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Comparison of efficacy of ozone and chlorine as antimicrobial agents against planktonic cells and biofilms… Nunez San Martin, Angela 1998

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COMPARISON OF EFFICACY OF OZONE AND CHLORINE AS ANTIMICROBIAL AGENTS AGAINST PLANKTONIC CELLS AND BIOFILMS OF SHEWANELLA PUTREFACIENS By Angela Nunez San Martin B. Sc. (Food Science), University of British Columbia, 1992 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF M A S T E R OF SCIENCE IN THE F A C U L T Y OF G R A D U A T E STUDIES Food Science We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA SEPTEMBER 1998 © Angela Nunez San Martin, 1998 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 scholarly 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 VqqJ SciPT)Ce The University of British Columbia Vancouver, Canada Da,e . ^ P f P f . a S /9U DE-6 (2/88) ii ABSTRACT The purpose of this study was to compare the efficacy of ozone and chlorine as antimicrobial agents against planktonic cells and biofilms of Shewanella putrefaciens. Shewanella putrefaciens is an intensive psychrotrophic spoilage organism of proteinaceous foods; in particular, fish. Because of the importance of this organism in the spoilage of seafood and the present use of ozone in some fishing boats, the antimicrobial tests were performed in cold environments characteristic of seafood processing plants and fish boats. For each planktonic and biofilm set of tests, there were 6 treatments to reflect a 3 x 2 factorial experiment. The factors were sanitizer type and temperature. The sanitizing agents employed were: no sanitizer (control), ozone and chlorine. The two experimental temperatures were 10° ± 0.5°C and 0.5° ± 0.5°C. Results of this study confirmed literature data that both chlorine and ozone are good biocidal agents. However, planktonic tests showed that ozonated distilled-deionized water at 0.25 ppm was more than 70 times more effective in inactivating Shewanella putrefaciens than distilled-deionized water containing 1.0 ppm available chlorine. Most of the microbial inactivation by either chlorine or ozone took place within the initial 30 sec. This inactivation was accompanied by a corresponding decrease in chlorine and ozone levels. After the initial decline, the surviving planktonic population remained stable. On the other hand, microbial inactivation data indicated that distilled-deionized water containing 1.0 ppm available chlorine was more effective than 0.25 ppm ozonated distilled-deionized water in inactivating Shewanella putrefaciens Ill biofilms grown on stainless steel chips. Biofilm populations exposed to either ozone or chlorine showed an initial sharp decline in their numbers within the initial 30 sec of exposure. After this period, the biofilm populations exposed to ozonated water declined at a slower rate. Biofilm populations exposed to 1.0 ppm available chloripe had a faster rate of cell inactivation as indicated by their steeper inactivation curves and final biofilm population. The tests also revealed that the indigo trisulfonate method may not be suitable to measure ozone concentrations in suspensions containing large number of organisms. Bacterial cells were shown to adsorb the indigo trisulfonate dye used to measure ozone concentration and thus cause incorrect determination of ozone concentrations. No mention of this problem had been found in searches of scientific literature. Thus, to overcome this major obstacle, the procedure used to determined ozone concentration had to be modified. In short, the interference caused by the bacteria and adsorbed indigo trisulfonate dye had to be removed prior to any determination of ozone concentration. This modified procedure to measure ozone concentration in microbial suspension was practical but it was also far from ideal. A new method must be derived Jhat has all the advantages of the indigo trisulfonate method but that does not interact with the microorganisms. This is very important given the reactive nature of ozone as well as its potential as a sanitizer not only in the food processing area but in fields such as aquaculture, wastewater and potable water treatments. IV TABLE OF CONTENTS Page Abstract i i Table of Contents iv List of Tables vii i List of Figures x Nomenclature xii i Acknowledgments xiv 1. INTRODUCTION 1 2. LITERATURE REVIEW 5 2.1 Basic Aspects of Ozone Chemistry 5 2.1.1 Ozone Generation 5 2.1.2 Chemical Nature of Ozone 7 2.2 Bacteriocidal Action of Ozone 11 2.3 Ozone Sanitation Studies 13 2.4 Sanitizing Efficiency of Ozone 17 2.5 Difficulties in the Determination of Ozone Concentration 18 2.6 Determination of Ozone Concentration by the Indigo Trisulfonate 25 Method 2.7 Basic Aspects of Chlorine Sanitation 26 2.7.1 Chemistry of Chlorine Sanitation 26 2.7.2 Hypochlorites 28 2.7.3 Free Available Chlorine 30 2.7.4 Inactivation of Bacteria by Active Chlorine Compounds 31 2.8 Sanitation by Chlorine Compounds 33 2.8.1 Chlorine Stability 33 2.8.2 Organic Matter 35 2.8.3 Concentration, pH and Temperature of Chlorine Solutions 35 2.8.4 Hard Water 37 2.8.5 Active Chlorine Compounds 37 2.8.6 Chlorine and Microbial Biofilms 42 2.8.7 Methods for the Determination of Available Chlorine 43 2.9 Microbial Resistance in Biofilms 44 2.10 Test Organism 51 2.10.1 Shewanella putrefaciens 51 V 3. MATERIALS A N D METHODS 57 3.1 Experimental Design 57 3.2 Experimental Procedures 60 3.2.1 Glassware 60 3.2.2 Water 60 3.2.3 Stainless Steel Chips 61 3.2.4 Planktonic Inoculum 62 3.2.5 Inoculation of Stainless Steel Chips 65 3.2.6 Growth of Biofilm 66 3.2.7 Ozone Generation and Measurement 68 3.2.8 Chlorine Generation and Measurement 69 3.2. 9 Planktonic Inoculum - Control 73 3.2.10 Planktonic Inoculum - Ozone 74 3.2.11 Planktonic Inoculum -Chlorine 79 3.2.12 Biofilm - Ozone and Chlorine 80 3.2.13 Planktonic Cells - Enumeration 81 3.2.14 Biofilm Cells - Enumeration 81 4. RESULTS A N D DISCUSSION 82 4.1 Decay Curves 82 4.2 Planktonic Suspensions - Control 82 4.3 Biofilm - Control 91 4.4 Planktonic Suspensions - Ozone 95 4.5 Biofilm - Ozone 100 4.6 Planktonic Suspensions - Chlorine 105 4.7 Biofilm - Chlorine 110 4.8 Statistical Analysis 117 5. CONCLUSION A N D RECOMMENDATIONS 131 6. REFERENCES v 136 7. APPENDIX 147 7.1 Problems Determining Ozone Concentration in Shewanella 147 putrefaciens Suspensions 7.2 Preliminary Ozone Sanitizing Tests - Table 13 147 7.2.1 Preparation of Shewanella putrefaciens Planktonic 147 Inoculum 7.2.2 Addition of Shewanella putrefaciens Planktonic 149 Inoculum to 0.5 ppm Ozonated distilled-deionized Water at 10° ± 0.5° C 7.2.3 Enumeration of Surviving Shewanella putrefaciens cells 149 Exposed to 0.5 ppm Ozonated distilled-deionized Water at 10° ± 0.5°C 7.2.4 Determination of Ozone Residual Concentration 150 7.2.5 Results from Preliminary Ozone Sanitizing Tests - Table 13 150 7.3 Preliminary Ozone Sanitizing Tests - Trouble-shooting 152 7.3.1 Preliminary Ozone Sanitizing Tests - Table 14 152 7.3.2 Preparation of Shewanella putrefaciens Planktonic 152 Inoculum 7.3.3 Addition of Shewanella putrefaciens Planktonic Inoculum 153 to distilled-deionized Water at 10° ± 0.5°C 7.3.4 Enumeration of Surviving Shewanella putrefaciens cells 154 Exposed to distilled-deionized Water at 10° ± 0.5°C 7.3.5 Absorbance Values at 600 nm 154 7.3.6 Results from Preliminary Ozone Sanitation Tests - Table 14 155 7.4 Preliminary Ozone Sanitizing Tests - Table 15 157 7.4.1 Preparation of Shewanella putrefaciens Planktonic 157 vii Inoculum 7.4.2 Addition oi Shewanella putrefaciens Planktonic Inoculum 158 to 0.5 ppm Ozonated distilled-deionized Water at 10° ± 0.5°C 7.4.3 Enumeration of Surviving Shewanella putrefaciens cells 158 Exposed to 0.5 ppm Ozonated distilled-deionized Water at 10° ± 0.5°C 7.4.4 Determination of Ozone Residual Concentration 159 7.4.5 Results from Preliminary Ozone Sanitizing 159 Test-Table 15 7.5 Variability Among Inoculated Stainless Steel Chips 162 LIST OF TABLES 1. C't values (mg.min/L) for 99 percent inactivation of microorganisms 16 by disinfectants at 5°C. 2. Review of some analytical methods for ozone. 21 3. Analytical methods for ozone in aqueous solutions. 23 4. Inactivation of microorganism by sodium hypochlorite. 29 5. Proposed mechanisms of chlorine antimicrobial activity. 34 6. Characteristics of chlorine compounds. 39 7. Some biocides in technical use in industrial water system. 52 8. Characteristics useful for identification of Shewanella putrefaciens. 54 biovars strains. 9. A N O V A for inactivation data of Shewanella putrefaciens suspensions 118 exposed to distilled-deionized water, ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min. 9a. Bonferroni adjustment on inactivation data of Shewanella putrefaciens 119 suspensions exposed to distilled-deionized water, ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min - Matrix of pairwise comparison probabilities. 10. A N O V A for inactivation data of Shewanella putrefaciens biofilms 120 ix exposed to distilled-deionized water, ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min. 10a. Bonferroni adjustment on inactivation data o f Shewanella putrefaciens 121 biofilms exposed to distilled-deionized water, ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min - Matr ix of pairwise comparison probabilities. 11. A N O V A for inactivation data of Shewanella putrefaciens suspensions 125 exposed to ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min. 11a. Bonferroni adjustment on inactivation data of Shewanella putrefaciens 126 suspensions exposed to ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min - Matrix of pairwise comparison probabilities. 12. A N O V A for inactivation data of Shewanella putrefaciens biofilms 127 exposed to ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min. 12a. Bonferroni adjustment on inactivation data of Shewanella putrefaciens 128 biofilms ozone (0.25 ppm) and chlorine (1.0 ppm) for 3 min - Matrix o f pairwise comparison probabilities. 13. Effect o f ozonated distilled-deionized water (10° ± 0.5°C) on 151 Shewanella putrefaciens suspension. 14. Effect o f distilled-deionized water (10° ± 0.5°C) inoculated with 156 Shewanella putrefaciens on indigo trisulfonate dye used to measure ozone concentration. 15. Effect o f ozonated distilled-deionized water ( 1 0 ° ± 0 . 5 ° C ) on 160 Shewanella putrefaciens suspension. LIST OF FIGURES 1. Schematic diagram of ozone generation by the corona discharge 6 procedure. 2. Electronic resonance structures of ozone. 8 3. Cyclic addition of ozone on unsaturated bonds. 10 4. Theoretical mechanisms of various stages of microbial attachment 46 and biofilm formation. 5. Experimental procedures for the preparation of Shewanella putrefaciens 64 planktonic inoculum. 6. Experimental procedures for the Preparation of Shewanella putrefaciens 67 biofilm inoculum. 7. Ozonation of distilled-deionized water used for sanitizing tests. 70 8. Steps used to prepare samples from planktonic suspensions 78 used in sanitizer tests for spectrophotometric determination of ozone concentration. 9. Decay of ozone dissolved in distilled-deionized water held at 83 10°±0.5°C. 10. Decay of ozone dissolved in distilled-deionized water held at 0.5°±0.5°C. 84 xi 11. Decay of chlorine dissolved in distilled-deionized water held at 85 10°±0.5°C. 12. Decay of chlorine dissolved in distilled-deionized water held at 86 0.5°±0.5°C. 13. Effect of distilled-deionized water (10° ± 0.5°C) on Shewanella 88 putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 14. Effect of distilled-deionized water (0.5° ± 0.5°C) on Shewanella 89 putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 15. Effect of distilled-deionized water (10°±0.5°C) on Shewanella 92 putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). 16. Effect of distilled-deionized water (0.5° ±0.5°C) on Shewanella 93 putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). 17. Effect of ozonated distilled-deionized water (10° ± 0.5°C) on 96 Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 18. Effect of ozonated distilled-deionized water (0.5° ± 0.5°C) on 97 Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). xii 19. Effect of ozonated distilled-deionized water (10° ± 0.5°C) on 101 Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). 20. Effect of ozonated distilled-deionized water (0.5° ± 0.5°C) on 102 Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). 21. Effect of chlorinated distilled-deionized water (10° ± 0.5°C) on 106 Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 22. Effect of chlorinated distilled-deionized water (0.5° ± 0.5°C) on 107 Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 23. Effect of chlorinated distilled-deionized water (10° ± 0.5°C) on 111 Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). 24. Effect of chlorinated distilled-deionized water (0.5° ± 0.5°C) on 112 Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests). NOMENCLATURE A N O V A Analysis of variance BSA Bovine Serum Albumin CFU/cm 2 Colony Forming Units/cm2 CFU/mL Colony Forming Units/mL d-d water distilled-deionized water EPA Environmental Protection Agency g gravitational force Ha alternative hypothesis Ho original hypothesis ID internal diameter min minute(s) nm nanometers mV millivolts OD outer diameter ppm parts per million rpm revolutions per minute (Agitation level) TSA Tryptic Soy Agar TSAc Tryptic Soy Agar + 0.05 mg/L congo red TSB Tryptic Soy Broth s seconds SSt Soluble Starch SS chips Stainless Steel chips u population mean um micrometer U.H.P Ultra High Purity ACKNOWLEDGMENTS xiv I would like to express my gratitude to my supervisor Dr. Skura for his patience, advice and encouragement throughout this project. I also want to thank the members of my supervisory committee Dr. Di l l , Dr. Durance and Dr. Li-Chan for their helpful comments and suggestions. Special thanks to Mr. S. Yee and Mrs V. Skura who helped me find that special piece of equipment, chemical and whatever else I needed in a hurry. I also want to thank all my classmates that extended a hand here and there when needed. Many thanks to R. Knap for his advice, support and for saving the day in so many occasions. Last but not least my appreciation and love to my parents for always being there for me with their support and encouragement. 