasan, J. Colloid Interface Sci., 166 (1994) 225. K. Koczo, L. Lobo and D.T.Wasan, J. Colloid and Interface Sci., 1992, 150, 492. Nikolov, A.D., Dimitrov, A.S. and Kralchevsky, P.A., Optica Acta, 33 (1986), 33. L.A. Lobo, A.D. Nikolov, A.S. Dimitrov, P.A. Kralchevsky and D.T. Wasan, Langmuir, 6 (1990) 995. L.A. Lobo and D.T. Wasan, Langmuir, 9 (1993) 1668. H. Beyer, Jenaer Rdsch., 16 (1971) 82. A.D. Nikolov, A.S. Dimitrov and P.A. Kralchevsky, Optica Acta, 33 (1986) 33. A.S. Dimitrov, P.A. Kralchevsky, A.D. Nikolov and D.T.Wasan, Colloids Surfaces, 47 (1990) 299. Y. Cao, E. Dickinson and D.J. Wedlock, Food Hydrocolloids, 4 (1990) 185. Y. Cao, E. Dickinson and D.J. Wedlock, Food Hydrocolloids, 5 (1991) 443. E. Dickinson and V.B. Galazka, in Food Polymers, Gels and Colloids, E. Dickinson (ed.), Royal Soc. of Chem. Spec. Publ. No. 82, pp. 494-497, 1991. R.D. Guenther, Modem Optics, John Wiley & Sons, pp. 361-431, 1990. J.P. Perl, C. Thomas and D.T. Wasan, J. Colloid Interface Sci., 137 (1990) 425. C. Thomas, J.P. Perl and D.T. Wasan, J. Colloid Interface Sci., 139 (1990) 1. J. Rudin and D.T. Wasan, J. Colloid Interface Sci., 162 (1994) 252. C. Thomas, PhD Thesis, Illinois Institute of Technology, Chicago, 1990.

zyx

Characterization of Food: Emerging Methods A.G. Gaonkar (Editor) 9 1995 Elsevier Science B.V. All rights reserved.

23

Chapter 2 A p p l i c a t i o n of state-of-the-art f l u o r e s c e n c e and i n t e r f e r o m e t r i c techniques to study c o a l e s c e n c e in food d i s p e r s i o n s D.C. Clark Institute of Food Research, Norwich Laboratory, Norwich Research Park, Colney, Norwich NR4 7UA, United Kingdom.

1. INTRODUCTION Coalescence is an important mechanism of destabilization of food foams and emulsions [1]. The coalescence process involves fusion of two adjoining gas bubbles in a foam or oil droplets in an oil-in-water emulsion by rupture of the thin aqueous film or foam lamella which keeps the dispersed phase separated. Foams generally contain a high phase volume of gas and the thin planar films form very rapidly as entrained liquid drains from the foam. In contrast, formation of thin films in emulsions is a much slower process. This is because the phase volume of the dispersed phase in emulsions is usually not as high as in a foam, the average droplet size is smaller than the average bubble size and there is generally a comparatively small difference in the density of the continuous and dispersed phases. A combination of these factors, coupled with the inclusion of stabilizing agents such as polysaccharide thickeners, means that the rate of creaming in emulsions is relatively slow. However, it is worth considering the thin films that may form between oil droplets in an emulsion when they pack together closely in the cream layer of an emulsion or during collision processes. Thin films are stabilized by two distinct mechanisms. The mechanism that prevails is dependent upon the molecular composition of the interface. Low molecular weight surfactants such as food emulsifiers or polar lipids congregate at the interface and form a fluid adsorbed layer at temperatures above their transition temperature (Figure l(a)). When a surfactantstabilized thin film is stretched, local thinning or dimple formation occurs in the thin film. This is accompanied by the generation of a surface tension gradient across the locally thin region. Surface tension is highest at the thinnest point of the stretched film, due to decreases in the surface concentration of emulsifier in the region of the stretch. Equilibrium surface tension is restored by lateral diffusion of surfactant in the adsorbed layer towards the region of highest surface tension. This surfactant drags interlamellar liquid into the thin region and contributes to the restoration of equilibrium film thickness. This process is often referred to as the Marangoni effect [2]. In contrast, the adsorbed layer in protein-stabilized thin films is much stiffer and often has viscoelastic properties [3]. These derive from the protein-protein interactions that form in the adsorbed layer (Figure 1(b)). These interactions result in the formation of a gel-like adsorbed layer in which lateral diffusion of molecules in the adsorbed layer is inhibited. Multilayer formation can also occur. This serves to further mechanically strengthen the adsorbed layer.

zy

24 When pure protein films are stretched, the change in interfacial area is dissipated across the film, due to the cohesive nature of the adsorbed protein layer and the deformability of the adsorbed protein molecules.

Figure 1. Schematic diagram showing the possible mechanisms of thin film stabilization. (a) The Marangoni mechanism in surfactant films; (b) The viscoelastic mechanism in proteinstabilized films; (c) Instability in mixed component films. The thin films are shown in cross section and the aqueous interlamellar phase is shaded.

25 Thin film instability can result in systems that contain both proteins and low molecular weight surfactants, as is the case in many foods. The origin of this instability may rest in the incompatibility of the two stabilization mechanisms; the Marangoni mechanism relyifig on lateral diffusion, the viscoelastic mechanism on immobilisation of the protein molecules that constitute the adsorbed layer. One can speculate that in a mixed system, competitive adsorption of low molecular weight surfactant could weaken or interfere with the formation of protein-protein interactions in the adsorbed layer thus destroying the integrity and viscoelastic properties of the adsorbed layer (Figure l(c)). This could be a Progressive process, with the presence of small quantities ~of adsorbed surfactant initially introducing faults in the protein film. Adsorption of more surfactant could induce the formation of protein 'islands' in the adsorbed layer, which were capable of slow lateral diffusion; too large to participate in a Marangoni type of stabilization. Adsorption of progressively more surfactant would reduce the size of the protein aggregates still further until the adsorbed protein was in its monomeric form. Ultimately, all the protein would be displaced from the interface by the surfactant. The properties of the adsorbed layers in thin films have been inferred from the results of many detailed studies of macroscopic air-water (a/w) or oil-water (o/w) interfaces. Whether such models accurately reflect the interfaces found in thin films is a matter of some contention. Certainly, the volume of bulk solution that is present beneath the adsorbed layer of a macroscopic interface is of infinitely larger volume than that found in the interlamellar region of a thin film. In the former case, surface tension has been shown to fall slowly over many tens of hours [4], consistent with conformational rearrangements of the adsorbed protein but also formation of multilayers of adsorbed protein. The timescale of such changes is irrelevant when compared, for example, to the lifetime of the foam that forms the head on a glass of beer. In addition, macroscopic interfaces are relatively insensitive to processes that can lead to the rupture of a foam lamella. For example, the adsorption of a lipid micelle and the subsequent spreading of lipid causes film rupture by a Marangoni effect (Figure 2). Interlamellar liquid associated with the polar head groups of the lipid is dragged away by the spreading lipid causing local thinning of the thin film and increasing the probability of film rupture.

Figure 2. A Schematic representation of the stages whereby a spreading particle causes local film thinning leading to film rupture [1].

26 Thus, there is a strong incentive to develop methods that allow controlled formation and characterisation of the adsorbed layer properties of thin liquid films.

2. PREPARATION OF AIR-WATER AND OIZ,-WATER THIN FILMS Although methods were available to prepare and investigate isolated air-suspended thin liquid films many years ago [5], they have only been developed further comparatively recently. The most extensive studies have been performed on surfactant-stabilized films using molecules such as sodium dodecyl sulfate [6]. Our apparatus has been developed from the film holders used by this Bulgarian group. Microscopic thin films [7,8] have generally been formed by introduction of a droplet of solution into a ground glass supporting ring or annulus (Figure 3). This device is crudely analogous to a miniaturized version of the loop children use to blow bubbles from soap solutions. However, rather than relying on gravity to drain off surplus bulk liquid as in the childs toy, film formation is initiated by withdrawal of liquid by applying controlled suction. This is achieved via a capillary sidearm that is connected to the film supporting ring. Liquid withdrawal is stopped once a thick horizontal planar film of appropriate diameter (e.g., 0.3mm) has been formed. Drainage of the thick film proceeds from this point mainly as a result of suction from the adjoining wedge shaped region that surrounds the film. This region is referred to as the Plateau border.

Figure 3. A schematic diagram of an air-water suspended thin liquid film held in a ground glass annulus.

27 Microscopic thin films are relatively fragile structures and are highly sensitive to changes in temperature, mechanical disturbance and evaporation. We have designed a dedicated chamber that allows the necessary control of the environment surrounding the film whilst not impeding observation of film drainage and measurement of equilibrium thickness or surface diffusion in the adsorbed layer. A photograph of our film chamber and a film support ring is shown in Figure 4. The film chamber is surrounded by a temperature-controllable brass housing. Crown glass optical windows allow observation of the thin film from either above or below the housing. Evaporation from the thin film, once formed in the chamber is controlled by the presence of a horseshoe-shaped trough which can be filled with the solution under investigation, prior to formation of the film in the chamber. The brass housing is suspended beneath an aluminium holder via a kinematic mount which allows levelling of the film using the micrometer adjusters. The aluminium holder fits directly onto the stage of an inverted microscope (Nikon Diaphot TMD) equipped with an epi-illumination attachment.

Figure 4. A photograph of the,thin~ film holder and temperature controlled chamber. More recently, we have developed a device that allows formation of thin films in a liquid bath [9,10]. The apparatus opens up many more opportunities for film formation under different conditions but is rather more difficult to operate than the film ring. The former apparatus, which can be used for formation of a/w or o/w thin films is shown in Figures 5 and 6. The liquid b a t h requires chemical treatment prior to introduction of the continuous phase solution. This involves thorough cleaning of the cell using chromic acid followed by drying,i Controlled silanation is used to create a highly localized hydrophobic patch on the optical window that forms the base of the cell. A 10 ~1 droplet of octadecyl trichlorosilane was found to bean effective silanation agent for this purpose [ 11]. Unreacted silanation reagent can be removed by washing with anhydrous heptane. The cell can then be filled with the continuous phase of interest which could be an aqueous protein solution, an oil-in-water emulsion or the separated continuous phase of an emulsion. An oil droplet or air bubble is then immediately introduced onto the hydrophobic patch by careful delivery using

28

Figure 5. A schematic diagram showing the apparatus used to form a thin aqueous film between oil droplets. Reproduced from reference [ 10] with the permission of Academic Press.

Figure 6. A photograph of an aqueous thin film formed between two droplets of ntetradecane. The laser beam illuminating the thin film (misaligned for clarity) can be used to measure lateral diffusion in the adsorbed layer or film thickness.

29 a hypodermic syringe and needle. The droplet will remain captive on the hydrophobic patch, provided it is of relatively small volume. The second droplet suspended from a concavetipped nozzle attached to a micrometer-controlled l ml glass syringe is then lowered into position above the captive droplet creating a thin aqueous film in the region of contact. Thin film drainage behavior can be viewed through the lower droplet by means of the inverted microscope in reflected light. It is relatively simple to convert the chamber to allow modelling of thin oil films between water droplets by application of the hydrophobic coating to all the inner surfaces of the chamber. A small hydrophilic spot can then be generated in the centre of the baseplate by judicious application of a small droplet of acid using a micropipette. In this case, captive and suspended water droplets are brought into close contact, thus providing a model for thin film formation in water-in-oil emulsions.

3. THIN F I L M DRAINAGE AND THICKNESS MEASUREMENTS Observation of the drainage process from thick film to equilibrium thin film can be very informative. Both a/w and o/w thin films have very poor contrast and are impossible to observe under the microscope under bright field (background) illumination conditions. However, it is possible to observe the films in reflected light mode using epi-illumination. This is possible because although the films are virtually transparent, a small quantity of illuminating light is reflected from both upper and lower interfaces (see inset Figure 9). This phenomenon is also exploited in the measurement of film thickness which is described later in this section.

Figure 7. A photographic sequence showing the drainage behavior of a thin film formed from 2 mM SDS in 2 mM sodium phosphate buffer, pH 7.0 containing 0.1 M NaC1. See text for description.

30 The drainage properties of surfactant-stabilized films [8], can easily be distinguished from protein-stabilized film drainage [12]. Surfactant-like drainage behavior is illustrated in the sequence of photographs shown in Figure 7. The sample shown in the figure is 2 mM sodium dodecyl sulfate in 2 mM phosphate buffer, pH 7.0 containing 0.1 M NaC1. The initial 20 seconds of drainage are characterized by rapid movement of regions of different thickness, distinguishable by their different bright colors, sweeping towards the periphery of the film (Figure 7(a)). This accurately conveys the fluid nature of the interfacial layer in these films. This phase of drainage terminates with the disappearance of colors leaving a white film which is 100-200 nm in thickness. Drainage proceeds for 2-3 minutes with darkening of the periferal region of the film to form thinner gray patches. The interlamellar liquid trapped in the central region of the film appears to be squeezed out into the Plateau border by the advancing thinner gray regions and forms white arcs around the edges of the gray regions (Figure 7(b)). This process continues in several discrete waves and the film darkens with each stage. This is followed approximately 3 minutes after film formation by spontaneous nucleation of black spots at random points in the film (Figure 7(c)). The black spots grow in size and coalesce (Figure 7(d) and (e)), and eventually approximately 5 minutes after formation the whole film becomes black (Figure 7(f)). This is termed the primary or common black film and has an equilibrium thickness of the order of 12nm. Most other low molecular weight surfactants, including polysorbate emulsifiers [ 13], sucrose esters [14], mono and diglycerides, lysolecithins [15] and lecithins, follow this type of drainage behavior with minor differences. Firstly, the surfactant must be above its transition temperature and in the liquid crystalline phase. Indeed, it is generally impossible to form stable films if the surfactant is in the gel state. Secondly, the chosen solution conditions may not favour drainage to the common black film stage. Quite often equilibrium thickness is achieved at some intermediate gray film stage or thicker. Alternatively, drainage can proceed to thickness regimes that are considerably less than that of the common black film. The observed equilibrium thickness represents the film dimensions where the attractive and repulsive forces within the film are balanced. This parameter is very dependent upon the ionic composition of the solution as a major stabilizing force arizes from the ionic double layer interactions between any charged adsorbed layers confining the film. Increasing the ionic strength can reduce the repulsion between layers and at a critical concentration can induce a transition from the primary or common black film to a secondary or Newton black film. These latter films are very thin and contain little or no free interlamellar liquid. Such a transition is observed with SDS films in 0.5 M NaC1 and results in a film that is only 5 nm thick. The drainage properties of these films follows that described above but the first black spot spreads instantly and almost explosively to occupy the whole film. This latter process occurs in the millisecond timescale. In contrast, protein-stabilized thin films display very different drainage characteristics [7]. Until recently, the work on protein-stabilized thin films was limited to preliminary measurements of equilibrium film thickness and determination of contact angle [16-19]. A sequence of photographs depicting stages in the drainage of a typical protein film are shown in Figure 8. The initial appearance of protein films immediately after formation is distinct from that of surfactant-stabilized films. The protein-stabilized thin film is characterized by a series of concentric white, black and brightly colored fringes or Newton's rings. These correspond to constructive and destructive interference patterns of light reflected from: the upper and lower interfaces of the film and interconnect regions of similar thickness. Initially, the fringes are closely spaced indicating that the film is thick. In addition, there is a steep

31 thickness gradient across the film (Figure 8(a)), which is thinnest in the central region. As drainage proceeds the fringes become more widely separated and the region bordering the contact line at the perimeter of the film lightens in color (Figure 8(b)). This is followed by darkening of the periphery of the film (Figure 8(c)) which eventually becomes black (Figure 8(d)) after approximately 10 minutes drainage (Figure 8(e)). Formation of a continuous black ring around the perifery of the film (Figure 8(0) traps liquid in the thicker central region and slows down the rate of drainage. Nevertheless, drainage proceeds, albeit at a slow rate due to the constriction at the perifery of the film and results in a shrinkage in the dimensions of the lighter central region of the film (Figure 8(g)). Eventually, the whole film becomes black (Figure 8(h)) but this may take in excess of 20 minutes. The drainage rate of the film is very sensitive to the time history of the interface. Aged interfaces generally result in films that are very slow to drain, due to their increased interfacial shear viscosity [3].

