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Thermal inactivation kinetics of lysozyme and preservative effect beer Makki, Farid 1996

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THERMAL INACTIVATION KINETICS OF LYSOZYME AND PRESERVATIVE EFFECT IN BEER By FARID MAKKI B.A.Sc. (Agric. Eng.), Institut National Agronomique Paris-Grignon (France), 1986 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE FACULTY OF GRADUATE STUDIES DEPARTMENT OF FOOD SCIENCE We accept this thesis as conforming toJhe_ranjired standard THE UNIVERSITY OF BRITISH COLUMBIA January 1996 © Farid Makki, 1996 In presenting this thesis in partial fulfilment of the requirements for an advanced degree at the University of British Columbia, I agree that the Library shall make it freely available for reference and study. I further agree that permission for extensive copying of this thesis for scholariy purposes may be granted by the head of my department or by his or her representatives. It is understood that copying or publication of this thesis for financial gain shall not be allowed without my written permission. Department of The University of British Columbia Vancouver, Canada DE-6 (2/88) ABSTRACT Thermal stability of lysozyme in aqueous buffer solutions was studied at different temperatures (73-100°C), pH values (4.2-9.0) and levels of sucrose (0, 5%, 15%) and sodium chloride (0, 0.1M, IM). The results, fitted to a first order model and expressed in terms of decimal reduction time (D), inactivation rate constant (k), decimal reduction temperature (z) and Arrhenius activation energy (EJ indicated that lysozyme was most stable at pH 5.2, and thermal stability decreased sharply as the pH increased to 9.0. A regression equation for prediction of k as a function of temperature and pH was derived, with the best fit obtained for the model in the pH range of 5.2-7.2 (adjusted multiple 1^=0.975). At pH values of 7.2 and 9.0, sodium chloride had a clear stabilizing effect against heat inactivation of lysozyme. Sucrose stabilized lysozyme against heat inactivation at 75°C but not at 91°C. Results of the study of the thermal inactivation kinetics of lysozyme proved to be practical and useful for prediction of residual lysozyme activity in the pH and temperature range studied. Thermal inactivation kinetics of lysozyme and its potential to prevent or delay microbial growth were also investigated in beer. Lysozyme at concentrations of 10 and 50 ppm appeared to delay growth of the spoilage bacteria L. brevis and P. damnosus in beer, but did not prevent growth of the bacteria. i i TABLE OF CONTENTS Page ABSTRACT ii TABLE OF CONTENTS iii LIST OF TABLES v LIST OF FIGURES vi ACKNOWLEDGEMENTS viii 1. INTRODUCTION 1 2. LITERATURE REVIEW 4 2.1 Chemical Characteristics of Lysozyme 4 2.1.1 Definition and Types of Lysozyme 4 2.1.2 Brief Description of the Lysozyme Molecule 5 2.1.3 Methods of Extraction and Quantitation of Lysozyme 8 2.2 Applications of Lysozyme as a Food Preservative 10 2.3 Thermal Stability of Lysozyme 14 2.3.1 Reversible Denaturation Measured by DSC 14 2.3.2 Factors Affecting Thermal Stability of Lysozyme 15 2.3.3 Irreversible Thermoinactivation 21 2.4 Use of Lysozyme as an Antimicrobial in Beer 26 i i i 3. MATERIALS AND METHODS 29 3.1 Thermal Inactivation of Lysozyme in Aqueous Buffers 29 3.1.1 Reagents 29 3.1.2 Substrate Preparation 29 3.1.3 Lysozyme Assay 29 3.1.4 Heat Treatment of Lysozyme 30 3.1.5 Calculation of Kinetic Parameters 33 3.2 Thermal Inactivation of Lysozyme in Commercial Beer 35 3.3 Inoculation of Beer With Lactic Acid Bacteria and Lysozyme 36 3.3.1 Bacterial Strains and Culture Conditions 36 3.3.2 Preparation of Stock Solutions of Lysozyme 38 3.3.3 Bottle Inoculation Procedure 38 3.3.4 Plating For Colony-Forming Units 39 4. RESULTS AND DISCUSSION 40 4.1 Effects of pH on Irreversible Thermal Inactivation of Lysozyme 40 4.2 Effects of Sucrose and Sodium Chloride on Thermal Inactivation of Lysozyme 58 4.3 Lysozyme's Thermal Inactivation Kinetics in Beer and Effect on Lactic Acid Bacteria 63 5. CONCLUSION 72 6. REFERENCES 74 i v LIST OF TABLES Table 1. Rate constants of irreversible inactivation of lysozyme at 100°C 23 Table 2. Buffer composition for different pH values 31 Table 3. Effect of pH on decimal reduction time (D) of lysozyme 47 Table 4. Reaction order with respect to time (n,) of the thermal inactivation of lysozyme, tested at intervals of 0.1 in the 0.5-2.0 range 48 Table 5. Effect of pH, sucrose and sodium chloride on the activation energy (E„) and decimal reduction temperature (z) of the thermal inactivation of lysozyme 52 Table 6. Regression output of the multiple regression analysis for the prediction of the reaction rate constant k of thermal inactivation of lysozyme as a function of pH and inverse temperature 56 Table 7. Confidence intervals (90%) for the decimal reduction temperature (z) and the activation energy (EJ of lysozyme at different pH values 57 Table 8. Effect of pH, sucrose and NaCl on thermal inactivation parameters D and k of lysozyme at 75°C and 91°C 59 Table 9. Stability of lysozyme in concentrated sucrose solutions 62 Table 10. Decimal reduction time at 4 different temperatures and reaction order of thermal inactivation of lysozyme in beer (pH 4.70) 65 Table 11. Effect of lysozyme on survival of Lactobacillus brevis B12 and Pediococcus damnosus B130 in beer 68 v LIST OF FIGURES Figure 1. The primary structure of hen egg-white lysozyme with the amino acids involved in the four disulfide bridges indicated 6 Figure 2. The time course of irreversible inactivation of lysozyme at 100°C as a function of pH and the enzyme concentration 22 Figure 3. Diagram of the reaction apparatus used for the serum bottle technique 32 Figure 4. Growth curves of Lactobacillus brevis B12 and Pediococcus damnosus B130 in beer 37 Figure 5a. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 4.20 ... 41 Figure 5b. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 5.20 42 Figure 5c. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 6.24 43 Figure 5d. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 7.20 44 Figure 5e. Thermal inactivation of dilute lysozyme in aqueous buffer atpH8.10 45 Figure 5f. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 9.00 46 Figure 6a. Effect of pH (4.20-6.24) on decimal reduction time (D) of lysozyme 53 Figure 6b. Effect of pH (7.20-9.00) on decimal reduction time (D) of lysozyme 54 Figure 7. Thermal inactivation of lysozyme in unpasteurized beer (pH4.70) 64 v i Figure 8a. Survival of Lactobacillus brevis B12 in unpasteurized beer in the presence of lysozyme (Trial 1) 69 Figure 8b. Survival of Lactobacillus brevis B12 in unpasteurized beer in the presence of lysozyme (Trial 2) 70 Figure 9. Survival of Pediococcus damnosus B130 in unpasteurized beer in the presence of lysozyme 71 v i i ACKNOWLEDGEMENTS I wish to thank my supervisor Dr. Timothy D. Durance for providing encouragement, guidance and support throughout this project. I would also like to extend my appreciation to the other members of the supervisory committe, Dr. S. Nakai, Dr. D.D. Kitts and Dr. E . Li-Chan for their helpful and constructive comments and suggestions on my research program. The second part of this project would not have been possible without the cooperation of Dr. T.M. Dowhanick (Labatt Breweries of Canada) who graciously provided the required bacterial strains, and Mrs. Jadwiga Sobczak (Labatt Breweries of Canada) for her expert advice. Finally I would like to thank Ms. Karoline Lee and Mr. Sherman Yee for providing technical advice and assistance. v i i i 1 1. INTRODUCTION The antirnicrobial activity of lysozyme ( muramidase; EC 3.2.1.17 ), known since the discovery of this enzyme in 1922 by Alexander Fleming, has stimulated considerable interest in its use as a "natural" food preservative. Mounting pressures to ban or reduce the use of chemical preservatives in foods, and the increasing consumer demand for "natural" preservatives and minimally processed foods, have further enhanced the potential of lysozyme for use in food manufacturing. Lysozyme has the ability to hydrolyze the 6-1,4-glycosidic bond between N-acetylmuramic acid and N-acetyl-D-glucosamine in polysaccharides which form the backbone of cell walls in certain bacteria. It has been shown to be effective in preserving a variety of foods such as fresh fruits and vegetables, tofu bean curds, meats, seafoods and wines, for which many Japanese patents have been granted (Cunningham et al., 1991). Lysozyme is currently used in Europe mainly as an alternative to nitrates, to combat the cracking of cheese wheels, a problem known as "late blowing" caused by Clostridium tyrobutyricum during the manufacture of semi-hard cheeses such as Edam and Gouda. In 1984, an estimated 100 metric tons of chicken egg white lysozyme were used in Europe for this purpose (Scott et al.,1987). The relatively high thermal stability of lysozyme also makes it attractive for use in pasteurized and heat-sterilized food products, possibly allowing reduced thermal processes, and therefore minimized nutritional and sensory quality loss (Fox and Morrissey, 1980). It may also find application as a safeguard against food poisoning caused by food pathogens, including thermophilic Clostridia, certain strains of C. botulinum and Listeria monocytogenes (Hughey 2 and Johnson, 1987; Hughey et al., 1989). The relative instability of most enzymes under operational conditions, which may involve high temperatures, extreme pHs and exposure to organic solvents or denaturing agents all leading to enzyme inactivation, is a major obstacle limiting their industrial use. Of these, thermal inactivation is by far the most important mode of enzyme inactivation (Klibanov, 1983). The suitability of lysozyme as a preservative in foods will, among other factors, be conditioned by its thermal stability in these products, as the latter undergo various processing operations. Many studies of thermal denaturation of lysozyme by differential scanning calorimetry (DSC) have been reported (Bull and Breese, 1973; Pfeil and Privalov, 1976a,b; Johnson et al., 1978; Back et al., 1979; Gekko, 1982). Although useful to predict trends, these results do not truly reflect inactivation events, and therefore are of little value for prediction of residual enzyme activity. The various factors that affect lysozyme activity such as temperature, pH, ionic strength, salts and alcohols have also been the subject of numerous investigations (Chang and Carr, 1971; Davies et al., 1969; Kravchenko et al., 1967; Yang and Cunningham, 1993). The work described in this thesis, which addresses the thermal inactivation kinetics of lysozyme in vitro and in a model beverage, was undertaken with the following research objectives: Part I- To evaluate the effect of pH over the range of 4.2-9.0, as well as of sucrose and sodium chloride, two commonly encountered ingredients in processed foods known to influence protein stability, on the thermal inactivation of lysozyme in aqueous buffer solutions. 