1 1 INTRODUCTION Sanitizers are a crucial part of a food plant's cleaning operations. Their application to food processing surfaces and equipment decreases the presence of pathogenic and spoilage microorganisms and consequently cross-contamination of incoming food material from soil and microorganisms remaining after regular cleaning operations (Marriott, 1989). Thus, sanitation procedures ensure that food products are safe and wholesome with a stable long shelf-life (Marriott, 1989). Many of the sanitizers currently in use by the industry have been found to be effective against microbial suspensions; however, their effectiveness against bacteria attached to contact surfaces and food processing equipment have not been fully evaluated. Bacteria can adhere firmly to contact surfaces in a short time. It is believed that they are able to bind to surfaces by producing exopolysaccharide glycocalyx polymers (Anwar et al., 1990). These polymers form complex matrices. Inside these matrices microcolonies several pells thick can develop and coalesce into what are referred to as biofilms (Anwar et al., 1990). Bacteria found in association with biofilms have been shown to exhibit greater resistance to antibiotics and sanitizers (Anwar et al., 1990). It has been proposed that this polysaccharide matrix helps organisms to bind to contact surfaces and also offer protection against antibacterial agents (Ronner and Wong, 1993). The food industry uses a variety of detergents and sanitizers to clean food contact surfaces. The choice of detergent and sanitizer is influenced by the type of soil to be removed; that is, inorganic soils such as calcium and magnesium carbonates or organic 2 soils such as food residues (Marriott, 1989). Water quality and water softness or hardness, also affect the efficiency of detergents and sanitizers (Marriott, 1989). In addition, the detergents and sanitizers must be compatible with equipment so that corrosion is minimized. Thus, a broad selection of detergents and sanitizers for various processing conditions are available. However, in the last decade, there have been increased environmental concerns about the chemical nature of many detergents and sanitizers used in various industries including the food industry. It can easily be foreseen that in the near future stricter control and fines will be exerted over sanitizers and other kinds of effluent disposed through sewage systems. Several of these sanitizers produce toxic by-products. Also, microbes develop resistance to some of them over time while others are not as effective against biofilms and some are too expensive. These are only a few of the concerns regarding the use of sanitizers. For example, one of the sanitizers often used for the elimination of planktonic cells and biofilms is chlorine. However, a problem that may arise with the use of chlorine is the formation of chlorinated organic compounds (holomethanes) by the reaction of chlorine with organic residues (Bott, 1991). These compounds are thought to be carcinogenic (Bott, 1991). Consequently, the use of chlorine as a sanitizer can pose problems of effluent disposal. In addition, surfaces and equipment sanitized by chlorine or chlorinated compounds may need to be rinsed in order to prevent food from picking-up these compounds as they are handled and/or processed (Bott, 1991). At the present time there is no such thing as the idea] detergent or sanitizer; they all have their benefits and drawbacks. Furthermore, the use of chlorine and other 3 commonly employed sanitizers may pose effluent disposal problems as government regulations become more strict. Thus, the food industry is or will be in need of environmentally friendly sanitizers that not only satisfy food plant waste requirements but also effectively eliminate or decrease the incidence of cross contamination of food material by pathogenic and spoilage organisms. Ozone is a substance that could be of interest as a substitute for chlorine and other sanitizers. Ozone is an unstable three-atom allotrope of oxygen formed by the excitation of molecular oxygen by ultraviolet radiation or an electrical discharge. A n allotrope has been defined as one of two or more different forms of an element in the same physical state (Ebbing, 1990) Recently, there has been an increased interest in the use of ozone as a sanitizing and sterilizing agent for a variety of applications. In fact, a study on the effect of chlorine and ozone on fresh water at a nuclear power plant showed ozone to be a stronger biocide than chlorine at similar time/treatment programs (Bott, 1991). In terms of ozone as a sanitizer in a food processing plant situation, Bott (1991) reported that ozone applied to achieve appropriate residual concentrations, with sufficient application time and turbulence can effectively kill and remove up to 99% of Pseudomonas in biofilms. Unfortunately, the majority of these studies utilized cell suspensions rather than biofilms and/or conditions not present on food contact surfaces and food processing equipment. In addition, it is difficult to compare these studies since they employed different experimental conditions and procedures. Also, in many cases, the authors did not clearly specify the conditions under which ozone was used; e.g. when recording ozone dosages, it was not clear whether these measurements referred to initial ozone concentrations or residual levels. Thus, there is a need for further research on the possible use of ozone as a sanitizing agent for food contact surfaces and processing equipment. It is worth noting that for many decades several European nations have been using ozone for the treatment of drinking water (Bablon et al., 1991a). Ozone has also been used in the removal of odour, colour and taste from foods, preservation of foods, control of algae growth as well as oxidation of inorganic and organic compounds in water and waste treatments (Wickramanayake, 1991). It is therefore not surprising that there has been increased interest in the use of ozone as a sanitizing and sterilizing agent for a variety of applications; including replacement of current sanitizing agents used in the food industry Therefore, in this study, a series of tests were performed to determine the effectiveness of ozone as a sanitizing agent against Shewanella putrefaciens. Shewanella putrefaciens was selected as the test organism because it is a well known spoiler of refrigerated proteinaceous foods; particularly fish and seafood. Tests were performed on both planktonic suspensions and biofilms of Shewanella putrefaciens. Biofilms were grown on stainless steel chips which were made of steel similar to that u$ed in the food industry. The effectiveness of ozone as a sanitizing agent was evaluated against chlorine and distilled-deionized water. Temperature effects were also assessed by performing all tests at 10° ± 0.5°C and 0.5° ± 0.5°C. Because Shewanella putrefaciens is a psychrotrophic spoilage organism, the antimicrobial tests were performed in cold environments characteristic of seafood processing plants and fish boats. 5 2 LITERATURE REVIEW 2.1 Basic Aspects of Ozone Chemistry 2.1.1 Ozone Generation Ozone, a strong oxidizing agent, is an unstable, subtle blue gas at room temperature. It is commercially produced by subjecting dry air or dry oxygen to ultraviolet radiation or an electrical discharge (Wickramanayake, 1991). The most common method used for commercial generation of ozone is the corona discharge method. Figure 1 illustrates the generation of 0 3 by the corona discharge method. In this method ozone is produced by passing oxygen or dried air through the field of a silent electrical discharge created by a high voltage alternating power source (Bott, 1991). This electrical discharge occurs in a gap between two electrodes separated by a ceramic dielectric medium (Bott, 1991). There are several mechanisms by which ozone is generated in a corona. However, the dominant mechanism is shown in equations 1.1-1.4. The ozone generation process is initiated when O2 is dissociated by free high energy electrons in the corona (Carlins and Clark, 1982). e-i + 0 2 - > 2 0 + e-i 1.1 This is followed by a three body collision involving atomic oxygen, O2 molecules and any other molecule (M) present in the gas mixture (Carlins and Clark, 1982). These collisions result in the formation of 0 3 . 0 + 0 2 + M —» O3 + M 1.2 Figure 1. Schematic diagram of ozone generation by the corona discharge procedure. Adapted from Wickramanayake, 1991. 7 However, oxygen can also be simultaneously produced by the reaction of ozone with atomic oxygen and electrons as shown in equations 1.3-1.4. O + 0 3 2 0 2 1.3 e-i + 0 3 0 2 + O + e-i 1.4 The final ozone concentration obtained from corona discharge generators depends on the sum of the above reactions (Carlins and Clark, 1982). These reactions are influenced by several factors such as oxygen content and temperature of the feed gas, contaminants in the gas, power density of the corona, etc. (Carlins and Clark, 1982). For example, i f air is used as the feed gas, ozone concentrations in the range of 1 to 3% can be produced. On the other hand, i f pure oxygen is used as the feed gas, ozone concentrations of 2-6% can be obtained (Wickramanayake, 1991). 2.1.2 Chemical Nature of Ozone Ozone is an unstable molecule composed of three oxygen atoms. It has a molecular weight of 48.00 grams/mole and a very low dipole moment of 0.49-0.58 debye (Rosenthal, 1974). It is a very strong oxidizing agent with an oxidation-reduction potential of + 2.7 volts. It melts at -192.5° ± 0.4°C and boils at -111.9° ± 0.3°C (Wickramanayake, 1991). The high reactivity of this triatomic allotrope of oxygen is the result of its electronic resonance structure shown in Figure 2. The nature of these resonance structures shows that the ozone niolecules can behave as dipole, electrophilic or nucleophilic agents (Bablon et al., 1991a). Two 0 0 n 0: :0 0: :0 :0 0; :0 Figure 2. Electronic resonance structure of ozone. Adapted from H. Rosenthal, 1974. 9 mechanisms are believed to be responsible for the high oxidative nature of ozone. These are: a. electrophilic addition of the ozone molecule to C=C double bonds and to lone electron pairs (Figure 3), and b. ozone-self decomposition through the formation of short living free radical like HO° and HO20, that initiate and propagate oxidative reactions (Rosenthal, 1974). The second mechanism is most prominent in aqueous solutions. In fact, there is a common consensus that the faster decomposition of ozone in water than in air is likely the result of a chain reaction initiated by hydroperoxide ions and the subsequent formation of highly reactive free radicals (Bott, 1991). Several mechanisms for ozone decomposition and the formation of free radicals have been proposed. These mechanisms have been described in greater detail by Gurol and Singer, 1982; Staehelin and Hoigne, 1982; Buhler et al., 1984; Staehelin et al., 1984; Tomiyasu et al., 1985; Sotelo et al., 1987 and Bablon et al, 1991a. The rate of ozone decomposition in aqueous systems is dependent on the nature of the systems being ozonated; that is, pH, organic load, temperature, turbulence and even ozone concentration (Wickramanayake, 1991). For example, at high pH, ozone decomposes faster and various oxidative agents are produced (Bablon et al., 1991b). Hence, ozone decomposes much more rapidly in salt water, which has a higher pH than fresh water (Rosenthal, 1974). However, at very high pH, the stability of qzone increases (Rosenthal, 1974). The effect of organic load on ozone decomposition depends on the 10 A5 )c=c( 0 o c ; b - c k 5+ 5-O 0 )c-c( a Figure 3. Cyclic addition of ozone on unsaturated bonds (a. primary ozonide; b. zwitterion; c. hydroxy-hydroperoxide). Adapted from Bablon et al., 1991a. 11 type and quantity of organic matter. Ozone becomes less soluble and stable as temperature increases (Bablon et al., 1991b). Similarly, increased turbulence decreases ozone stability. 2.2 Bactericidal Action of Ozone The mechanism by which ozone acts as a sanitizer against microorganisms, particularly bacteria, is not clear. It is common knowledge that ozone is an excellent biocide against bacteria, viruses, amoebae cysts and yeast; however, how ozone affects the cell constituents of these organisms and leads to their inactivation has been difficult to determine (Wickramanayake, 1991). These difficulties arise from the analytical techniques needed to quantify the cell components to be ozonated and the products of their ozonation. In addition, the many different reaction sites on cell constituents may result in different direct ozone reaction and radical reaction mechanisms (Bablon et a l , 1991a and b). In short, some of the main chemical components of microbial cell constituents are carbohydrates, proteins, lipids and combinations of these compounds such as glycolipids, glycoproteins, lipoproteins, etc.. Other important components are the nucleobases adenine, guanine, uracil, thymine and cytosine. The reactivity of ozone and its sites of attack are different for each of these classes of compounds. For example, ozone reactivity with polysaccharides leads to the breakage of glucosidic bonds and production of monosaccharides (Bablon et al., 1991b). This is followed by oxidation of the alcoholic sites of the liberated monosaccharides and the eventual formation of aliphatic 12 acids and aldehydes (Bablon et al., 1991b). However, this set of events proceeds slowly even though both direct ozone and its hydroxyl radicals are involved in the reaction sequence. On the other hand, ozone readily reacts with amino acids at neutral or basic pH. The preferred sites of attack are the nitrogen atom or the R group consisting of alkyl sulfur or unsaturated bonds (Bablon et al., 1991b). Ozone can attack either site or both at the same time. Therefore, the effect of ozone on polypeptides and proteins will depend on the nature of their constituent amino acids. Saturated fatty acids react very slightly with ozone. However, the reactivity increases significantly when ethylenic bonds are present in the carbon chain. The products of such reactions are aldehydes, acids and hydrogen peroxide. The nature of these by-products indicates that ozone most likely interacts by a 1-3 cyclo addition on double bonds (Bablon et al., 1991b). Ozone also reacts rapidly with nucleobases, particularly thymine, guanine and uracil as well as their nucleotides. From the observed reactivity of ozone with organic compounds present in cell constituents possible sites of action that may lead to the inactivation of bacteria and other organisms can be foreseen. In simple terms, the cell is composed of a cell wall, a cytoplasmic membrane and the cytoplasm containing the genetic material and other organelles. The cell wall consists mainly of polysaccharides, phospholipids and amine sugars. It has been suggested that the observed rapid action of ozone on microorganisms with associated cell lysis results from damage to cell walls and membranes, followed by attack on cytoplasmic components. The first site of attack on cell walls and membranes would most likely be the double bonds of its lipid components (Bablon et al., 1991b). The 13 cytoplasmic membrane which is composed of many proteins would also provide a site for ozone reactions. If ozone crosses these membranes, the cytoplasm and genetic material will be the next sites of attack since ozone rapidly degrades nucleic acids (Bablon et al., 1991b). Ozone also affects bacterial enzymatic processes by reaction with the sulfhydryl groups of some enzymes. In fact, it has been found that ozonated bacteria lose the ability to breakdown some sugars and produce gas. The quickness of oxidation and inactivation of bacteria and other organisms when exposed to ozone suggests that ozone behaves differently from chlorine. It has been proposed that chlorine oxidizes particular enzyme systems while ozone behaves as a general oxidizing agent (Bablon et al., 1991b). 2.3 Ozone Sanitation Studies A number of studies have investigated the role of ozone as a sanitizing agent; however, most of those studies have focused on drinking water and wastewater treatment rather than the food industry. Studies by Bott (1991) and Greene et al. (1993) evaluated the efficiency of ozone in environments resembling food contact surfaces. The study by Bott et al. (1991) revealed that ozone at appropriate residual concentrations, time and turbulence could effectively kil l and remove up to 99% of Pseudomonas in biofilms. Greene et al. (1993) compared the effectiveness of ozone and chlorinated sanitizers in the removal of biofilm from heavily contaminated stainless steel surfaces. They found that ozone was as effective as chlorine in decreasing bacteria by >99%. Yet, optimum ozone contact time and residual concentration for biofilm removal were not determined. In addition, some experimental procedures were not clearly defined. 