Figure 8. A photographic sequence showing the drainage behavior of an a/w thin film formed from BSA in distilled water, adjusted to pH 8 containing 25 mM NaC1. See text for description. Reproduced from reference [12] with the permission of Academic Press. Equilibrium film thickness can be measured by interferometry [7,8] using an apparatus of the type shown in Figure 9. When studying a/w films, we use an interferometer mounted above the stage of the inverted microscope. The interferometer comprises an interrogating beam from a 3 mW helium-neon laser (632.8 nm) which is passed through an optical chopper and is directed down onto the surface of the film by means of a beam splitter. The beam is focused onto the thin film using an extra long working distance objective lens (Nikon M-plan, magnification x20 or x40). The diameter of the illuminated spot on the film surface is 25 50/~m. The majority of the incident light is transmitted through the film and care must be taken with the inverted microscope to ensure that appropriate barrier filters are fitted to the

32 eyepieces to avoid injury to the operator. A small fraction of the incident light is reflected from both the upper and lower interfaces of the film and passes back up the optical axis of the interferometer (inset Figure 9). These reflected beams are transmitted by a 633 nm narrow pass filter, positioned above the beam splitter and the combined signal is detected at a photodiode. The detected intensity fluctuates between the two extremes of totally constructive and totally destructive interference, thus producing in the photodetector output a varying signal that is a measure of film thickness. The signal from the photodiode is amplified (AMP) and fed into a phase sensitive detector (PSD) referenced to the chopper frequency. The PSD, which reduces signal noise and improves signal stability, is set up with a constant negative offset to compensate for background reflected light from the chamber windows etc. The signal is output via a chart recorder.

Figure 9. A schematic diagram of the interferometer used to measure thin film thickness. The inset shows that light is both transmitted and reflected by the thin film. Reproduced from reference [7] with the permission of the Royal Society of Chemistry. The equilibrium film thickness (h) is calculated using the expression:

~,

[

h . . . . . sin "l ~ 27rn [

I/Ira 1 + [4R/(1-R)2].[1-I/Im]

] 0.5 } J

(1)

zyxwvu 33

where (n-l) 2 R

-

(2)

(n+l) 2

and k is the wavelength of the laser, n is the refractive index of the film, I is the intensity of light at the photodiode at equilibrium, and Im is the intensity at the last maximum. In the ideal situation the chart output resembles the interferograph shown in Figure 10, and this can be achieved relatively easily with protein films by careful positioning of the spot near the central region of the film. Often it is more difficult to achieve such output with surfactant films due to their fluid nature and the fact that regions of differing thickness sweep across the interrogating beam of the interferometer. Accurate determination of I and Im can be difficult due to uncertainty in the position of the minimum. We find that the most reproducible results are obtained if the minimum value is taken as the signal observed after removal of the film annulus at the end of the experiment.

Figure 10. Interferograph from a draining a/w suspended thin film showing the calculated change in film thickness with time. Reproduced from reference [20] with the permission of the Institute of Brewing.

34 If a complete interferograph is obtained it is possible to construct a drainage curve for the film (Figure 10) since the film thickness at each maximum and minimum can be calculated from equation (1). In the case of aqueous thin films between oil droplets (Figure 5), the interferometer beam is brought into the microscope through the epi-illumination attachment whereby the objective lens is used to both observe the film and focus the interferometer beam. The contrast of the observed image is much improved in stray light is minimized by positioning a pinhole at the image plan of the epi-illumination device. The thickness calculations remained the same as for the a/w films as the refractive index of the aqueous thin film was the same in both cases.

4. SURFACE DIFFUSION MEASUREMENTS BY FRAP Observations of film drainage behavior provides an indication of the structural properties of the adsorbed layers. It is simple to distinguish between protein or surfactant-stabilized films. However, most food systems contain mixtures of both proteins and low molecular weight surfactants. Detailed study of the thin film properties of protein solutions containing increasing levels of surfactant reveals a corresponding decrease in film thickness with increasing surfactant concentration [ 10,13,15,21 ]. In addition, at certain critical molar ratios of the two components, we have observed a transition in drainage behavior to a intermediate type of drainage [15,22]. The latter possess features of both surfactant-like and protein-like drainage. Typically, distorted Newton's rings are observed as the once rigid protein stabilized interface becomes more fluid. Clearly, important changes in the adsorbed layer structure and surface rheological properties are occurring but it is difficult to identify a method that would allow their direct investigation in the thin film. Certainly such delicate structures would not be amenable to study by conventional surface shear or surface dilational methods. Indeed, these methods are still not widely available for the investigation of macroscopic interfaces. A radical alternative was sought and found. A technique referred to by several different names including fluorescence recovery (or redistribution) after photobleaching (FRAP), fluorescence microphotolysis and fluorescence photobleaching recovery (FPR) was first developed in the mid 70's and has proved a useful technique for the study of lateral diffusion processes in biological cell membranes and the cytoplasm [23,24] and has been reviewed recently [25]. The previous use of the technique in interfacial studies was limited to investigation of the lateral diffusion of lipid at the a/w interface of a Langmuir trough [26]. Several variations of the FRAP technique exist but the simplest are based on the principals outlined in the schematic diagram shown in Figure 11. The method requires that the molecular species of interest is fluorescent labelled or alternatively that an independent fluorescent probe molecule is partitioned into the environment of interest. In the case of thin films, the surface diffusion properties of a given protein in the adsorbed layer can be measured by forming a thin film (diameter 100-200 #m) as described above which includes fluorescent-labelled protein. An attenuated laser beam is used to illuminate a small spot (approximate diameter 5 #m) on the surface of the thin film, eliciting fluorescence from labelled molecules contained within the spot, which is recorded (Figure l l(a)). These fluorescent molecules are then irreversibly photodestroyed (bleached) by increasing the intensity of the laser beam approximately 1000x for a few milliseconds (Figure 1 l(b)), before returning the laser intensity to its attenuated level. Fluorescence returns to the bleached spot only if the bleached molecules are free to diffuse laterally away from the spot to be replaced

35 by non-bleached molecules in the surrounding film diffusing into the spot (Figure 1 l(c) and(d)). Measurement of the time dependence of this process and knowledge of the dimensions of the bleached spot, allows calculation of the surface diffusion coefficient.

Figure 11. A schematic diagram showing the various stages in typical spot FRAP experiment. See text for explanation The design and construction of a FRAP apparatus has been recently reviewed [27]. The purpose of the majority of the optical components is to deliver a well-defined, microscopic Gaussian or uniform circular cross section beam, that can be rapidly modulated, to the sample. A schematic of our FRAP apparatus is shown in Figure 12. The light source used is an Argon ion laser (Coherent Innova 100 - 10). We have examined three different beam modulation arrangements during the course of our studies. The first device we tried was an acousto-optic modulator (Coherent 304A) which contains a crystal, that diffracts the input laser beam into a number of secondary beams. The intensity of the secondary beams can be modulated by application of an RF signal to the crystal [28]. The disadvantage of this device is that the output beam was of ellipsoidal rather than circular cross section and therefore did not have a true Guassian intensity profile. Since the uniform circular beam is obtained by projecting the Guassian beam from the laser through a microscopic pinhole, it was not possible to deliver a uniform circular or Gaussian cross section bleach pulse to the sample as required. The second modulation method involved the positioning of an LCD light valve (Displaytech) between two crossed Glan Thompson polarizers. Application of a DC voltage to the light valve caused rotation of the plane of polarisation of the laser beam from that of the first polarizer such that it was no longer extinguished by the second polarizer. Although this method produced acceptable beam profiles, the LCD had a rather limited lifetime before laser-induced damage significantly reduced its performance resulting in a reduction in the intensity difference between the monitor and bleach beams. Our preferred modulation method is one of the first described [26], and involves generation of an attenuated beam by reflection off glass flats as shown in Figure 12. When the fast electronic shutter (c) is closed, only the monitoring laser beam (a), which has been attenuated by multiple reflection illuminates the

36 sample. When the shutter is open the intense bleaching beam (b) which is transmitted through two of the glass fiats passes through to the sample. The crucial factor with this modulator arrangement is beam alignment to ensure that both attenuated and bleach beams are coincident at the sample.

Figure 12. A schematic diagram of a FRAP apparatus. See text for a key to the abbreviations. The beam provided by the modulator passes through a beam monitor (beam splitter and photodiode), the signal from which is used to electronically compensate for minor fluctuations in the laser beam intensity. The beam is then launched through a pinhole aperture (A~) located at the image plane, at the entrance port of the epi-illumination attachment of the fluorescence microscope. Our apparatus can be used with both upright (Nikon Optiphot) or inverted (Nikon Diaphot TMD) microscopes but the latter is most convenient for thin film measurements. The filter block in the epiillumination attachment is selected to match the laser line used for excitation andthe emission peak of the fluorescent probe. The 488nm line is the most popular for FRAP measurements with the Argon ion laser, as it can be used to excite a number of different fluorophores including fluorescein, 4-chloro-7-nitrobenz-2-oxa-l,3-diazole (NBD chloride) and members of the carbocyanine family. The use of the well-defined laser-line for

37 excitation renders the short band pass interference filter usually supplied with the filter block for wavelength in the filter block redundant. A 510 nm dichroic mirror (DM), mounted at an angle of 45 ~ is suitable for reflection of the 488 nm excitation beam used for the fluorophores above, through the extra long working distance objective (Nikon) lens (magnification x40 or x20) of the microscope and onto the sample. This will also allow acceptable transmission of the emitted light from the above fluorescent labels. A 520 nm long pass filter (LBF) removes stray excitation light and prevents it reaching the photon counting photomultiplier tube (PMT; Thorn-EMI 9816B) positioned at the cine camera port of the inverted microscope. The PMT is protected during the bleaching pulse by an electronic gating circuit and a mechanical shutter (MS). Prior to entering the detector, the emitted light beam passes through a second aperture (A2) again positioned at the image plane. The two apertures have equivalent diameters (e.g., 200/~m) and serve to make the apparatus confocal. This feature is not so important in the case of thin films, since these can be considered 2dimensional systems once they have drained to equilibrium thicknesses. However, the confocal arrangement is most useful if diffusion measurements are planned using 3-dimensional systems (e.g., probe diffusion in a gel etc). System timing and control, data acquisition and data analysis are performed using a VME microcomputer system (Motorola 68020). The diameter of the focused laser spot on the sample was measured using a beam profile measuring device (BeamScan, Model 2180; Photon Inc.). FRAP data were analysed by a non-linear least squares fit to an expression [8,23,29], defining the time dependence of the fluorescence recovery (F(t)). The apparatus as described above delivered a laser spot of uniform circular cross sectional intensity to the sample and the recovery curves obtained could be analysed with the expression:

F(0 = Fo~

zyxwvutsr

1 -2(1 - Fo/F ~)[O.5(rD/t)e2~D It(Io(2 ro/t) 100

(m + 1)!(2m + 2)! + I2(2rD/t)) +

/.

(-rD/t)m+2] }

(3)

m!2(m + 2)! 2

m=l

where F0 is the fluorescence intensity after the bleach, F o~ is the fluorescence intensity to which the signal recovers and I0 and 12 are modified Bessel functions. The lateral diffusion coefficient, D, is given by

D = w2/4ro

(4)

where w is the radius of the circular spot and ro is the characteristic diffusion time. It is simple to modify the apparatus to include a spacial filter that provides a Gaussian beam profile at the sample. T h e recovery curves obtained with such an experimental arrangement could be analysed by a much simpler expression [29] of the form:

38

zyxwvutsrq zyxwv

F0 + Foo(t//3ro) F(t) = --

(5)

1 + t/~r D

where/3 is related to the proportion of bleach, P (i.e., the prebleach fluorescence intensity F0 divided by the prebleach fluorescence intensity). In practice, the value of/3 is obtained from a lookup table in the analysis programme.

4.1. Fluorescence-labelling of samples Measurements of surface diffusion in thin liquid films by the FRAP method requires the presence of fluorescent molecules in the adsorbed surface layer. The low molecular weight of surfactant molecules and absence of a reactive side groups makes fluorescent-labelling difficult. In addition, conjugation with a fluorophore is likely to significantly alter the surface active properties of the molecule. Therefore, it is preferable to adulterate the surfactant solution of interest with trace quantities of fluorescent lipid or surfactant analog. A range of molecules are commercially available [31] and we have had considerable success using samples such as negatively charged, 5-N-(octadecanoyl)-aminofluorescein (ODAF), positively charged, 3,3'-dioctadecyl oxacarbocyanine perchlorate (DiO), neutral NBD-dihexyldecylamine and phospholipid analogs such as NBD-phophatidylethanolamine. FRAP measurements of protein diffusion at interfaces can be achieved in one of several ways (Figure 13). One option involves controlled covalent labelling of the protein molecule of interest with a reactive fluorescent molecule as in Figure 13(a), [12,32]. An alternative and in some cases simpler approach which we have used recently involves direct addition of trace quantities of a fluorescent lipid analog (Figure 13(b)). For example, we have shown that an amphipathic fluorescent molecule such as ODAF, which has low solubility in water, when added in very low concentrations preferentially partitions into the adsorbed layer, where it can be used to probe the global surface viscosity [8,33]. One major advantage of this approach is that it eliminates the requirement to isolate a protein of interest from a complex mixture (e.g., /3-1actoglobulin from whey protein isolate), the covalent labelling and reconstitution of the system by adding back the labelled protein. This could alter the properties of the total system. The third possible approach, which has been widely used in FRAP studies in cell biology, involves indirect, selective fluorescent-labelling of the protein of interest by interaction with a fluorescent-labelled antigen binding fragment of an antibody raised against the protein (Figure 13(c)). This opens up the opportunity of selective labelling of a protein in a complex mixture, without the need to isolate, label and then reconstitute the system. Considerable care needs to be exercised during fluorescence-labelling of proteins to avoid alteration of the surface properties of the protein. Many reactive fluorescent derivatives are now available from most major chemical companies and specialist suppliers such as Molecular Probes Inc [31]. The isothiocyanate derivative of fluorescein (FITC) has been widely used in our work to label the e-amino group of lysine residues in proteins. Efficient labelling is achieved if the pH of the protein solution is raised to approximately 9.2 to ensure significant deprotonation of the amine groups on the protein surface. Under such conditions, effective labelling of BSA can be achieved in the presence of a 2-fold excess of FITC. The predominant product obtained under these conditions is singly labelled FITC-BSA [12]. Covalent reaction of this fluorophore will occur at lower Ph [32], but the reduction in rate of reaction means it is necessary to add higher molar ratios of fluorophore to the stock

39 protein solution. Labelling under less alkaline conditions is necessary in the case of ~lactoglobulin, since this protein undergoes an irreversible denaturation under our normal labelling conditions [34].

Figure 13. A schematic diagram showing three different approaches to introducing a fluorescent label into thin films to measure surface diffusion in the adsorbed layer. Fluorescein is highly fluorescent at neutral pH. The quantum yield of this fluorophore is very significantly reduced as the pH is reduced below neutrality. This is caused by the protonation of a negatively charged carboxylic acid group on the fluorescein molecule. Thus, labelling of a protein by a single FITC molecule results in the loss of a positively charged

40 amino group and the introduction of a negative charge at neutral pH, a change in net charge of 2. Therefore, it is important to ensure that extent of labelling is minimized and that the properties of the mildly labelled protein do not differ significantly from those of the unlabelled protein. We have examined the foaming properties, thin film drainage and thickness properties of labelled BSA, ~-lactoglobulin, ot-lactalbumin and ~-casein and have not identified significant alteration in their properties provided the prepared conjugate contains on average less than 1 mole of fluorophore per mole of protein.

zyxw

4.2. Surface concentration measurements by fluorescence The FRAP apparatus can also be used in a semi-quantitative manner to measure the surface concentration and subsequent competitive displacement of adsorbed labelled species, such as the fluorescent-labelled protein in the adsorbed layer of a/w or o/w thin films [ 10]. This can be achieved by focusing the low power 488 nm beam on the film and detection of the emitted fluorescence using the FRAP photon counting photomultiplier. The detected fluorescence signal is proportional to the amount of adsorbed protein at the interfaces of the thin film provided that the incident laser intensity is kept constant. Calculations have proved that the contributions from non-adsorbed protein molecules in the interlamellar region of the film are negligible [12].

4.3. FRAP measurements of surface diffusion in surfactant or lipid-stabilized thin films Thin films stabilized by SDS were selected as the test system during the construction and commissioning of our FRAP apparatus [8]. Most measurements were performed on samples containing 1 mole of ODAF per 150 moles of SDS. Results obtained using lower concentrations of ODAF ( > two-fold) confirmed that the data were not influenced by the presence of ODAF. Surface excess measurements were performed using a modification of the method of Weil [35]. The two-fold increase in concentration of ODAF between solution and collected foam showed that it was preferentially associated with the a/w interface although not as effectively as SDS which showed a five-fold increase in concentration. The solution conditions chosen were appropriate for formation of common black films and measurements were undertaken once the films had reached equilibrium thickness. Fluorescence recovery was rapid necessitating use of a very short bleach pulse [8]. The signal-to-noise ratio of individual curves was quite poor and acceptable data curves were only obtained after summation of 10 or more experimental curves. A typical curve is shown in Figure 14(a). An average surface diffusion coefficient of 6.8x10 7 cm2/s was obtained for ODAF in SDS-stabilized films. Two different spot sizes were used to determine whether ODAF mobility was due to lateral diffusion or linear flow in the thin film. These phenomena can be distinguished since the characteristic recovery time is proportional to the increase in the spot diameter under conditions of flow and to the increase in spot diameter squared when diffusion is the dominant process [29]. The results obtained supported the conclusion that the surface molecular mobility observed in these films resulted from diffusion rather than flow. The observed lateral diffusion coefficient was dependent upon the positioning of the laser spot in the film. A 25 % reduction in the diffusion coefficient was observed in the region within 25-50 #m of the periphery of the film. This may result from the presence of thinner regions at the film periphery or other competing processes such as marginal regeneration. Increasing interlamellar viscosity by addition of glycerol reduced the rate of thin film drainage and decreased the lateral diffusion coefficient.