3 The kinetics of lysozyme inactivation were described in terms of decimal reduction times (D), decimal reduction temperatures (z), inactivation rate constant (k) and Arrhenius activation energy (EJ. These data are expected to allow approximate prediction of thermal stability of lysozyme in specific food processing situations. Part II- To evaluate the thermal inactivation kinetics of lysozyme in a beverage typically undergoing pasteurization, such as beer, and to investigate lysozyme's potential to prevent or delay growth of lactic acid bacteria, responsible for spoilage in beer. 4 2. LITERATURE REVIEW 2.1 Chemical Characteristics of Lysozyme 2.1.1 Definition and Types of Lysozyme Lysozymes are defined as 1,4-B-N-acetylmuramidases cleaving the glycosidic bond between the C-l of N-acetylmirramic acid and the C-4 of N-acetylglucosamine in the bacterial peptidoglycan (Jolles and Jolles, 1984). They are found in a variety of human tissues and secretions (tears, milk, serum, saliva), as well as in those of various other vertebrates, invertebrates and in bacteria, phages and plants. The ubiquitous nature of these proteins coupled with their bacteriolytic property have led to the widely accepted hypothesis that lysozyme plays a significant part in the defence mechanism of various organisms against bacterial infection. However, despite extensive immunological and evolutionary studies devoted to lysozymes, the biological role of the latter is still not exactly known (Jolles and Jolles, 1984). The most thoroughly investigated type of lysozyme, which also happens to be the most commonly encountered, is the hen egg-white lysozyme, also referred to as the c-type (chicken-type) lysozyme. The early determination of the primary structure of hen egg-white lysozyme by Jolles et al., (1963) and Canfield (1963), and the elucidation of its 3-dimensional structure by X-ray crystallographic analysis by Blake et al., (1965) paved the way for research on the relationships between amino-acid sequence and 3-dimensional conformation of proteins, using lysozyme as a model. The complete primary structures of many other c-type lysozymes have 5 since been established . These include, among others, lysozymes isolated from quail (Kaneda et al.,1969; Prager et al., 1972), turkey (La Rue and Speck, 1970) and duck (Hermann and Jolles, 1970; Kondo et al., 1982) egg-whites, as well as from human milk (Jolles and Jolles, 1971), baboon (Hermann et al.,1973), rat (White et al., 1977) and bovine stomach (Dobson et al., 1984; Jolles et al., 1984). Another type of lysozyme, first discovered in 1967 by Dianoux and Jolles, is the goose egg-white lysozyme referred to as the g-type (goose-type) lysozyme. The g-type lysozymes have a molecular weight of around 21,000 daltons instead of 14,000 for the c-type lysozymes, and do not cross react immunologically with the latter. The complete primary structure of g-type lysozymes from black swan (Simpson et al., 1980), ostrich (Schoentgen et al.,1982) and Emden goose (Simpson and Morgan, 1983) have so far been determined. Other distinct types of lysozyme which differ from the c and g types on the basis of structural, catalytic and immunological criteria have also been reported, such as bacteriophage (T4) lysozymes (Inoue et al., 1970; Szewczyk et al., 1982), plant lysozymes (Smith et al., 1955) and fungal lysozymes (Fouche and Hash, 1978). 2.1.2 Brief Description of the Lysozyme Molecule Hen egg-white lysozyme is a relatively small molecule consisting of a single polypeptide chain of 129 amino acid residues. This basic protein has a molecular weight of 14,300 daltons and an isoelectric point of 10.7 (Powrie and Nakai,1986). The enzyme is cross-linked by four disulfide bridges between residues 64-80, 76-94, 6-127 and 30-115, as illustrated in Figure 1. The molecule has roughly the shape of a spheroid with a deep cleft on one side which is the active site of the enzyme. The polypeptide chain is folded on itself with Figure 1. The primary structure of hen egg-white lysozyme with the amino acids involved in the four disulfide bridges indicated ( Source: Proctor, 1991). 7 the first 40 amino acid residues from the N-terminal end forming a compact globular domain characterized by a hydrophobic core trapped between two oc-helices (residues 4-15 and 24-35). Residues 40-85 form a second more hydrophilic domain composed partly of a fl-pleated sheet, one surface of which protrudes toward the surface while the other lines one side of the active site cleft. The remainder of the polypeptide chain partially fills the gap between the two domains and builds up the other side of the active site cleft with mainly hydrophobic residues (Blake etal., 1978). Two carboxyl groups (Glu-35 and Asp-52) have been identified as the functional groups involved in the activity of hen egg-white lysozyme. The carboxyl group of Glu-35 in its non-ionized form serves as a proton donor, and that of Asp-52 in its ionized form stabilizes, by electrostatic interaction, the oxocarbonium ion intermediate (Inoue et al., 1992). According to the generally accepted mechanism of hydrolysis proposed by Blake et al., (1967) lysozyme first attaches itself through hydrogen bonds and electrostatic forces to the polysaccharide, comprised of alternate units of N-acetylglucosamine (NAG) and N-acetylmuramic acid (NAM). In the process, the ring of the NAM residue becomes distorted contributing to the formation of a carbonium ion at the C, of this residue, and to the consequent weakening of the C, carbon-oxygen bond. Glu-35 acts as a proton donor facilitating the formation of a hydroxyl group with the bridge oxygen atom, and the release of the adjacent NAG residue. Heterolysis of the Q carbon-oxygen bond produces a carbonium ion which is stabilized by interaction with the negative charge of the side chain of Asp-52. Finally, a water molecule attacks the carbonium ion thus completing the hydrolysis (Vernon, 1967). 8 2.1.3 Methods of Extraction and Quantitation of Lysozyme Numerous methods for the isolation and purification of lysozyme from hen egg-white have been reported in the literature. The classic procedure, developed by Alderton and Fevold (1964) involves direct crystallization at pH 9.5 in the presence of 5% sodium chloride, with lysozyme yields of 60-80%. The major disadvantage of this method is that it requires several crystallization steps involving the addition of salts and additives which affect the functionality of the remaining egg-white. Since then, many chromatographic procedures for the isolation of lysozyme, based on ion exchange or affinity chromatography have been proposed. One of the first adsorbent materials used to isolate lysozyme by affinity chromatography was chitin (Cherkasov and Kravchenko, 1967), which can be partially hydrolyzed by lysozyme. Glucochitin, obtained by deamination of chitin from crab shells was found to be an even more suitable material (Cherkasov and Kravchenko, 1969), since reduced ionic charge due to deamination also results in reduced non-specific binding of the deaminated chitin due to loss of ion-exchange properties. Weaver et al., (1977) found that squid deaminated chitin (DECH) had an even higher specificity for lysozyme than crab DECH, along with good stability and allowing fast flow rates. The major limitations of affinity methods which have hindered their adoption for large scale isolation of lysozyme from egg-white are the high cost of the affinity support materials, poor flow rates of undiluted egg white through affinity resins and short useful life of affinity resins compared to ion exchange resins. Currentiy the most efficient and widely used method for lysozyme separation involves ion exchange chromatography combined with isoelectric precipitation. Egg white is mixed with a suitable cation exchange resin such as carboxymethyl cellulose (CMC), where lysozyme 9 binds specifically to the latter. Once the residual egg-white has been filtered off and recovered, the resin is placed in 5% sodium chloride causing a release of the bound lysozyme into solution. The lysozyme solution is then adjusted to pH 9.5 which causes a precipitation of the lysozyme. The protein is then redissolved, desalted, concentrated and freeze dried (Durance, 1987). A similar procedure was reported by Ahvenainen et al., (1979) who instead of CMC used a commercially available cation exchange resin Duolite C-464, with good yield and purity of lysozyme. Their batch method however required a rather complex resin regeneration process. Although column ion exchange offers better results than bulk ion exchange in terms of yield and purity of isolates, suitable resins with the macroporous structure and mechanical rigidity necessary for column chromatography were not available until relatively recently (Durance, 1994). A convenient and efficient column chromatography method using cation exchange was reported by Li-Chan et al., (1986). Among several different resins that were tested and compared for lysozyme adsorption capacity, recovery of lysozyme, egg-white flow rates and potential for clogging, Duolite C-464 proved to be the most suitable support material, with lysozyme recovery in the 90-95% range. The simplicity of the process, based on a cycle consisting of egg-white application, weak buffer rinse and strong buffer elution of lysozyme, makes it suitable for operation in an automated, continuous mode. Since the discovery of lysozyme, numerous methods have been devised for its quantitation including turbidimetric, lysoplate assay, immunoassay and other methods. One of the oldest and most widely used procedures is the turbidimetric assay, based on spectrophotometric measurements of the clearing of a turbid suspension of Micrococcus 10 lysodeikticus in the presence of lysozyme (Smolelis and Hartsell, 1949, 1952; Parry et al.,1969; Gorin et al., 1971). The clearing phenomenon is a complex process connected only indirectly with lysozyme's lytic action. Turbidimetric methods have the advantages of simplicity, rapidity and high sensitivity (Grossowicz and Ariel, 1983). 2.2 Applications of Lysozyme as a Food Preservative Most of the work on the use of lysozyme as a food preservative has been performed in Japan, where numerous patents have been granted for processes using lysozyme to extend shelf life of fresh fruits and vegetables, seafood, meat (Kanebo Ltd., 1973), tofu bean curd (Taiyo Food Co. Ltd., 1972), dried milk (Morinaga Milk Industry Co. Ltd., 1970), wine and sake (Eisai Co. Ltd., 1971), to name a few. However, the limited lytic spectrum of lysozyme, mostly effective against gram-positive bacteria, has prevented its use on a wide basis, as a general antimicrobial agent in foods. Currently, the most developed case of commercial use of lysozyme in food is its utilization by European cheese manufacturers to prevent "late blowing" in several varieties of semi-hard and hard cheeses of the Edam and Gouda type. Late blowing, a recurring problem of cheese production in many parts of Europe, is a defect caused by the butyric acid fermentation brought about by the germination and growth of clostridial spores, especially those of Clostridium tyrobutyricum, present in the cheese milk. Grass and maize silages, common feeds for dairy cattle have been recognized as the major sources of contamination of raw milk with the pasteurization-resistant spores of C. tyrobutyricum (International Dairy Foundation, 1987). 