14 Restaino et al. (1995) tested the antimicrobial efficacy of ozonated deionized water produced by a recirculating ozone reactor on a variety of foodborne pathogens, spoilage organisms and coliforms. The tests were performed with ozonated deionized water with no added organic matter and ozonated water containing 20 ppm of soluble starch (SSt) or 20 ppm of bovine serum albumin (BSA). The results from those tests indicated that ozonated water was effective in killing both gram positive and gram negative food related organisms as well as yeast cells. The gram-negative organisms; S. typhimurium, E. coli, Pseudomonas aeruginosa and Yersinia enterocolitica showed more sensitivity to ozonated water with or without organic matter added than the gram positive organisms. Listeria monocytogenes was an exception showing more than 5 log reductions compared to the 3 log reductions observed for S. aureus, E. laecalis and B. cereus. Yeast cells of Candida albicans and Zygosaccharomyces bailii showed 4.5 log reduction. However, fungal spores of Aspergillus niger were considerably more resistant; after 5 min exposure only 1 log reduction was observed. It was also noted that the kind rather than the amount of organic matter significantly affected the killing efficiency of ozonated deionized water. BSA significantly decreased ozone levels in deionized water and thus the death rates in the presence of BSA were considerably lower. No significant differences in the killing efficiency of ozonated water were observed by the addition of SSt (Resteino et al., 1995). Thus, research data indicates that the sensitivity of bacteria to ozone varies considerably. E. coli is one of the most sensitive organisms. On the other hand, gram-positive cocci such as Staphylococcus and Streptococcus species and gram-positive 15 bacillae and the mycobacteria are much more resistant to ozone. As would be expected, spores are more resistant to ozone than vegetative cells. Similarly, viruses are also more resistant to ozone than vegetative cells but not more resistant than sporular forms of mycobacteria. Parasites are also more resistant to ozone than viruses and vegetative bacteria (Bablon et al., 1991b). In general, ozone is a better or at least as good a sanitizer as those currently used in the food industry. For example, Table 1 illustrates the high killing capacity of ozone compared to chlorine based sanitizing agents. These comparisons are based on O t values (Concentration • time); values derived from subjecting a specific microbial population to a constant sanitizing agent concentration (C) for a specific time (Bablon et al., 1991b). The results from the few studies on ozone as a sanitizing agent for the food industry against undesirable microorganisms are encouraging. However, most data including ozone tests, were derived from treatments of planktonic suspensions under more or less ideal circumstances. It is true that in many cases this is unavoidable due to practical problems. Nevertheless, this does leave a void in our knowledge of sanitizer efficiency when trying to compare a new alternative sanitizing agent or procedures, etc. to those currently in use. c PS o e o N O 1) "2 o o u M3 o Q 2 c CL o c 1 o O 6 OH a PL, cu C o u o c ca OX) l -o o CN O o © CN O o o o © o r- r--d vd N CN CN ©' d d ON o o o d C N un oo © ^ IT) CM o oo Ti-en vo oo cn o o O 1-H i n O d d o o vn m A oo o o © d o © rr m &3 S s 17 2.4 Sanitizing Efficiency of Ozone The rate of inactivation of microorganisms is believed to increase with increasing temperature. Temperature is thought to influence the rate at which sanitizing agents diffuse through microorganisms' protective membranes and their reaction with cell constituents. An increase in temperature of 10°C is usually accompanied by an increase in the reaction rate by an order of 2 or 3 (Bablon et al., 1991b). However, even though ozone reactivity with various substrates increases with increasing temperature, its solubility and stability in aqueous systems decreases (Bablon et al., 1991b). Nevertheless, several studies have shown that although increasing temperatures from 0° -30°C decreases the solubility of ozone and increases its decomposition rate, it does not affect microbial death rate as determined by total bacterial count (Bablon et al., 1991b). One of the problems that arise from this situation is that the Ot values for ozone are less reliable than for other sanitizers because of volatility and reactivity as well as the difficulties in maintaining constant residual ozone concentration. The effect of pH on the biocidal efficiency of ozone is not as important as for other sanitizers such as chlorine. Several studies have indicated that there are relatively minor changes in ozone efficiency at pH values ranging from 5.8-8. The changes that are observed with different pH were related to the rate of decomposition of ozone (Bablon et al., 1991b). At higher pH, ozone decomposes faster and various oxidizing agents are produced. However, studies performed at different pH while maintaining a constant ozone residual concentration, showed no differences in the extent of microbial inactivation (Bablon et al., 1991b). 18 Ozone inactivation of microorganisms is also influenced by the state in which the organisms are present and the quality of the water; that is, if they are in a free state, as aggregates, attached to the surface of organic or mineral matter or cell debris, etc. (Bablon et al., 1991b). However, the degree of turbidity of the aqueous system is not as important as the type of turbidity present in inhibiting the biocidal effect of ozone. For example, unoxidizable mineral materials seem to have little influence on the efficiency of ozone. On the other hand, organic materials tend to use up considerable amounts of ozone forming compounds of decreased biocidal activity and consuming active species derived from ozone decomposition (Bablon et al., 1991b). Therefore, the water utilized in ozonation of microorganisms must be of optimum quality in terms of the amount of turbidity and dissolved organic matter in order to ensure adequate killing of microorganisms. 2.5 Difficulties in the Determination of Ozone Concentration Ozone concentrations in aqueous solutions are difficult to determine due to rapid ozone decomposition in aqueous solutions, ozone volatility and reactivity with many organic and inorganic compounds present in aqueous media, reagents, unsterilized glassware, etc. As a result, ozone measurements may be subjected to considerable error. Furthermore, many analytical techniques used to determine ozone concentrations are based on modified methods for chlorine determination even though chlorine chemistry is quite different from ozone chemistry (Gordon et al., 1988). In addition, most of these modified methods rely on the total number of oxidants present in solution for their 19 determination of ozone levels (Wickramanayake, 1991). Thus, many studies that have looked at the sanitizing and sterilizing ability of ozone using ozone measurement techniques such as iodometric methods may be over estimating their values by measuring compounds other than ozone. As previously described, ozone decomposition produces very reactive and catalytic intermediates such as H 0 2 " (hydroperoxide ion), 0 2 (superoxide radical), H 0 2 (hydroperoxide radical), 0 3 " (ozonide radical ion), OH (hydroxyl radical) and H 2 0 2 (hydrogen peroxide). These reactive species greatly affect the stability of ozonated solutions, degree of interference by oxidizable compounds as well as the precision and accuracy of several analytical methods used to determine ozone residues (Gordon et al., 1988). Thus, it is likely that ozone in aqueous samples will be decomposing and reacting from the time samples are removed for measurement and subjected to measuring steps. The difficulties of measuring ozone concentration is well illustrated by iodometric methods. For example, in the 1985 standard method for ozone determination in an aqueous system, iodide ions in an unbuffered potassium iodide solution are oxidized by ozone to iodine. The solution is subsequently adjusted to a pH of 2 by the addition of sulfuric acid. The released iodine is then titrated with sodium thiosulfate to a starch end point (Bablon et al., 1991a; Gordon et al., 1988). The reaction between ozone and potassium iodide should be as described by equations 2 and 3. 0 3 + 21" + H 2 0 -> I 2 + 0 2 + 20H" The released iodine is titrated with sodium thiosulfate: 2 20 I 2 + 2 S 2 0 3 2 - 21- + S 4 0 6 2 " ^ Ideally the ozone/iodine stoichiometry for this reaction should be 1 to 1 but extensive studies have determined that the stoichiometric ratio ranges frpm 0.65 to 1.5. This ratio is influenced by pH, buffer composition, buffer concentration, iodide ion concentration, sampling techniques and reaction time (Bablon et al., 1991a; Gordon et al., 1988). The ozone/iodine stoichiometry has been shown to be altered by the pH of the ozone/iodide ion reaction and the pH during iodine measurement. Iodate ions and hydrogen peroxide formed during this reaction are thought to affect the ozone/iodine stoichiometry (Gordon et al., 1988). Several modifications to the iodometric method have been made in an attempt to obtain the desired 1 to 1 stiochiometry. However, when this ratio is obtained, it is a result of balances among the various reactions involved in this method rather than ozone-iodine ratio (Bablon et al., 1991a). Similar interferences or drawbacks are also found in other methods, thus an "ideal method" that deals effectively with the reactive nature of ozone and whose values are a direct representation of the actual ozone in the system is required. Such an ideal method should also be quick, easy to perform, free from interferences including those from products of ozone decomposition and oxidized by-products, and easy to automate (Gordon et al., 1988). Tables 2 and 3 list and review several analytical methods for ozone in aqueous solutions. 21 c o N O Vi T 3 O » S "3 o OS C o CN I > '3 T3 O o o <0 s o T3 O 4) s o & O OH U 43 > O "o -a o o o s <L> l—I .s 13 60 C w H U <J [in 2 2 d o 1 '6 1^  « , > 3 CT <D > .a a d 12 d o c3 d l-o lo 60 s o VI -a CD -a •8 _g o 15 d o d <u o d o o 0) 3 .s c o o H -d d o >, d ' N , «J 12 03 d 'C >, loo <D 3 O 60 d <D o -t-> o <u ^ ID a) b 9 -° > o (D k 'C E ^ S o w »-, 3 I ID -4-» E o <D T3 CD vi 3 o a. 23 I 1 60 u o c W ri o o ,1 l l I t O u •a I a o U ; S g § ,5,5 8 8 o o u I o O T3 u s in u T3 a -a «5 c/3 (0 a 9 t> B o H =» § « ° " S ^ o a g '3 £ < & 8 00 vo >© vd vd ^ ^ 0 1 ^ ^ 0 1 CM CN CN o4 CN tn « M H H W .JJ -H -S CN CN CN CN CN f% f> I—i i— 1— UH & & 8- « rf « g g I 60 60 60 O O O O O i B fl g •— — '~ "~ A •a -g ; | a £ £ | .§ ^ 'S '5? *X CN CN CN CN Ol ^ O O O u u o u O u CN CN i n m >n o •n © © ' © <n •n © IT) i n © © CN CN <o 1 1 © ' •n </-> •n <n - i i n © © i n <n CN © © © © © i — i 1 1 • • • i • 1 • r o © i n CN © © c o © *—i © © © © © " © © © © A © © ' A © CN CN i — l © © CN © © c o © © © © © © © ©_ © © © © © © © ' © © © ' I & O 3 u 3 3 3 •a -a « (3 O O U U CN CN + , H i t "of 2 M A A CN CN a; © © •n © m CN © S o JO T3 O < S Q o -c o, o o <L> CL 00 O 60 o i s o Q O > > - J < c o T 3 "I o « o 04 O O CN V CN O z c o i u > o CN o C4 -Z CN Pi t~~ © CN i n © 3 © © a § ^ s & o Q 6 03 Pi Z c4 Z CtJ z C4 z © v o 60 u .S 24 oo on eo a 4> O S c§ M-H u | T 3 ti £ 2 O 'GO 8^ ' (1) o ca w <3 w 60 l l a ? e 60 S3 <l> Q +-* »—] T3 O s •a I oo a s: S ^ s i : <u o o CM o 4> a. 00 60 ti T 3 O 13 o •s o a o u W oo .2 'o o 9< 00 .1° N '•3 "3 O Pi z LO CO ^ -a -o — i 3 3 O I I « o o o uua! P4 ps! pi Z Z Z £4 z z z z o o 2 o 2 1 J-8 pi z &H H i Pi z z z CM I N O ft! ©' ° Z u O is o * 1 u fa S3 | CQ S 13 •43 a <o 2 00 <D 3 (3 O O Pi Z z Pi z pel Z o o 3 g D. U U oo I 1 & 3 oo s a O oo u w g OH g Z B oo c o o o ti u a o a, o •a 00 > B o Z O S3 5 13 60 VPS c a • 3 <u 6 !r! ^c2 o ea Q Q Pd tr> Z © Pi ^ Z o Pi o Z o o o o 6b. o 5 © o O o 00 00 <L> •— a 13 o o PH OH I 1 § c s oo >? C c3 C _o o 1U o e 2 c N a . 2 ^ A <« "3 JL £i <u eg "o o > o o 3 ••— (U o H J < T3 t: o ON CCS C3 C CCS 6 ccS CCS X) U T) ta ( H & 3 > . o 00 &0 25 2.6 Determination of Ozone Concentration by the Indigo Trisulfonate Method Of all the ozone concentration methods that have been investigated, the indigo trisulfonate method appears to be the closest to the "ideal method". This method offers the following advantages; direct measurement of ozone, simplicity, good precision, sensitivity, selectivity and rapidity (Bablon et al., 1991a). The indigo method is also less susceptible to interference by substances such as manganese ions, chlorine, H 2 0 2 , ozone decay products and organic substances produced by ozonolysis that are often found in ozonated water and which may interfere with measurement of ozone concentration. For example, the indigo trisulfonate method incorporates procedures that allows for the determination of ozone concentration in the presence of chlorine and manganese (Bader andHoigne, 1982). The effectiveness of this method is due to the direct stoichiometric reaction (1:1) of ozone with the double bond of indigo trisulfonate resulting in the decolouration of the indigo trisulfonate under acidic conditions (Bader and Hoigne, 1982). This stoicheometric decolouration reaction allows for the spectrophotometric measurement of ozone concentration at 600 nm. At 600 nm, a decrease in absorbance is linearly correlated with increasing ozone concentration (Bader and Hoigne, 1982). For measuring purposes an indigo trisulfonate stock solution is first prepared according to the procedures described by Bader and Hoigne (1982). This stock solution consists of distilled water, concentrated analytical grade phosphoric acid and potassium indigo trisulfonate. The indigo stock solution is the main component in the preparation 26 of diluted versions of itself which are formulated depending on the anticipated range of ozone concentration (Bader and Hoigne, 1982). Ozone concentration is determined by taking the difference between two samples; a standard consisting of indigo trisulfonate dye and ozone demand free distilled-deionized water and the test sample containing the indigo dye and the ozonated distilled-deionized water (Bablon et al., 1991a). Ozone concentration (mg 03/L) = (AAbs • 100) / (f • b • v) 4 where b = path length of the cuvette (cm) v = volume of the sample added (mL) f = slope of the calibration curve at 600 nm (0.42 + 0.1 cm"1 per mg/L, with molar absorptivity = 20,000 M • cm'1) 2.7 Basic Aspects of Chlorine Sanitation 2.7.1 Chemistry of Chlorine Sanitation Chlorine and chlorine compounds have found a large niche in the food industry as effective and inexpensive sanitizers. Five main groups of chlorine compounds are used as sanitizing agents: liquid chlorine, hypochlorites, inorganic chloramines, organic chloramines and chlorine dioxide (Odlaug, 1981). Although all of these compounds have antimicrobial activity; some are stronger biocidal agents (Odlaug, 1981). The use of chlorine compounds as sanitizing agents dates back to nineteenth century Europe when chlorinated lime was used for the treatment of London's sewage and disinfection and 27 deodorization of hospital wards (Dychdala, 1991). Since then, active chlorine compounds have become the sanitizer of choice for the purification of water and for sewage treatment. Different formulations at different concentrations are available for both industrial and domestic use. Chlorine's success as a sanitizer is due to its rapid and fairly wide spectrum of activity against microorganisms. It is also easy to use and is relatively non-toxic at recommended usage levels (Marriott, 1989). Chlorine is most commonly available as a sanitizing agent as hypochlorite in the form of Na, K , Ca and M g salts. When chlorine dissolves in water it hydrplyzes to form HOC1. HOC1 will further dissociate to form H + and OC1" as illustrated in the succeeding reactions (Dychdala, 1991). C l 2 + H 2 0 -> HOC1 + H+ + CI" 5 Ca(OCl)2 + H 2 0 C a ^ + H 2 0 + 20C1 6 Ca(OCl)2 + 2H 2 0 - » Ca(OH)2 + 2HOC1 7 HOC1 <-> H+ + OC1- 8 The relative concentration of HOC1 to OC1" in solution will be dependent on pH, temperature and ionic strength (Odlaug, 1981). It has been observed that HOC1 is the agent with the greatest microbial activity. Consequently, chlorine based sanitizers are more effective at lower pH where there is an abundance of HOC1 (Odlaug, 1981). As the pH rises, OC1", a very weak biocidal agent becomes dominant and the antimicrobial action of the chlorinated solution is diminished. However, i f the pH declines below 4, HOC1 decomposes and C l 2 becomes the dominant specie (Elliott, 1980). Neither OC1" nor C l 2 are very effective against microorganisms. In the case of 28 chlorine dioxide, which does not hydrolyze in water, the entire molecule appears to have biocidal activity (Marriott, 1989). Once chlorine compounds react with microorganisms and organic matter they are reduced to inactive chlorides such as NaCl (Gardner and Peel, 1986). 