41

Figure 14. Typical FRAP data curves obtained with (a) 2 mM SDS in 2 mM sodium phosphate buffer, pH 7.0 containing 0.1 M NaC1 and 14/zM ODAF; (b) FITC-BSA (0.5 mg/ml) in distilled water, pH 8.0, at an equilibrium film thickness of 83 nm; (c) FITC-BSA (0.2 mg/ml) in 50 mM Na acetate buffer, pH 5.4 at an equilibrium common black film thickness of 14 nm. This study comprised the first reported direct experimental measurement of surface diffusion in air-suspended thin liquid films.

4.4. Surface diffusion measurements in protein-stabilized films The solution diffusion properties ofFITC-labelled BSA were measured by FRAP [12]. The results showed that the protein diffused freely in solution with a diffusion coefficient of approximately 3x10 7 cm2/s. This was in reasonable agreement with previously published values [36]. FRAP measurements were also made on thin films stabilized by FITC-BSA. The films were allowed to drain to equilibrium thickness before measurements were initiated. Thin films covering a range of different thicknesses were studied by careful adjustment of solution conditions. BSA stabilized films that had thicknesses up to 40 nm showed no evidence of surface diffusion as there was no return of fluorescence after the bleach pulse in the recovery part of the FRAP curve (Figure 14(c)). In contrast, experiments performed with thin films that were > 80 nm thick showed partial recovery (55 %) of the prebleach level of fluorescence (Figure 14(b)). This suggested the presence of two classes of protein in the film; one fraction in an environment where it was Unable to diffuse laterally, as seen with the films of thicknesses < 45 nm, and a second fraction that was able to diffuse with a calculated diffusion coefficient of l x l 0 -7 cm2/s. This latter diffusion coefficient was 3 times slower than that

zy

42 observed for FITC-BSA in solution. Care needs to be exercised in the interpretation of these data. Firstly, the slow drainage of the protein films especially once the perimeter of the film reaches black thicknesses suggests that these films contain very little interlamellar liquid. Therefore, it is reasonable to assume that the vast majority of the fluorescence signal from the < 45 nm thick films originates from protein in the adsorbed layer. The complete immobility of the fluorescent-labelled protein in these structures over the timescale of our experiments suggests that diffusion in the interlamellar liquid region is very restricted or highly compartmentalized. Indeed, it is possible that protein molecules bridge between the two interfaces [37]. The partial recovery observed in films > 80 nm thick (Figure 14(b)), is consistent with abolition or a significant reduction in the impediments to diffusion in such films. However, the diffusion coefficient is significantly lower than that observed in aqueous solution. Calculations predict a significant enhancement (several orders of magnitude of concentration ) of protein in the adsorbed layer compared to the interlamellar solution. Therefore, it is necessary to define a mechanism that can account for an increase in protein concentration in the interlamellar space to explain the observed 55 % recovery, whilst impeding protein diffusion compared to bulk solution. One hypothesis involves a low affinity interaction and exchange of protein adsorbed in the secondary layers with that in the interlamellar space. This would be consistent with a previous FRAP result of mobile and immobile fractions of BSA bound at a macroscopic quartz-water interface [38]. In this study, partial recovery was attributed to adsorption/desorption processes in the adsorbed multilayers.

zyxwvutsr

5. CHANGES IN THIN FILM PROPERTIES AS A FUNCTION OF INCORPORATION OF L O W M O L E C U L A R W E I G H T SURFACTANT IN THE ADSORBED PROTEIN LAYER

5.1. Air-water thin films We have predicted that transitions in surface diffusion behavior will be observed under certain conditions in mixed protein/low molecular weight surfactant systems (Figure 1). The behavior of these systems depends on the protein and surfactant type. The effect of protein type may be studied separately by examining the effect of a given surfactant, for example the polysorbate emulsifier, Tween 20 (polyoxyethylene (20) sorbitan monolaurate) on different proteins. This is a non-ionic emulsifier which is water soluble, has a critical micelle concentration of approximately 35 /zM and has a bulky polar headgroup [39]. In our experiments, we have formed films from a range of samples composed of a fixed concentration of the protein of interest but containing increasing levels of surfactant. To facilitate comparison, the data obtained are uniformally presented in terms of the molar ratio of Tween 20 to protein (R). We have been able to categorise the effect of this emulsifier on a range of proteins into three classes of behavior.

5.1.1. Type I: Globular protein with surfactant binding activity. ~-lactoglobulin (~-lg) and Tween 20 is a classic example of a mixed component system that displays Type I behavior [13,22]. A summary of film thickness, surface concentration of FITC-/~-lg and FITC-~3-1g surface diffusion is given in Figure 15. All these data were obtained at a protein concentration of 0.2 mg/ml.

43

Figure 15. A summary of the film thickness (o), surface concentration of FITC-B-lg (x) and FITC-/~-lg surface diffusion (A) as a function of the molar ratio of Tween 20 to protein (R) at the interfaces of a/w thin films. Reproduced from Faraday Discussion 98 with the permission of the Royal Society of Chemistry. Fluorescence measurements reveal that the displacement of FITC-~-lg from the a/w interfaces occurs in several distinct steps, The initial phase of FITC-~-lg displacement begins at R = 0.1 [10]. Paradoxically, this coincideswith an increase in film thickness. Observation of the films reveals the appearance of coexisting regions of two distinct thicknesses in the thin film in the R value range of 0.4 - 0.8 [13,22]. We interpret the pseudo plateau in the displacement data as an indication that no further displacement of protein occurs in certain regions of the film (e.g., in parts of the thicker regions) in this R value range. However, further displacement of FITC-/3-1g is observed in the R value range of 0.8 to 1.0, which shows good correlation with the onset of surface diffusion in this a/w thin film system. Only minor displacement is observed between R = 1.0 and 2.0, which corresponds to the concentration ratio where the surface diffusion coefficient is increasing sharply. Further gradual displacement is observed at higher R values. Major changes in all three measured parameters in Figure 15 occur at R = 0.9 - 1.0. It is significant that this is also the point where instability is first observed in the bulk foam [13,22]. Thus, there is strong evidence that suggests a link between changes in the adsorbed

44 layer structure in the thin films and bulk foam stability in this system. Complete understanding of the behavior of this system is only possible with knowledge of the surfactant binding properties of the protein. We have measured the binding of Tween 20 by ~-lg, and found it to be characterized by a dissociation constant (Kd) for the complex of 4.6 ~M [10]. This has allowed calculation of the relative concentrations of free/3-1g, Tween 20 and ~-lgTween 20 complex present in a given solution of these two components. From the binding data, it is evident that at R = 1, the solution contains effectively equivalent amounts of all three species, free Tween 20,/3-1g and complex [40]. Using these data and further evidence [22,40], we have been able to construct an elaborate mechanism that explains different stages in the breakdown of the adsorbed layer structure in this system, which is shown in Figure 16.

Figure 16. A schematic representation of the change in interactions and composition of the adsorbed layer at the a/w interface in solutions of/~-lg containing increasing levels of Tween 20. Reproduced from reference [40] with the permission of the Royal Society of Chemistry. In this schematic, the B-lg molecules are depicted as jigsaw puzzle pieces, since the proteinprotein interactions in the interface generated by this particular protein are very strong [3]. The experimental evidence is consistent with the complex formed between/3-1g and Tween 20 being unable to interact with B-lg or other molecules of the complex. It is convenient to depict the complex in our schematic model by shielding the protein-protein interaction site on the icon with the hydrophilic polyoxyethylene chain of the Tween 20 molecule. This is

45 reasonable since the complex has been shown to have a much larger hydrodynamic radius than /3-1g alone or the/3-1g/Span 20 complex [22]. Span 20 is sorbitan monolaurate and therefore lacks the polyoxyethylene side chains that are present on the Tween 20 molecule. This inability to interact may explain the film thickening (Figure 15) observed at low R values (0.1 - 0.6), since the complex may become trapped in the adsorbed layers or interlamellar space of the draining film. More importantly, at R > 0.4, the presence of the complex appears to induce loss of multilayers from the film and the appearance of local thin regions (Figure 15). As R reaches 0.9, complex begins to appear in the primary adsorbed layer, breaks the cohesive nature of the adsorbed/3-1g layer and causes the onset of surface diffusion. The sharp increase in surface diffusion coefficient of the FITC-/3-1g/Tween 20 complex is superseded by a more gradual rate of increase at R > 1.3, as the appearance of free Tween 20 in the primary adsorbed layer decreases surface viscosity by diluting the adsorbed complex. Finally, at R > 5, the complex is almost completely displaced from the interface. Several other proteins that bind emulsifiers follow the general trends of this model. For example, the properties of the lipid binding protein from wheat called puroindoline has broadly similar properties [15]. 5.1.2. Type II: Globular protein which does not bind surfactant. Solutions containing mixtures of c~-lactalbumin (o~-la) and Tween 20 are a classic example of a two component system that displays Type II behavior [21]. A summary of foam stability, film thickness and FITC-c~-la surface diffusion is given in Figure 17. Alone, a-la produced less stable foams than/3-1g, and it was necessary to increase the stock protein concentration to 0.5 mg/ml (35.4 #M).

Figure 17. A summary of the bulk foam stability (Fq), equilibrium thin film thickness (o), and FITC-a-la surface diffusion (zX)as a function of molar ratio of Tween 20 to protein (R). The concentration of a-la was 0.5 mg/ml (35.4/zM). Reproduced from reference [41] with the permission of VCH Verlagsgesellschaft.

46 Tween 20 was considerably more effective at reducing the stability of foams of o~-la than was the case with/3-1g. There was a significant decrease in o~-la foam stability in the presence of Tween, at R values as low as 0.05. Minimal foam stability was observed at R = 0.15. There was no observed change in film drainage behavior or onset of surface diffusion in the adsorbed protein layer up to this R value. The only observed change was a progressive decrease in film thickness. Therefore, it is likely that disruption of adsorbed multilayers is responsible for a reduction in the structural integrity of the adsorbed protein layer and that this increases the probability of film rupture. An improvement in foam stability was observed as R was increased to > 0.15 (Figure 17). This was accompanied by the onset of surface diffusion of c~-la in the adsorbed protein layer. This is significantly different compared to our observations with/3-1g, where the onset and increase in surface diffusion was accompanied with a decrease in foam stability. Fluorescence and surface tension measurements confirmed that a-la was still present in the adsorbed layer of the film up to R = 2.5. Thus, the enhancement of foam stability to levels in excess of that observed with o~-la alone supports the presence of a synergistic effect between the protein and surfactant in this mixed system (i.e., the combined effect of the two components exceeds the sum of their individual effects). It is important to note that Tween 20 alone does not form a stable foam at concentrations < 40 ~M [22]. It is possible that o~-la, which is a small protein (Mr = 14,800), is capable of stabilizing thin films by a Marangoni type mechanism [2] once ot-la/o~-la interactions have been broken down by competitive adsorption of Tween 20. A schematic model showing the Tween 20-induced change in the structure of the adsorbed layer of c~-la is shown in Figure 18. In this schematic diagram, the o~-la molecules are depicted as shapes that interact together (Figure 18(a)), but in a much weaker fashion than the /3-1g molecules in Figure 16. This is consistent with the lower interfacial viscosity observed with this protein [3]. In this simpler two component system, competitive adsorption of low levels of Tween 20 (0< R < 0.2), may cause faults to occur in the primary adsorbed layer of protein (Figure 18(b)). One can envisage the presence of large regions or plates of interacting ot-la molecules at the interfaces of the thin film, which are not capable of independent surface diffusion on the timescales of the FRAP experiment. However, the ability of such thin films to withstand thermal or mechanical stretching would be significantly reduced by faults or weaknesses in the adsorbed layer due to the presence of low levels of Tween 20. Incorporation of higher levels of Tween 20 into the adsorbed layer would progressively increase the breakdown of ot-la interactions such that at R = 0.2, surface diffusion of FITC-o~-la is observed (Figure 18(c)). Ultimately, as the concentration of Tween 20 is increased further (R = 2.5), the protein is completely displaced from the interface. 5.1.3. Type III: Random protein that does not bind surfactant. /3-casein (/3-cas) and Tween 20 is an example of a mixed component system that displays Type III behavior [42]. A summary of foam stability, film thickness and FITC-/3-cas surface diffusion is given in Figure 19. All these data were obtained at a/3-cas concentration of 0.5 mg/ml. The foam stability of /3-cas foams progressively decreased with added Tween 20. In contrast, there was a very sharp transition in equilibrium film thickness at R = 0.5. Surprisingly, surface diffusion of/3-cas was not detected at any R value in these films. This was unexpected since it has been reported that adsorbed layers of 13-cas are characterized by a very low surface viscosity [3], signifying that protein-protein interactions in/3-cas films are very weak. We had expected to observe surface diffusion either in the films stabilized by

47

Figure 18. A schematic representation of the change in interactions and composition of the adsorbed layer at the a/w interface in solutions of c~-la containing increasing levels of Tween 20. protein alone or in the presence of low quantities of Tween 20. The data suggest an alternative mechanism of destabilization operates which involves phase separation of the/3-cas and Tween 20 in the adsorbed layer. A very speculative schematic representation of this mechanism is shown in Figure 20. Evidence in support of this model comes from observations of the coexistence of two regions of differing thickness in the thin films at R = 0.5 which coincided with the observed transition in film thickness. Photographs of thin films depicting this condition are shown in Figure 21.

48

Figure 19. The foam stability, film thickness and surface diffusion of adsorbed ~-cas as a function of the concentration of added Tween 20 in a/w thin films. The ~-cas concentration was held constant at 0.2 mg/ml (8.33 /~M). Reproduced from reference [41] with the permission of VCH Verlagsgesellschaft.

Figure 20. A schematic representation of the change in interactions and composition of the adsorbed layer at the a/w interface of solutions of/3-cas containing increasing levels of Tween 20.

49 Fluorescence measurements revealed that the concentration of adsorbed protein was much reduced in the thinner regions, but high in the thicker regions. There was only sufficient protein adsorbed to allow FRAP measurements to be made in the thicker regions of the film. The results showed that the protein present in the thicker region was immobile. The absence of significant fluorescence from the thinner regions of the film suggested that these regions contained very little protein. However, /3-cas must be able to influence the interface in these regions since the stability of the foams was still minimal even at high concentrations of Tween 20. If the protein had been totally displaced the concentration of Tween 20 alone should have been sufficient to stabilize the foam.

Figure 21. Photographs of the drainage behavior and coexistence phenomena in thin films formed from solutions of/3-cas and Tween 20 of composition R = 0.5. (a) Early stages of drainage of the thin film showing protein-like (concentric rings) and surfactant-like (distortions) features; (b) A sample showing a few dark (thin) regions in a predominantly gray film; (c) A sample showing light (thick) regions in a predominantly dark film. In summary, three different types of emulsifier-induced transitions in thin film behavior have been observed. The mechanisms of destabilization depend on the strength of proteinprotein interactions in the adsorbed layer. The stronger the interactions, the more emulsifier is needed to destabilize the thin film. Knowledge of the mechanism of destabilization allows the formulation of scientific strategies for control of stability. Preliminary results have shown that enhancing the interactions in the adsorbed layer through the addition of natural crosslinking agents is a promising approach [43]. Alternatively, introduction of a component capable of selective binding of the low molecular weight destabilizing agent (e.g., lipid) is another possibility [15,44].

zyxwvut

5.2 Oil-water thin films We have used our thin film techniques to compare the behavior of the protein adsorbed layers of a/w and o/w thin films [10,45]. The results revealed significant differences between these two related systems. Data from film thickness, FRAP and surface concentration

50 measurements from o/w thin films stabilized by mixtures of ~-lg and Tween 20 are presented in Figure 22.