11 Edam cheese prepared with 500 U/ml (about 20 ppm) of lysozyme from 25L of milk containing approximately 103 C. tyrobutyricum spores/ L , had after five weeks of ripening resisted late blowing, whereas the control prepared without lysozyme was no longer suitable for consumption (Wasserfal et al., 1976). Wasserfall and Teuber (1979) demonstrated that lysozyme concentrations of 500 U/ml (about 20 ppm) killed 99% of 5x10s resting vegetative cells of C. tyrobutyricum within 24 hours of incubation at 25°C. Spores were completely resistant but the overall germination of spores into vegetative cells was delayed by 1 day in the presence of lysozyme. The authors concluded that some steps in the conversion of spores to vegetative cells must be sensitive to lysozyme. Birkkjaer et al., (1982) reported that addition of lysozyme at a dose of 1-2 g/lOOL milk was as effective as potassium nitrate added at lOg/lOOL milk for preventing late blowing of Danbo cheese due to C. tyrobutyricum. They reported that 99% of the lysozyme added to the cheesemilk was precipitated with the casein and found its way into the cheese, where it was not inactivated during the ripening process. The addition of lysozyme to cheesemilk prior to rennet addition, at the current usage levels of 20-25 mg/L milk (20-25 ppm), although sufficient to inhibit outgrowth of Clostridia does not normally interfere with the growth of the lactic inoculum, but in some instances, strains of suitable culture resistant to lysozyme must be selected (Scott et al., 1987). Lysozyme at levels of up to 50 ppm has been shown to be more detrimental to the growth of spoilage and pathogenic bacteria in milk than to that of the lactic-acid producing bacteria (Bottazzi et al., 1978; Akashi, 1972). In some European countries which have prohibited or restricted the use of nitrates in foods due to health concerns, the use of lysozyme has proved to be an efficient alternative in 12 the control of late blowing. It is estimated that about 20-25% of the Emmental cheese made in France is produced with lysozyme addition at 25mg/L (Cretin-Maitenaz, 1985). The major obstacles to a wider use of lysozyme in the countries which have authorized its use in cheese, seem to be its cost (Wasserfall and Teuber, 1979; Banks et al., 1986) and its limited effectiveness at current authorized usage levels (maximum 25-30 mg/L of cheesemilk), to prevent late blowing when the C. tyrobutyricum spores content is too high, exceeding levels of 103 per litre of cheesemilk (International Dairy Federation, 1987). The antimicrobial activity of hen egg-white lysozyme against bacteria involved in foodbome diseases has also been reported by several researchers. Hughey and Johnson (1987) showed that certain strains of C. botulinum and four strains of Listeria monocytogenes were effectively killed by hen egg-white lysozyme in culture media and in buffer. Their study demonstrated that the food spoilage thermophiles Clostridium thermosaccharolyticum and Bacillus stearothermophilus were also highly susceptible to lysozyme. In an extension of this study, Hughey et al., (1989) showed that 100 ppm lysozyme killed or prevented growth of L. monocytogenes Scott A in several foods inoculated with 104 cells/g of food sample. Lysozyme was more active in vegetables (fresh corn, green beans, cabbage, lettuce and carrot) than in animal-derived foods (fresh pork sausage and Camembert cheese) that were tested. In both of the previous studies it appeared that lysis of the pathogenic and spoilage bacteria was enhanced when lysozyme was used in conjunction with EDTA. This was particularly apparent with certain strains of C. botulinum, many of which were completely resistant to lysozyme alone but were inhibited and lysed by the combined use of lysozyme and EDTA. 13 Bactericidal properties of lysozyme alone and in combination with other antimicrobials were also studied by Proctor (1991), in-vitro and in hotdogs and hamburger against the pathogens Salmonella, L.monocytogenes and Staphylococcus aureus. Lysozyme or nisin alone were found to be bactericidal against L. moncytogenes in-vitro, and their use in tandem had a synergistic bactericidal effect in hotdogs. Lysozyme or nisin alone did not reduce S. aureus counts on inoculated hotdogs, but their combined use did show a bactericidal effect, which was further enhanced with the addition of EDTA. Although the mechanism of the observed interaction of lysozyme with chelators such as EDTA has not yet been elucidated, two explanations have been suggested: (i) the chelator may slow growth by binding an important metal necessary for growth, thus allowing lysozyme to digest the cell wall faster than it can be synthesized, or (ii) it may disrupt the cell wall structure thus enabling better attack by lysozyme (Hughey et al., 1989). The higher effectiveness of using egg-white lysozyme in conjunction with other chemical or physical preservative methods has also been recognized by Yang and Cunningham (1993), as well as by Banks et al., (1986). The latter conclude in their review, that the control of the homeostasis of all the organisms present in a food's mixed flora is rarely achievable by one bactericidal agent alone, and rather recommend the combined use of two or more antimicrobial agents because of their synergistic effects. 14 2.3 Thermal Stability of Lysozyme 2.3.1 Reversible Denaturation Measured by DSC A protein is ordinarily in an equilibrium between the folded (N) and unfolded (U) states. This equilibrium, which under physiological conditions strongly favors the folded state, shifts towards the unfolded state at high temperatures, extreme pHs and high concentrations of denaturants (Yamada et al., 1993). With an increase in temperature, all enzymes eventually lose their catalytic activity. This thermal inactivation however, may be either reversible or irreversible, depending on whether return to ambient temperature is accompanied by a return of enzymatic activity within a reasonable period of time (Klibanov, 1983). The overall process can be depicted by the classical scheme of Lumry and Eyring (1954): N~ U - I where N is the native enzyme, U the reversibly unfolded and inactive enzyme, and I the irreversibly inactivated enzyme. The mechanisms involved in the first step, referred to as thermal denaturation or melting of proteins are thoroughly investigated and well understood (Kauzmann, 1959; Tanford, 1968; Privalov, 1979). However, the pathways and mechanisms of irreversible thermoinactivation of enzymes, remain obscure, due to the involvement of numerous chemical reactions (Tanford, 1968), and because of severe conceptual and experimental problems encountered in their investigation (Ahern and Klibanov, 1985). Most of the information available on the thermal stability of lysozyme has been obtained through thermodynamic studies of thermal unfolding of this enzyme. Among techniques available for studying the thermodynamics of protein stability, differential scanning calorimetry (DSC) is the one that has been the most widely used. 15 DSC, which is a technique of thermal analysis, is defined by the International Conference for Thermal Analysis (ICTA) as: "a technique in which the difference in energy inputs into a substance and a reference material is measured as a function of temperature while the substance and reference material are subjected to a controlled temperature programme" (MacKenzie, 1979). Thermal denaturation of proteins is a highly cooperative phenomenon, which is recorded as a differential heat flow displayed as an endothermic peak on the DSC thermogram, since the disruption of intramolecular hydrogen bonds that occurs during the transition from the native to the unfolded state is an endothermic reaction (Ma et al., 1990). Two important parameters are obtained from the analysis of a DSC thermogram: temperature of denaturation which can be estimated from the transition, generally as the temperature of maximal excess heat flow ( T ^ or the peak temperature (Tp), and enthalpy of denaturation (AH). The AH value, calculated from the area under the transition peak, is actually a net value resulting from the combination of endothermic reactions, such as the disruption of hydrogen bonds, and exothermic reactions, including protein aggregation and the breakup of hydrophobic interactions (Ma and Harwalkar, 1991). The enthalpy of denaturation provides an estimate of the thermal energy required to denature the protein. 2.3.2 Factors Affecting Thermal Stability of Lysozyme The various parameters that affect the thermal stability of lysozyme have been studied in terms of changes in the thermodynamic parameters of the unfolding process. The major role played by hydrophobic interactions in the stabilization of the native conformation of proteins has been known for many years (Kauzmann,1959; Tanford, 1968; Dill, 1985; Dill et al.,1989). 16 After determining the melting points of lysozyme and 13 other proteins, Bull and Breese (1973) compared them to calculated hydrophobic indices and to average residue volumes of the same proteins. They observed a strong positive correlation between the denaturation temperatures and the calculated parameters, and suggested that the hydrophobic index might be associated with the unfolding transition of the proteins since hydrophobic residues tend to have large volumes and therefore increase the free energy associated with the unfolded state. With the development of genetic engineering techniques, further insights into the role of hydrophobic interactions in protein stabilization have been provided by studies on different mutant proteins including lysozyme, obtained by site-directed mutagenesis. Matsumura et al., (1988, 1989a) observed a direct relationship between thermal stability and hydrophobicity of amino acids substituted at position 3 in the lysozyme of bacteriophage T4. In the wild type enzyme position 3 is occupied by an De residue which contributes to a major hydrophobic core of the C-terminal domain and links it to the N-terminal domain. Any change in the hydrophobicity of the amino acid at this position is therefore expected to affect the overall stability of the protein. Thermodynamic analysis of the thermal unfolding of 13 variants at position 3 revealed a linear relationship between the stability of the enzyme and the hydrophobicity of the substituted amino acids. In a scanning calorimetric study of the thermal denaturation of the lysozyme of phage T4 and of its mutant form Arg96-His, Kitamura and Sturtevant (1989) showed that at any temperature within the range -20 to 60°C, the free energy of unfolding of the mutant form is more negative than that of the wild type by 3-5 kcal.mol'1. The possible causes of the decrease in the free energy of unfolding brought about by the replacement of arginine by histidine were 17 discussed in detail by Weaver et al., (1989). These authors attribute the apparent destabilization to the loss of a helix-dipole interaction between the side chain of Arg96 and the C-terminal end of helix 82-90, and to significant strain caused by the introduction of the imidazole ring of histidine. Alber et al., (1987) compared the structure and the thermodynamic stability of phage T4 lysozyme and 13 of its mutants in which the Thr at position 157 had been substituted with other amino acids. In the wild type lysozyme the Thr-157 forms a hydrogen bonding network around position 157, as revealed by high-resolution X-ray analysis. A positive correlation was observed between thermal stability of the different variants and the ability of the substituted amino acids to maintain these hydrogen bonds. The most stable variants (Asn, Ser, Asp) were found to be those containing hydrogen bonds similar to those of the wild type, whereas a significant decrease in thermal stability was observed in the variants where the substituted amino acid did not form hydrogen bonds. A good example of the latter is the variant in which the substitution of Thr at position 157 by He, resulted in a decrease of the free energy of unfolding by 12.1 kJ.mol"1 and a reduction of the melting temperature by 11°C, as compared to the wild type lysozyme (Alber et al., 1987). A rational approach to protein stabilization by introducing specific amino acid substitutions which decrease the configurational entropy of unfolding was proposed by Matthews et al., (1987). The two amino acid substitutions, Gly-77- Ala and Ala-82- Pro in Phage T4 lysozyme, which are expected to restrict the number of configurations available to the backbone of the unfolded molecule, were shown to stabilize the enzyme by contributing each about 4.18 kJ.mol"1 to the free energy of stabilization. The contribution of new disulfide bonds introduced by site-directed mutagenesis to the 18 thermal