2.7.2 Hypochlorites Hypochlorites such as NaOCl and Ca(OCl) 2 are among the most commonly employed chlorine based sanitizers. They can inactivate both gram-negative and gram-positive microorganisms in relatively short contact times. A 90% decrease in vegetative bacterial cell population can be obtained in 1.5 to 100 sec at relatively low levels of free available chlorine (Dychdala, 1991). Table 4 summarizes results from some studies in which NaOCl was used as sanitizing agent. Although different methodologies were employed in each study, it was generally found that the test microorganisms were very sensitive to hypochlorite at low levels of free available chlorine (Odlaug, 1981). Some organisms; however, appeared to be more resistant to hypochlorite, requiring not only higher levels of free available chlorine but also longer contact times (Odlaug, 1981). Acid-fast bacteria are also moderately susceptible to chlorine compounds (Gardner and Peel, 1986). On the other hand, some spores are considerably more resistant to chlorine, requiring free available chlorine concentrations 10 to 1000 times higher than vegetative cells (Odlaug, 1981). It has been suggested by Phillips (1952) and Odlaug (1981) that the increased resistance of spores to chlorine is due to changes in the molecular configuration of proteins protecting the sulfhydryl groups of essential enzymes, while the same groups appeared to be unprotected in vegetative cells (Dychdala, 1991). Chlorine 29 CO D O e i-,VD CL) o 03 o 1 « 2 8 IP £ < ft" OH c 03 60 SO ON NO ON SO O N r~-SO „ O N —j ^  ON Co ^ i— i CD C3 cJ ^ CvJ S ffi H Us ^ "S C3 03 £ GO ^CO T3 T3 C 03 C o3 o3 O u O g j » ' § CD CD • c ° t ; o S o r ~ o SO so so ON O N ON 03 ON ON co O N ON o3 o3 *"0 -t-> C CD CD CD o3 co ffi ffi ffi O H 2? & 03 C N O N oo oo "O vo so cs O N O N M ~ ~ % _T _T T3 03 03 C +-> 03 <U <U W O O j>> M H H <o o <n O N <o T f * - H C N o sp m' i - H ro ro r-m V >—i </-> </-> K o r-H O O N C N O O A <o SO XO NO C N co ^ © C N o s© vo so ro C N o T j - t o o o oci r-.' t-- r~- oo iri oo r~- oo r~-' so' so' oo . " 3 "C) ^ ^ * •»•- v j Q .=2 O O v j v» I o C» .?3 Q v5 J < W .5 CD CD ^ ft ft ^ >> O > 15 & § o S Ss. v\> -s o vU i vi) O v\> C ^ U c j ^ - O - C , ESv, C ^ ^ ^ t ^ ^ ^ C < j ^ ^ ^ ^ D Q Q Q O v J s j o o C N O C N o 3 "o cS cC =3 = T 3 T3 e o T3 " 9^ «•§ .22 =3 <U CD oi J3 O m O -C N —' S 2 2 H I I * & & & j2 ^ < t o t o <U 4) U O . o 30 has also been found to be moderately effective against viruses and fungi (Gardner and Peel, 1986). 2.7.3 Free Available Chlorine The term "free available chlorine" noted in the previous section is a means of expressing the oxidizing power of the chlorine compound based on the equivalent amount of elemental chlorine present in the compound in question (Dychdala, 1991). Thus, the concentration of a chlorine compound like NaOCl can be referred to as "available chlorine" by determining the electrochemical equivalent amount of chlorine in NaOCl as illustrated by equations 9 and 10 (Dychdala, 1991). C l 2 + 2e- -> 2C1- 9 OC1- + 2e" + 2BT - » CI" + H 2 0 10 Equation 9, shows two moles of chlorine reacting with 2 electrons to produce inert chloride. Chloride can also be produced by the reaction of one mole of OC1" and 2 electrons (eq. 10) (Dychdala, 1991). Consequently, it can be said that one mole of hypochlorite is electrochemically equivalent to one mole of chlorine and thus have 70.91 g of available chlorine, the molecular weight of chlorine (Dychdala, 1991). Therefore, since NaOCl contains one mole of hypochlorite, it can also be said to contain 70.91 g of chlorine per mole. Since the molecular weight of NaOCl is 74.5, NaOCl provides about 95.18 weight percent of available free chlorine (Dychdala, 1991). When water is chlorinated some of the chlorine will be consumed by organic and inorganic impurities present in the water. The remaining unconsumed chlorine is thus 31 referred to as residual available chlorine (Dychdala, 1991). This residual available chlorine can be present as free or combined chlorine. Free available chlorine, as defined by Weidenkopf (1953), is a term usually given to chlorine found in water in the form of elemental chlorine, HOC1 and OC1" (Dychdala, 1991). Combined available chlorine, on the other hand, refers to chlorine which has reacted with ammonia or other nitrogenous compounds resulting in the formation of chloramines or N-chloro compounds (Dychdala, 1991). 2.7.4 Inactivation of Bacteria by Active Chlorine Compounds Many studies designed to elucidate the mechanism(s) by which chlorine exerts its antimicrobial effect have been reported. Several mechanisms have been proposed but they have not all been proven experimentally. Initially, it was thought that active chlorine compounds like HOC1 inactivated microorganisms by direct reaction of some chlorine species with the organism's protoplasm resulting in the formation of toxic N -chloro compounds (Baker, 1926). Later studies by Knox et al. (1948) proposed that chlorine inhibited sulfhydryl enzymes and other enzymes sensitive to oxidation. The inhibition of enzymes essential for life, e.g. aldolase, by chlorine compounds was thought to be responsible for bacterial death (Knox et al., 1948). This is supported by the observed correlation between inhibition of glucose oxidation and percentage of bacteria killed (Green and Stumpf, 1946; Knox et al. 1948). Work done by Friberg (1956) using radioactive 36C1 found that vegetative cells took-up CI as free available chlorine but not as combined available chlorine. The amount of free available chlorine taken up by the cells 32 increased with both time and concentration of free available chlorine. Thus, Friberg (1956) concluded that the formation of toxic N-chloro compounds by chlorine with the cell's protoplasm does not appear to be responsible for the rapid inactivation of bacterial cells. Instead, it is the first contact oxidation reactions occurring prior to chlorine accumulation and formation of N-chloro compounds within the cell that results in the organism's death (Green and Stumpf, 1946; Knox et al. 1948). Friberg (1957) also used 3 2P to demonstrate that small amounts of chlorine can cause damage to the permeability of bacterial cell membranes which was confirmed by the leakage of the 3 2P from nucleoproteins of bacterial cells (Odlaug, 1981). Further support of changes in bacterial membrane permeability were provided by Camper and McFeters (1979). They treated E. coli with 0.5 ppm available chlorine and observed that membrane functions, in particular those involved in the transport of extracellular nutrients, were affecteci (Camper and McFeters, 1979). They stated that it would not be surprising to find that the cell membrane, which is in direct proximity to environmental stresses, would be the site of chlorine induced chemical interactions. Thus, it would be improbable that chlorine would react with intracellular sulfhydryl groups and not interact witty the bacterial envelope (Camper and McFeters, 1979). It has also been suggested that chlorine inactivates spore cells by first stimulating spore germination and then attacking the germinated spore (Wyatt and Waites, 1973). Kulikovsky et al. (1975) treated Bacillus cereus spores with 0.25% NaOCl at 20°C. After 19.5 min there was a 99% loss of spore viability. They also found, through chemical and ultrastructural studies, that chlorine caused degradation of the spore 33 integuments. Degradation of the outer spore coat resulted in disruption of the permeability barriers and thus leakage of Ca + 2 , dipicolinic acid, ribonucleic and deoxyribonucleic acids (Kulikovsky et al., 1975). Table 5, lists some of the proposed mechanisms for the antimicrobial activity of chlorine. 2.8 Sanitation by Chlorine Compounds 2.8.1 Chlorine Stability The presence of one or a combination of the following factors will affect the stability of free chlorine and in turn the biocidal properties of chlorine solutions; these factors are: chlorine concentration, pH, temperature, presence and concentration of catalysts, presence and concentration of organic matter and U V irradiation (Dychdala, 1991). Free available chlorine solutions are more stable at lower concentrations, higher pH and lower temperatures. Copper, nickel or cobalt are strong catalysts of chlorine decomposition (Dychdala, 1991). Organic matter decreases the oxidizing properties and hence the sanitizing ability of chlorine by reducing it to inert chloride (Gardner and Peel, 1986). The destabilizing effect of U V can easily be stopped by storing chlorine solutions in the dark and in closed containers. Thus by controlling the presence of certain components as well as regulating environmental conditions such as pH, temperature and storage of chlorine, the half-life of chlorine can be enhanced (Dychdala, 1991). The stability of chlorine compounds can also be improved by the addition of stabilizers which will convert some of the free chlorine to combined chlorine in the forrn of N-chloro compounds. These compounds in turn will slowly release some of the cornbined chlorine 34 Table 5. Proposed mechanisms of chlorine antimicrobial activity. 1) disruption of protein synthesis 2) oxidative decarboxylation of amino acids into nitriles and aldehydes 3) reaction with nucleic acids, purines, and pyrimidines 4) interference with metabolism by oxidation of key enzymes 5) promotion of D N A lesions with the subsequent loss of DNA-transforming capability 6) inhibition of oxygen uptake and oxidative phosphorylation accompanied by leakage of some macromolecules 7) creation of toxic N-chloro derivatives of cytosine 8) generation of chromosomal aberrations Source: Marriott, 1989. 35 into the solution. Unfortunately, the biocidal activity of these compounds is also slower. Other more stable combined chlorine compounds such as inorganic and organic chloramines work in a similar manner (Dychdala, 1991). 2.8.2 Organic Matter Chlorine compounds easily react with all types of organic matter resulting generally in the consumption of available chlorine and decrease in antimicrobial activity. Some compounds like carbohydrates and starches consumed or combined with chlorine without decreasing its biocidal properties (Troller, 1983). On the other hand, other types of organic matter appear to protect cells from chlorine (Troller, 1983). Thus, it is recommended that sanitizing with chlorine compounds be performed after thorough cleaning. An available chlorine concentration sufficiently high to leave an adequate residue after the organic demand of the solution and microorganisms have been met, is necessary (Gardner and Peel, 1986). Chlorine also reacts with nitrogenous compounds forming organic chloramines. Some of these chloramines retain some biocidal activity but they are eventually destroyed by oxidative reactions (Dychdala, 1991). 2.8.3 Concentration, pH and Temperature of Chlorine Solutions It is reasonable to presume that an increase in chlorine concentration will be accompanied by increased biocidal activity. This presumption is true as long as other variables such as pH, temperature and concentration of organic matter remain constant. 36 Under these circumstances, it has been observed that as the chlorine concentration is increased the rate of kill increases as well (Weber and Levine, 1944) As previously mentioned, pH plays a critical role in the effectiveness of chlorine compounds as sanitizers. As the pH increases the biocidal activity of chlorine decreases. An early study performed by Rudolph and Levine (1941) which used 25 ppm available chlorine to obtain a 99% kil l of B. metiens spores showed that as the pH was increased the killing time also increased; 2.5 min at pH 6, 3.6 min at pH 7, 5 min at pH 8, 19.5 min at pH 9, 35.5 min at pH 9.35, 131 min at pH 10 and 465 min at pH 12.86 (Dychdala, 1991). Other combined chlorine compounds such as chloramines also behave in a similar manner. The increased activity at lower pH is believed to be associated with the higher concentration of HOC1 (Gardner and Peel, 1986). However, at pH lower than 4 the HOC1 decomposes and liberates chlorine gas (Gardner and Peel, 1986). Therefore, it has been recommended that chlorine solutions be in a pH range of 6 to 8; in this range corrosion and irritation is minimized while HOC1 concentration is high enough for sanitizing purposes (Walker and LaGrange, 1991). Many reactions proceed faster at higher temperatures. The same seems to be true about the biocidal activity of chlorine compounds in solution. The research carried out by Rudolph and Levine (1941) also evaluated the effect of temperature on the biocidal activity of a 25 ppm Ca(OCl) 2 solution at pH 10. They found that killing of the microorganisms required 121 min at 20°C; 65 min at 30°C; 38.5 min at 35°C and 9 min at 50°C. Other studies, e.g. Weber and Levine (1944), also observed that the increased biocidal activity at higher temperature seemed to be only slightly influenced by pH. The 37 increased effectiveness of chlorine compounds at higher temperatures have been put to use in hot cleaning and sanitation in the food and dairy industries (Gardner and Peel, 1986). 2.8.4 Hard Water Calcium and magnesium ions do not seem to affect antimicrobial activity of chlorine compounds. Tests performed by Shere (1948) found no difference in the activity of a 5 ppm available chlorine sodium hypochlorite solution at 20°C using 0 or 400 ppm hardness. However, ferrous or manganous cations, and nitrite or sulphide anions can reduce HOC1 to inert chloride (Gardner and Peel, 1986). On the other hand, small quantities of potassium bromide may enhance the performance of hypochlorites (Gardner and Peel, 1986). 2.8.5 Active Chlorine Compounds Chlorine and chlorine based compounds are among the most widely used sanitizers in the food industry. They were first utilized by the dairy industry for the sanitation of milk bottles (Walker and LaGrange, 1991). Further research on the benefits of chlorine as a sanitizer resulted in its incorporation in water used for washing and rinsing of equipment and utensils during routine cleaning and sanitation (Walker and LaGrange, 1991). In 1931, canneries began adding chlorine to cooling water to minimize spoilage of canned food by microorganisms introduced into sealed cans by leakage of contaminated water through apparently intact seams (Scott, 1937). The addition of 38 chlorine to water used in food plants resulted in a decrease in the number of organisms in the water, equipment and final processed food. Thus, it not surprising that many food plants now add chlorine to their entire water supply; a procedure referred to as in-plant chlorination (Walker and LaGrange, 1991). This ensures that water used in the plant has 5 to 7 ppm available chlorine (Walker and LaGrange, 1991). Chlorine residual concentrations are increased to 15 to 20 ppm during cleaning operations (Walker and LaGrange, 1991). Presently, chlorine based sanitizers are used in a wide range of food industries, e.g. dairy, soft drink and sugar beet industry, canneries, vegetable packers, slaughterhouses and many more (Walker and LaGrange, 1991). Some of the main chlorine compounds used as sanitizers in the food industry are listed in Table 6. The concentration of available chlorine used in food plants ranges from 5 ppm to 5000 ppm; the higher amounts are used to control mold growth. However, the most common concentration employed is 200 ppm (Gardner and Peel, 1986). As early as 1929, Prucha (1929) recommended that all utensils and equipment should, be thoroughly clean before being sanitized. He also recommended that freshly prepared chlorine solutions should be used for sanitation. Chlorine solutions to be used for sanitizing utensils and large equipment should have a minimum concentration of 50 to 100 ppm available chlorine. On the other hand, chlorinated solutions used for spraying large equipment should consist of 200 ppm available chlorine (Dychdala, 1991). He also emphasized that chlorine solutions like hypochlorites were most efficient when used on already cleaned surfaces for at least 10 sec so that sufficient time was allowed for inactivation of microorganisms (Walker and LaGrange, 1991). In the Grade A 39 CO C O O H e o o CD _c o CD C+H O CO O CD O C3 c3 O so CD 13 o <+H 'S <=> 8 O to 2 > CD o H H .S o a e o o 13 o <L> J 3 o o o OO 08 3 <2 13 o a CD u 0 S o o xr O O kH u T3 x? o CN 0 CO =) O CD CO 03 O o 13 O CO U O > CO o x? <jx O O < 1 J H & .2 u o > CO o Z 4 3 e>x X ? O O 1) i* O OH w o > CO u o "ccf u X? CN x? O O CD O v? ov V CN O CZ) X so U U m 53 40 c o o (D o 3 O PH 'c3 o ^ 3 13 o <+H '3 => 8 s O . f l • f l 1 3 o u U H .S o o 13 o -5 rd oo to fl fl fl o o o Z Z Z eb 60 60 c . f l _C o *+H o H W Q o ca J D CD & 3 3 2 O 43 o JO 53 53 55 So 55 0 S-6s-C N is O O H 6s-C N EC U o u u z o u u z >» o 11 I * o o o O O H z c*% u u o o w | 1 O o o O O H 0 S-Z C N O EC CO O o In O 1 T3 .fl t- g "3 5 ^ '3 M S 5* .2 -g _2 -3 -2 ••- 2 fl o o 2 i - J O CO 00 00 O " V H o s o o ^ H o o u C N O H o u fl <u o 3 2 U -3 41 Pasteurized Milk Ordinance, 1985 recommendations of the U.S. Public Health service, it was stated that hypochlorite solutions containing 50 ppm available chlorine be used for at least 1 min at a minimum temperature of 24°C for the disinfection of utensils and equipment (Dychdala, 1991). When chlorine compounds are used as sprays, a minimum of 100 ppm available chlorine should be used (Dychdala, 1991). Hypochlorites have also been used for direct sanitizing of food items. Fish and shellfish caught in polluted waters may be treated with hypochlorites to minimize health risks (Dychdala, 1991). Available chlorine concentrations may range between 200 to 6000 ppm depending on the level of organic contamination of the fish or shellfish (Dychdala, 1991). Eggs are also treated with chlorine solutions because they too are exposed to high levels of organic contamination and pathogens (Walker and LaGrange, 1991). Another example of direct sanitizing of food products is the addition of chlorine at 200 ppm to the water in chill tanks use to cool poultry carcasses (Walker and LaGrange, 1991). The chlorinated water helps lengthen the shelflife of poultry meat as well as decrease cross-contamination by microorganisms washed from the carcasses (Walker and la Grange, 1991). Fruits and vegetables may also be decontaminated by washing with approximately 4 to 5 ppm available chlorine (Dychdala, 1991). However, direct cleansing of food items with chlorine may also result in only marginal improvements to the microbiological quality of a food product; it has been found that treatment of beef, poultry, lamb and brussels sprouts with chlorinated water at 200 ppm only resulted in less than a two fold decrease in bacterial counts (Walker and LaGrange, 1991). 