Figure 22. A summary of the film thickness (o), surface concentration of FITC-/~-lg (x) and FITC-~-lg surface diffusion (zX) as a function of the molar ratio (R) of Tween 20 to protein at the interfaces of o/w thin films. Reproduced from Faraday Discussion 98 with the permission of the Royal Society of Chemistry. These data can be compared with those for a/w films shown in Figure 15. Such comparison suggests that there is substantially less protein at the interface in o/w thin films, indeed almost five times less. However, care needs to be exercised when equating surface concentration to fluorescence intensity. It is possible that the fluorophore is located in different environments in the two types of thin film and that the difference in fluorescence intensity is a fluorescence quantum yield effect. However, this is unlikely since the surface concentration, as judged by the surface fluorescence signal at which surface diffusion is first observed, in both a/w and o/w films is very similar at approximately 600 counts per channel. It is reasonable to assume that the structure of the adsorbed layer is similar at the point where surface diffusion is first observed. The presence of similar surface counts indicates that the quantum yield of fluorescence is similar at both o/w and a/w interfaces. Thus, this strongly supports the

51 presence of multilayer structures in the adsorbed layers of/3-1g in a/w thin films. These multilayers need to be removed by competitive adsorption of Tween 20 before surface diffusion is observed. However, in the case of the o/w thin films, the surface concentration data confirms that these films are initially comprised of an adsorbed monolayer. Displacement of the protein from the adsorbed layer in o/w thin films shows very different behavior from its a/w counterpart. Although displacement of protein from the o/w interfaces initiates at approximately the same solution composition (i.e., R = 0.1), there i s little evidence for the stepwise displacement observed in the a/w thin films. This observation is further confirmation of the monolayer versus multilayer structure at the o/w and a/w thin films. The displacement of/3-1g has also been investigated in oil-in-water emulsions of ntetradecane [46,47]. In these reports it was shown that the protein was not completely displaced until R = 10, which was considerably higher than R = 1 - 2 in Figure 22. This will be discussed further below. The onset of surface diffusion of adsorbed FITC-/3-1g in both a/w and o/w film coincides with the initiation of displacement of protein from the monolayer (or primary adsorbed layer). However, the R value at which this occurs is different for the a/w and o/w systems. The origin of this difference is not clear, particularly if the onset of surface diffusion is explained by the adsorption of complex as is the case with the a/w films (Figure 16). The experimental results shown in Figure 22 were obtained from thin films prepared by adsorption from aqueous solutions containing 0.2 mg/ml/3-1g and appropriate concentrations of Tween 20, to captive oil droplets (as in Figure 5). Under these solution conditions, only 6 % of the/3-1g was in the complexed form rather than the 50% present at the point where surface diffusion is first observed in the a/w thin films. It is possible that this small amount of complex is sufficient to disrupt the monolayer in the o/w thin film allowing surface diffusion of the remaining adsorbed protein. An alternative explanation involves the emulsifier (/3-1g and Tween 20) concentration to interfacial area ratio. Our o/w thin film experiments involved a protein concentration of 0.2 mg/ml and an interfacial area of approximately 6x10 -5 m 2. This amounts to a protein load per unit area 200x greater than used in previous studies of emulsions of these two components [46,47]. In the latter, complete displacement of/3-1g required the presence of a 10-fold higher Tween 20 concentration than reported in our thin film experiments. Thus a considerably larger fraction of the total protein present was adsorbed in the emulsion experiments. Therefore, at an equivalent R value there was more Tween 20 present in the thin film system relative to the amount of adsorbed protein, than in the emulsion. This could explain the displacement of/3-1g at lower R values in the thin film experiments. We have tested this hypothesis in some recent o/w thin film experiments [45]. It was not practical to reduce the protein load per unit area of interface to that found in the emulsion experiments, since the very low concentrations required would have been very slow to reach equilibrium adsorption. We circumvented this problem in a unique way. Rather than adsorb emulsifier mixtures from aqueous solution, we formed the oil droplets and the thin film in a preformed emulsion. Therefore, the adsorbed layers on the captive droplets formed by adsorption of surfactant from the continuous phase of the emulsion. The results are shown in Figure 23, where surface diffusion data of FITC-/3-1g in o/w and a/w thin films as a function of added Tween 20 are summarised.

52

Figure 23. The lateral diffusion coefficient of adsorbed FITC-/3-1g in thin films as a function of added Tween 20. (9 o/w thin films formed from aqueous non-homogenized solutions of /3-1g at 3 mg/ml; (s), o/w thin films formed from 10% v/v n-tetradecane emulsion or emulsion subnatant samples of FITC-/3-1g, initial protein concentration 3 mg/ml; (.), a/w thin films formed from aqueous non-homogenized solutions of 13-1g at 0.2 mg/ml. The diffusion behavior of the protein in the o/w thin films formed from emulsified samples was completely different from that observed from non-homogenized samples used to form o/w or a/w thin films, since the onset of diffusion occurred at R = 4. This was in closer agreement with previous reports [46,47] since some displacement of/3-1g by Tween 20 has been reported to occur at R = 4 but not R = 0.1. However, again the transition point at R = 4 does not comply with our destabilizati0n model shown in Figure 16. It is now evident that at least part of the shift to R = 4 is explained by a shear-induced conformational change in/3-1g. This complicates matters further by introducing a second class of free protein into the solution with altered Tween 20 binding capacity. Nevertheless, the results are beginning to converge towards equivalent solution compositions being required to induce surface diffusion in both o/w and a/w systems.

6. THE RELATIONSHIP OF FRAP MEASUREMENTS OF SURFACE DIFFUSION AND OTHER SURFACE R H E O L O G I C A L MEASUREMENTS The FRAP data described above report molecular self diffusion in the adsorbed layers at the interfaces of thin films. The measurements are sensitive to the strength of interactions

53 between the molecules at the interface. Once the strong protein-protein interactions have been weakened or destroyed by competitive adsorption of low molecular weight surfactant, surface diffusion ensues. The magnitude of the measured diffusion coefficient of adsorbed protein will be dependent upon the density of molecular packing at the interface, since this will influence the surface viscosity.

Figure 24. A comparison of the data obtained from a range ot surface rheological measurements of samples of/3-1g as a function of Tween 20 concentration. (R), The surface diffusion coefficient of FITC-/3-1g (0.2 mg/ml) at the interfaces of a/w thin films; (X), the surface shear viscosity of/3-1g (0.01 mg/ml) at the o/w interface after 5 hours adsorption; (o), the surface dilational elasticity and (o) the dilational loss modulus of/3-1g (0.2 mg/ml). It was of interest to compare the results obtained with the FRAP technique with those obtained with classical surface rheological techniques. Our detailed knowledge of properties of solutions of/3-1g containing Tween 20 made this an ideal system on which to compare the methods. Firstly, surface shear viscosity measurements were performed on the Tween 20//3-1g system [47] using a Couette-type torsion-wire surface rheometer as described previously [3,48]. All the experiments were carried out at a macroscopic n-tetradecane-water interface at a fixed protein concentration of 0.01mg/ml. In the absence of Tween 20, the surface shear

54 viscosity of the adsorbed interfacial film of/3-1g rapidly increased to over 500 mN.m 1 during the first hour of adsorption, followed by a more gradual increase to 720 mN.m -1 after 20 hours. Samples containing Tween 20 in the concentration range between R = 0 to 1, showed a time dependent increase in surface shear viscosity, but values obtained at a given time were always significantly lower than that observed with/3-1g alone. The surface shear viscosity data obtained with samples of/3-1g containing Tween 20, 5 hours after formation of the o/w interface are shown in Figure 24. The data show that increasing the concentration of Tween 20 caused a progressive drop in the observed surface shear viscosity. At R = 1, the surface shear viscosity was too low to measure accurately, without using a much finer torsion wire. There is a clear complementarity between the surface shear viscosity and FRAP measurements; the former is sensitive when the surface viscosity is high and molecular diffusion is zero due to protein-protein interactions, the latter is sensitive when the surface viscosity is very low due to the abolition of interactions. The surface rheological properties of the/3-1g/Tween 20 system at the macroscopic a/w interface were examined by a third method, namely surface dilation [40]. Sample data obtained are presented in Figure 24. The surface dilational modulus, (E) of a liquid is the ratio between the small change in surface tension (AT) and the small change in surface area (AlnA). The surface dilational modulus is a complex quantity. The real part of the modulus is the storage modulus, e' (often referred to as the surface dilational elasticity ,Ea). The imaginary part is the loss modulus, E", which is related to the product of the surface dilational viscosity and the radial frequency (~Tao~). Experiments with the/3-1g/Tween 20 system were performed at a macroscopic a/w interface at a/3-1g concentration of 0.2 mg/ml [40]. The data obtained relate to the properties of the interface 20 minutes after formation. Up to R = 1, the storage modulus (dilational elasticity) was large and relatively constant, whereas the loss modulus (dilational viscosity) increased with increasing R. As R was increased to higher values there was a marked decrease in the storage modulus (dilational elasticity) and a gradual increase in the loss modulus (dilational viscosity). In summary, the data show the presence of a transition in surface dilational behavior in this system at a solution composition of approximately R = 1. At this point, there is a transformation in the adsorbed layer properties from elastic to viscous. The results of these studies show that all three surface rheological methods give results that correlate with each other. In addition, the results add further evidence in support of our model for Tween 20-induced changes in adsorbed layers of/3-1g (Figure 16). The three surface rheological techniques described here are very complementary, each providing different but related data and providing sensitivity across different ranges of interfacial layer stiffness. However, it is important to note that only the FRAP method can be applied to both macroscopic and thin film samples.

7.

FUTURE PROSPECTS

The progress made using the methods described in this report has opened up a number of opportunities. There are consumer and health pressures to reduce the consumption of synthetic emulsifiers used in processed foods. Therefore, a need exists to identify alternative 'natural' replacement emulsifiers. One approach is to develop 'natural', biodegradable emulsifiers through biosynthetic routes using enzymes. Alternatively, more widespread use of proteins as emulsifiers would be an option if their functional properties were more

55 predictable. We have identified a number of mechanisms of destabilization of proteinstabilized foams involving competitive adsorption of surfactant and the breakdown of proteinprotein interactions in the adsorbed layer. The knowledge that this approach allows development of scientific strategies for controlling destabilization in protein-stabilized systems. Currently, we are examining two different approaches, The first involves inclusion or exploitation of existing natural ingredients in the food system of interest, which are capable of enhancing interactions in the adsorbed layer by crosslinking. It has proved possible to test candidate molecules for this role using one of our characterized systems (e.g., /3-1g/Tween 20). Preliminary studies with this system have identified that the beer foam stabilizing activity of the iso-a-acid fraction in hop extract operates through a protein crosslinking mechanism [43]. Our knowledge of the mechanisms of destabilization in foams allows strategic targeting of a number of other natural compounds (e.g., divalent ions, bifunctional acids e.g., tartaric acid, phenolics and mixtures of polysaccharides or proteins). The second approach is most effective when the destabilizing agent is only present at low concentrations (e.g., egg yolk lipid in meringue). Here, removal of the destabilizing component by selective binding has potential. Preliminary work has demonstrated the effectiveness of exploiting the lipid binding properties of the protein, puroindoline from wheat flour [15], for selective removal of destabilizing components (e.g., lipid) from model and real beverage systems [44]. However, the effectiveness of this protein is not explained by binding alone and needs further study [15]. Our understanding of the influence of competitive adsorption on emulsion stability is less secure. Recent work has identified several marked differences between the adsorbed layer properties at air/water and oil/water interfaces (e.g., multilayer versus monolayer formation). Advancing our knowledge of the stabilization of emulsions by protein merits further investigation, since emulsions comprise a major sector of processed foods. If competitive adsorption of surfactants influences the stability of protein emulsions in a similar manner to foams, use of the strategies outlined above may be appropriate for controlling destabilization. If we are successful, food processors specialising in the preparation of food dispersions (e.g., foaming and sparkling beverages, salad dressings, sauces, ice cream etc) will benefit from the results of this research. The work provides the underpinning knowledge that will allow food ingredient manufacturers to supply 'natural' emulsifier proteins and functionality enhancing components to meet the legislative demands for food ingredients in the future, whilst satisfying consumer demands for the elimination or reduction of use of synthetic additives in foods.

ACKNOWLEDGEMENTS The author would like to acknowledge the involvement of Alan Mackie, Peter Purdy and Dr. Andrew Pinder in the design, construction and continued development of the FRAP apparatus. The experimental results described in this paper were obtained in collaboration with Mark Coke, Peter Wilde and David Wilson. The author wishes to thank AM and PW for discussions relating to the text and PW for his assistance in preparing the figures. This work was funded by the AFRC.

56

zyxwvutsrq

REFERENCES

.

3. .

5. .

7.

.

10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22.

23. 24. 25. 26. 27. 28.

P. Walstra, in 'Foams: Physics, Chemistry and Structure' A.J. Wilson (ed.), Springer Series in Applied Biology, Springer-Verlag, London, 1989, p. 1. W.E. Ewers and K.L. Sutherland., Aust. J. Sci. Res. Ser.A., 5 (1952) 697. J. Castle, E. Dickinson, B.S. Murray and G. Stainsby, ACS Symp. Ser., 343 (1987) 118. D.E. Graham and M.C. Phillips, J. Colloid Interf. Sci., 70 (1979) 403. K. Mysels, K. Shinoda and S. Frankel in 'Soap Films' Pergamon Press, New York, 1959. A. Sheludko, Adv. Colloid Interf. Sci., 1 (1967) 391. D.C. Clark, A.R. Mackie, L.J. Smith and D.R. Wilson in 'Food Colloids', R.D. Bee, P. Richmond and J. Mingins (eds.), Royal Society Special Publication No.75, Cambridge, 1989, p. 97. D.C. Clark, R. Dann, A.R. Mackie, J. Mingins, A.C. Pinder, P.W. Purdy, E.J. Russell, L.J. Smith and D.R. Wilson, J. Coll. Interf. Sci., 138 (1990) 195. D.C. Clark and P.J. Wilde in 'Gums and Stabilisers for the Food Industry - 6', Oxford Press, Oxford, 1992, p. 343. P.J. Wilde and D.C. Clark, J. Coll. Interf. Sci., 155 (1993) 48. C.J. Brock and M. Enser, J. Sci. Food Agric., 40 (1987) 263. D.C. Clark, M. Coke, A.R. Mackie, A.C. Pinder and D.R. Wilson, J. Coll. Interf. Sci., 138 (1990) 207. M. Coke, P.J. Wilde, E.J. Russell and D.C. Clark, J. Coll. Interf. Sci., 138 (1990) 489. D.C. Clark, P.J. Wilde, D.R. Wilson and R. Wustneck, Food Hydrocolloids, 6 (1992) 173. P.J. Wilde, D.C. Clark and D. Marion, J. Agric. Food Chem., in press. Z. Lalchev, K. Kristov and D. Exerowa, Coll. Polym. Sci., 257 (1979) 1248. D.N. Platikanov, G.P. Yampol'skaya, N.I. Rangelova, Zh.K. Angarska, L.E. Bobrova and V.N. Izmailova, Kolloidn Zh., 42 (1981) 893. P.R. Musselwhite and J.A. Kitchener, J. Coll. Interf. Sci., 24 (1967) 80. D.N. Platikanov, G.P. Yampol'skaya, N.I. Rangelova, Zh.K. Angarska, L.E. Bobrova and V.N. Izmailova, Kolloidn Zh., 43 (1981) 177. D.C. Clark, Ferment, 4 (1991) 370. D.C. Clark, P.J. Wilde and D.R. Wilson, Coll. Surf., 59 (1991) 209. D.C. Clark, M. Coke, P.J. Wilde and D.R. Wilson in 'Food Polymers, Gels and Colloids', E. Dickinson (ed.), Royal Society Special Publication No.82, Cambridge, 1990, p. 272. D. Axelrod, D.E. Koppel, J. Schlessinger, E. Elson and W.W. Webb, Biophys. J., 16 (1976) 1055. R. Peters, Naturwissenschaften, 70 (1983) 294. R. Peters and M. Scholtz in 'New Techniques of Optical Microscopy and Microspectroscopy', R.J. Cherry (ed.), Macmillan, London, 1990, p. 199. K. Beck and R. Peters, in 'Spectroscopy and Dynamics of Molecular Biological Systems' P.M. Bayley and R.E. Dale (eds.), Academic Press, London, 1985, p. 177. D.E. Wolf, in 'Fluorescence Microscopy of Living Cells in Culture. Part B.,' D.L. Lansing Taylor and Y. Wang (eds.), Academic Press, San Diego, 1989, p. 271. P. Garland, Biophys. J., 33 (1981) 481.