stability of T4 lysozyme has also been the subject of numerous studies (Perry and Wetzel, 1984, 1986; Wetzel, 1987; Wetzel et al., 1988; Matsumura et al., 1989b). The wild type enzyme contains no disulfide bonds but has two free Cys residues at positions 54 and 97. The substitution of an He residue at position 3 with a Cys, results in the formation of a disulfide bond between the latter and Cys-97. The engineered Cys3-Cys97 disulfide bond was reported to stabilize the enzyme against reversible thermal unfolding and against irreversible thermal inactivation, although the mechanisms of stabilization seemed unrelated (Wetzel et al., 1988). Mutants which in addition to the introduced disulfide bond also contained temperature-sensitive mutations and were therefore less stable against reversible thermal unfolding, compared to the wild type or other uncross-linked variants, were actually more stable against irreversible thermal inactivation (Wetzel, 1987; Wetzel et al., 1988). Wetzel (1987) concluded that the thermodynamic stability provided by a disulfide bond can not be confined to the conformational entropy effect, since strained disulfides, although tolerated within the structure of globular proteins, can reduce or overwhelm the stabilization resulting from the decrease in conformational entropy. Furthermore, the stabilizing effect of a disulfide bond against irreversible inactivation is not always directly correlated to its stabilizing effect against reversible unfolding. A disulfide bond reducing the free energy of stabilization might stabilize the enzyme against thermoinactivation by reducing the rate of irreversible processes, such as aggregation, whereas another disulfide bond with a different bonding geometry, could provide free energy of stabilization to an ideal unfolding equilibrium without stabilizing against irreversible inactivation processes (Wetzel, 1987). New disulfide bonds introduced by site-directed mutagenesis at four different locations 19 in a Cys-free T4 lysozyme by Matsumura et al., (1989b) produced different effects on the thermal stability of the enzyme illustrating the importance of the exact bonding geometry, as stated by Wetzel (1987). Two of the variants, which were linked by a disulfide bond at positions 9-164 and 21-142 were reported respectively to have melting temperatures 6.4 and 11.0°C higher than the wild type enzymes, whereas the two mutants with disulfides between positions 127-154 and 90-122 either destabilized or had no effect on the stability of the enzyme (Matsumura et al., 1989b). Treatment of proteins with bifunctional reagents have been shown to result in the formation of either intermolecular or intramolecular cross-links with a stabilizing effect on the proteins, which has been attributed to the introduction of additional physical constraint to unfolding and the rigidification of the native conformation (Klibanov, 1979; Mozhaev and Martinek, 1984). Johnson et al., (1978) determined the thermodynamic parameters of the thermal unfolding of iodine-treated lysozyme, which was characterized by a cross-link in the form of an ester bond created between Glu-35 and Tip-108. The melting temperature of the enzyme was reported to have increased by 29.4°C, as a result of the formation of the cross-link, which corresponded to a 21.8 kJ.mol1 increase in the free energy of denaturation. The effect of additives such as salts, sugars and polyhydric alcohols on thermal stability of lysozyme have also been the subject of numerous DSC studies. Back et al., (1979) investigated the effects of various sugars and polyols on the temperature (T^ of the maximum rate of denaturation of lysozyme and 3 other proteins. Addition of a sugar or polyol at 50% w/w raised Tm of all four proteins, with the magnitude of the stabilizing effect (ATm) depending on both the nature of the protein and the nature of the sugar or polyol. At pH 3, 20 sucrose, glucose, sorbitol and glycerol raised T m of lysozyme respectively by 15.0, 17.0, 18.5 and 8.5°C. The dominant mechanism by which sugars and polyols stabilize proteins against heat denaturation was suggested to be through their effect on the structure of water, which in turn determines the strength of hydrophobic interactions (Back et al., 1979). Gekko (1982) determined the thermodynamic parameters for the thermal unfolding of lysozyme in the presence of different polyols. He reported an increase of the melting temperature and of the free energy of unfolding with increasing concentration of polyols, and suggested that the protein stabilization was caused by a solvent medium effect or solvent ordering effect. Studying the effects of polyethylene glycol (PEG) on preferential hydration and thermal stability of lysozyme and 3 other proteins, Lee and Lee (1987) found that lysozyme's thermal transition temperature was decreased in the presence of PEG. This result was explained by the hydrophobic characteristics of PEG. Being partially hydrophobic in nature, PEG would interact favorably with hydrophobic side chains made available when the protein is unfolded, leading to a preferential stabilization of the unfolded state (Lee and Lee, 1987). The magnitude of the decrease in the thermal transition temperature of proteins induced by PEG was found to be related to the hydrophobicity of the proteins involved, as evidenced by a linear relation found between AT m and average hydrophobicity. In a series of papers Pfeil and Privalov (1976a, 1976b) presented a thermodynamic description of lysozyme using a direct method for obtaining thermodynamic standard functions of single states of proteins. Standard functions of enthalpy (H), entropy (S) and the Gibbs free energy (G) of native and denatured lysozyme were reported in the range of 0-100°C and pH 1.5-7.0. The change in the Gibbs energy of denaturation reached 16 kcal.moi"1 at conditions 21 of maximal protein stability (0°C, pH 4.5-7.0) and 14.5 kcal.moi"1 at 25°C and neutral pH (Pfeil and Privalov, 1976b). 2.3.3 Irreversible Thermoinactivation Factors which help stabilize a protein against reversible unfolding will also make it more resistant to irreversible denaturation by shifting the equilibrium toward the native conformation (Kristjansson and Kinsella, 1991). In recent years, numerous studies have been undertaken in an attempt to elucidate the mechanism of irreversible thermal inactivation of lysozyme (Ahern and Klibanov, 1985; Volkin and Klibanov, 1987; Tomizawa et al., 1994, 1995a, b). Ahern and Klibanov (1985), determined inactivation rate constants of hen egg-white lysozyme at 100°C in the pH range of 4.0-8.0. In order to distinguish between conformational mechanisms of inactivation and those of a covalent nature, they used concentrated solutions of strong denaturants such as guanidine hydrochloride or acetamide, which are known to disrupt noncovalent interactions in proteins and to reduce disulfide bonds. Irreversible thermal inactivation of lysozyme followed first order kinetics at pH 4, 6 and 8 (Figure 2), and the time course of inactivation was independent of the initial enzyme concentration over several orders of magnitude (Ahern and Klibanov, 1985). The processes causing irreversible inactivation of lysozyme at 100°C were reported to be deamidation of asparagine residues, hydrolysis of peptide bonds on the carboxyl side of aspartic acid residues, destruction of disulfide bonds and formation of incorrect (scrambled) structures, the relative contributions of which depend on the pH. At pH 4, hydrolysis of peptide bonds at Asp residues and deamidation of Asn residues 22 T i m e lhours ) Figure 2. The time course of irreversible inacuvation of lysozyme at 100°C as a function of pH and the enzvme concentration. (Curve a) pH 4 (0.1M sodium acetate), (curve b) pH 6 (0 .01M sodium cacodvlate), and (curve c) pH 8 (0.1M sodium phosphate). The concentrations (A*M) of Ivsozvme were O. 1000: • . 100: • . 50: • . 10: ±. 5: * . ! ; « . 0.5 (Ahem and Klibanov, 1985). 23 Table 1. Rate constants of irreversible inactivation of lysozyme at 100°C ( Ahern and Klibanov, 1985). Rate constant fhour'h Irreversible thermoinactivation pH4 pH6 pH8 Directly measured overall process* 0.49 4.1 50 Due to individual mechanisms Deamidation of Asn residues1" 0.45 4.1 18 Hydrolysis of Asp-X peptide bonds* 0.12 0 0 Destruction of cystine residues5 0 0 6 Formation of incorrect structures1 0 0 32 *Determined from Figure 2. TJetermined by nonequilibrium polyacrylamide gel electrophoresis. TJetermined by SDS gel electrophoresis of the reduced enzyme. 'Determined by titration of SH groups formed upon reduction of cystine residues. TJeterrnined as the difference between the time courses of irreversible thermoinactivation of lysozyme in the absence and in the presence of 8M acetamide. 24 accounted alone for the thermal inactivation at 100° C (Table 1), and deamidation which was the sole inactivation process at pH 6 showed an increase in rate with increasing pH. Tomizawa et al., (1994) reexamined the mechanism of irreversible inactivation of lysozyme at pH 4, 100°C and concluded that irreversible thermoinactivation was the result of the accumulation of several chemical reactions including isomerization of Asp-Gly, deamidation of Asn, racemization of Asp and Asn, and cleavage of Asp-X peptide bonds. Although Ahern and Klibanov (1985) had reported that deamidation of Asn and/or hydrolysis of Asp-X peptide bonds were the causes of the irreversible inactivation of lysozyme at pH 4, 100°C, Tomizawa et al., (1994) did not completely agree with their conclusion. By comparing the extent of deamidation of denatured and non-denatured lysozyme, using continuous acid PAGE in the presence of 8M urea they found no gap that would indicate a difference in the extent of deamidation, with both fractions showing only some monodeamidated bands. Similarly, a comparison of cleavages of peptide bonds, carried out using SDS-PAGE after reduction of the non-denatured and denatured lysozymes with mercaptoethanol, indicated that hydrolysis of Asp-X peptide bonds could not be the single cause of irreversible inactivation of lysozyme at pH 4, 100°C (Tomizawa et al., 1994). Tomizawa et al., (1995a) also re-examined the mechanism of irreversible inactivation of hen egg-white lysozyme at neutral pH (6 and 8) and 100°C, and investigated effects of additives (copper ions and organic reagents) on the inactivation. They concluded that the inactivation of lysozyme at neutral pH and 100°C was mainly caused by intermolecular and intramolecular disulfide exchanges and the production of irreversibly denatured lysozyme, which was destabilized by multiple chemical reactions other than disulfide exchange. In 25 addition, Tomizawa et al., (1995a) reported that independently of the latter, deamidation slightly affected the inactivation by causing a decrease of electrostatic interaction between positive charges of lysozyme and negative charges of the bacterial cell wall. As for the effects of additives on the thermal inactivation, the stabilizing effect of a small amount of copper ion was found to be due to the suppression of disulfide exchange by catalyzing air oxidation of heat-induced trace amounts of free thiols. Organic reagents (acetamide, ethanol, and glycerol), in addition to suppressing intermolecular disulfide exchange by decreasing hydrophobic interactions, were also reported to change the mechanism of the inactivation to that under acidic conditions by shifting the pK„ values of dissociable residues. Finally, precipitate production due to protein aggregation and the rate of loss of enzymatic activity, monitored over a wide range of lysozyme concentration (0.2-4.0 mg/mL), were shown to increase with increase in protein concentration. Having elucidated the mechanism of irreversible inactivation of lysozyme at pH 6 and 100°C, in another study Tomizawa et al., (1995b) investigated the possibility of stabilizing lysozyme against irreversible thermoinactivation by the addition of various salts and trifluoroethanol (TFE), which were expected to suppress chemical reactions such as racemization of Asp and/or Asn as well as deamidation of Asn. It was confirmed that salts and TFE suppressed lysozyme inactivation at pH 6 and 100°C, and was found that the addition of phosphate ions accelerated lysozyme inactivation as well as the rates of deamidation of Asn-Gly and racemization of Asp and/or Asn in lysozyme. As illustrated in this literature review, most if not all of the studies on the irreversible thermoinactivation of lysozyme were motivated by the elucidation of the mechanisms 26 involved, which have not yet been as thoroughly investigated and well understood as the pathways of reversible thermal denaturation of lysozyme. Since lysozyme has a rather high thermal stability, and the covalent processes suspected to be involved in irreversible inactivation are more likely to occur at higher temperatures, all of the previously cited studies were carried out at 100°C. Proteins can be exposed to a wide range of pH conditions and/or high temperatures during processing and storage of food. The objective of this study was to determine the irreversible thermal inactivation kinetics of lysozyme rather than the details of the mechanisms involved, and a rather wide range of pH (4.2-9.0) and temperature (73-100 °C) conditions were chosen for carrying out our experiments. 