42 2.8.6 Chlorine and Microbial Biofilms The biocidal activity of chlorine compounds is decreased in the presence of biofilms. Studies performed by Cadwell (1990) demonstrated that sodium hypochlorite at low concentrations of available chlorine (0.5 to 5 ppm) were only inhibitory to cells in biofilms and once the biofilm was removed from contact with the chlorine solution, the cells and biofilm began to grow again. A 50 ppm available chlorin^ solution will decrease biofilm formation and only temporarily stop cell growth i f chlorine is removed from the system (Dychadala, 1991). Bolton et al. (1988) looked at strains of Staphylococcus aureus which appeared to be resistant to routine cleaning and sanitizing procedures. These strains tend to colonize defeathering equipment and can be found throughout the processing plant. These Staphylococcus strains were found to be eight times more resistant to hypochlorite (at a 5 ppm level) than isolates from the natural skin flora of poultry. The increased resistance of strains of Staphylococcus aureus to chlorine was attributed to the ability of strains of Staphylococcus aureus to grow in macroclumps buried in a protective extracellular slime layer (Bolton et al., 1988). Another study by LeChevallier et al. (1988) showed that bacterial attachment to surfaces provides a greater resistance to sanitizers. They observed that unencapsulated Klebsiella pneumoniae grown in a rich nutrient medium and attached to glass slides had a 150 fold increase in their resistance to sanitizing agents. Other factors influencing resistance to sanitizing agents include age of the biofilm, bacterial encapsulation, production of extracellular polymers and growth conditions such as growth medium and growth temperature (LeChevallier et al., 1988). Resistance afforded by these factory can have a 43 multiplicative effect, thus increasing the resistance to chlorine from 2 to 10 fold (LeChevallier et al., 1988). This may explain the survival of bacteria in chlorinated water supplies (LeChevallier et al., 1988). Therefore, in a food processing environment, one must be aware of the effects of food soil, type of surface, cleaning routines, sanitizing agent and concentration will have on the effectiveness of sanitizing procedures. 2.8.7 Methods for the Determination of Available Chlorine. Several methods exist to check the available chlorine present in chlorinated solutions. For example, in the Iodometric method, free chlorine reacts wjth an acidified test solution of potassium iodide (KI) resulting in liberation of iodine which is then titrated with a standard solution of sodium thiosulfate to a starch end point (Dychdala, 1991). Another titration method involves the titration of a chlorinated solution with a standard sodium arsenite solution that uses Kl-starch as the indicator (Dychdala, 1991). The concentration of dilute chlorine solutions (ppm level) can be determined by the orthotolidine (OT) method. Orthotolidine is a colourless reagent that changes to a yellow-orange-red colour when added to dilute chlorine solutions. The concentration of available chlorine is determined by comparing the intensity of this yellow-orange-red colour to a standard (Dychdala, 1991). Palin's DPD method makes use of the N , N -diethyl-p-phenylene-diamine (DPD) reagent to test dilute chlorine solutions (ppm level). The dilute chlorine solutions will change the DPD reagent to a pink-reddish colour based on the concentration of chlorine (Dychdala, 1991). Both the OT and D P p methods can 44 be adjusted to determine not only free available chlorine but total available chlorine, combined chlorine, and chloramines. The amperometric method is a electrometric titration method where a current is passed through a titration cell holding a dilute chlorine solution which acts as the oxidizing agent and a standard phenylarsene oxide (PAO) as the titrating reducing agent. The available chlorine (ppm level) is calculated using the volume of PAO used during the titration (Dychdala, 1991). Available chlorine can also be determined by the used of polarographic membrane techniques (Dychdala, 1991). Residual chlorine electrodes also provide a way for easy and rapid analyses of residual chlorine concentrations (ppm level) without the need of large samples of chlorinated solutions often required by titration methods. For example, the Orion -Model 97-70 Residual Chlorine Electrode permits the measurement of all types of available chlorine including free chlorine, hypochlorites and chlorine bound to nitrogenous compounds in an easy, quick and accurate way (Orion Research, Inc. - V W R Scientific Products). It can detect chlorine concentration of 0.01 to 20 ppm at an optimum temperature range of 0° to 50°C. It also has a pH range of 0 to 14 and a reproducibility of ± 2% between chlorine concentrations of 0.2 to 20 ppm (Orion Research, Inc. - V W R Scientific Products). 2.9 Microbial Resistance in Biofilms Biofilm is the term given to biological deposits that form on any surface. These deposits consist of microorganisms and their extracellular products, mainly polysaccharides and glycoproteins (Mattila-Sandholm and Wirtanen, 1992). These 45 extracellular components are not exclusively produced by biofilm cells but they are produced in much larger quantities than by their planktonic counterparts. Biofilms are found on almost any surface exposed to moisture, regardless of the nutritional levels (Wilderer and Characklis, 1989). In fact, it has been observed that biofilm formation increases when microbes are exposed to low nutrient levels. Hence, it is believed that biofilms serve to protect microorganisms from hostile environments and entrap nutrients from their surroundings (Mattila-Sandholm and Wirtanen, 1992). Given the right conditions, most microorganisms can form biofilms; however, some microbes are better producers of biofilms than others. Some of the most notorious biofilm producers are from the genera Pseudomonas, Enterobacter, Flavobacterium, Alcaligenes, Staphylococcus, and Bacillus (Mattila-Sandholm and Wirtanen, 1992). Biofilm forming microorganisms form these matrices when they adhere to surfaces. Micrporganisms are able to bind to surfaces with the aid of specialized appendages called fimbriae or flagella and exopolysaccharide glycocalyx polymers which form the adhesive matrices (Zottola, 1994). Attached microorganisms will grow and multiply inside these matrices adhering not only to the surface but other cells thus creating microcolonies several cells thick (Zottola, 1994). A biofilm will be formed when these growing colonies, become large enough that they can attract and capture debris, nutrients and other microorganisms (Zottola, 1994). Figure 4 illustrates theoretical mechanisms of microbial cell attachment and biofilm formation postulated by a number of scientists (Zottola, 1994). This figure presents microbial attachment as a two step process; a reversible stage and an irreversible 47 stage that leads to colonization. In the reversible stage, the microbial cell manages to hold on to the contact surface by electrostatic and Van der Waals forces (Busscher and Weerkamp, 1987). Van der Waals forces are active at separation distances > 50 nm. At these distances, the non-specific macroscopic cell surface properties also play a crucial role in the ability of the cell to adhere to contact surfaces. At separation distances between 10 and 20 nm, both Van der Waals and electrostatic interactions are active (Busscher and Weerkamp, 1987). At this stage, adhesion is still reversible but as time progresses it eventually become irreversible as the microbial cell surface becomes able to re-arrange itself and form specific short-range interactions. However, in order for these short-range interactions to occur, water has to be removed from in between the microbial cell and the contact surface (Busscher and Weerkamp, 1987). Thus, hydrophobic surface components of the microbial cell are likely to play an active role in removing this water film. At separation distances of < 1.5 nm, Van der Waals forces, electrostatic interactions and specific interactions are all at work. At very small distances (< 1.5 nm) specific interactions can occur i f the cell is capable of extruding adhesion probes and hydrophobic groups are able to dehydrate the contact surface (Busscher and Weerkamp, 1987). These specific interactions can lead to irreversible adhesion and eventual colonization. The formation of biofilm can be both beneficial and detrimental to the food industry. For example, in the production of vinegar, biofilms containing acetic acid producing bacteria are essential for the efficient generation of vinegar from substrate. Microbial biofilms are also found in bioreactors employed in the production of fermented 48 foods and in filters used in food waste water treatments (Zottola, 1994). However, microbial biofilms can also cause many problems. The presence of biofilm on heat exchange surfaces increases the resistance to heat transfer and energy losses. Filters can be clogged and rendered useless. Biofilm in pipes can increase fluid friction resistance. Equipment may deteriorate more rapidly due to corrosion (Wilderer and Characklis, 1989). Microbial biofilms attached to food contact surfaces and food also serve as a reservoir of possible spoilage and pathogenic organisms which can contaminate in-coming food material and already processed foods (Zottola, 1994). Thus, the presence of microbial biofilm on food contact surfaces has been a concern and the focus of increased research in the food industry. This increased interest has in part come from the emergence of several foodborne pathogens. A number of studies on L. monocytogenes have examined their ability to form biofilms over a temperature range of 10° to 35°C and the protective effect of biofilms against cleaning and processing treatments (Mattila-Sandholm and Wirtanen, 1992). Work by Herald and Zottola (1988) showed that L. monocytogenes can attach to food contact surfaces. This attachment was accompanied by the abundant production of exopolysaccharides. Other researchers have observed similar behaviour, thus further confirming the ability of L. monocytogenes to bind to food contact surfaces and form biofilms (Zottola, 1994). These studies have also demonstrated that L. monocytogenes and other organisms in biofilms have an increased resistance to sanitizers and processing treatments such as heat (Zottola, 1994). Frank and Koffi (1990) reported that L. monocytogenes in biofilm was more resistant to benzalkonium chloride than planktonic cells. L. monocytogenes in 49 biofilm required exposures of 12 to 20 min whereas 30 sec exposure to benzalkonium was required to kill unattached cells. There are also indications that a proportion of cells attached to biofilms can survive heat treatments of 70°C for 5 min. Mattila-Sandholm and Wirtanen (1992) found that P. fragi, L. monocytogenes, Enterococcus hirae, and Bacillus subtilis present in 24-96 hr old biofilms held at 25°C were more resistant than their planktonic counterparts to five different sanitizers. However, other studies also indicated that the resistance of microorganisms in biofilms to adverse conditions increases with biofilm age (Mustapha and Liewen, 1989; Frank and Koffi, 1990; LeChevallier et al., 1988; Mattila-Sandholm and Wirtanen, 1992). The actual hazard to safety presented by biofilms, given implementation of adequate cleaning and sanitation procedures is not yet fully known. But reasons for concerns are justifiable. It must be remembered that biofilms often consist of more than one organism and their extracellular products. The formation of biofilms by one or more organisms may attract others. Sasahara and Zottola, (1993) observed that a pure culture of L. monocytogenes attached sparsely to glass coverslips while P. fragi (an exopolymer producing organism) formed converging layers of cells. However, when L. monocytogenes and P. fragi were grown together, attachment and colony formation by L. monocytogenes improved (Sasahara and Zottola, 1993). In addition, biofilms are not static systems; they are continually undergoing cell growth and sloughing in a flowing system. Krysinski et al. (1992) evaluated the effectiveness of several cleaning and sanitizing agents on E monocytogenes attached to stainless steel and plastic surfaces for 24 hr at 25°C. Resistance of the attached cells varied according to the surface material. Greater resistance to cleaners and 50 sanitizers were observed on polyester/polyurethane surfaces followed by polyester and stainless steel. Sanitizers had little effect in removing the cells even after exposure time was increased to 10 min. On the other hand, unattached cells underwent a 5 log decrease in only 30 sec (Krysinski et al., 1992). Cleaners were more efficient in inactivating attached cells than sanitizers. This is to be expected since cleaners are meant to break the polysaccharide matrix and remove soil composed of fats, carbohydrates, proteins and minerals. If sanitizers are used alone, they can be inactivated by the soil and prevented from reaching the targeted organisms (Zottola, 1994). Sanitizers may be efficient against the first layers of the biofilm; however, these layers of microorganisms and their extracellular components may also act as a barrier to the diffusion of the sanitizing agent and consequently limit its effectiveness (Zattola, 1994). But i f cleaners are used first and they remove the soil, the organisms will be exposed and inactivated by the sanitizers. Thus, this confirms the appropriateness of conventional procedures that make use of cleaners followed by sanitizers in the removal and inactivation of microorganisms (Zottola, 1994). If the cleaning and sanitation procedures are inadequate, contamination of raw and processed foods is possible. The type of surface material used in the design of equipment and processing lines is also important. Smooth surfaces in good condition with no cracks, dead corners, scratches, etc. are easier to clean. On the other hand, bends, pockets and extended joints are areas that offer physical protection and favourable growth conditions (Mattila-Sandholm and Wirtanen, 1992; Carpentier and Cerf, 1993). For example, gaskets in equipment tend to accumulate dirt, nutrients and other particles; thereby, creating a good environment for biofilm growth (Mattila-Sandholm and 51 Wirtanen, 1992). Thus elimination of biofilms from food contact surfaces and equipment should be based on a systematic and comprehensive plan that incorporates good hygiene, sanitation practices and a HACCP plan that includes good design practices. Table 7 describes the advantages and disadvantages of some common biocides available to the food industry. 