57 29. D. Axelrod, in 'Spectroscopy and Dynamics of Molecular Biological Systems', P.M. Bayley and R.E. Dale (eds.), Academic Press, London, 1985, p. 163. 30. J. Yguerabide, J. Aschmidt and E.E. Yguerabide, Biophys. J., 39 (1982) 69. 31. R.P. Haugland, in 'Handbook of Fluorescence Probes and Research Chemicals', Molecular Probes Inc. 1992. 32. G.E. Means and R.E. Feeney in 'Chemical modification of Proteins' Holden Day, San Francisco, 1971. 33. I.S.K. Craig, P.J. Wilde and D.C. Clark, Coll. Surf. B:Biointerfaces, in press. 34. H.F. Swaisgood, in Developments in 'Dairy Chemistry - 1', P.F.Fox (ed.), Applied Science Publishers, London, 1982, p. 1. 35. I. Weil, J. Phys. Chem., 70 (1966) 133. 36. G. Barisas and M.L. Leuther, Biophys. Chem., 10 (1979) 221. 37. O.D. Velev, A.D. Nikolov, N.D. Denkov, G. Doxastakis, V. Kiosseoglu and G. Stanlidis, Food Hydrocolloids, 7 (1993) 55. 38. T.P. Burghardt and D. Axelrod, Biophys. J., 33 (1981)455. 39. D.C. Clark, Encyclopaedia of Food Science, Food Technology and Nutrition, Academic Press, 1993, p. 1577. 40. D.C. Clark, P.J. Wilde, D. Bergink-Martens, A. Kokelaar and A. Prins in 'Food Colloids and Polymers: Structure and Dynamics', E. Dickinson and P. Walstra (eds.), Royal Society of Chemistry Special Publication No. 113, Cambridge, 1993, p. 354. 41. D.C. Clark, A.R. Mackie, P.J. Wilde and D.R. Wilson, in 'Food Proteins Structure and Functionality', K.D. Schwenke and R. Mothes (eds.), 1993, p. 263. 42. D.R. Wilson, P.J. Wilde and D.C. Clark in 'Food Colloids and Polymers: Structure and Dynamics', E. Dickinson and P. Walstra (eds.), Royal Society of Chemistry Special Publication No. 113, Cambridge, 1993, p. 415. 43. D.C. Clark, P.J. Wilde and D.R. Wilson, J. Inst. Brew., 97 (1991) 169. 44. D.C. Clark, P.J. Wilde and D. Marion, J. Inst. Brew., in press 1993. 45. A.R. Mackie, P.J. Wilde, D.R. Wilson and D.C. Clark, Royal Chem. Soc. Faraday Trans. 89 (1993) 2755. 46. J-L. Courthaudon, E. Dickinson, Y. Matsumura and A. Williams, Food Struct., 10 (1991) 109. 47. J-L. Courthaudon, E. Dickinson, Y. Matsumura and D.C. Clark, Coll. Surf., 56 (1991) 293. 48. E. Dickinson, B.S. Murray and G. Stainsby, J. Colloid Interf. Sci., 106 (1985) 259.

This Page Intentionally Left Blank

Characterization of Food: Emerging Methods A.G. Gaonkar (Editor) 9 1995 Elsevier Science B.V. All rights reserved.

59

Chapter 3 M e t h o d s for characterization of structure in whippable d a i r y - b a s e d emulsions Niels M. Barfod Grindsted Products 38, E d w i n Rahrsvej DK-8220 Brabrand Denmark

1.

INTRODUCTION

This chapter describes some methods to study physical characteristics and ingredient interactions in whippable dairy-based emulsions. The story of whippable emulsions begins with natural dairy cream. From this starting point a range of dairy-type whippable emulsions has been developed over the years. In unhomogenized dairy cream the natural phospholipids contribute to the whipping properties of the cream. However, after homogenization the particle size of the fat globules decreases, and the total fat surface area increases. This means that the interracial concentration of polar lipids decreases because milk serum proteins adsorb at the newly formed interfaces, and the whipping properties are lost. Consequently, additional polar lipids or emulsifiers are needed to obtain good whipping properties in most industrially manufactured products. This chapter will deal with the following types of whippable emulsion: 9 9 9 9

Whipped topping Natural and imitation whipping cream Ice cream Aerated desserts

The formulations of these products vary greatly, and therefore only principal and important aspects of product stability and functional properties will be discussed. Most studies have dealt with ice cream, because commercially this product is the most important whippable emulsion. Thus, methods for characterization of ice cream will be highlighted more than other types of whippable emulsions. There is a fundamental problem which must be solved when dealing with whippable emulsions. Before use the emulsion must be sufficiently stable. On the other hand, it must b e p o s s i b l e to destabilize the emulsion by mechanical treatment combined with air incorporation (whipping, air pressure, cooling, freezing). The partly destabilized fat globules in the whipped emulsion are important for the stability of the foam structure. There is a

zy

60 delicate balance between emulsion stability and instability. If the emulsion is too stable, it will not whip, if it is not stable enough, the foam formed will collapse after a short whipping time. The ingredient composition and manufacturing process are important for the different types of whippable emulsions. In many industrially produced whippable emulsions, functional ingredients, such as food emulsifiers and hydrocolloids are used to improve functionality and product stability.

zyxwvutsrq

Product description of whippable emulsions Toppings are spray-dried emulsions made from sodium caseinate, vegetable fat such as palm kernel or coconut fat, and emulsifiers with low polarity, such as acylated (acetylated or lactylated) monoglycerides or propylene glycerol monostearate. Whipping cream (30-40% butterfat) can be made from natural cream, but its whipping properties may be improved by changing the manufacturing process or by using additional milk protein fractions and/or food emulsifiers. Whipping cream with reduced fat content (25 % or less tat) can be made by incorporating food emulsifiers and hydrocolloids ~. Imitation whipping creams are made from skimmed milk powder or sodium caseinate, vegetable fats, and emulsifiers as described above for toppings. More polar emulsifiers in low dosage may be incorporated to ensure storage stability. Sodium alginate may be used to prevent syneresis of the foam after whipping 2. In general, the emulsifier dosage in imitation whipping cream is 10 tilnes less than that in toppings. Ice cream is made from skimmed milk, condensed skimmed milk or skimmed milk powder in combination, and dairy cream, butter or butter oil. In some countries vegetable fat is used to replace dairy fat. Usually, monoglycerides or mono-diglycerides are used, but other more polar emulsifiers can also be used. The emulsifier dosage is similar to that used in imitation cream. Ice cream also contains sugar and hydrocolloids, which mainly influence the freezing behaviour of the ice cream mix. Aerated desserts are products with a stabilized foam structure based on dairy ingredients or dairy analogues. They may be based on neutral, acidified (yogurt-type) or concentrated milk, and are typically low in tat ( < 10%) and high in sugar (8 to 15 %). The foam structure may be stabilized by selected emulsifiers/hydrocolloids for different products and different manufacturing processes 2.3. The main difference between aerated dessert products and other whippable emulsions is the gelation of the continuous water phase. The most common hydrocolloids used for this purpose are gelatine, alginate and carrageenan. Aerated desserts may be whipped in continuous aerators (cold-stored products) or in ice cream freezers (frozen products). 1.1.

General mechanisms for stabilizaiion of whippable emulsions The subject of this chapter is whippable emulsions, and some background theory on foams may be appropriate. To produce a foam, stable or metastable, it is necessary for surface-active molecules 1.2.

61 such as emulsifiers or proteins to be adsorbed at the gas-liquid interface of the air bubbles to build a stabilizing film. The hydrophobic residues of the molecules will be attracted towards the air phase and the hydrophilic residues will be attracted towards the water phase. The main factor determining the stability of such foams is the rate and extent of drainage from the thin liquid film. In general, this type of foam is relatively unstable. The stability may be enhanced by increasing the viscosity of the liquid by increasing the dry matter content or adding certain hydrocolloids. The foam stability may also be enhanced with hydrocolloids, in particular microcrystalline cellulose. In addition to surface-active molecules, the foam of whippable emulsions contains particles in the form of tat globules trapped in the continuous phase. During whipping, fat globules penetrate and partially replace the protein fihn at the air-water interface 4. The foam stability is affected by the degree of aggregation of fat globules in the vicinity of the air-water interface. The tat composition, and in particular its crystallization behaviour, exerts a dominant influence on the quality of whippable emulsions. Adsorption of fat globules to the air bubbles depends on the hydrophobicity of the fat globules. The hydrophobicity depends on the amount of protein bound on the surface of the fat globules. In general, proteins act as emulsion stabilizers whereas certain food emulsifiers induce controlled emulsion destabilization during whipping. Later in this chapter it will be shown how emulsifiers, such as polar lipids, control protein binding to the surface of fat globules and thus the aggregation and whipping properties of these products. Hydrocolloids may be used to increase viscosity and inhibit syneresis of the foam and gel the water phase in whippable emulsions. In frozen systems such as ice cream, hydrocolloids have the additional effect of inhibiting the growth of ice crystals thus enhancing foam stability and improving texture.

zyxw

1.3.

Methods for characterization of whippable emulsions In the food industry a range of practical or descriptive tests are used to evaluate product quality and the stability of whippable emulsions. Using such methods a number of reliable and commercially valuable whippable emulsions have been developed over the years. To develop new whippable emulsion systems which are more difficult to stabilize, i.e. primarily low-fat products, lnore advanced physical methods have been used to elucidate the fundamental mechanisms behind the behaviour of whippable emulsions. In this chapter the physical methods for analyzing whippable emulsions are divided into analyses of 1) the emulsified fat phase 2) the fat-water interface and air-water interface, and 3) the continuous water phase. The descriptive tests are mentioned at the end of the chapter as it is easier to explain the meaning of these tests after the fundamental mechanisms have been described.

2.

EMULSIFIED FAT PHASE

As already mentioned above, the flmctional properties of whippable emulsions depend largely on the properties of the tat globules they contain. The fat globules form the skeleton of the foam. The crystallization behaviour inside the fat globules of whippable emulsions is decisive for the stabilization of the foam structure after aeration. It is a well-known fact in the food industry that whippable emulsions made with liquid fats are totally devoid of functionality.

62 The quality of the fat crystallization in whippable emulsions is important, e.g. crystallization rate, and shape, form and size of the fat crystals formed. In some creams, needle-shaped crystals at the oil-water interface have been shown to be associated with partial coalescence resulting in defects in whipping properties s,6. The fat phase in many oil-water emulsions is in a supercooled state, since nucleation followed by crystal growth is greatly reduced if fat is present in a large number of isolated droplets with small particle size7. Nucleation may be enhanced with emulsifiers present in the fat phase by increasing the number of nucleation centres, resulting in the formation of many small fat crystals. This will result in improved functional properties. The various aspects of the importance of fat crystallization with regard to the functional properties are beyond the scope of this chapter, but methods to analyze this important phenomenon in whippable emulsions are described below. 2.1.

Thermal analyses Differential Thermal Analysis (DTA) and Differential Scanning Calorimetry (DSC) are techniques used to measure the heat changes which occur in a small sample (1 to 30 mg) subjected to heating or cooling at a known linear rate (typically 1 to 30~ per minute) 8. One example of this type of analysis is described. Ice cream emulsions (mixes) are normally subjected to a cooling period of several hours at 0 to 5~ before freezing and whipping. During this treatment several physical changes take place 9. These changes are described later in this chapter. One change is the crystallization of tat globules in the mix, which can be followed by DSC analysis as shown in Figure 1.

Figure 1 Melting enthalpy of bulk fat and emt, lsified fat of ice cream mix with (+E) and without (-E) emulsifier alter cooling at 5~ measured by DSC.

The melting of crystalline tat took place between 15~ and 40~ and was analyzed at a heating rate of 10~ ~~ In non-emulsified state fat crystallizes very quickly. However, in emulsified state a reduced crystallization rate is observed. In the presence of emulsifier (saturated mono-diglyceride) the crystallization rate is enhanced, and the supercooling is reduced. The melting enthalpy in Figure 1 is expressed per gram of fat in the sample analyzed.

zyxwvutsr 63

Solid Fat Content (SFC) by Low Resolution NMR (wide-line or pulse) Analysis of solid tat content by NMR has replaced older techniques such as dilatometry ll. The material may be studied in an equilibrium without melting. The analytical time is less than l0 seconds using NMR in contrast to more than 15 minutes using DSC, and the amount of sample material is about 100 times higher when using NMR than when using DSC. This is important if the sample material is not completely homogenous. One drawback with NMR is that the liquid signal from water in o/w emulsions has to be subtracted to obtain the true SFC. This can be done by analyzing emulsions and fat blends with no tendency to supercooling and making calibration curves. Another possibility is to measure reference samples without fat and calculate the true SFC by subtracting the signal from the water and water-soluble components 9. 2.2.

Figure 2 shows the same experiment as in Figure 1, but analyzed by NMR. The same information is obtained, but the NMR method is easier and quicker to use.

Figure 2 Fat crystallization of bulk fat and emulsified fat of ice cream mix with (+E) and without (-E) elnulsifiers after cooling to 5~ measured by pNMR.

The NMR method may also be used to study supercooling phenomena in spray-dried topping emulsions 12. Figure 3 shows that the %solids content is lower in topping powders than in the corresponding simple dry mixtures. With effective emulsifiers (propylene glycol monostearate (PGMS)) supercooling is slightly reduced, and with ineffective emulsifiers (glycerol monostearate (GMS)) hardly any supercooling takes place. Supercooling increases with increased protein (sodium caseinate) content due to both reduction in fat particle size and

64 to increased lipid-protein interaction. Sodium caseinate with 3 % peptide bonds hydrolysed results in hardly any supercooling, but a similar fat particle size as intact sodium caseinate. Other globular food proteins tested have been found to be less effective than sodium caseinate. The lipid-protein interaction is specific to high lauric fats such as hardened coconut oil or palm kernel oil, and is not evident with other fats such as partially hydrogenated soybean oil, fish oil or normal butterfat ~3.

Figure 3 Per cent solids of topping powders and corresponding dry mixtures measured by pNMR at temperatures from 5~ to 25~ Reprinted from reference 12, courtesy of the American Oil Chemists' Society.

Crystallization of supercooled fat in topping powders may be studied by NMR afterreconstitution in heavy water. Below room temperature spontaneous fat crystallization takes place under isothermal conditions in the presence of effective emulsifier (PGMS) but not with ineffective emulsifiers or without emulsifiers (Figure 4).

65

zy

Figure 4 Crystallization of supercooled fat at 15~ measured by pNMR in the absence or presence of emulsifiers (PGMS or GMS).

The time scale of tat crystallization is much shorter for topping powders than for ice cream mix as presented in Figure 2. This is due to the much higher emulsifier content in topping powder. The induction of tat crystallization in whippable emulsion systems is due to interfacial protein desorption from the tat globules of the emulsion mediated by the emulsifiers. This phenomenon is described in section 3.1. Other methods to study fat crystallization in whippable emulsions may be used, e.g., a recently developed technique using ultrasonic velocity ~4. 2.3.

X-ray diffraction This technique is particularly suited for studying polymorphism of fats and ordered structures of emulsifier in water, e.g., liquid crystals or 'gel' phases. The concentration of emulsifiers in food emulsions is often too low to allow the formation of multi-layered liquid crystals at oil-water interfaces ~s. In systems in which the emulsifier concentration is sufficiently high, such as toppings, the formation of 'gel' phases appears to play a role. Studies of topping tat phases by X-ray diffraction analysis show that the triglycerides from hydrogenated coconut oil do not co-crystallize with the PGMS emulsifier added 16. The coconut fat crystallizes in a beta-prime form with long spacings of 36 A, whereas the emulsifier crystallizes in alpha-form with long spacings of 49 A. After contact with water at 5~ the long spacings of emulsifier in the tat phase increases from 49 A in the bulk phase to 56 A in the interfacial phase. This increase of 7 A can only be due to the penetration of water into the polar regions of the emulsifier caused by the so-called hydration force ~7. Water absorption into the tat phase of the topping results in interfacial protein

66 desorption, and with regard to crystallization results in a more stable foam structure. For further details see reference 16. A model of the emulsifer-water gel structure formed near the oil-water interface of the emulsion is shown in Figure 5. Similar results on toppings have been presented by Westerbeek and Prins l~.

Figure 5 A schematic model of the formation of lipid gel phase by hydration of the polar groups in crystalline regions of emulsifiers, d = interplanar Bragg spacing; d~ = thickness of lipid bilayer; dw = thickness of water layer. Redrawn from reference 15, courtesy of Marcel Dekker Inc.

In a standard topping formulation 10 to 20% of the emulsifier in the fat phase is used to produce the desired foam stability and overrun after whipping. This is due to protein desorption and tat crystallization during whipping in cold water. Practical tests have shown that low-fat topping powders (down to 80% fat reduction) may be produced by a concomitant increase in emulsifier concentration (up to 50%) in the fat phase 19. A higher concentration of emulsifier tacilitates enhanced formation of alpha crystalline gel structure, which is obviously important for the whipping properties and foam texture of low-tat topping products. 2.4.

Electron microscopy (EM) Various electron microscopy techniques have been used to study the structures of whippable emulsions such as normal and cryo-scanning electron microscopy or transmission electron microscopy using various preparation methods such as freeze fracturing, freeze etching, etc. The literature is quite extensive, and only a few important papers will be discussed in this chapter. EM studies of whipped cream show that the air bubbles are completely surrounded by a layer of tat globules which protrude partially into the air bubbles. These parts of the fat globules no longer have their original membrane layer, but exhibit surface layers of crystallized tats. The fat globules adsorbed around the air bubbles are bonded together with

67 coalescent tat. The cross-linking of fat globules adsorbed to the adjacent air bubbles by chains of coalescent globules establishes a stabilizing infrastructure in the foam 4'2~ Although the lipid phase in powdered toppings is finely dispersed in the form of small globules (< 1 /zm), strong destabilization takes place after reconstitution in cold water, resulting in platelet-like crystal agglomerates (Figures 6 and 7).