2.4 Use of Lysozyme as an Antimicrobial in Beer Despite the poor nutrient status of beer, its low pH (3.8-4.7), the antiseptic action of hops and the presence of ethanol, a wide range of bacteria and wild (non-brewing) yeasts still have the ability to grow in finished beers. Growth of these "non-brewing" microorganisms can result in physical, chemical and organoleptic alterations of beer, rendering the latter unsuitable for consumption. The lactic acid bacteria of the genera Lactobacillus and Pediococcus are the most troublesome of the beer spoilage bacteria. These bacteria, which thrive during the oxygen deprived stages of fermentation and storage, usually come from the malt dust or the air in the brewery (Dowhanick, 1990). Brewery lactobacilli are gram-positive, heterofermentative microorganisms that produce lactic acid, acetic acid, carbon dioxide, ethanol and diacetyl. Spoilage by lactobacilli is characterized by a "buttery" (butterscotch) flavor of diacetyl and 27 a "silky" turbidity (Priest, 1987). Regarding spoilage of beer by Pediococci, P. damnosus is undoubtedly the most common and feared Pediococcus encountered in North American breweries. Unlike lactobacilli, it is interesting to note that this organism is apparently only found in beer, brewing yeast and wines, and not in brewing raw materials or plant materials, suggesting considerable adaptation to the particular habitat provided by beer (Priest, 1987). Spoilage by P. damnosus is characterized by a "buttery" aroma of diacetyl and ropiness in beer, due to the production of extracellular slime consisting of a complex heteropolymer made of glucose, mannose and nucleic acid (Hough et al.,1982). In order to destroy all expected biological contaminants in filtered beer, brewers rely on pasteurization either before or after bottling. Although the temperature/time units of beer pasteurization are relatively low compared to those for milk or canned foods, pasteurization does alter the organoleptic attributes of beer. Thus many breweries have switched to sterile filtration, using membranes with very small pore sizes (0.45 ^m) to remove the bacteria. Sterile filtration of beer has become possible due to significant developments in membrane technology in recent years. However, according to Reid et al., (1990) the advantages gained by the sterile filtration of beer as opposed to pasteurization, such as improved flavor, are to some extent outweighed by the additional controls that are necessary to ensure microbiological stability and filterability. But the two most important drawbacks of sterile filtration are the high cost of membranes and their relatively low load capacity. Since the lactobacilli and pediococci responsible for spoilage of beer are gram-positive, and since lysozyme has the ability to lyse a wide range of bacteria, particularly gram-positive 28 bacteria, the idea of using lysozyme to prevent or delay spoilage in beer seems to be quite appealing. Although there is no information in the literature on the use of lysozyme in beer, a number of studies and patents suggest that lysozyme can be quite valuable in the preservation of wine, sake and mirin. In a Japanese study by Uchida et al., (1972) quoted by Cunningham et al., (1991) on the inhibitory effect of egg-white lysozyme on growth of lactobacilli from mirin, the growth of Lactobacillus heterohiochii, L. fermenti, L. plantarum and L. casei was completely inhibited by 20 ppm lysozyme. In another study by Yajima et al., (1968) quoted by Proctor and Cunningham (1988), lysozyme was shown to have a strong inhibitory effect on hiochi bacteria (sake putrefying lactic acid bacteria) in sake. The minimum concentrations of lysozyme needed to inhibit growth of bacteria were reported to be 10 ppm for Lactobacillus heterohiochii, 20 ppm for L. homohiochii, 1.25 ppm for L. fermenti and more than 100 ppm for L. acidophilus. As for the stability of lysozyme in the presence of ethanol, in a series of experiments on the stability of lysozyme over time in various chemical, food and pharmaceutical products, Proctor (1991) reported that lysozyme in light beer (Coors light, pH 4.48) remained fairly stable, losing only 20% activity in one year. In the light of the information presented here, it appeared that beer, as a product that typically undergoes pasteurization, would be an ideal candidate in which to test lysozyme's potential to prevent or delay the growth of spoilage bacteria. 29 3. MATERIALS AND METHODS 3.1 Thermal Inactivation of Lysozyme in Aqueous Buffers 3.1.1 Reagents Lysozyme from chicken egg white was purchased from Sigma Chemical Co. (St. Louis, MO). The 3X crystallized, dialysed and lyophilized lysozyme had a specific activity of 39,000 U/mg solid (41,400 U/mg protein). Lyophilized cells of Micrococcus lysodeikticus were also obtained from Sigma Chemical Co. Sucrose, sodium chloride, sodium hydroxide, potassium chloride, potassium phosphate (KH2P04), disodium hydrogen orthophosphate (Na2HP04), citric acid monohydrate (C 6H 80 7.H 20) and boric acid used for preparing the different buffers were all of reagent grade. 3.1.2 Substrate Preparation Fresh M. lysodeikticus solutions (about 1.5 mg lyophilized cells/10 mL 0.066M KH 2 P0 4 buffer, pH 6.24, initial A450=0.675) were made just prior to carrying out the lysozyme assays, since the turbidity of a micrococcus suspension tended to drop quickly upon storage at 3°C. 3.1.3 Lysozyme Assay Lysozyme activity was determined by one of the oldest and most widely used lysozyme assay methods, based on the reduction in turbidity of a suspension of dried M. lysodeikticus 30 cells in the presence of lysozyme. The procedure followed was the one described by Sigma Chemical Co.fTiuUetin 11-77 for Lysozyme Prod. No. L-6876), with few modifications. The lysozyme samples and the micrococcus suspension were equilibrated to 25° C in a water bath, prior to the assay. For each assay, O.lmL of the enzyme sample was added to 2.5mL of a micrococcus suspension, quickly but gently mixed by inversion, and placed immediately in the spectrophotometer. The time lag between the addition of the lysozyme sample and time "zero" was 23 seconds. Decrease in A 4 5 0 versus the reference buffer solution was recorded at 5 sec intervals for 50 sec, with a lag of 10 sec, using a Shimadzu UV 160 spectrophotometer. One unit of activity was defined as the amount of enzyme producing a change in A 4 5 0 of 0.001/min under these conditions. Lysozyme samples were assayed at least in triplicate. 3.1.4 Heat Treatment of Lysozyme Buffer composition and molarity are listed in Table 2. Buffer pH was determined at 25°C. Ionic strength (I) of buffers was not kept constant, but all lysozyme samples were assayed at pH 6.24 (1=0.07). Stock solutions of the enzyme were made by dissolving 4.3 mg of lyophilized lysozyme in 50 mL of appropriate buffer for each pH, corresponding to a specific activity of about 3400 Units/mL or 86 ppm. The initial concentration of lysozyme was doubled for the experiments carried out at pH 9. Heat treatments were carried out using a serum bottle technique similar to the one described by Mikolajcik and Rajkowski (1980). For each heat treatment, 9 mL of the appropriate buffer was pipetted into a 10 mL Supelco glass serum vial (Figure 3). The bottle was then closed with a Supelco rubber septum, sealed with a Supelco aluminum cap using a Table 2. Buffer composition for different pH values. pH Buffer 4.20 0.06M C 6H 80 7 (citric acid)/ 0.08M Na.HP04 5.20 0.04M C 6H 80 7 (citric acid)/ 0.1 IM Na^HP04 6.24 0.066M KH2P04 7.20 0.014M C 6H 80 7 (citric acid)/ 0.1M Na,HP04 8.10 0.05M Tris/ HC1 9.00 0.05M KC1/ 0.05M H3B03/ 0.02M NaOH 32 ALUMINUM SEAL RUBBER SEPTUM SERUM BOTTLE HEATING MENSTRUUM TCP VIEW OF ALUMINUM SEAL Figure 3. Diagram of the reaction apparatus used for the serum bottle technique. A top view of the pre-punctured aluminum seal is also shown. 33 Wheaton ( 224303 ) hand-operated crimper (Wheaton, Millville, NJ), and placed into an agitated water bath (Blue M Electric Company, IL) set at the temperature of the experiment. Temperature inside the vial was monitored with a 0.02 inch diameter (24 gauge) copper-constantan thermocouple probe (OMEGA Engineering Inc, Stanford, CT.) connected to a Dorec Minitrend 205 datalogger (Emerson Electronic Company, San Diego, CA.). Once the content of the vial reached the required temperature, 1 mL of lysozyme solution was injected into the vial through the rubber stopper, using a single-use Becton Dickinson sterile syringe fitted with a 25-gauge needle. Although this briefly lowered reaction temperature about 5°C, nominal temperature was re-established within 2 minutes. At selected time intervals, about 0.5 mL of sample was taken from the vial, using a separate sterile syringe and injected into an Eppendorf micro test tube placed in a water/ice bath. Samples were assayed for activity either the same day, or were stored at 3°C until assay time. For each time-temperature only one sample was taken to be assayed at least in triplicate. There were no replicates for the heat treatment since the heating of the medium in the agitated glass vials was homogeneous. 