2.10 Test Organism 2.10.1 Shewanella putrefaciens Shewanella putrefaciens is a gram-negative, rod-shaped, facultative anaerobe approximately 0.5-1.0 by 1.1-4.0 pm. It is a non-spore former, non-pigrnented, motile, by a single polar flagellum, organism usually associated with aquatic or marine environments (MacDonell and Colwell, 1985). When grown on agar, S. putrefaciens forms convex, circular, slightly viscous or mucoid colonies approximately 2-3 mm in diameter. These colonies are usually red-brown or salmon-pink in colour. If grown on blood agar, hemodigestion will appear in areas of heavy growth (Gilardi, 1991). The optimum growth temperature of S. putrefaciens is about 20°-25°C, although some strains are capable of growth at refrigerated temperatures (Lee, 1979). Table 8 illustrates some characteristics of S. putrefaciens useful to its identification. Different biovars of S. putrefaciens can be noted by their ability to oxidize disaccharides, grow in 6.6% NaCl and on Salmonella-Shigella medium (Gilardi, 1991). Biovar 1 contains mainly strains from environmental sources such as dairy and fishery. They are one of the major psychrotrophic food spoilage organisms of refrigerated protein rich foods such as fish, 52 Table 7. Some biocides in technical use in industrial water system. Chlorine Advantages: broad spectrum of activity; residual effect; advanced technology available; can be generated on site; active in low concentrations; destroys biofilm matrix and supports detachment. Disadvantages: toxic by-products; degradation of recalcitrant compounds to biodegradable products; development of resistance; corrosiveness; reacts with extracellular polymer substances (EPS) in biofilms; low penetration characteristics in biofilms; oxidizes S2- to elemental sulfur [extremely difficult to remove from surfaces]. Hypochlorite Advantages: cheap; effective; destabilizes and detaches the biofilm matrix; easy to handle; useful for biofilm thickness control. Disadvantages: poor stability; oxidizing; rapid aftergrowth observed; toxic by-products; corrosive; does not control initial adhesipn. C10 2 Advantages: can be generated on site; activity less pH dependent; less sensitivity against hydrocarbons; effective in low concentrations. Disadvantages: explosive gas; safety problems; toxic by-prpducts. Chloramine Advantages: good penetration of biofilms; reacts specially with microorganisms; less toxic by-products; higher residual effect because of lower reactivity with water ingredients. Disadvantages: less effective than chlorine against suspended bacteria; resistance observed. Bromine Advantages: very effective against a broad microbial spectrum. Disadvantages: toxic by-products; development of resistance. Ozone: Advantages: efficiency similar to that of chlorine; decomposes to oxygen; no residues; weakens biofilm matrix. Disadvantages: oxidizes bromide in seawater; reacts with organics and can form epoxides; degrades humic acids and makes them available; corrosive; short half-life; sensitive to water ingredients. H 2 0 2 Advantages: decomposes to water and oxygen; relatively non-toxic; can easily be generated in situ; weakens biofilm matrix and supports detachment and removal. Disadvantages: high concentrations (>3%) necessary; resistance commonly observed; corrosive. 53 Table 7. (Continue) Peracetic acid Advantages: very effective in low concentrations; broad spectrum; kills spores; decomposes to acetic acid and water; no toxic by-products known; penetrates biofilms. Disadvantages: corrosive; rather unstable; increases DOC (Dissolved organic carbon). Formaldehyde Advantages: low costs; broad antimicrobial spectrum; stability; easy application. Disadvantages: resistance in some organisms; toxicity; suspected of promoting cancer; reacts with protein-fixing biofilms on surfaces; legal restrictions. Glutaraldehyde Advantages: effective in low concentrations; cheap; nonoxidizing; noncorrosive. Disadvantages: does not penetrate biofilms well; degrades to formic acid; increases DOC (Dissolved organic carbon). Isothiazolones Advantages: effective at low concentrations; broad antibiotic spectrum. Disadvantages: problems of compatibility with other water ingredients; inactivation by primary amines. Quaternary Advantages: effective in low concentrations; surface activity supports ammonium biofilm detachment; relatively non-toxic; adsorbs to surfaces and prevent compounds biofilm growth. Disadvantages: inactivation by low pH; Ca 2 + ; M g 2 + , development of resistance. Source: Mattila-Sandholm and Wirtanen, 1992. 54 Table 8. Characteristics useful for identification of S. putrefaciens biovar strains. Signb Characteristic3 Biovar 1 Biovar 2 Biovar 3 red-brown or pink color + + + Indophenol oxidase + + + Motility + + + Polar monotrichous flagella + + + Sucrose, maltose, acid (OFBM) + - -Hydrogen sulfide (Kligler iron agar) + + + Ornithine decarboxylase (DBM) + + + DNase + + + Growth on Salmonella-Shigella agar - + -Growth on 6.5% NaCl agar - + -a. Abbreviation: OFBM, OF basal medium, D B M , decarboxylase base Moeller medium. b. +, 90% or more positive within 2 days; -, no reaction (90% or more); + or -most strains positive; - or +, most strains negative. Source: Gilardi, 1991. 55 meat and poultry (Gilardi, 1991). This biovar has also been isolated from human clinical cases but it is not considered to be of clinical significance (Gilardi, 1991). The remaining biovars have been mainly isolated from human clinical cases and in some instances from environmental sources. S. putrefaciens was previously known as Alteromonas putrefaciens and Pseudomonas putrefaciens. Previously, these two earlier species were considered to be one until the study by Levin (1972) separated them into two species based on the guanine-plus-cytosine content of their deoxyribonucleic acid (Parker and Levin, 1983). The group with a relatively low, 43% to 48%, guanine + cytosine content in their D N A was classified as A. putrefaciens (Stenstrom and Molin, 1990). The other group with a guanine + cytosine content of 58% remained classified as Pseudomonas putrefaciens. Later studies showed that the genetic differences between A. putrefaciens and other Alteromonas species were significant enough to give A. putrefaciens an independent genus (Stenstrom and Molin, 1990). Later, A. putrefaciens as well as A. hanedai and two abyssal marine strains were classified under the genus Shewanella (Stenstrom and Molin, 1990). In the food industry, S. putrefaciens is known to be an intensive psychrotrophic spoilage organism of proteinaceous foods; in particular, fish. Thus, its control is of special importance to the seafood industry given the high perishability of fish and the ability of this organism to flourish under low storage temperatures (Venugopal, 1990). During ice or refrigerated storage of fish, the microbial flora of fish changes. As storage time increases, psychrotrophic, gram-negative organisms such as Alteromonas, 56 Pseudomonas and Shewanella become the dominant species (Venugopal, 1990). Fish spoilage by these organisms produce the typical volatile odour bearing compounds such as volatile basic nitrogen compounds, volatile acids, H 2 S and mercaptans (Venugopal, 1990). Stenstrom and Molin (1990) regarded S. putrefaciens as the most powerful fish spoiling organism because of its ability to produce hydrogen sulphide and reduce trimethyl amine oxide. S. putrefaciens was the dominant organism in 38% of the fish samples examined by Stenstrom and Molin (1990). S. putrefaciens was also the dominant H 2 S producing organism isolated from ground beef samples held at 20°C (Parker and Levin, 1983). Because of the extensive handling of fish and seafood from the time it is caught to processing, the factors which influence microbial contamination are various and may include catching methods, on board-handling, fishing vessel sanitation, processing and storage conditions (Venugopal, 1990). Contamination at the processing plant may well be the result of cross- and post-contamination of cut fish fillets and cleaned seafood products such as shellfish from biofilm present on food contact surfaces, cleaning and shucking equipment, containers, knives, etc. Once again, adequate hygienic and sanitation procedures are necessary at each step from the initial catch all the way to the storage of the processed product (Venugopal, 19^0). Thus, it is of interest to the food industry, in particular the seafood sector, to observe the effect of ozone as a antimicrobial against S. putrefaciens biofilms. In fact, refrigerated ozonated water and ice have been used on fishing vessels to preserve fish quality until the arrival of the fish at the processing plant. Ozone has also been shown to increase the storage life of rockfish (Kotters et al., 1997). 57 3 MATERIALS AND METHODS 3.1 Experimental Design Objective: To compare the efficacy of ozone and chlorine as antimicrobial agents against planktonic cells and biofilms of Shewanella putrefaciens. The data obtained from the antimicrobial tests were placed into two groups (planktonic tests and biofilm tests) for their corresponding Analysis of Variance. Design: The design of the experiment was a completely randomized design modeling the effects on lethality of different sanitizers, temperatures, and physical states (planktonic or biofilm) of the test organism. The statistical model is shown below: Y = M + E where: Y = sample observation M = sample mean E = error For each planktonic and biofilm set of tests, there were 6 treatments to reflect a 3 x 2 fjactpriaj experiment. The factors were sanitizer type and temperature. The sanitizing agents were: no sanitizer (control), ozone and chlorine. The two experimental 58 temperatures were 0.5° ± 0.5°C and 10° ± 0.5°C. The following illustrates the factorial experiment: 3 N 0 3 Cl2 2 TjJ N T ! 0 3 T i CI2T1 T 2 | N T 2 0 3 T 2 C 1 2 T 2 Therefore NT j 0 3 T ! C1 2 T! N T 2 0 3 T 2 C 1 2 T 2 P | P N T i PO3T1 PC1 2 T! P N T 2 P 0 3 T 2 PC1 2 T 2 and N T ! 0 3 T i C l 2 T i N T 2 0 3 T 2 C 1 2 T 2 B | B N T i B 0 3 T i B C l 2 T i B N T 2 B 0 3 T 2 B C 1 2 T 2 59 where: N = no sanitizing agent (control) O3 = ozone CI2 = chorine T l = temperature 1 T 2 = temperature 2 P planktonic cells of S. putrefaciens B = S. putrefaciens' biofilm Each treatment was replicated 3 times. Experimental data from the research was collected and analyzed by Analysis of Variance with Minitab using the General Linear Program and the Bonferroni Pairwise Comparisons (Minitab for Windows, release lOXtra. Minitab Inc., State College, PA). Hypothesis: The hypothesis for the planktonic and biofilm tests are as follows: HOplanktonic . There is no difference between treatments means; that is, u, =u 2 = u 3 ... = U6 Hapianktc,™: There is a difference between the means of at least two treatments HObiofta: There is no difference between treatments means; that is, U, =U2 = U 3 ... =1^ Harlan: There is a difference between the means of at least two treatments 60 3.2 Experimental Procedures 3.2.1 Glassware A l l glassware used in the experiment was first cleaned with tap water and left to soak overnight in a 2.0% solution of FL-70 detergent (Fisher Scientific). The glassware was subsequently thoroughly rinsed with tap water, followed by distilled-deionized water. The cleaned glassware was then filled with distilled-deionized water and placed in a plastic pail which was filled with distilled-deionized water until all the glassware was completely submerged. The glassware in the container (holding approximately 10 litres of water) was ozonated for at least 20 min until an ozone residual level >2.0 ppm was achieved. Following ozonation, the glassware was removed, wrapped in aluminum foil and dry heat sterilized at 180°C for 7 hr. This lengthy cleaning procedure derived from Domingue et al. (1988) ensured that no oxidizable residuals that could interfere with the measurement of ozone or its functionality as a sanitizing agent, remained on the glassware. A l l glassware used with ozone, chlorine or no sanitizer was cleaned and sterilized in this manner. 3.2.2 Water Due to the variability of components making up tap water from day to day, distilled-deionized water was used for all sanitizing tests. Distilled-deionized water was used for the ozonation and chlorination of both planktonic cells and the stainless steel chips coated with biofilms. Distilled-deionized water was also used in the preparation of indigo trisulfonate stock solutions and reagents and in any other circumstance where 61 water was required. The distilled-deionized water used in all tests had a pH of 6.5 ± 0.2. Distilled-deionized water required for ozone measurement purposes, was first rendered ozone demand free by ozonating it for 20 min until residual levels >1.0 ppm were obtained. This was followed by boiling the ozonated distilled-deionized water for an hour to remove any residual ozone. Once the ozone demand free water had cooled, it was measured spectrophotometrically at 600 nm using the indigo trisulfonate dye and the procedures indicated by Bader and Hoigne (1982). No trace of ozone was found in ozone demand free water made in this manner. Water used for the preparation of chlorine standards was rendered free of chlorine or chlorine demand by the addition of potassium permanganate to distilled-deionized water followed by distillation (Model 97-70 Residual Chlorine Electrode Instruction Manual, 1991). 