Figure 6 a. Structure of topping powder in the dry state with close-packed globular fat particles (f) (diameter less than 0.5 txm). b. Crystallization and transformation of the globular structure of fat particles into thin layers of crystal platelets (c). Reprinted from reference 21, courtesy of Verlag Th. Mann.

During whipping these crystalline lipid platelets accumulate at the air-water interface of the air bubbles and also form bridges between them 22'23. The structure of whipped topping is thus completely different from that of whipped dairy or liquid imitation creams. In the latter systems the air bubbles appear to be covered in a monolayer of fat globules, which are rarely deformed and which protrude with a substantial part of their volume into the air phase of the bubbles. If large fat crystals are present, they are considered detrimental to foam stability, in contrast to whipped toppings 6 (Figure 7).

68

Figure 7 Left: Surface of air bubbles (a) in whipped topping emulsion is completely covered with a thick layer of plate-shaped tat crystals (c). w = water phase. Right: Surface of air bubbles (a) of whipped imitation cream is covered with a monolayer of only slightly destabilized globules (f). Reprinted from reference 22, courtesy of Scanning Microscopy International.

The flocculated fat globules of whipped cream contain fewer contact points, and the foam is therefore not as stiff as in toppings in which the aggregated crystal platelets have a large surface area, many contact points and thus increased stiffness. This means that an acceptable topping foam may be tbrmed at a much lower tat content than is the case with liquid whipping creams. Ice cream is a partially frozen fat-stabilized foam ~. The interaction of the fat globules (0.5 to 1.0/~m) of the ice cream mix and the air bubbles in the finished product depends to a great extent on the presence of emulsifiers such as mono-diglycerides. In the absence of emulsifiers, micellar casein adsorbs strongly to the dispersed fat phase, thus preventing adsorption of tat globules to the air bubbles. In the presence of emulsifiers, interfacial protein layers are desorbed during cold storage (ageing), and during whipping and freezing in the ice cream machine, thereby facilitating the binding of partially crystalline fat globules to air 23. This air bubble stabilization is important for the physical properties of this product (Figure 8).

69

zyxwvut

Figure 8 Air bubbles in ice cream (a). Interface (arrows) between a large air bubble (A) and water phase (W) in an ice cream sample without emulsifier. There is very little adsorption of fat globules to the air-water interface, which is stabilized by a thin protein film only. (b) Corresponding structures in an ice cream with emulsifier (saturated lnono-diglycerides). Fat globules interact strongly with the air-water interface. Reprinted from reference 23, p 242, courtesy of Marcel Dekker Inc.

2.5.

Particle size distribution The characterization of emulsions by particle size distribution analysis has been facilitated in recent years by a range of new instruments. Most of these instruments employ laser light diffraction principles, and have replaced older spectrophotometric methods. To obtain optimal functional properties, the range of average fat particle size of most dairy type whippable emulsions is from about 0.5/xm to 1.0/xm. If the average fat particle size increases, the emulsion will not be stable in the long term and the whipping properties will not be as good. This is most critical for systems which are to be stored for a long time before whipping, e.g., UHT-treated imitation cream. Particle size analysis may also be employed to detect the degree of fat globule agglomeration. In liquid imitation whipping crealn weak agglomeration of fat globules before whipping is beneficial for the whipping properties. The degree of tat globule agglomeration,

70 as well as the size and mechanical stability of the aggregates formed after whipping the emulsions, may be estimated by particle size distribution analysis as shown in Figure 9. The Figure shows analysis of frozen ice cream melted at 0~ to 5~ using a Malvern 2600 Particle Sizer.

Figure 9 Particle size distribution analysis of air bubbles stabilized with aggregated fat globules in ice cream atier thawing and dilution 100 x with water at 0~ to 5~ Effect of emulsifier concentration (MDG = saturated mono-diglyceride) on air bubble size and stability. Upper row: Samples analyzed without degassing. Lower row: Samples analyzed after degassing for 5 minutes.

No stable aggregates form in the absence of emulsifiers. The agglomeration of fat globules can be observed in tile presence of emulsifiers (0.3 and 1.5 % mono-diglycerides). The stability of air bubbles with aggregated tat globules may be tested by evacuating the samples, which will expand all unstable air bubbles and induce a breakdown. If the amount of emulsifier is overdosed (1.5%), large aggregates forln and the air bubble stability is reduced, resulting in a reduction of aggregate size after degassing due to the collapse of the

71 air bubble structure. When the dosage of emulsifiers is optimal (0.3 %), smaller, more stable air bubbles form. These bubbles do not collapse after degassing. This type of analysis requires careful sample preparation and experience to get reproducible results. It is only possible to analyze relatively stable air bubbles with this technique. Large unstable air bubbles may break down during dilution before analysis. Particle size and particle aggregate size distribution is now being used for monitoring product stability and functional properties in a range of food emulsion systems 24. A new foam analyzer has recently been developed in Holland 25. The foam is illuminated by continuous light from an opto-electronic unit, and the light reflection is measured by an optical glass fibre probe, which is moved down through the foam at a known speed. More light is reflected when the probe tip is in a gas than when it is in a liquid. The reflected light is converted into an electronic analogue signal from which the bubble size distribution in the foam is calculated by a computer 25. The advantage of this method is that samples can be studied without dilution, and it is quicker than electron microscopy methods. It is believed that the method will provide valuable information on foams in the fixture. Using this method also makes it possible to measure the rate of foam drainage and collapse, as well as the gas fraction in the foam. 2.6.

Light microscopy This technique does not provide information as detailed as the above-mentioned methods, but may be used as a rough quality check of whippable emulsions. The method is suitable for detection of the presence of large tat globules in the emulsion. Fat globule agglomeration may be distinguished from tat coalescence by using a combination of phase contrast and polarized light illumination. The detection of small fat globules with quick Brownian motions may be made easier by diluting the sample with a polar solvent with a higher viscosity than water, e.g., glycerol 26.

zyxwvu

2.7.

Free Fat Esthnate (FFE) FFE is an extraction method using heptane which measures the churning out of fat during emulsion stabilization 9. High protein and low fat content reduce destabilization, whereas the presence of emulsifiers, cold treatment at 5~ and mechanical treatment (whipping, possibly combined with freezing) increases destabilization. The FFE method is of great practical use to verify the level of mechanical treatment applied to ice cream mix during aeration and freezing. It also provides an indication of the storage stability and creaminess of the product tested 27. The total fat content in whippable emulsions may be estimated by the Gerber method 28 or by the gravimetric method 29.

3.

INTERFACIAL EFFECTS

The interfacially bound protein layer on fat globules is influenced greatly by the emulsifier and hydrocolloid content as well as by processing conditions. During homogenization of whippable emulsions at high temperatures, emulsification is facilitated by emulsifiers, whereas protein binding to the fat globules acts as an emulsion

72 stabilizing mechanism 15. However, the effect of emulsifiers on the long-term properties of emulsions is far more important than their influence on particle size distribution during homogenization. In general, protein-fat binding is weakened in emulsions containing emulsifiers 3"'31. The effect is temperature-dependent and increases at low temperatures (5~ to 10~ 3-~. In whippable emulsions, such as ice cream mix, toppings and homogenized creams, weakening the protein-tat binding by emulsifiers results in an improvement of the whipping properties 9,12,33,34.

zyxwvuts

3.1.

Protein-fat binding The amount of protein bound to fat globules is usually estimated by high-speed centriti~gation followed by quantitative protein analysis (e.g. Kjeldahl method) of the isolated cream layer or fat-free water phase 9"35,36. This phenomenon may be studied in greater detail by fast protein liquid chromatography 37, or by confocal scanning laser microscopy 38. It is recommended that the temperature in these types of studies be strictly controlled, as protein-tat binding is highly dependent on temperature. The effect of temperature and whipping on three whippable emulsion systems is shown in Table 1. For further details and results, see references 9'12'13'16. Table 1 Protein-fat binding in three whippable emulsion systems % Fat-adsorbed protein

System

Control 25~ 5~

Foam

With Emulsifiers 25~ 5~ Foam

Imitation cream 1~ Ice cream Topping

83 38 42

69 162~ 22

80 30 34

1) 2)

80 21 24

38 12 1

4 42) 0

The amount of adsorbed protein is initially high due to the high fat content (approx. 30%) in this system Analyzed after defrosting the frozen foam at 0~

Low temperatures, whipping, and the presence of emulsifiers all increase protein desorption. Protein desorption in toppings takes place very quickly after reconstitution in cold water, due to the high emulsifier content, but in liquid systems such as ice cream mix the process is much slower, and takes many hours. This is the primary reason why a long ageing period at 5~ in ice cream production is required Q. Protein desorption in ice cream mix during ageing at 5~ with and without saturated mono-diglycerides is shown in Figure 10. In the presence of emulsifiers protein desorption is accelerated.

73

Figure l0 Protein desorption from the fat globules into the water phase during ageing of ice cream lnix with (+E) and without (-E) emulsifier (saturated mono-diglyceride).

Figure 11 Protein binding to fat globules in ice cream mix at various temperatures and after ice cream production (I.C.). The latter analyzed at 5~ after thawing ice cream at 0~ Effect of hydrocolloid blend and emulsifier.

Milk protein desorption at low telnperature is due to stronger hydrophilic and weaker hydrophobic forces, and is caused mainly by dissociation of beta-casein 39. Hydrocolloids are used in ice cream to increase viscosity and inhibit ice crystal growth. In general, hydrocolloids also increase the protein load on the fat globules during the manufacture of emulsions 4~ This may be due to direct protein-polysaccharide binding at the o/w interface and/or protein-polysaccharide incompatibility in the water phase41. This

74 phenomenon has not been fully recognized in ice cream and should be studied in greater detail since it may give rise to important functional effects. Figure 11 shows the results of an ice cream mix containing a commercial hydrocolloid blend in combination with monodiglycerides. The protein load increases in the presence of hydrocolloids. In the presence of additional emulsifiers a very effective desorption of protein takes place during whipping and freezing in the ice cream machine. Effective protein desorption is facilitated by the increased viscosity of the mix due to increased surface shear forces, which makes the ice cream continuous freezer work better. The desorption of thick protein layers from fat globules of ice cream mix containing emulsifiers and hydrocolloids during ageing and mechanical treatment may also be observed by transmission electron microscopy (Figure 12). The protein bound to the surface of fat globules is desorbed as a thick coherent skin 23.

Figure 12 Transmission electron microscopy study of protein desorption in ice cream mix containing emulsifiers and hydrocolloids. (a) Immediately after homogenization the fat globules (t) are stabilized by adsorbed partially dissociated casein micelles (arrows). (b) During ageing the mix at 5~ the previously adsorbed protein film is released in the form of coherent protein layers (arrows) into the water phase (w). (c) After mechanical treatment in the ice cream freezer, desorbed protein layers are seen more often in the water phase without association to tat globules (arrows). From reference 48, courtesy of Dr. W.Buchheim, Kiel, Germany.

75

3.2. Interracial protein hydration The ageing at 5~ of whippable emulsions such as ice cream mix will enhance the hydration of milk proteins in the system. This is due to a property of casein micelles in milk. At low temperatures, the hydration or voluminosity of casein increases. The voluminosity is the volume of hydrated protein per gram of protein. This can be studied by analyzing the protein and water content in the sedimented casein pellet after centrifugation of skimmed milk. The increased hydration at low temperature is due to lower protein content in the pellet owing to dissociation of protein from the micelle (mainly beta-casein), and corresponds to data from the literature 42. During ageing of the mix, interfacial milk protein hydration also increases simultaneously with protein desorption from the fat globules. The water content of the isolated cream layers after centrifugation of ice cream mix can be analyzed by Karl Fischer titration. From such analyses, interfacial protein hydration can be calculated (Figure 13).

Figure 13 Desorption and hydration of protein bound to fat globules of ice cream mix during ageing at 5 ~ The voluminosity or hydration of interfacially bound protein may be calculated from the amount of water bound per gram of fat divided by the amount of protein bound per gram of fat. This corresponds to the volume of water per gram interfacial protein. Calculations show that emulsifiers facilitate interfacial protein hydration. This property is probably connected with their ability to desorb protein from the interface (Figure 14).

zy

76

Figure 14 Effect of low temperature on hydration of bovine casein micelles and of interfacially bound protein in ice cream mix with (+ E) and without (-E) emulsifier (saturated mono-diglyceride).

Figure 15 Effect of temperature on average particle size of ice cream mix with and without emulsifier.

The increased interfacial hydration in the presence of emulsifiers gives rise to a slight increase in the particle size of the fat globules in the ice cream mix (Figure 15). The volume of cream layers after centrifugation also increases up to 100% when lowering the temperature

77 from 30 oC to 5 oC. The increased interfacial hydration at 5~ described below.

gives rise to an increased mix viscosity as

3.3.

Interfacial tension Interracial tension analysis may be used to study the interaction of emulsifiers and milk protein at the oil-water interface of whippable emulsions. The interfacial activity of proteins is affected only slightly by temperature changes. In general, emulsifiers can reduce interfacial tension much more than protein, and this effect is especially pronounced at low temperatures. The relationship between surface tension and temperature in emulsifiers was observed two decades ago by Lutton et al. 43. They explained that this relationship is due to a transition from a liquid-expanded type of monolayer existing at high temperatures (above 40~ to a solid condensed monolayer existing at a lower temperature (below 20~ In solid condensed monolayers the molecular packing of the emulsifier molecules is much denser than in the liquid expanded monolayers, and these differences result in lower or higher surface tension, respectively. Models of such surface films are shown in Figure 16. Emulsifier molecules are packed more closely in the solid condensed film than in the liquid condensed film.

B

Water

Solid condensed film Surface areaJmol" 20-25/k 2

Water

Liquid condensed film Surface area/mol: 35-60/k 2

Figure 16 A schematic model of a solid condensed surface film (A) at temperatures below the melting point of the emulsifier, and of a liquid condensed fihn (B) at high temperatures (adapted froln reference 43). Such types of study may be performed using the Wilhelmy plate as a measuring device for interfacial tension analysis. This makes it possible to measure interracial tension continuously during temperature changes in the sample vessel controlled by external heating and cooling equipment 9. It is important to use a very pure triglyceride oil which is liquid down to 0~ to avoid disturbance of the analysis due to triglyceride crystallization. The connection between interfacial activity and emulsifier crystallization is easily

78 demonstrated in a system with a high emulsifier concentration in the oil phase, such as in toppings. Figure 17 shows measurements of interfacial tension between sunflower oil containing 5% emulsifier (propylene glycol monostearate) and distilled water. Separate samples of the oil phase containing emulsifier were analyzed for solid fat content.

Figure 17 Crystallization of the oil phase (sunflower oil) during cooling from 50~ to 0~ and interfacial tension ('7) between 5 % propylene glycol monostearate in sunflower oil and distilled water.

At temperatures above 25~ the presence of emulsifier results in only a slight reduction in interracial tension compared to a pure oil-water interface ('7 - 2 5 mN/m). When the temperature is decreased further, a significant drop in interfacial tension (,7) is registered due to interfacial crystallization followed by crystallization of emulsifier in the bulk oil phase below 15~ The increase in 7 observed at temperatures below 10~ is artificial being caused by a viscosity increase due to the crystal network which has formed. Interracial tension studies in relation to ice cream were also carried out using model two-phase systems similar to those mentioned above in connection with whipped toppings 9. These studies were carried out to analyze the interplay of emulsifiers and milk proteins at the oil-water interface. Emulsifiers were dissolved in sunflower oil, and protein in the water phase. With increasing amounts of saturated mono-diglycerides in the oil phase, increased interfacial activity was observed at low temperatures. At a concentration of 0.1%, which is usual in ice cream mix, the drop in interracial tension starts just below room temperature (15~ At this concentration no visible crystallization of emulsifier takes place in the oil phase. When both skimmed milk proteins and emulsifiers are present, a mixed film of both types of surtace active species forms at 40~ (Figure 18). When cooled, the emulsifier

79 crystallizes and dominates the interfacial tension. This will accelerate protein desorption from the oil-water interface. After reheating, the emulsifier melts and gives rise to readsorption of protein previously repelled from the interlace.

Figure 18 Interfacial tension of sunflower oil/water with and without protein (0.25% skimmed milk) in the water phase, and with and without 0.1% emulsifier (saturated monodiglyceride) in the oil phase. The two-phase systems were heated to 40~ for 1 hour, cooled to 5~ and reheated again to 40~ Symbols O = Oil; W = Water; P = Protein; E = Emulsifier. Reproduced from reference 44, courtesy of The American Institute of Chemical Engineers9 1990 AIChE. All rights reserved.

Reversible interfacial effects are also observed in ice cream emulsion systems as regards protein desorption and readsorption. The interfacial interaction of milk proteins and emulsifiers during temperature changes is believed to be the keystone in explaining the physical changes which take place in ice cream mix during ageing. Protein desorption, fat crystallization, and flocculation of fat globules appear to correlate with the interfacial activity of emulsifiers during cooling. In the absence of emulsifiers, the physical changes at low temperature appear to be reduced considerably 9. 3.4.