3.1.5 Calculation of Kinetic Parameters. Reaction order with respect to time (n,) of thermal inactivation of lysozyme at constant temperature was tested at intervals of 0.1 by the method of Durance et al., (1986). For predictive purposes, thermal destruction of lysozyme activity was treated as a first order reaction with respect to time. Decimal reduction times (D) were calculated from the negative inverse of the slope of the linear regression line of log lysozyme activity versus time at a 34 particular temperature: log(N0/N)= (t-to)/D (1) where N=activity at time t, N0=initial activity, t=time and t^ixritial time. Decimal reduction temperatures (z) were calculated from the negative inverse of the slope of log(D) versus temperature: log(D0/D)= (T-T0)/z (2) where D0=decimal reduction time at the initial temperature T 0 and D corresponds to the decimal reduction time at temperature T. D values were converted to thermal inactivation rate constants (k) at the same temperature: k= 2.303/D (3) The Arrhenius activation energy (EJ of the reaction at each pH and solute concentration was calculated, using the Arrhenius relationship: lnk= InA - (Ea/R)(l/T) (4) where R is the universal gas constant (1.987 cal.deg"1.mole1), E a is the activation energy (kcal.moi1), T is the temperature in °K and A is the frequency factor. Coefficients of determination (r2) were adjusted for sample size n as: adjusted r 2 = 1- ( l - r ^ n - l ) / ^ ) The effect of pH on thermal inactivation rates of enzymes has been described in theory as follows: lnk= ln(ko) - c(pH) (5) where ko is an intercept value, representing the reaction rate at pH=0 ( Stauffer, 1989). By combining (4) and (5), we obtained the following equation, allowing the prediction of k 35 at any pH and temperature T: ln(k)pHT=(E./R)((T-346.16)/346.16*T) + ta(kbW - cfeH) (6) By rearranging (6) we obtained: ln(k)= a + 6,/T + BjpH where: a= ES/(R*346.16) + lnfo)^^ (7) and fi,= -E./R (8) The reference temperature of 73°C (or 346.16°K) was chosen, because this was the lowest temperature used in our study. The statistics were performed using SYSTAT statistical program version 5.03. 3.2 Thermal Inactivation of Lysozyme in Commercial Beer Decimal reduction times (D) were determined at 4 different temperatures for lysozyme in beer, following the same procedure as in the first part of the project when using buffer solutions. The stock solution of lysozyme was made in beer, instead of buffer, and for each heat treatment 9 mL of beer was sealed into a serum type vial, to which 1 mL of the stock solution of lysozyme was added once the required temperature was reached in the vial. The brand of beer used was the local unpasteurized Okanagan Spring lager, with 5% alcohol and a pH of 4.7. 36 3.3 Inoculation of Beer With Lactic Acid Bacteria and Lysozyme 3.3.1 Bacterial Strains and Culture Conditions The bacterial strains used for this study were L. brevis strain B-12 and P. damnosus strain B-130, both from the Labatt Bacterial Culture Collection. The strains were shipped in MRS broth from the Labatt Brewing Research Department in London, Ontario. For maintaining the strains, upon arrival 0.8 mL of each strain in the original MRS broth was immediately stored at -86°C in 20% glycerol. A sampling of each strain was also subcultured in beer (Okanagan Spring lager, 5% alcohol) and incubated at 3 different temperatures ( 26°, 28° and 30° C ) for 24 hr, after which 0.8 mL aliquots of each subculture were also stored at -86°C in 20% glycerol. In order to revive the strains, primary cultures of L. brevis and P. damnosus were prepared by moculating few ice crystals from the freezer stocks into 40 mL of sterilized beer, and incubating at 28°C for 120 hr, as suggested by Roche et al. (1993). Secondary cultures were then prepared by moculating 1 mL of each primary culture into 40 mL of sterilized beer and incubating at 28°C. As can be seen in Figure 4, the two strains appeared not only to survive in beer at 28°C, but also to proliferate. They both exhibited typical microbial growth curves, characterized by the succession of a lag phase, during which they adapt to their new environment, an exponential growth phase, a stationary phase and a phase of decline (Jawetz etal., 1989). Sterilization of all equipment, microbiological media and beer was conducted at 121.1°Cfor 15 min. 37 Figure 4. Growth curves of Lactobacillus brevis B-12 and Pediococcus damnosus B-130 in beer. 38 3.3.2 Preparation of Stock Solutions of Lysozyme Three levels of lysozyme were tested: 0, 10 and 50 ppm. For each trial a fresh stock solution of lysozyme was prepared by dissolving 342 mg of lyophilized chicken egg-white lysozyme into 10 mL of sterilized beer. For the 50 ppm level, 0.5 mL of this stock solution was inoculated into each beer bottle with a final volume of 342 mL (341 mL beer + 0.5 mL lysozyme inoculum + 0.5 mL bacterial inoculum). For the 10 ppm level, the previous stock solution was diluted 5 times, and 0.5 mL of the diluted solution was used to inoculate each bottle. 3.3.3 Bottle Inoculation Procedure A standard curve of CFU/mL vs. Absorbance (Abs) at 600 nm was obtained for each strain, in order to help monitoring the growth of the cultures. For Abs. measurements, all dilutions were done in sterile 0.1 % peptone (Difco) water. Samples of the secondary cultures of L. brevis and P. damnosus were regularly taken for Abs. measurements as well as for agar plating. Once the counts reached 106-107 CFU/mL bottles were inoculated (Dowhanick and Sobczak, 1994). For each strain/day/lysozyme level a separate bottle of beer was used. Each bottle (at room temperature) was inoculated first with 0.5 mL of the appropriate lysozyme solution (except for lysozyme level 0), and then with 0.5 mL of the appropriate bacterial inoculum. The bottles were then partially deaerated by tapping, capped and inverted to ensure proper distribution of the microorganisms in the beer (Dowhanick and Sobczak, 1994). For day 0, 3 bottles (1 for each lysozyme level)/ strain were opened 15 min after being inoculated and 39 samples were taken for plating. The remaining bottles were transferred to an incubator and were incubated at 28°C until removal for plating on the appropriate day. 3.3.4 Plating for Colony-Forming Units On the appropriate days the bottles were gently swirled to resuspend the cells. After opening under aseptic conditions, an aliquot of 5mL was taken from each bottle, and serial dilutions were done in sterile 0.1 % peptone water. Plate counts were estimated by the drop plating method, dividing each plate into six equal sections and plating 20 /*L of sample per section. Each plate was done at least in triplicate. Incubations were on Universal Beer Agar (UBA, Difco ) + 0.1% polyoxyethylene-sorbitan monoleate ( Tween 80, Difco ) under microaerophilic conditions at 28°C, using Oxoid anaerobic jars model HP 11 with BBL CampyPak gas generating envelopes (Becton Dickinson). Incubation periods of 3 days for L. brevis plates and 5 days for P. damnosus plates, allowed the colonies to reach a size where they would be clearly visible, without too much aggregation taking place. 4 . RESULTS AND DISCUSSION 40 4.1 Effects of pH on Irreversible Thermal Inactivation of Lysozyme Lysozyme heat treatments usually ranged from a few minutes (e.g. 5 min, pH 9, 91 °C) to a few hours (e.g. 330 min, pH 5.2, 82°C). However the 75°C treatments at pH 4.2 and 5.2 required a 24 hr period, because of the remarkably high thermal stability of lysozyme at these pH values. The relationship between logarithm of lysozyme activity and incubation time at different temperatures appeared to be linear (Figures 5a-5f). Results were consistent with a pseudo first order reaction with respect to time (Table 4), although in most cases we did not continue the reaction to 80 to 93% completion, the range necessary to confidently claim an estimate of reaction order (Durance et al., 1986; Arabshahi and Lund, 1985). More complete reactions would have required longer heat treatments and preferably higher initial lysozyme concentrations than is typical of food applications (Proctor and Cunningham, 1988). Although first order did not always yield the best fit by regression of the linearized data, in most cases the best estimate of reaction order was close to 1 with an r2 for the regression very close to the one of a first order reaction. A substantial deviation from linearity did occur at pH 5.2, 75°C, where activity remained almost constant for 600 min before begixining to decline. This reaction was not sufficiently complete to meaningfully test reaction order. Stability at pH 4.2 and 5.2 was so high as to make precise estimates of slope difficult (Table 3). Experiments of several days would be required for better estimates. In preh'minary experiments at pH 6.24, lysozyme activity was measured immediately 41 Time (min) Figure 5a. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 4.20. 42 Figure 5b. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 5.20. 43 10 0 50 100 150 200 250 Time (min) Figure 5c. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 6.24. 44 Figure 5d. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 7.20. 45 Figure 5e. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 8.10. 46 Figure 5f. Thermal inactivation of dilute lysozyme in aqueous buffer at pH 9.00. 47 Table 3. Effect of pH on decimal reduction time (D) of lysozyme. pH Temperature (°C) D(min) ar* n k (IO*3 min1) 4.20 75 2.63x10s 0.97 37 0.88 82 1.08X103 0.95 38 2.14 91 5.56X102 0.89 17 4.15 100 1.86X102 0.97 22 12.4 5.20 75 7.14x10s 0.73 35 0.32 82 1.54x10s 0.91 29 1.50 91 4.29X102 0.84 15 5.37 96 3.95X102 0.95 17 5.83 6.24 73 1.47x10s 0.71 13 1.57 81 5.68X102 0.89 10 4.05 84 3.75X102 0.82 10 6.15 93 1.38X102 0.99 10 16.7 95 8.32x10' 0.95 36 27.7 7.20 75 4.20X102 0.98 18 5.48 82 1.42X102 0.98 25 16.2 91 6.27x10' 0.98 40 36.7 95 3.93x10' 0.98 27 58.6 8.10 75 9.81x10' 0.98 14 23.5 82 8.77x10' 0.86 23 26.3 91 6.00x10' 0.94 38 38.4 95 3.58x10' 0.98 28 64.3 9.00 75 1.40x10' 0.92 19 165 80 1.43x10' 0.96 29 161 85 9.73 0.98 22 237 91 4.30 0.84 30 536 a r adjusted for sample size n. Table 4. Reaction order with respect to time (n,) of the thermal inactivation of lysozyme, tested at intervals of 0.1 in the 0.5-2.0 range. pH Temperature (°C) Best estimate of nt "i^forn^l Reaction completion ( regression "r2) (%) 4.20 75 0.8-0.9 (0.975) 0.974 75 82 1.0-1.1 (0.950) 0.950 52 91 0.5 (0.904) 0.895 39 100 1.4-1.6 (0.983) 0.975 77 5.20 75 2.0 (0.780) 0.741 50 82 1.9-2.0 (0.920) 0.911 37 91 2.0 (0.889) 0.853 47 96 1.4-1.8 (0.957) 0.954 50 6.24 73 0.5-0.8 (0.731) 0.730 27 81 2.0 (0.919) 0.899 40 84 1.6-1.7 (0.846) 0.841 48 93 0.8 (0.993) 0.990 86 95 1.5 (0.966) 0.952 80 7.20 75 0.5-0.7 (0.985) 0.981 63 82 0.7-0.8 (0.986) 0.981 86 91 0.9-1.0 (0.981) 0.981 90 95 0.8-0.9 (0.981) 0.977 95 8.10 75 1.0 (0.982) 0.982 94 82 1.3 (0.888) 0.864 96 91 1.1 (0.940) 0.938 90 95 0.9-1.0 (0.984) 0.984 94 9.00 75 1.2 (0.942) 0.929 95 80 1.1 (0.961) 0.958 93 85 1.1 (0.982) 0.979 96 91 1.1 (0.853) 0.848 97 a r2 of linearized reaction equation 49 following heat treatment, and after refrigerated storage of 1 and 2 days. Activity did not change over 2 days of storage, from which we concluded that the reaction under study was essentially irreversible. The overall protein inactivation reaction has been described as: native protein « denatured protein - aggregated protein where the second step is irreversible (Myers, 1990). Denaturation of proteins, which is only the first step in heat inactivation, is a reversible phenomena and it is important to distinguish between denaturation and inactivation if one wishes to predict residual enzyme activity. Many DSC studies of the effects of pH and additives on thermal denaturation of lysozyme have been reported. But as mentioned earlier, these results are of limited practical value for prediction of residual lysozyme activity. While partial unfolding of the native protein seems to be the first universal step in enzyme thermoinactivation, the subsequent events are highly specific for individual enzymes (Klibanov, 1983). These events, which are responsible for the irreversibility of thermal inactivation of enzymes can be divided into two groups: covalent and non-covalent. Covalent changes are usually involved when extreme conditions (e.g. pH, temperature) have been used to inactivate an enzyme (Feeney, 1980; Nashev et al., 1977). Protein aggregation has often been suggested as the sole non-covalent mechanism responsible for irreversible thermal inactivation of enzymes. It occurs as unfolded protein molecules interact with each other through their exposed hydrophobic regions, in order to reduce the free energy of the unfolded molecules which are