3.2.3 Stainless Steel Chips Stainless steel chips (SS chips) of the following specifications: type 304 steel, finish No 4, 30x30 mm 2 area, 22 gauge thickness and a 3.175 mm (1/8") diameter perforation in one corner, were prepared for biofilm development and sanitizer studies by a modification of the method employed by Mustapha and Liewen (1989). Thus, the SS chips were cleaned by soaking them overnight in a 2.0% solution of FL-70 detergent (Fisher Scientific), followed by thorough scrubbing with a deluxe scrubbing sponge (Royal Sponge Mfg. Ltd.) , rinsing with tap water and distilled deionized water. The SS chips were then wrapped in aluminum foil and autoclaved at 121°C for 20 min. Sterile 62 forceps and a stainless steel rod with a hook at one end were used to aseptically handle the SS chips during their use; that is the reason for the perforation in one of the corner of each SS chip. 30 mm 3.175 mm diameter 22 gauge thickness 3.2.4 Planktonic Inoculum S. putrefaciens (A. T. C. C. No 8071, Rockville, MD) cultures frozen at -80°C in 1 mL of 50% glycerol solution were activated by consecutive transfers into TSB and TSA as described by Prahst (1994). The frozen vials were thawed and 1 mL transferred into a test tube (Length: 150 mm, ID: 15 mm) holding 5 mL of sterile TSB broth. These cultures were incubated for 24 hr at 10°C in a rotatory-type shaking water bath agitated at 100 rpm. One loop of these cultures was subsequently streaked onto TSA plates which were incubated at 25°C for 48 hr or longer until distinct individual colonies were obtained. A colony from a TSA plate was then transferred to two test tubes (Length: 150 mm, ID: 15 mm) containing 5 mL of TSB and incubated at 10°C for 24 hr in the rotatory-63 type shaking water bath at 100 rpm. One mL of each of these suspensions was then transferred to another set of test tubes containing 5 mL of TSB and incubated again at 10°C for 24 hr in the rotatory-type shaking water bath (Orbit Shaker Bath Model No 3543, Lab-Line Instruments, Inc., Melrose Park, 111.) at 100 rpm. This step was repeated one more time. After these series of inoculations and incubations, an individual 5 mL culture was transferred to each of two 250 mL erlenmeyer flasks containing 100 mL of TSB. These consecutive transfers were required to give the bacterial culture enough time to grow and adapt to the new incubation temperatures. The 100 mL microbial suspensions destined for experimental trials at 0.5° ± 0.5°C and 10° ± 0.5°C, were incubated for approximately 48 hr at 10°C in a rotatory-type shakipg water bath maintained at 120 rpm. The culture in the two flasks containing 100 mL of the microbial suspension were then transferred to a sterile centrifuge bottle and pelleted by centrifugation at 9000 g for 20 min at 10°C. The resulting supernatant was discarded while the bacterial pellet was re-suspended in 200 mL sterile, distilled-deionized water and centrifuged one more time at 9000 g for 20 min at 10°C. Once again the supernatant was discarded, the bacterial pellet re-suspended in 27 mL of sterile distilled-deionized water. This 27 mL bacterial suspension was then used as the inoculum for planktonic sanitizer tests. These steps are shown in Figure 5. In order to obtain approximately similar starting bacterial populations in each final 27 mL bacterial inoculum, a standard curve was made showing S. putrefaciens CFU/mL versus absorption at 600 nm over 4 days. This & putrefaciens suspension was cultured in the same manner as the planktonic inoculum; except that one of the final 5 64 1 colony Stock solution held @-80"C 1 mL 1 mL Centrifuge @ 9000g for 20 min @ 10"C I Bacterial pellet resuspended in 200 mL sterile distilled-deionized water Centrifuge @ 9000g for 20 min @ 10'C Bacterial pellet resuspended in 27 mL sterile distilled-deionized water Resuspended pellet used as planktonic inoculum either 1 L distilled-deionized water 1 L ozonated d-d water 1 L chlorinated d-d water I 1 mL samples taken every 30s. were diluted in 9 mL 0.194 peptone water containing 15 mg/L sodium sulfite Plated (hydrophobic grid-membrane method) on TSAc, TSAc + 0.5% and + 2, ,0% NaCl Figure 5. Experimental procedures for the preparation of planktonic inoculum. 65 mL of TBS used to inoculate a 100 mL of TBS was added to a 1000 mL erlenmeyer flask containing 400 mL of TBS. The 400 mL TSB suspension was incubated for 4 days at 10°C in the rotatory shaking water bath held at 120 rpm. Every 24 hr, three 10 mL samples were aseptically transferred to sterile test tubes. One mL from each sample was diluted in a series of 9 mL 0.1% peptone water test tubes and enumerated using the hydrophobic grid membrane method. The sterile Iso-grid 0.45p membrane filters (QA Life Sciences, Inc., San Diego, CA) employed in this method were placed on TSA containing 0.05 g/L congo red (Sigma Chemical, St. Louis, MO) and incubated at 25°C until distinct colonies were visible. Four mL from each sample were then used for spectrophotometric readings at 600 nm. Sterile TSB was used to standardize the spectrophotometer. Thus, prior to sanitizing tests, 4 mL samples of each 100 mL TSB suspensions grown for planktonic inoculum were measured at 600 nm. If the spectrophotometric values were close to the desired value of 0.180 ± 0.020 as determined previously by the standard curve, the 100 mL suspensions were centrifuged and the planktonic inoculum prepared as discussed in the previous paragraph. 3.2.5 Inoculation of Stainless Steel Chips The bacterial suspensions used to inoculate the SS chips were prepared in the same manner as the planktonic inocula; with the exception that only one of the 100 mL of TBS S. putrefaciens suspensions was prepared and incubated for approximately 24 hr at 10°C in the rotatory-type shaking water bath at 120 rpm. This suspension was not centrifuged but added to a 1000 mL erlenmeyer flask containing 300 mL of sterile TSB 66 which was in turn incubated under the same conditions for 48 hr. Twenty five mL of this suspension was then added to sterile 40 mm x 130 mm flat bottom, rimless culture tubes (Corning Glass Works, New York) containing 20 mL of sterile TSB. A stainless steel chip was subsequently added to each 45 mL TSB suspension used for each set of biofilm sanitizer tests. These test tubes containing the SS chips were incubated for 10 days at 10°C in a rotatory shaking water bath at 120 rpm. During every 48 hr of these 10 days incubation period, each SS chip was transferred aseptically to fresh 45 rnL sterile TSB tubes. After incubation, the SS chips were removed and rinsed by dipping and shaking the SS chips in 80 mL sterile distilled deionized water for about 15 sec. This step was repeated 3 times. One mL of the last rinse was surface plated on TSAc agar and incubated for 48 hr at 25°C to determine i f all non-attached cells were removed. This procedure was a modification of the method used by Krysinski et al. (1992). These steps are illustrated in Figure 6. 3.2.6 Growth of Biofilm Prior to any sanitizing tests involving biofilms, biofilms were grown under two different conditions. This was done to determine variability among biofilms grown from the same starting inoculum and those from different starting inocula. Thus, 15 SS chips were inoculated with S. putrefaciens as described in the previous section. A l l 25 mL S. putrefaciens suspensions used to inoculate each chip came from the same 300 mL inoculum. The SS chips were incubated for 10 days at 10°C in a rotatory shaking water bath at 120 rpm. During every 48 hr of these 10 days incubation period, each SS chip 67 1 colony 10 wide-flat bottom test tubes containing 20 mL sterile TSB i Test tubes containing the stainless steel chips were incubated for 10 days at 10°C in a rotatory shaking water bath at 120 rpm. The stainless steel chips were transferred aseptically to fresh sterile TSB media every 48 hr. I Stainless steel chips removed and rinsed by dipping and shaking in 80 mL sterile distilled-deionized water for 15 s. This step was repeated 3 times / lmL of the last rinse was plated on TSAc agar to determine i f all non-attached cells were removed Sanitation tests Figure 6. Experimental procedures for the preparation of biofilm inoculum. 68 was transferred aseptically to fresh 45 mL sterile TSB tubes. At the end of the incubation period, the SS chips were washed in 80 mL 0.1% peptone water for 15 s. The washing step was repeated three times. The SS chips were then transferred to 40 mm x 130 mm Flat bottom, rimless culture tubes (Corning Glass Works, New York) containing 45 mL of 0.1% peptone water containing 0.15 mg/L sodium sulfite (Sigma Chemical Company, St. Louis, MO) and 2 g of microscopic 100pm diameter glass beads (Xymotech Biosystems Inc., Quebec). The SS chips were then vortexed for 2 min with the aid of a Baxter S/P Vortex Mixer, Serial No 037882 (Baxter Diagnostics Inc., Deerfield, IL). One mL of these suspensions were enumerated using the hydrophobic grid membrane method. Another set of 15 SS chips were also inoculated with S. putrefaciens. However, each SS chip was inoculated with 25 mL of a Shewanella suspension derived from different starting 300 mL inocula of S. putrefaciens. These SS chips were incubated and enumerated in the same manner described above. 3.2.7 Ozone Generation and Measurement A Sander 301 ozonator (Sander, Uetze-ltze, Germany) was employed to produce ozone using pure, U.H.P grade oxygen containing < 3 ppm of water as the feed gas (Praxair, Delta, B.C). Ozone in the gas phase was monitored with a Erwin Sanders ozone gas phase monitor - model 451. The generated ozone was dispensed to the test solutions through Norprene tubing having an ID of 1/4", OD of 3/8" and a wall diameter of 1/16" (Fisher Scientific). A stainless steel diffuser was attached at the dispensing end of the tubing in order to deliver small uniform gas bubbles. A 50 mm magnetic rpd stirrer set at 69 565 rpm was used to thoroughly mix the ozone. A l l procedures involving generation of ozone and its application were carried out in a fume hood. This setup is illustrated in Figure 7. The desired ozone concentration of approximately 0.25 ppm was obtained by bubbling ozone for about 15 to 20 sec into a graduate cylinder (Length: 265 mm, ID: 80 mm) containing 1000 mL distilled-deionized water held at 10° ± 0.5°C or 0.5° ± 0.5°C. The ozone concentration in the gas stream was approximately 1 g 0 3 /m 3 with a 5 L/hr flow rate. The ozone was generated using 6.4 volts and a pressure of 1.35 atm. Ozone concentration was determined using the Indigo method as described by Bader and Hoigne (1982). The following formula was used to determine ozone concentration spectrophotometrically: Ozone concentration (mg 03/L) where b v f (AAbs • 10)/(f-b -v) path length of the cuvette (cm) volume of the sample added (mL) slope of the calibration curve at 600 nm (0.42 + 0.1 cm"1 per mg/L, with molar absorptivity = 20,000 M" 1 - cm - 1) 3.2.8 Chlorine Generation and Measurement Chlorine at a concentration of approximately 1.0 ppm, was prepared by diluting concentrated sodium hypochlorite (5.56% commercially produced JavexTm Bleach, 70 t Ozone delivery Ozone containing ^ -water Stainless steal aerator o • o 9 * * 0 • « 0 \*t * „ * ^ 0 o »0 * Bacterial inoculum E D * Biofilm covered stainless steel chips _^ Magnetic stirrer Figure 7. Ozonation of distilled-deionized water used for sanitizer tests. 71 Colgate-Palmolive, Toronto, ON) with distilled-deionized water until the desired concentration was reached. Chlorine concentration prior to and during sanitizing tests were determined using a residual chlorine (Cl 2) ion selective electrode model 97-70 (Orion Research, Inc. -V W R Scientific Products, Boston, MA) . The electrode was connected to a Corning pH/mV meter (Type N E 1604). Chlorine readings were obtained as millivolts. In order to convert chlorine concentration in millivolts to ppm, a standard curve had to be constructed. This standard curve constructed on semilogarithmic paper had millivolt values on the linear axis and concentration values as ppm chlorine on the logarithmic axis. At least two points were needed to make a chlorine calibration curve. These two points were obtained by making up a 1.0 ppm standardizing chlorine solution and by determining the slope of a tenfold change in chlorine concentration. The first point used was the millivolt value for the standardizing 1.0 ppm available chlorine solution. This 1.0 ppm standardizing solution was made by adding 1 mL residual chlorine standard (100 ppm as Cl 2 ) , 1 mL acetic acid reagent and 1 mL iodide reagent to 150 mL beaker and allowing it to stand for 2 min for complete reaction. After 2 min, 99 mL distilled-deionized chlorine demand free water was added. The residual chlorine ion electrode was then used to take the millivolt reading of the standardizing solution. The millivolt value of this solution was equivalent to 1.0 ppm available chlorine. Thus, the first point, 1.0 ppm available chlorine was obtained. The second point was obtained by determining the slope of a tenfold change in chlorine concentration. Thus, a solution of « 100 ppm chlorine was prepared by diluting 72 1 mL of a 5.56% hypochlorite solution (5.56% commercially produced JavexTra Bleach, Colgate-Palmolive, Toronto, ON) with distilled-deionized chlorine demand free water. In a 150 mL beaker, a 100 mL distilled-deionized chlorine demand free water was combined with 1 mL iodide reagent and 1 mL acetic acid reagent. One rnL of the * 100 ppm chlorine solution previously prepared was then added to the 150 mL beaker. The contents of the beaker were stirred gently for 2 min to allow a complete reaction. Then the millivolt value of the solution was taken. Thereupon, another 10 mL of the chlorinated solution was added to the solution in the 150 mL beaker. This new solution was again stirred gently for 2 min and its millivolt reading taken. The difference between these two potential readings is considered to be the slope of the electrode (tenfold change in C l 2 concentration). The second point in the graph was therefore obtained by adding this potential difference (« 29.5 mV/decade) to the value obtained for the standardized 1.0 ppm available chlorine solution. This value was equivalent to « 10.0 ppm available chlorine and was therefore used to construct the standard curve for available chlorine. This electrode method required 100 mL samples and the addition of two reagents, 1 mL of acetic acid reagent and lmL of iodide reagent which were purchased from Orion Research Inc. (Orion Research. Inc., Beverly, MA) . It is worth noting that the EPA has approved this electrode method for chlorine determination in drinking water and wastewater (EPA No 330.5) 73 3.2.9 Planktonic Inoculum - Control In order to determine the initial S. putrefaciens population in each 27 mL inoculum, three 1 mL samples were transferred to three test tubes containing 9 mL of 0.1% peptone water. These samples were serially diluted. One mL of these series of dilutions were then enumerated using the hydrophobic grid membrane method. The remaining 24 mL inoculum was then added to 1000 mL of distilled-deionized water held at either 10° ± 0.5°C or 0.5° ± 0.5°C. The suspension was thoroughly mixed with a 50 mm magnetic stirrer set at 565 rpm and the aid of Corning Laboratory Stirrer/Hot Plate, Model PC-320. The bacteria were thoroughly dispersed through the water in approximately 20 sec. One mL samples were then retrieved every 30 sec during a 3 min period. Each 1 mL sample was diluted with 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI) containing 15 mg/L sodium sulfite (Sigma Chemical Company, St. Louis, MO). Anhydrous sodium sulfite was used to neutralize both chlorine and ozone residuals (Domingue et al., 1988). Every 9 mL test tube of 0.1% peptone water used for microbial dilutions contained 15mg/L sodium sulfite. The temperature of the distilled-deionized water was maintained at 10° ± 0.5°C or 0.5° ± 0.5°C by placing the graduate cylinders in large plastic beakers containing iced water during the whole duration of the test run. The temperature of the distilled-deionized water and the iced water were monitored throughout the test run. 74 3.2.10 Planktonic Inoculum - Ozone Prior to any sanitizer tests involving ozone, tests were performed to determine i f ozone concentration remained constant during the time required for the sanitizer tests. Thus, ozone decay curves of ozonated distilled-deionized water at initial concentrations of «0.25 ppm were performed at 10° ± 0.5°C and 0.5° ± 0.5°C. During these tests, samples of ozonated water were taken for ozone measurements every 20 s for 10 min. Four replicates of ozone decay curves were performed at each temperature. The addition of the planktonic inocula to ozonated water was performed in the same manner as described in section 3.2.9. However, several problems were encountered when attempting to determine residual ozone concentrations at 30 second intervals using the indigo trisulfonate method as indicated by Bader and Hoigne (1982). First, the ozone readings as determined by the indigo trisulfonate method gave inconsistent results. These inconsistencies were believed to be due to interference of ozone concentration measurements by the presence of Shewanella putrefaciens cells in the suspension. The presence of suspended cells gave the impression of a decreased ozone concentration. Secondly, there also appeared to be an interaction taking place between the bacteria and /or its slime and the indigo trisulfonate. Centrifugation of the suspension resulted in the formation of a blue pellet. Thus, the indigo trisulfonate seemed to have been either absorbed into the bacterial cell and/or attached to its surface. When this suspension was tested for residual ozone, the suspended organisms with the attached indigo trisulfonate dye gave the appearance of a higher ozone concentration than was actually present. 