Surface tension In whippable emulsions with a high fat content, the air-water interface of the foam after whipping is dominated by adsorbed deproteinated fat globules. In whippable emulsions with a low fat content other foam stabilizing lnechanisms come into play, such as proteinhydrocolloid and protein-emulsifier interactions. The former subject may be studied by

80

zyxwvutsr

spectrophotometric analysis, the latter by various surface monolayer techniques 45,46 Increased emulsifier and hydrocolloid content is necessary to obtain stable foams when the fat content is reduced. Figure 19 shows results from practical tests of ice cream systems47.

Figure 19 Recommended dosages of commercial integrated emulsifier/hydrocolloid blend (CREMODAN'"SE 47) in ice cream mix. There are several reasons for this relationship. First, smaller fat globules with increased surface area tbrm in a low-tat recipe due to the use of higher homogenization pressure in such systems. Second, the protein-fat ratio is higher in low-fat recipes, resulting in stronger and thicker protein coverage on the tat globules which is more difficult to desorb. Third, the emulsifier takes over the function of tat in low-fat recipes, and will concentrate at the interface between air and serum, i.e., the emulsifiers will stabilize the air cells in a similar way to that of agglomerated tat. The adsorption of emulsifier to the air-water interface can be detected clearly by surface tension measurenaents because emulsifiers result in far greater depression of surface tension than proteins. Such analyses may also give intbrmation regarding the binding mechanisms of emulsifier in low-fat ice cream mix described below. Very surface-active emulsifiers (high HLB value) are capable of forming micelles in water. The latter is in equilibrium with emulsifiers at the air-water interface. At a certain concentration (= critical micelle concentration, CMC) the surface will be saturated with emulsifier and no further reduction in surface tension will be observed. The CMC can be found by surface tension measurenaents according to Figure 20.

81

Figure 20 A schematic figure showing how to find critical micelle concentration (CMC) from surface tension analysis at varying emulsifier concentrations. Monoglycerides and mono-diglycerides have low HLB values and cannot form micelles. They build up a multi-layer at the surface, resulting in a constantly decreasing surface tension as their concentration increases. However, in systems with proteins such as fat-free ice cream mixes, these emulsifiers behave as if they have a CMC. A possible explanation for this observation is that the unbound emulsifiier in the fat-free mix is in equilibrium with the protein-bound emulsifier. Above a certain concentration of emulsifier in the mix, any surplus of emulsifier will adhere to the protein in the water phase after the surface has been saturated. The unadsorbed emulsifier is seen as very small crystals less than 200 nm by electron microscopy analysis 4s. Without proteins the emulsifier will normally adsorb quickly to the surface, but in the presence of proteins adsorption takes up to 1 hour at 25 ~ (Figure 21).

Figure 21 Effect of protein, fat (oil) and emulsifier on surface tension of low-fat ice cream mix at 25 ~

82 Increased fat and increased protein content in the rnix delay adsorption of emulsifiers to air. Low temperature also has an inhibiting effect on this phenomenon. Surface tension analysis may be used to measure dosage effect in low-fat ice cream mixes. Such studies show that on a weight basis emulsifier is bound 10 times more strongly to tat than to milk protein in the nlix 49. As little as 1% fat in the mix has a very strong effect on the stability of the final ice cream (mentioned later under Descriptive Tests). Due to this strong effect the fat phase is believed not to be in a globular but in a more expanded crystalline state in such systems. This would give better possibilities for covering the air bubbles in the foam. This theory is highly speculative, and requires ti~rther studies for clarification.

zyxw

4.

W A T E R PHASE

The properties of the water phase in whippable emulsions are important for product stability. The water phase is influenced by the soluble components of the systems, i.e., sugars, proteins and hydrocolloids. Interfacial hydration may also influence the properties of the water phase, particularly in high-fat systems. 4.1.

NMR Pulse NMR techniques, both low-field and high-field, were applied to study the properties of water in food systems. All three possible nuclei, ~H, 2H and 170, were probed, and various models for data interpretation were developed. An extensive review of the subject may be found in Schmidt and Lai 5~ Most of the data were collected probing the ~H nucleus owing to high sensitivity, although problems of data interpretation due to chemical exchange and cross-relaxation are under debate -s~. These types of analysis are most useful in monitoring changes or trends in hydration. T2-relaxation analysis may be used to study the effect of ingredient composition on the properties of water in whippable emulsions ~6. In food systems non-exponential relaxation curves are often found. This can be accounted for by the presence of 2, 3 or more recognizable components representing species of hydrogen atoms with different mobility 51. Figure 22 is an example of such an analysis of ice cream mix. A data program from Bruker was used to resolve relaxation curves into two components. From such analyses the relative abundance (%) of each hydrogen species and their corresponding T2-values may be calculated. The figure shows the effect of emulsifier (E) and hydrocolloids (H) on the properties of H atoms with short T2 (usually called bound water).

zyxwv

83

Figure 22 Effect of emulsifiers (E) and hydrocolloids (H) on properties of bound water in ice cream mix (T2 time and percentage of hydrogen atoms with low T2). Both hydrocolloids and emulsifiers increase the water-binding capacity in the mix (increased % of hydrogen atoms with low T2 and decreased T2 values). A synergistic effect is observed when both ingredients are present. From studies described earlier in this chapter, the effect of hydrocolloids is assumed to be due to simple water binding and increased thickness of protein layers around the fat globules, whereas the effect of emulsifiers may be due to the increased hydration of interfacially bound protein as well as increased hydration of polar groups of emulsifier at the oil-water interface. Water crystallization in frozen whippable emulsions such as ice cream or aerated desserts, may be analysed by the NMR technique similar to that described for solid fat content analysis. Again, this technique is best used for only relative studies on the effects of ingredient composition on freezing/melting behaviour. 4.2.

Thermal analysis Differential scanning calorimetry is a very suitable method to study the behaviour of melting and freezing of water in frozen food systems. Using this technique it is also possible to measure the glass transition temperature. However, this may be of minor interest because the glass transition temperature in traditional ice cream is much lower than the storage temperature in ordinary freezing cabinets 52. Freezing point determination A successful calculation of the freezing points of ice cream mixes was made using the freezing points observed for sucrose solutions after correction for effects of lactose and milk proteins. Good agreement was obtained between the calculated and observed freezing point values in a series of experimental mixes 53. This is due to the fact that fat, protein and hydrocolloids in general have a negligible effect on the freezing point of the water solutions in which they are dispersed. Freezing point analysis then makes it possible to calculate the amount of water that will be frozen at any particular temperature during freezing, hardening,

4.3.

zy

84 and storage of ice cream. For details see Doan and Keeney 53. The characteristic freezing curve for ice cream can be used to explain why relatively low freezer drawing temperatures help facilitate a smooth-textured ice cream.

Figure 23 A typical freezing curve for ice cream showing the percentage of water frozen at various temperatures. Redrawn from reference 53.

More than 50% water is converted into ice crystals in ice cream at -5~ to -6~ which is the common drawing temperature for correctly operated continuous freezers. This portion of the water freezes very rapidly, often in less than one minute. Fast freezing induces the formation of small ice crystals, a critical prerequisite for smooth ice cream. At slightly higher temperatures (such as -4~ which is the common drawing temperature for batch freezers), less than 40% water is frozen and the freezing time will be longer. This is one of the reasons why ice cream frozen continuously is smoother in texture than batch-frozen products. A coarse texture may also develop as a result of heat shock, which involves alternate thawing and freezing of the water in the ice cream owing to temperature fluctuations in the hardening and storage cabinet. This results in a reduction of the textural quality of the ice cream. 4.4.

Size distribution of ice crystals Microscopic analysis is the only method available for estimating ice crystal size in ice cream. Light microscopy, equipped with cold stage and image analysis, may be used for this purpose 54. Low temperature scanning electron microscopy may also be used 55. Apart from the processing conditions discussed in section 4.3, hydrocolloids are important ingredients for controlling ice crystal growth in ice cream 56. Despite considerable scientific research in this area, the mechanism of this action remains obscure 57'58. Hydrocolloids do not influence the amount of water frozen or the glass transition point in ice cream which was believed to be involved in the stabilizing effect of hydrocolloids52. When ice cream starts to freeze, ice nucleation begins and water will freeze out of the solution in the form of pure crystals. As water is removed from the mix in the form of ice, the concentration of dissolved solids in solution increases. The unfrozen portion of the mix becomes increasingly concentrated as freezing continues, and contains dissolved sugars, milk

85 proteins, salts, and the hydrocolloids. During freeze concentration, the viscosity of the unfrozen phase becomes very high, primarily due to the increased hydrocolloid concentration, and this is believed to restrict the diffusion of water to existing ice crystals during fluctuations in temperature, or simply slowing down the latter process 52. Numerous hydrocolloids have been used in ice cream to inhibit ice crystal growth during distribution and storage. Useful hydrocolloid combinations and concentrations have been found for various ice cream products 59. The air cell stabilizing effect of agglomerated fat globules, promoted by emulsifiers and the ice-crystal-growth-controlling eft'ect of hydrocolloid stabilize the foam structure of ice cream to a great extent. This is evident by melt down analysis (see section 5.2) of ice cream exposed to heat shock.

4.5.

zyxwvuts

Wheying-off test In addition to their role in primary stabilization related to viscosity increase, some hydrocolloids (particularly carrageenan) are traditionally used as secondary stabilizers. Many of the primary stabilizing hydrocolloids, including locust bean gum and carboxy methyl cellulose induce precipitation of the milk proteins in the mix. This phenomenon in ice cream mix is known as wheying-off, and may be due to direct protein-polysaccharide binding and/or protein-polysaccharide incompatibility in the water phase 4~ The latter phenomenon may be due to decreased 'solvent quality' due to the competition between protein and polysaccharide for solubilisation. Carrageenan can prevent this wheying-off from occurring. Carrageenan binds directly to milk proteins forming a gel network which will protect the proteins from precipitation by the other hydrocolloids. Carrageenan is usually used at a much lower concentration than other hydrocolloids. This combined use of carrageenan and other hydrocolloids is very important in the stabilization of pasteurized chill-stable and UHT-treated ice cream premixes in softserve ice cream production. The effect of carrageenan is magnified in the freeze-concentrated aqueous phase of deep frozen ice cream, resulting in firm, cohesive gelation 6~ The wheying-off preventing activity may be estimated by making ice cream mix with locust bean gum as the main stabilizing hydrocolloid. The test carrageenan is added in different concentrations and the mixes are heated to 70~ for 30 minutes, cooled to 25~ with occasional stirring, and kept for 16 to 20 hours at 5~ The concentration at which wheying-off starts is estimated by visual inspection of graduated cylinder, and compared to a standardized carrageenan. From such studies the relative strength of the carrageenan being tested can be calculated 61.

5.

DESCRIPTIVE TESTS

A range of methods are used to test the textural quality of whippable emulsions. These methods are used to quantity the mechanical properties of the various products.

5.1.

Viscosity The viscosity range varies, depending on the whippable emulsion system in question. In whipped toppings viscosity increases as soon as the topping powder is reconstituted in cold water. This is due to the tbrmation and aggregation of hydrated fat crystals which will

86 stabilize the foam during whipping ~2. In UHT imitation whipping cream, a low viscosity of the emulsion before whipping is essential. The undesirable increase in viscosity during storage of cream is due to aggregation of fat globules, and this will reduce the pourability. If the agglomeration is too strong, the whipping properties will also be reduced. The viscosity of cream may be kept low by incorporating a sufficient amount of milk proteins and ionic emulsifiers, which will improve the emulsion storage stability before whipping. Fat globule aggregation is also minimized by quick cooling of the hot emulsion immediately after homogenization. In frozen whipping cream products, hydrocolloids are often used for ice crystal control 59. This will, of course, give higher emulsion viscosity. The viscosity of ice cream mix is important for processing in the ice cream freezer and must be within certain limits. Factors which may increase viscosity are increased %solids content, particularly hydrocolloids and protein, and low drawing temperatures in the freezer. Viscosity is usually measured on a simple comparison basis using a 50 to 100 ml capacity pipette, marked at an arbitrary place below the bulb. The flow time required to discharge the sample to the lower mark may be determined for water and then for the sample being tested for comparative purposes, and recorded in seconds62. More sophisticated rotation viscometers may also be used. The viscosity effect of hydrocolloids on ice cream mix is due to several factors. Hydrocolloids have a direct viscosity effect in binding large amounts of free water in the mix. Some hydrocolloids, such as kappa-carrageenan, form a gel network in the mix by binding to the milk proteins 6~ In general, hydrocolloids increase the thickness of the interfacial protein layer around the tat globules, and increased interfacial hydration is also obtained (see sections 3.1 and 3.2). Increased interfacial hydration is correlated to increased viscosity of mixes made with different hydrocolloids (Figure 24).

Figure 24 Viscosity of ice cream mix with different hydrocolloid types determined by the pipette method (flow time in seconds). Relation to interfacial hydration of fat globules

(%H20).

87 The viscosity effect increases exponentially when the ice cream is frozen 33. The effect of hydrocolloids becomes particularly dominant as free water crystallizes out during freezing 6~ The viscosity of aerated dessert mixes should be sufficiently low to withstand pasteurization, homogenization and ageing. On the other hand, the viscosity should be sufficiently high at low temperatures to stabilize the foam structure of the products. The foam should not gel or set before it is tapped, and should remain stable for several weeks without collapsing or showing signs of syneresis 2. Comlnon types of hydrocolloids for aerated desserts are gelatine, alginate and carrageenan. These hydrocolloids are all lnore or less shear-reversible gelling agents and are therefore suitable for use in aerated desserts 2. Only gelatine, which is acid-stable, can be used in low-pH desserts (yogurt-type desserts). When gelatine is used, the ageing temperature must be above 20~ and the mix must be agitated continuously to prevent the mix from gelling before it enters the aerator~. If hydrocolloids are used in sufficient quantities to enable them to gel the mix, then they will also be able to tbrm a stable foam when whipped. Starch and emulsifiers can also be used to provide aerated desserts with more body and a creamier consistency 2. 5.2.

Rheology of whipped emulsions After whipping whippable elnulsions obtain more solid-like properties. This means that ordinary viscometry measurements are not useful. The solid-like properties may be measured by non-destructive dynamic rheology analysis or by destructive methods using a Penetrometer, Jelly Tester, Instron instruments, or other types of texture analyzers. The latter methods are the most useful due to their simplicity and speed. Texture analysis of whippable emulsion must always be compared with the amount of air incorporated into the foam, which is known as percentage overrun and is calculated as follows: %

Where

Overrun

=

W1

-

W2

x

i00

1 -Weight of a given volume of whippable emulsion before whipping W2 = Weight of the same volume of whippable emulsion after whipping

W

Other useful parameters are whipping time and estimation of syneresis (serum separation from the foam). In ice cream the percentage of overrun is controlled in the ice cream machine, where the mix is whipped and frozen to a certain predetermined overrun. Only very few studies regarding the rheology of frozen ice cream are reported 63'64. This area should be studied in further detail to relate organoleptic and visual evaluations to instrulnental analysis. 5.3.

Melt-down analysis To test the melt-down properties, a rectangular block of ice cream of defined size is taken from the storage cabinet (e.g., at -20~ and is placed on a wire gauze (mesh size. e.g., 4 ram) at a controlled temperature between 15 and 25~ The melting may be followed

88 by weighing the melted ice cream collected in a beaker below the wire gauze. The time until the first drop falls, the amount of ice cream melted after 60 minutes, and the shape (stand-up quality) of the ice cream remaining at the top of the gauze are often used for evaluation 60,65. An example of melt-down analysis is shown in Figure 25. As little as 1% fat gives an enormous quality improvement in the texture of fat-reduced ice cream which has been properly stabilized by emulsifiers and hydrocolloids 66.

zy

Figure 25 Melting resistance of non-fat and low-fat ice cream (redrawn from reference 65)

In most countries consumers regard good ice cream melting properties as being synonymous with minimuln drip loss and good shape retention on melting. By contrast, in North America the retention of shape in melted ice cream is regarded as a defect 67.

5.4.

Organoleptic evaluation Organoleptic evaluation and product stability are usually assessed by a small expert panel trained to evaluate product appearance and ice cream consistency including smoothness, firmness, creaminess, sandiness, body, icy texture, and other properties. For a review of common body and texture defects, scoring and grading see Arbuckle 62. Although organoleptic evaluation is basically the most important analysis in practical ice cream product development, it is difficult to use that as the basis for exact conclusions. Despite these difficulties, it is always organoleptic analysis which has the highest priority due to its direct relationship with consumer acceptance. This argument is also valid for other types of whippable emulsions.

REFERENCES .

2. .

4.