thermodynamically unstable (Kristjansson and Kinsella, 1991). In our experiments however, we did not observe any protein aggregation. This can probably be explained by the low level of lysozyme used, since aggregation is highly unlikely 50 in very dilute solutions of enzymes (Klibanov, 1983). The thermal inactivation of lysozyme occurring under relatively mild conditions of pH and temperature, where covalent transformations are unlikely, can be explained by the concept proposed by Klibanov and Mozhaev (1978). According to this model, upon cooling unfolded enzyme molecules can refold into new structures that are more stable thermodynamically, but that lack the catalytic activity of the native conformations. Experimental D values (Table 3) confirmed previous findings that lysozyme is more heat stable in acidic solutions (i.e. far from its isoelectric point of 10.7). Tables 3 and 8 show the dramatic effect of pH on the decimal reduction time of lysozyme. At 75°C, the time required to destroy 90% of the lysozyme activity rose from 2632 min at pH 4.2 to a maximum of 7143 min at pH 5.2, then dropped sharply to 14 min as the pH increased to 9.0. As discussed in the literature review, irreversible thermoinactivation of lysozyme is a rather complex process involving conformational (non-covalent) changes such as protein aggregation and formation of incorrect structures, as well as a host of various covalent reactions, the relative importance of which was shown to depend on the pH (Ahern and Klibanov, 1985; Volkin and Klibanov, 1987; Tomizawa et al., 1994, 1995a, 1995b). It is therefore difficult for us to relate the observed increase in the rate of irreversible inactivation as pH increased from 5.2 to 9.0, to specific chemical reactions, since we did not investigate the inactivation mechanisms that were involved. Ahern and Klibanov (1985), observed a similar pattern for the irreversible inactivation of lysozyme at 100°C, with inactivation rate constants of 0.49, 4.1 and 50 hr" \ respectively at pH 4, 6 and 8. These results are comparable to the results we obtained at pH values of 4.20 (observed k at 100°C of 0.74 hr"1) and 6.24 51 (estimated k at 100°C of 2.79 hr"1), but differ by one order of magnitude for the higher pH value since our estimate of k at pH 8.10, 100°C was 4.60 hr"1. Since for each pH condition Ahern and Klibanov also detennined the rate constants of specific chemical reactions that were suspected to be involved in the inactivation process, by comparing the latter to the rate constants of the overall inactivation reaction they were able to determine the relative importance of each indivdual mechanism. Another complementary explanation to help us interpret the present results, is provided by the observation that the formation of incorrect structures leading to irreversible enzyme inactivation was most prevalent as the pH of the solution approached the isoelectric point of the enzyme. This was suggested to reflect reduction of net charge on the protein that would otherwise hinder the incorrect refolding of the enzyme molecule (Ahern and Klibanov, 1986; Klibanov and Ahern, 1987). The Arrhenius relationship provides a means of summarizing the impact of temperature changes on reaction rate. Arrhenius activation energies (EJ of the thermal inactivation of lysozyme, as monitored by the lytic reaction of lysozyme on M. lysodeikticus, at different pH values, sodium chloride and sucrose concentrations are presented in Table 5. In thermal bacteriology, the alternative z concept is often used to describe destruction kinetics. When logarithm of D was plotted against temperature, the relationship was approximately linear (Figures 6a,6b). Therefore the decimal reduction temperature or z can be used to predict the impact of changing temperatures on D values. Back et al., (1979) indicated a DSC transition temperature (TJ of 71°C at pH 7 for lysozyme. With a z of 20.1 C° (Table 5) and equation 2, our estimate of the D at 71 °C was 664 min. This illustrates the limitations of using T m values from DSC studies to predict residual activity of enzymes. 52 Table 5. Effect of pH, sucrose and sodium chloride on the activation energy (EJ and decimal reduction temperature (z) of the thermal inactivation of lysozyme. pH Eab(kcal/mole) zb ( °C) Reference buffers 4.20 26.4 22.5 5.20 36.1 16.4 6.24 32.0 18.2 7.20 29.2 20.1 8.10 12.0 48.7 9.00 18.8 30.5 NaCrO.lM 5.20 30.0 19.3 7.20 32.1 18.1 9.00 5.24 111 NaCl'l.OM 5.20 12.5 46.5 7.20 41.8 13.9 9.00 18.9 30.8 Sucrose* 5% 5.20 30.1 19.2 7.20 36.3 16.0 9.00 27.6 21.0 Sucrose* 15% 5.20 42.0 13.8 7.20 41.1 14.1 9.00 21.6 26.9 * Solutes added to reference buffers. b Based on k or D at 4 different temperatures, except for reference buffer at pH 6.24 (5 temperatures) and solutes added to reference buffers (2 temperatures). 53 Figure 6a. Effect of pH (4.20-6.24) on decimal reduction time of lysozyme. 54 Figure 6b. Effect of pH (7.20-9.00) on decimal reduction time of lysozyme. 55 According to Stauffer (1989), plots of ln(k) versus pH and ln(k) versus 1/T should both be linear for heat inactivation of enzymes. Equation (6), a combination of equations (4) and (5), can be quite useful for predicting k within the pH and temperature bounds of this study. We found the relationship between ln(k) and the two independent variables 1/T and pH was close to linear (adjusted multiple 1^=0.975) in the pH range of 5.2-7.2, but less so above or below that range (Table 6). The regression equation obtained for this pH range and temperatures between 73°C and 100°C was: ln(k) p H >T= 32.90- 1.62xl04/T + 1.19(pH) It is possible to calculate E a from equation (8). A value of 32.38 kcal/mole was obtained, very close to our experimental average of 32.43 kcal/mole over the pH range of 5.2 to 7.2. Residual lysozyme activity after any thermal processing treatment within the pH and temperature bounds of our study can be estimated by either of these essentially equivalent procedures: equations (1) and (2) or equation (7) plus the Arrhenius relationship. However, since our experiments were performed in aqueous buffer solutions rather than in specific food systems, the data show only general trends without considering potential interactions between lysozyme and typical ingredients, other than sodium chloride and sucrose, present in foods. Proteins in particular may affect stability. Both z and E a values should therefore be used with caution. Given that the parameters were based on only four or five temperatures, confidence intervals are quite wide (Table 7). T-tests indicated no significant difference in either parameter from pH 4.2 to 7.2, although a significant increase did occur at pH 8.1 (P<0.05). Between pH 4.2 and 7.2 we recommend use of the average z of 19.3 C°. 56 Table 6. Regression output of the multiple regression analysis for the prediction of the reaction rate constant k of thermal inactivation of lysozyme as a function of pH and inverse temperature. Regression coefficients pH range n adjusted r2 constant pH l(fx(l/T) 5.2-7.2 13 0.975 32.90 1.19 -1.62 5.2-8.1 17 0.924 27.40 1.01 -1.39 5.2-9.0 21 0.923 23.00 1.24 -1.28 4.2-8.1 21 0.879 29.40 0.76 -1.40 4.2-7.2 17 0.875 34.00 0.75 -1.56 4.2-9.0 25 0.868 25.30 0.99 -1.29 57 Table 7. Confidence intervals (90%) for the decimal reduction temperature (z) and the activation energy (EJ of lysozyme at different pH values. pH n z(°C) confidence interval Ea(kcal/mol) confidence interval 4.20 4 22.5 M 18.6-28.5 26.4 20.7-32.1 5.20 4 16.4 M 10.7-35.0 36.1 18.1-54.1 6.24 5 18.2 M 16.5-20.5 32.0 28.2-35.9 7.20 4 20.1 0 5 16.1-26.6 29.2 22.7-35.7 8.10 4 48.7 * 27.3-222 12.0 2.30-21.7 9.00 4 30.5" 16.3-248 18.8 1.60-36.1 ns - Not significantly different (p>0.10) * p<0.05 58 In our study we did not investigate the effects of protein concentration and of the buffer salt on the thermal inactivation kinetics of lysozyme. Tomizawa et al., (1995a) showed that acetamide decreased the rate of lysozyme inactivation at pH 6 and 100°C, whereas Ahern and Klibanov (1985), indicated that acetamide did not affect the time course of irreversible thermoinactivation of lysozyme at pH 6 and 100°C. According to Tomizawa et al., (1995a) these opposite results could be partly explained by the differences in buffer type and protein concentrations. In their experiments they used 0.05 M phosphate buffer and a wide range of lysozyme concentrations (0.2-2.0 mg/mL), whereas Ahern and Klibanov investigated the inactivation under low lysozyme concentrations (< 0.7 mg/mL) in 0.01 M sodium cacodylate. Furthermore, Tomizawa et al., (1995a) reported that loss of lysozyme activity was dependent on the protein concentration, and did not show first-order kinetics, whereas according to Ahern and Klibanov (1985) lysozyme inactivation followed first-order kinetics with a time course independent of the initial enzyme concentration over several orders of magnitude. A more thorough investigation of the irreversible thermal inactivation kinetics of lysozyme should therefore include the effects of initial protein concentrations and of buffer salts. 4.2 Effects of Sucrose and Sodium Chloride on Thermal Inactivation of Lysozyme The effects of sucrose and sodium chloride on D and k values of lysozyme inactivation, are shown in Table 8 for the pH range of 5.2-9.0. At 75°C at pH vlaues of 7.2 and 9.0, increasing concentrations of sucrose had a stabilizing effect, which is consistent with DSC studies (Back et al., 1979; Kristjansson and Kinsella, 1991), in which sugars and polyols stabilize proteins against heat denaturation. Stabilization was ascribed to strengthening of the 59 Table 8. Effect of pH, sucrose and NaCl on thermal inactivation parameters D and k of lysozyme at 75°Cand91°C. 75°C 91°C pH solute D (min) •r5 k (lO^min-1) n D (min) k (lO^min-1) n 5.2 None 7.14x103 0.73 0.32 35 4.29x102 0.84 5.37 15 NaCl 0.1M 3.57X103 0.90 0.64 65 5.32X102 0.73 4.33 40 1.0M 1.89X103 0.80 1.22 65 8.55X102 0.57 2.69 35 Sucrose 5% 3.13X103 0.80 0.74 88 4.61X102 0.84 5.00 27 15% 5.88X103 0.75 0.39 60 4.08X102 0.87 5.64 22 7.2 None 4.20xl02 0.98 5.48 18 6.27x10' 0.98 36.7 40 NaCl 0.1M 5.32X102 0.96 4.33 21 6.92xl0l 0.98 33.3 33 1.0M 1.79X103 0.76 1.29 21 1.26X102 0.95 18.3 33 Sucrose 5% 5.78X102 0.97 3.98 30 5.76x10* 0.97 40.0 32 15% 8.00X102 0.95 2.88 23 5.87x10' 0.98 39.2 33 9.0 None 1.40xl0l 0.92 165 19 4.30 0.84 536 30 NaCl 0.1M 2.26x10' 0.83 102 23 1.62x10' 0.81 142 24 1.0M 4.24x10' 0.90 54.3 20 1.28x10' 0.93 180 20 Sucrose 5% 2.28x10' 0.74 101 22 3.96 0.81 582 22 15% 3.32x10' 0.87 69.4 21 8.44 0.95 273 15 * r adjusted for sample size n. 60 interactions between hydrophobic regions of lysozyme, induced by the effect of sucrose on the structure of water. According to Oakenfull and Fenwick (1979), hydrophobic interaction in aqueous-organic mixed solvents depends on solvent structure, with maximum hydrophobic interaction occuring in those solvent mixtures in which the 3-dimensional hydrogen-bonded structure of water is most developed. Timasheff and co-workers (Lee and Timasheff, 1981; Arakawa and Timasheff, 1982; Gekko and Timasheff, 1981a,b) who have studied extensively the stabilizing effect of sugars and polyhydric alcohols on proteins, explain the stabilizing effect in terms of preferential hydration of the protein in the presence of these additives. These authors relate the stabilizing effect of sugars to their effect on the surface tension of water, and suggest that the major factor in preferential hydration is the free energy required to form a cavity in the solvent, needed for accomodating the protein molecule. Since such a cavity would be larger for the unfolded protein because of larger surface area, the free energy associated with the unfolded state would be higher and therefore the folded