Mann, E.J., Dairy Industries International 52 (1987) 15. Groven, S." Application of Emulsifiers and Stabilisers in Selected Dairy Products. Grindsted Technical Paper 215 (1989). Nielsen, H: Aerated Desserts. Grindsted Technical Paper 220 (1993). Brooker, B.E., M. Anderson & A.T. Andrews, Food Microstructure 5 (1986) 277.

89

,

Van Boekel, M.A.J.S and Walstra. P., Colloids Surf. 3 (1981) 109. zyxwvutsrqponmlkjihgfedcbaZYXWVUTSRQPON

6.

7. 8. ,

10.

11. 12. 13.

14. 15.

16.

17. 18.

19. 20. 21. 22. 23.

24. 25. 26. 27. 28. 29. 30.

Brooker, B.E., Food Structure 9 (1990) 223. Skoda, W. and Van den Tempel, M., J. Colloid Science 1___88(1963) 5687. Berger, K., in "Food Emulsions", second edition, K. Larsson and S.E. Friberg (Eds). Marcel Dekker, New York (1990) 367. Barfod N.M., Krog N., Larsen G. and Buchheim W., Fat Sci. Technol. 9.3 (1991) 24. Barfod N.M., Effect of emulsifiers on fat crystallisation in ice cream emulsions, in Proceedings from 15th Scandinavian Symposium on Lipids, Rebild Bakker, Denmark, V.K.S. Shukla and G. H6hner (Eds.), Lipidforum, G6teborg (1989) 133. van Putte, K. and van den Enden. J., J. Am. Oil Chem. Soc. 5 (1974) 316. Barfod, N.M., and Krog, N., J. Am. Oil Chem. Soc. 64.4(1987) 112. Krog, N., Barfod N.M. and Buchheim W., Protein-fat-surfactant interactions in whippable emulsions, in "Food Emulsions and Foams", E. Dickinson (Ed.), Royal Society of Chemistry, London (1987) 144. McClements, D.J. and Povey, M.J.W., Int. J. Food Sci. Technol. 2__33(1988) 159. Krog, N., Food emulsifiers and their chemical and physical properties, in 'Food Emulsions', second edition, K. Larsson and S.E. Friberg (Eds.), Marcel Dekker. New York (1990) 127. Barfod N.M., Krog, N. and Buchheim, W., Lipid-protein emulsifier-water interactions in whippable emulsions, in "Food Proteins", J.E. Kinsella and W.G. Soucie (Eds.), Am. Oil Chem. Soc., Champaign, Illinois (1989) 144. Le Neveu, D.M., Rand, R.P., Parsegian, V.A. and Gingell, D., Biophys. J. 1__88(1977) 209. Westerbeek, J.M.M. and Prins, A., Function of alpha-tending emulsifiers and proteins in whippable emulsions, in "Food polymers, gels and colloids", E. Dickinson (Ed.), Royal Society of Chemistry, Cambridge (1991) 147. Bern, M.B., Topping powder, internal Grindsted report (1992). Buchheim, W., Gordian 7___88(1982) 184. Buchheiln, W., Kieler Milchwirtschafte Forschungs Berichte 4__33(1991) 247. Buchheim, W., Barfod, N.M. and Krog, N., Food Microstructure 4 (1985) 221. Buchheim, W. and Dejmek, P., Milk and dairy-type emulsions, in "Food Emulsions", second edition, K. Larsson and S.E. Friberg (Eds.). Marcel Dekker, New York (1990) 203. Anonymous: Unilever uses Mastersizer to monitor particle size. Ice Cream and Frozen Confectionery. March 1993, 189. Bisperink, C.G.J, Ronteltap, A.D. and Prins, A., Adv. Colloid Interface Sci. 3___88(1992) 13. Anonymous: Phase contrast microscopy of ice cream mix, Technical Memorandum 217, Grindsted Products (1993). Andreasen, T., Grindsted system for stick novelties, paper presented at the INTER-EIS Seminar 1987, Solingen, Technical Paper 214. Grindsted Products. Anonymous: Determination of fat in ice cream (ice cream mix) according to the Gerber method, Technical Memorandum 214, Grindsted Products (1993). Anonymous: Determination of fat in ice cream - gravimetric. Technical Memorandum 215. Grindsted Products (1993). de Feijter, J.A., Benjamins, J., Tamboer, M., Colloids Surf. 27 (1987) 243.

90 31. 32. 33. 34. 35. 36. 37. 38. 39.

40. 41.

42. 43. 44. 45. 46. 47. 48.

49 50.

51. 52. 53.

54. 55. 56. 57. 58.

Darling, D.F., Birkett, R.J., Food Colloids in Practice, in "Food Emulsions and Foams", E. Dickinson (Ed.), Royal Society of Chemistry, London (1987), 1. Dickinson, E. and Tanai, S., J. Agric. Food Chem. 4__9_0(1992) 179. Keeney, P.G., Food Technol. 3__6_6(1982) 65. Towler, C. and Stevenson, A., New Zealand J. of Dairy Sci. and Technol. 2__33(1988) 345. Goff, H.D., Loboff, M., Jordan, W.K. and Kinsella, J.E., Food Microstructure 6 (1987) 193. Oortwin, H. and Walstra, P., Neth. Milk Dairy J. 3_33(1979) 134. Dickinson, E., Rolfe, S.E. and Dalgleish, D.G., Food Hydrocolloids 3 (1989) 193. Heertje, E., Nederlof, J1, Hendrickx, H.A.C.M. and Lucassen-Reynders, E.H., Food Structure 9 (1990) 305. Reimerdes, E.H., Changes in the proteins of raw milk during storage, in "Developments in dairy chemistry", vol. 1, P.F. Fox (Ed.), Applied Science Publishers, London (1982) 271. Tolstoguzof, V.B, Food Hydrocolloids 4 (1991) 429. Dickinson, E. and Euston, S.R., Stability of food emulsions containing both protein and polysaccharide, in "Food polymers, gels, and colloids", E. Dickinson (Ed.), Royal Society of Chemistry, Cambridge (1991) 132. Bloomfield, V.A. and Morr, C.V, Neth. Milk Dairy J. 2__7_7(1973) 103. Lutton, E.S, Stauffer, E., Martin, J.B. and Fehl, A.S, J. Colloid Interface Sci. 3___00 (1969) 283. Krog, N. and Barfod N.M., AIChE Symposium, Series 8___66,No. 277 (1990), 1. Rahman, A. and Sherman, P., Colloid Polym. Sci., 260 (1982) 1035. La Libert6, M.-F., Britten, M. and Paquin, P., Can. Inst. Food Sci. Technol. J. 2__.!_1 (1988) 151. Anonymous, CREMODAN*"SE 47, Product Description 214, Grindsted Products (1988). Buchheim, W., Structures and interactions in ice cream mixes. In Proceedings of the Penn State Ice Cream Centennial Conference, M. Kroger (Ed.), Pennsylvania State University, College Park, PA, (1992) 281. Barfod, N.M., Unpublished results. Schmidt, S.J. and Lai, H.-M., Use of NMR and MRI to study water relaxations in foods. In "Water relationships in foods", H. Levine and L. Slade (Eds.), Plenum Press, New York (1991) 405. Brosio, E., Altobelli, G. and DiNola, A., J. Food Technol. 1___99(1984) 103. Goff, H.D. and Caldwell, K.B., Modern Dairy 7__Q0(1991) 14. Doan, F.J. and Keeney, P.G., Frozen dairy products. In "Fundamentals of Dairy Chemistry", B.H. Webb and A.H. Johnson (Eds.), AVI Publishing Co., Westport, Conn. 1965, 771. Donhowe, D.P., Hartel, R.W., and Bradley, R.L., J. Dairy Sci. 7__44(1991) 3334. Caldwell, K.B., Goff, H.D. and Stanley, D.W., Food Structure 1__!1(1992) 1. Caldwell, K.B. Goff, H.D. and Stanley D.W., Food Structure 1_1_1(1992) 11. Muhr, A.H. and Blanshard, J.M.V., J. Food Technol. 2__!_1(1986) 683. Buyong, N. and Fennema, O., J. Dairy Sci. 7__!_1(1988) 2630.

91 59. 60. 61. 62. 63. 64. 65. 66. 67.

Knightly, W.H., J. Food Technol. 22 (1968) 73. Dea, I.C.M., Int. Food Ingred., No. 1 (1991) 9. Anonymous. GENU Control Method C306-1, The Copenhagen Pectin Factory Ltd. (Hercules Inc.) (1978). Arbuckle, W.S., Ice Cream. Third Edition. AVI Publishing Company Inc., Westport, Conn., 1977. Shernlan, P., J. Food Sci., 30 (1965) 202. Windhab, E., ZFL 5 (1989) 242. Larsen, G., "The principle of homogenisation of an ice cream mix", paper presented at the INTER-EIS Seminar 1988, Solingen, Technical Paper 216, Grindsted Products. Christensen, E.S., "hnprovement of creaminess in non-fat and low-fat frozen desserts", paper presented at INTER-EIS Seminar, Solingen 1991. Mahdi, S.R. and Bradley, R.L, J. Dairy Sci. 5_]_1(1968) 931.

This Page Intentionally Left Blank

Characterization of Food: Emerging Methods A.G. Gaonkar (Editor) 9 1995 Elsevier Science B.V. All rights reserved.

93

Chapter 4 U l t r a s o n i c c h a r a c t e r i z a t i o n of foods D.J. McClements Department of Food Science, University of Massachusetts, Amherst, MA 01003, USA

1. I N T R O D U C T I O N Ultrasound is the study and application of sound waves whose frequency is too high to be detected by the human ear, i.e., above about 16 kHz [1]. This is a purely arbitrary cut-off point, determined by the limitations of the human ear. The physics describing the propagation of ultrasonic waves is the same as that describing the propagation of sound waves. Ultrasound is already an established technique for characterizing the physical properties of many biological and non-biological materials [ 1]. It is routinely used in medicine to detect tumors and to determine the health and sex of fetus' in the womb [ 1]. In materials testing it is used to characterize the position and size of cracks in metals and plastics [2]. Oceanographers use acoustics to map the contours of the sea-bed, and to determine the location, number and size of fish swimming in shoals [1,3]. The chemical processing industry uses ultrasound to determine the concentration of solutes in aqueous solutions and to determine the flow rate of liquids and particulates in pipes [2]. It is not suprising therefore that ultrasound can also be used to characterize food materials. The possibility of using ultrasound to characterize foods has been realized for over half a century [5-8]. The wide variety of different applications investigated during this period (see Section 5), reflects the diversity and complexity of food materials, as well as the versatility of the ultrasonic technique. Even so, there are still few areas in the food industry where ultrasound is recognized as an established technique for characterizing foods, with perhaps the exception of the inspection of meat quality. This situation will almost certainly change in the near future, and ultrasound will become as important a tool as NMR for characterizing foods. Advances in microelectronics have made available sophisticated electronic instrumentation capable of making accurate ultrasonic measurements at relatively low-cost. The interaction between ultrasound and microheterogeneous materials is fairly well understood, and there are mathematical formulae available for interpreting ultrasonic measurements in a number of systems relevant to the food industry. Finally, ultrasound offers a number of advantages over alternative techniques used to characterize food: it is capable of rapid and precise measurements, it is non-intrusive and non-invasive, it can be applied to systems which are concentrated and optically opaque, it is relatively inexpensive and it can easily be adapted for on-line measurements. There are two distinct types of applications of ultrasound in the food industry: high and low intensity [7]. High intensity ultrasound is used to physically alter the properties of a material through which it propagates. It utilizes relatively high power levels (> 1 W cm -2) and low frequencies (< 0.1 MHz). Typical applications of high intensity ultrasound are cleaning, homogenization, cell disruption, promotion of chemical reactions and extraction [7]. Low intensity ultrasound is used to provide information about the physical properties of materials. The power levels used are lower than those used in high intensity applications (< 0.1 W cm -2) and the frequencies higher (0.1 - 100 MHz). Low intensity ultrasound does not alter the

zy

94

zyxwvut zyxwv

properties of a material and is therefore non-destructive. Only low intensity applications are reviewed in this chapter. The objectives of this chapter are to introduce the basic concepts of ultrasonic propagation in materials, to describe some of the most important methods for measuring and interpreting ultrasonic measurements, and to outline existing and possible applications of the technique in the food industry. 2. U L T R A S O N I C P R O P A G A T I O N IN M A T E R I A L S

2.1. General considerations Ultrasound is used to obtain information about the properties of a material by measuring the interaction between a high frequency sound wave and the material through which it propagates. This interaction depends on the frequency and nature of the ultrasonic wave, as well as the composition and microstructure of the material. The parameters most commonly measured in an ultrasonic experiment are the velocity at which the wave travels and the extent by which it is attenuated. To understand how these parameters are related to the properties of foods it is useful to consider the propagation of ultrasonic waves in materials in general.

Figure 1. Ultrasonic compression and shear waves generated by the application of a sinusoidal force F(t) to the material. An ultrasonic wave can propagate through a material in a number of different ways. Consider a material to consist of a series of imaginary layers of particles (Figure 1). If a force is applied to one end of the material it will act throughout the material due to restoring forces between the layers. When an oscillating mechanical wave is applied perpendicular to the surface of the material a compression wave is generated, which m o v e s t h r o u g h the material as a series of expansions and compressions. The oscillation of the layers is in the same direction as the propagation of the ultrasonic wave. If the ultrasonic wave is applied parallel to the surface of the material a shear wave is generated. In this case the layers move perpendicular to the direction of propagation of the ultrasonic wave. Other types of wave are also possible, e.g., surface or lamb waves [2], although these are seldom used in the food industry at present. There is no net movement of the particles in a material: each layer

95

Figure 2. Dependence of the displacement of a particle from its equilibrium position on the time and distance the wave has traveled. simply oscillates around its equilibrium position and returns to this position when the energy stored as ultrasound is dissipated. An ultrasonic wave is represented graphically by considering the displacement (~ of the layers of particles from their equilibrium positions (Figure 2). The displacement varies with the distance (x) traveled by the wave and the time (t). The amplitude of the particle displacement decreases with distance because of attenuation of the ultrasonic wave (see later). The important characteristics of an ultrasonic wave are the amplitude and frequency (f), which are chosen by the investigator, and the wavelength (;~) and attenuation coefficient (a), which are characteristic of the material. The ultrasonic velocity (c) is simply related to the wavelength and frequency: c = Z.f, so that it is also a characteristic of the material. Measurement of the ultrasonic velocity (or wavelength) and attenuation coefficient is the basis of the ultrasonic testing of materials. A mathematical description of an ultrasonic wave must describe the dependence of the particle displacement on distance and time, and the reduction of its amplitude with distance traveled through the material. For plane sinusoidal waves the following equation is appropriate: 9 27rx 2nt - ~ oe'(T-T)

e-aX

zyxw (1)

The first term describes the sinusoidal variation of the particle displacement with distance, the second term the variation with time, and the final term describes the attenuation of the wave. In most text books equation 1 is written in the following form:

- ~o ei(~-~

(2)

Here co is the angular frequency ( = 2rcf) and k is the wave number ( = o~/c + M), which contains information about the ultrasonic properties of the material, i.e., the velocity and

96 attenuation coefficient. The oscillatory variations in the particle displacement are accompanied by variations in the velocity of the particle, and the local pressure, temperature and density of the material [ 1]. Variations in these quantities can be described by equations with a similar form to that for particle displacement and are the starting point for the derivation of most mathematical formulations used to describe ultrasonic propagation in materials. In practice ultrasound is usually propagated through materials in the form of pulses rather than continuous sinusoidal waves. Pulses contain a spectrum of frequencies, and so if they are used to test materials that have frequency dependent properties the measured velocity and attenuation coefficient will be average values. This problem can be overcome by using Fourier Transform analysis of pulses to determine the frequency dependence of the ultrasonic properties.

zyxwvutsrq

2.2 Relationship between the ultrasonic and physical properties of a material A simple relationship can be derived between the ultrasonic properties of a material and its physical properties by a mathematical analysis of the propagation of plane waves in a material. A general wave equation can be derived by differentiating equation 2 twice with respect to distance and twice with respect to time: d2

2

(3)

zyxwvut

This equation is applicable to the propagation of electromagnetic waves, as well as to ultrasonic waves, although the terms in the wave number have different meanings. It is fairly straight forward to derive an equation which describes the propagation of high frequency sound waves in a material by considering the restoring forces acting on an element of the material as the wave passes through [ 1]:

d2~

p d2~

dx 2

E dt 2

(4)

Here E is the appropriate elastic modulus (which depends on the physical state of the material and the type of wave propagating) and p is the density. By combining equations 3 and 4 the physical properties of a material (E and p) can be related to its ultrasonic properties (c and a).

(5) Thus a measurement of the ultrasonic properties can provide valuable information about the bulk physical properties of a material. The elastic modulus and density of a material measured in an ultrasonic experiment are generally complex and frequency dependent and may have values which are significantly different from the same quantities measured in a static experiment. For materials where the attenuation is not large (i.e., a