state would be energetically favored. According to this theory sugars are preferentially excluded from the domain of the protein, which leads to unfavorable interactions between the protein and the solvent raising the free energy of the system, but more so for the unfolded state. We did not find a stabilizing effect of sucrose at 91" C in that there was no significant change of D values at the two levels of sucrose compared with no sucrose at all, except in one instance at pH 9.0 with 15% sucrose. This is logical, since hydrophobic forces are reported to weaken above 80°C (Myers, 1990). Sucrose levels were deliberately chosen at 5% and 15% to reflect levels found in processed foods. The DSC results were obtained using 50% w/w sucrose solutions. References to the impact of sugars on thermal inactivation are harder to find 61 but Yoshitake and Akabori (1977) as cited in Proctor (1991), also found increased thermal stability of lysozyme in sucrose syrups, with inactivation rates decreasing as the pH was lowered from 6.0 to 3.2 (Table 9). As for the effect of sodium chloride, our results show an increase in D values with increasing concentrations, except in one instance (pH 5.2 at 75°C), where sodium chloride caused a drop in D. A repeat experiment confirmed the original result at pH 5.2, 75°C. DSC studies of denaturation of lysozyme also indicate increased thermal stability with sodium chloride at "salting in" concentrations of 0.1 M to 1.0M, again suggesting that hydrophobic interactions contribute to increased stability (Myers, 1990). Salt at this concentration would also be expected to increase solubility, suppress aggregation and therefore increase stability even at temperatures not conducive to hydrophobic interactions. The effects of solutes on z and E a (Table 5) could not be tested statistically as these estimates were based on only two D (or k) values each, making conclusions hazardous. 62 Table 9. Stability of lysozyme at 20 ppm in concentrated sucrose solutions*. Temperature Time (min) Remaining lysozyme activity (%) pH 6.0 pH 3.2 20% sucrose 60% sucrose 20% sucrose 60% sucrose 15 89 98 100 100 60°C 30 85 93 100 100 60 80 92 100 100 15 53 100 100 100 80°C 30 53 100 100 100 60 35 94 100 100 15 16.5 32 82 98 100°C 30 0 15 67 79 60 0 6.8 25 46 * Adapted from Yoshitake and Akabori, 1977, quoted by Proctor, 1991. 63 4.3 Lysozyme's Thermal Inactivation Kinetics in Beer and Effect on Lactic Acid Bacteria The results of thermal inactivation of lysozyme in beer were consistent with a pseudo first order reaction with respect to time (Figure 7), showing an almost perfect linear relationship between logarithm of lysozyme activity and incubation time (r2 above 0.95). Decimal reduction times (D) were calculated from the negative inverse of the slope of the linear regression line of log lysozyme activity versus time at each temperature, and are presented in Table 10. It is mteresting to note that the D values found for lysozyme in beer, are slightly lower than the values which could be predicted in an aqueous buffer solution at a similar pH of 4.7, suggesting a greater sensitivity of lysozyme to thermal inactivation in a beer type environment. Proctor (1991), also reported a higher stability of lysozyme activity in ethanol solutions (5-15 % ethanol) than in ethanol containing food products such as light beer, white wine and cooking wine over a one year period. This illustrates the fact previously mentioned, that results obtained in buffer should not be simply extrapolated to foods, since food systems are much more complex in terms of environment. The decimal reduction temperature (z) of lysozyme in beer, calculated from the negative inverse of the slope of log(D) versus temperature was 23.15 C°. The results of CFU viability analyses of L. brevis and P. damnosus in beer containing lysozyme are presented in Table 11. Plating results for L. brevis, shown in Figure 8a seemed to indicate an initial drop in survival in bottles containing 10 and 50 ppm lysozyme. But after day 3, the survival trend in bottles with 10 ppm lysozyme was almost identical to the trend observed in bottles with no lysozyme, with counts reaching 106 CFU/mL on day 9, whereas 64 Figure 7. Thermal inactivation of lysozyme in unpasteurized beer (pH 4.70). 65 Table 10. Decimal reduction time at 4 different temperatures and reaction order of thermal inactivation of lysozyme in beer (pH 4.7). Temperature (°Q D'Cmin) n Best estimate of ntc (regression V) 82 179 0.958 21 0.8 (0.967) 87 113 0.952 19 0.8 (0.961) 91 86 0.960 22 0.9 (0.961) 95 47 0.965 23 0.8 (0.974) * Based on a first order reaction b r2 of linearized reaction equation, adjusted for sample size n c Reaction order with respect to time, tested at intervals of 0.1 over the 0.5-2.0 range. 66 the counts in the bottles with 50 ppm lysozyme remained at the day 3 level (103 CFU/mL) throughout the rest of the experiment. These results were suggesting that although the 10 ppm level of lysozyme did not have much inhibitory effect on L. brevis, the 50 ppm level seemed to exert some degree of inhibition. The results of a second trial (Figure 8b) suggested that in fact even 50 ppm lysozyme might not have the suspected inhibitory effect on L. brevis, since after a sharp initial drop in the survival curve, the CFU/mL counts in bottles with 50 ppm lysozyme were up again after day 12 and reached even higher levels than in control bottles with no lysozyme in the following days. However it seems that although the levels of lysozyme chosen in our study are too low to inhibit growth of L. brevis, they do delay spoilage of beer due to these bacteria. Similar conclusions were made by Bottazzi et al., (1978) who studied the effect of lysozyme on thermophilic lactic acid bacteria in milk. They found thatL. helviticus in milk was inhibited by greater than 50 ppm lysozyme. As for the results obtained with P. damnosus (Figure 9), it seems that there is little difference between the survival trend observed in the bottles with 10 ppm lysozyme and the one in the bottles with 50 ppm lysozyme. The two levels of lysozyme seem however to exert a slight inhibition on the growth of P. damnosus in beer. A more complete investigation of the effects of lysozyme on the beer spoilage bacteria used in this study, would obviously require the use of a wider range of lysozyme concentrations preferably including a much higher upper limit. There are a couple of possible explanations for the initial drop in CFU/mL counts followed by the sharp increase after a couple of days that was observed with both strains at 67 10 and 50 ppm lysozyme levels. Bacteria might be fixing itself preferentially on the glass surface. As a result of this microbial attachment, most of the bacterial growth would initially occur close to the bottle surface, and little or no growth would be detected in samples taken from other parts of the bottle, in spite of the fact that each bottle was shaken before sampling in an attempt to properly mix the beer. Once the colonies reach a critical mass, cell detachment from the biofilm results in the release of cell aggregates in the liquid medium. This would explain the sharp increase in CFU/mL counts observed after the initial drop. In order to minimize the biofilm effect, further experiments should be carried out in plastic containers instead of glass bottles, since microorganisms are less likely to form biofilms on plastic surfaces. Lysozyme could also have caused bacterial injury rather than death of the bacteria. This should be tested in future experiments by using a more challenging incubation medium for detection of uninjured bacteria. Table 11. Effect of lysozyme on survival of L. brevis B-12 and P.damnosus B-130 in beer. Counts (CFU/mL )* Lysozyme 0 ppm 10 ppm 50 ppm Day 0 1 3 9 16 P. damnosus B-130 1.48X103 (0.19X103) 2.80xl03 (0.57X103) 9.17x10" (0.85x10") 1.87x10s (0.31x10s) 2.20xl03 (0.25xl03) 1.50x10" (0) 1.18X104 (0.19xl04) 2.50x10" (0.71x10") 1.70x10s (0.25xl06) 3.33x10" (0.62x10") 1.33x10s (0.19x10s) 5.33x10" (1.25x10") 1.37xl04(0.17xl04) 2.33x10" (0.24x10") 1.42x10* (0.22X104) Day 0 1 3 9 16 L. brevis B-12 (Trial 1) 3.92x10s (0.13xl03) 5.37x10s (0.56x10s) 4.07x10s (0.06x10s) 1.00x10" (0) 4.22x10s (0.06x10s) 2.90x10s (0.11x10s) 5.37xl06 (0.28xl06) 5.52x10s (0.92x10s) 3.13x10s (0.49x10s) 3.25x10s (0.22x10s) 5.47x10s (0.56x10s) 0.50x10" (0) 2.42x10s (0.12x10s) 7.67x10" (1.70x10") 1.58x10s (0.12x10s) Day 0 3 6 9 12 15 21 I. brevis B-12 (Trial 2) 1.93X104 (0.18xl04) 1.23X104 (0.03X104) 1.08x10s (0.14x10s) 2.13x10" (0.74x10") 1.80xl06 (0.38xl06) 2.15x10s (0.11x10s) 2.05x10s (0.33x10s) 4.74X104 (0.24X104) 2.11x10s (0.22x10s) 3.08x10s (0.06x10s) 1.55x10s (0.15x10s) 2.20x10s (0.30x10s) 7.29X104 (0.32X104) 6.90X104 (0.32X104) 5.17x10s (0.47x10s) Nd' Nd* Nd* 8.87x10s (0.31x10s) 4.39xl06 (0.57xl06) 1.74x10* (0.19x10s) " Mean and standard deviation values (n^3) * Not detected 69 Figure 8a. Survival of Lactobacillus brevis B12 in unpasteurized beer in the presence of lysozyme (Trial 1). 70 8.00 Days Figure 8b. Survival of Lactobacillus brevis B12 in unpasteurized beer in the presence of lysozyme (Trial 2). 71 8.00 6.60 h 5.20 3.80 h 2.40 h 1.00 8 12 16 20 Days Figure 9. Survival of Pediococcus damnosus B130 in unpasteurized beer in the presence of lysozyme. 72 5. CONCLUSION Thermal inactivation of hen egg-white lysozyme in aqueous buffer solutions was investigated in the pH range of 4.2-9.0, and the inactivation kinetics were described in terms of decimal reduction times (D), decimal reduction temperatures (z), inactivation rate constant (k) and Arrhenius activation energy (EJ. Loss of lysozyme activity over time at different temperatures showed pseudo first order kinetics, although a more confident claim of reaction order in some cases would have required more complete reactions. Lysozyme appeared to be most stable at pH 5.2, and thermal stability decreased sharply as the pH increased to 9.0. A regression equation for prediction of k as a function of temperature and pH was derived, with the best fit obtained in the pH range of 5.2-7.2 (adjusted multiple r*= 0.975). At pH values of 7.2 and 9.0, low levels of sodium chloride (0.1M and 1.0M) clearly stabilized lysozyme against heat inactivation, while sucrose at levels of 5% and 15% had a stabilizing effect only at the lowest temperature, 75°C. Studies of thermal inactivation of lysozyme such as the one described here, provide a more practical approach for prediction of residual activity of lysozyme subjected to heat treatment than DSC studies, the results of which do not truly reflect inactivation events. Thermal inactivation kinetics of lysozyme and its potential to prevent or delay growth of spoilage bacteria were also studied in beer. Loss of lysozyme activity in beer followed pseudo first order kinetics with respect to time, and the decimal reduction temperature (z) of the enzyme was found to be 23.15 C°. 73 Lysozyme at 10 and 50 ppm did slightly delay growth of the spoilage bacteria L. brevis and P.damnosus in beer, but did not prevent growth. A more thorough investigation of lysozyme's potential to prevent growth of lactic acid bacteria in beer would require the use of a wider range of enzyme concentrations preferably including a much higher upper limit. 74 6. REFERENCES Ahern, T J . and Klibanov, A . M . 1985. The mechanism of irreversible enzyme inactivation at 100°C. Science. 228:1280-1284. Ahern, T J . and Klibanov, A . M . 1986. Why do enzymes irreversibly inactivate at high temperatures? In "Protein Structure, Folding and Design", D.L. Oxender (Ed), UCLA Symp. Moi. Biol. Cell. Biol., Vol. 39, p. 283, Liss, New York. Mvenainen, R., Heikonen, M . , Linko, M. and Linko, P. 1979. 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