75 Several modifications were made to correct this problem (Appendix 7.1.3). However, in most cases, it was found that ozone sensitivity to organic and inorganic materials interfered with attempts to measure it. The best alternative was to make a standard in which the bacterial cells interacting with the dye were removed. These standards would then be compared to an ozonated suspension of S. putrefaciens cells in which the bacterial cells interacting with the dye were also removed. Therefore, the difference between the standards and the test ozone solutions would give us an estimation of the decrease in ozone concentration over 3 min. The following are the modifications made to measure ozone concentration in planktonic suspension over the 3 min test runs. A. Two inocula of S. putrefaciens were prepared as described in section 3.2.4 of Materials and Methods. The 100 mL bacterial suspensions were first centrifuged separately. Each resulting pellets was resuspended in a 100 mL of sterile water and combined together. This 200 mL suspension was centrifuged once more and the resulting final pellet was re-suspended in 54 mL of sterile distilled-deionized water. This 54 mL suspension was thoroughly mixed and divided into two 27 mL aliquots that served as the two starting bacterial inocula. Three mL from each inoculum were used to determine the initial bacterial population. The remaining 24 mL were used for the sanitizing tests. B. Two graduate cylinders containing 1000 mL ozonated and distilled-deionized water were employed for the ozonation tests. 76 C. One of the inocula was added to a 1000 mL of distilled-deionized water. The distilled-deionized water and bacteria were agitated by a 50 mm magnetic stirrer set at 565 rpm. Once the bacteria were thoroughly dispersed through the water, 5 mL samples were taken and added to 3 mL of indigo trisulfonate solution. These indigo trisulfonate and suspension mixtures were then diluted with 2 mL of ozone demand free water. These 5 mL samples were taken every 30 sec. The final 10 mL indigo-bacterial suspensions were vortexed and then centrifuged at 9000g for 20 min. Supernatants from 10 mL suspensions were removed with a syringe attached to a 0.45 u filter unit (Millipore Products Division, Bedford, M A ) with a 16G 11/2 sterile disposable needle (Becton, Dickinson and Company, Rutherford, NJ). The filter removed any suspended organisms still present in the supernatant as well as the indigo dye attached to the cells. The filtered supernatants were placed into two cuvettes for spectrophotometric reading at 600 nm. They served as standard references for each 30 second interval during the 3 min sanitizer test using ozone. D. The second inoculum 24 mL, was added to another graduate cylinder containing approximately 1000 mL of 0.25 ppm ozonated distilled-deionized water. Every 30 sec for 3 min, a 5 mL sample was taken. The 5 mL sample was carefully added to 3 mL of indigo solution. The pipette tip was dipped into the indigo trisulfonate solution before the ozonated suspension was released. This prevented ozone loss to the atmosphere. For the same reason, the ozonated samples were released carefully so that bubbles were not formed and ozone lost. Once the indigo trisulfonate and the ozonated samples had reacted, 2 mL of ozone demand free water was added to each sample. The final 10 mL suspensions were vortexed and then centrifuged at 9000 g for 20 min. The supernatants were removed with a syringe attached to a 0.45 p filter unit (Millipore Products Division, Bedford, M A ) with a 16G 11/2 sterile disposable needle (Becton, Dickinson and Company, Rutherford, N.J.). The filtered supernatant contained only the decoloured indigo trisulfonate solution that reacted with the ozone. A l l S. putrefaciens cells and the indigo trisulfonate dye attached to them were removed. These filtered supernatants were used to measure ozone concentrations using the standards in part C. The filter did not appear to interact or interfere with ozone readings. These procedures were able to remove most of the interference caused by the presence of the bacteria and its adsorption of the indigo dye. Figure 8 illustrates the steps used for determining residual ozone concentration in the presence of S. putrefaciens. At the same time samples were being taken for ozone residual measurements, 1 mL samples were also taken every 30 sec for 3 min to enumerate S. putrefaciens survival. These samples were diluted in 9 mL of 0.1% peptone water containing 15 mg/L sodium sulfite. Sodium sulfite served to inactivate any remaining sanitizing agent. These diluted samples were further serially diluted and 1 mL aliquots used for enumeration of S. putrefaciens using the hydrophobic grid membrane method. 78 24 mL Shewanella putrefaciens inoculum 5 Standard 1000 mL distilled-deionized water Shewanella putrefaciens suspension 5 mL suspension 3 mL indigo solution 2 mL d-d water 1 sample retrieved every 30 s 1 vortex i Centrifuge all samples at 9000g for 20 min at 10°C I Filter supernatants I Absorbance at 600 nm value of std solution at 30 s value of std solution at 60 s etc... -4 24 mL Shewanella putrefaciens inoculum 1> Standard 1000 mL distilled-deionized water Shewanella putrefaciens suspension tf m ism 8*1 5 mL suspension 3 mL indigo solution 2 mL d-d water 1 sample retrieved every 30 s I vortex Centrifuge all samples at 9000g for 20 min at 10°C I Filter supernatants I Absorbance at 600 nm value of test solution at 30 s value of test solution at 6,0 s etc... Figure 8. Steps used to prepare samples from planktonic suspensions used in sanitizer tests for spectrophotometric determination of ozone concentration. 79 3.2.11 Planktonic Inoculum - Chlorine Prior to any sanitizing tests involving chlorine, tests were performed to determine if chlorine concentration remained constant during the time required for chlorine sanitation. Thus, decay curves of «1.0 ppm chlorinated d-d water were performed at 10° ± 0.5°C and 0.5°C ± 0.5°C. One hundred mL samples of » 1.0 ppm chlorinated d-d water were taken every minute for 10 min. Four replicates of chlorine decay curves were performed at each temperature. During the sanitizing tests, three 1 mL samples were taken from the 27 mL bacterial inoculum for the enumeration of the initial S. putrefaciens population per mL of inoculum. The remaining 24 mL was added to 1000 mL of 1.0 ppm chlorinated distilled-deionized water. The suspension was thoroughly mixed with a 50 mm magnetic stirrer set at 565 rpm. One mL samples were retrieved every 30 sec during a 3 min period. Each 1 mL sample was diluted with 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI) containing 15 mg/L sodium sulfite (Domingue et al., 1988). These dilutions were further serially diluted and 1 mL aliquots used for the enumeration of S. putrefaciens survival in 1.0 ppm chlorinated d-d water. Enumeration was done using the hydrophobic grid-membrane method. During these 3 min, a 100 mL aliquot was also being removed every minute to measure the chlorine concentration with a residual chlorine electrode. Unlike ozone, S. putrefaciens cells did not interfere with available chlorine concentration measurements. 80 3.2.12 Biofilm - Ozone and Chlorine After the biofilm-coated SS chips were rinsed, three SS chips were set aside and added to 45 mL of 0.1% peptone water neutralizing solution containing 15 mg/L sodium sulfite and 2 g of sterile microscopic 100pm diameter glass beads (Xymotech Biosystems Inc., Quebec). These SS chips were used as standards for the enumeration of S. putrefaciens present on SS chips prior to being submerged into distilled deionized water or either of the two sanitizers. The remaining SS chips were suspended from a stainless steel frame into a graduate cylinder containing 1000 mL of distilled-deionized water or either 0.25 ppm ozone or 1.0 ppm sodium hypochlorite solutions. The water in the graduate cylinder was stirred with a 50 mm magnetic stirrer set at 565 rpm using a Corning Stirrer/ Hot Plate (Model PC-320). Every 30 sec, a chip was removed from each of these solutions and placed in sterile 40 mm x 130 mm Flat bottom, rimless culture tube (Corning Glass Works, New York) containing 45 mL of 0.1% peptone water, 15 mg/L sodium sulfite and 2 g of sterile microscopic 100pm diameter glass beads. From the ozonated solutions, 5 mL samples were taken every 30 sec for ozone concentration measurements. In the case of the chlorinated solutions, 100 mL samples were retrieved every minute using a 100 mL syringe (Becton, Dickinson and Company, Rutherford, NJ). It was observed during the ozonation tests involving the SS chips that the microorganisms in the biofilm did not interfere with the measurement of residual ozone concentrations. Thus a standard as in the case of the planktonic suspensions was not required. 81 3.2.13 Planktonic Cells - Enumeration The dilution tubes containing the 1 mL test samples for the sanitizer tests were vortexed and serially diluted. One mL of these suspensions (or more i f required) were used to determine survival of S. putrefaciens using the hydrophobic grid-membrane method. The cell suspensions were filtered using 0.45p iso-grid sterile membranes (QA Life Sciences, Inc., San Diego, CA) which were later transferred to TSAc, TSAc + 0.5% NaCl and TSAc + 2.0% NaCl. The plates were incubated at 25°C for 48 hr or until visible colonies were obtained. 3.2.14 Biofilm Cells - Enumeration S. putrefaciens survival in biofilms exposed to the three sanitizing treatments was determined using an adaptation of the method employed by Oh and Marshall (1995). The SS chips exposed to the different sanitizers were placed in the 45 mL 0.1 % peptone solution containing sodium sulfite and the microscopic glass beads and vortexed for 2 min. After vortexing, 1 mL from each 30 sec sample was removed and added to 9 mL of 0.1% peptone water. These dilutions were further diluted in corresponding sterile 9 mL of peptone water. One mL of these dilutions (or more i f required) were then enumerated by the hydrophobic grid membrane method using 0.45 p iso-grid sterile membranes (QA Life Sciences, Inc., San Diego, CA). These membranes were plated on TSAc, TSAc + 0.5% NaCl and TSAc + 2.0% NaCl. A l l these plates were incubated at 25°C for 48 hr or until visible colonies were observed. 82 4. RESULTS AND DISCUSSION 4.1 Decay Curves Prior to sanitizing tests, decay curves for 0.25 ppm ozonated and 1.0 ppm chlorinated distilled-deionized water at 10° ± 0.5°C and 0.5° ± 0.5°C were performed. The decay curves are shown in Figures 9 to 12. Decay curves were done in order to determine i f both dissolved ozone and chlorine concentrations remained constant during the time frame used for sanitizer tests. As illustrated in these figures there was very little change in ozone concentration during the 10 min trials. Chlorine concentration remained the same during this time. Temperature did not appear to be have an effect on either ozone or chlorine decomposition during the testing period. The stability of ozone and chlorine during this first 10 min is important since it permits us to put aside the effects of ozone and chlorine decomposition during the first few minutes used for sanitizer tests. Thus, during sanitizer tests involving ozone and chlorine any decrease in ozone and chlorine concentration during the crucial testing period can be assumed to be the result of ozone or chlorine-microbial interactions and not due to the natural decay of ozone or chlorine. 4.2 Planktonic Suspensions - Control Figures 13 and 14 illustrate the effect of distilled-deionized water held at 10° ± 0.5°C and 0.5° ± 0.5°C on S. putrefaciens suspensions. S. putrefaciefis populations whether in the planktonic or biofilm forms were enumerated on three media; TSAc, TSAc + 0.5% NaCl and TSAc + 2.0% NaCl. TSAc served as the non-selective media 83 84 (wdd) uojjejjueouoo euozo 85 M l Hi * <= 5 5 •= I * * t o O LO o -H o o o o (O (LUfJd) U0|}BJ1U90U00 8UU0|LjQ o CM O CD <D Hi * * Hi * * o CO CO CO CD Hi * * Hi * * jD CO -a N 'c o CD 1 "O _gj CO X3 C T3 CD _> O CO CO T3 CD c Hi * * o O CD O CD Q Hi * * 44* c • s c 2 2 2 2 to "g "2 5 T - CM CO o o 00 4 4 * o CM *** o CD CO OT ^ CD OT E ^ o o CO * 4 * o CM o CD o o o co o o CM (wdd) uoiiejiuaouoo a u u o m o 87 while TSAc + 0.5% and 2.0% NaCl were used as the selective media. Every point on the graph is an average of three points from 3 replicate runs. In Figure 13, the number of S. putrefaciens CFU/mL present in the water when enumerated with TSA showed relatively little change from the starting population of 1.1 x 106 CFU/mL. The average population over the 3 min was 1.1 x 106 CFU/mL + 3.3 x 104 CFU/mL. This means that there was, on average, a 2.9 % difference between samples taken every 30 sec over the 3 min testing period. 5*. putrefaciens did not grow as well in TSAc + 0.5% NaCl when compared to counts on TSAc. The average starting population on TSAc + 0.5% NaCl was 8.7 x 105 CFU/mL. The average population over the 3 min testing period was 8.4 x 105 CFU/mL ± 3.9 x 104 CFU/mL. Thus, there was a 4.6 % variation in CFU/mL from samples taken every 30 sec over the 3 min testing period. The lower counts on this medium were likely due to some of the organisms being sensitive to 0.5% NaCl. In this particular case, it appeared that about 23.2 % of the organisms could not grow on TSAc + 0.5% NaCl. S. putrefaciens counts on TSAc + 2.0% NaCl were even lower. The average starting population on TSAc + 2.0% NaCl was only 5.4 x 103 CFU/mL. Thus, approximately 99.5% of the cells could not grow on TSAc + 2.0% NaCl. The average population over the 3 min exposure was 5.2 x 103 CFU/mL ± 1.4 x 102 CFU/mL; a 2.6 % variation in CFU/mL from samples taken every 30 sec over the 3 min testing period. Figure 14 shows similar results as those observed with S. putrefaciens suspensions exposed to 10° ± 0.5°C distilled-deionized water. On TSAc, the initial population prior to exposure to distilled-deionized water at 0.5° ± 0.5°C was 1.2 x 88 1.00E+07 1 .OOE+06 £ 1.00E+05 + 1.00E+04 + E B LL o CD O 1.00E+03 + 1.00E+02 + 1.00E+01 + 1.00E+00 -•—TSAc + no NaCl -B—TSAc + 0.5% NaCl -A—TSAc + 2.0% NaCl 30 60 90 120 150 180 Time (s) Fig. 13. Effect of distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests). 89 1.00E+07 1 .OOE+06 1.00E+05 + 1.00E+04 + Z) o Ui o 1 .OOE+03 + 1.00E+02 + 1 .OOE+01 + 1 .OOE+00 •4--•—TSAc + NaCl - « — TSAc + 0.5% NaCl -A—TSAc + 2.0% NaCl 30 60 90 Time (s) 120 150 180 Fig. 14. Effect of distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests) 90 106 CFU/mL. The average population during the 3 min testing period was 1.2 x 106 CFU/mL ± 6.2 x 104 CFU/mL. This represented a 5.1% variation among the samples taken very 30 sec. The initial population on TSAc + 0.5% NaCl was 9.9 x 10s CFU/mL. The average population during the 3 min test period was 9.5 x 105 CFU/mL ± 5.2 x 104 CFU/mL. Thus, there was a 5.5% variation among samples taken over a period of 3 min. Finally, the initial population on TSAc + 2.0% NaCl media was 2.1 x 103 CFU/mL. The average population over 3 min was 2.1 x 103 CFU/mL ± 2.2 x 102 CFU/mL. This is a 10.7 % variation among samples retrieved during the 3 min test. S. putrefaciens did not grow as well on TSAc + 0.5% NaCl and TSAc + 2.0% NaCl. It appeared that 19.4 % of the organisms could not grow on TSAc + 0.5% NaCl and 99.8% could not grow on TSAc + 2.0% NaCl. The percentage of organisms that did not grow on either media were similar in both planktonic treatments. The degree of variation among samples taken every 30 sec was relatively small considering these values were the averages of three replicates. In addition, mixing of the suspension, as was done in this case, does not guarantee uniform distribution of the organisms. Subsequent dilutions may have also increased these differences. The larger differences observed among these samples as the NaCl concentration was increased from 0.5 to 2.0% may have been due to the increasing sensitivity of the organisms to NaCl. A l l considered, S. putrefaciens populations exposed to distilled-deionized water at 10° ± 0.5°C and 0.5° ± 0.5°C remained relatively constant throughout the 3 min testing period. The data showed, that d-d water at 10° ± 0.5°C and 0.5° ± 0.5°C had little or no effect on the S. putrefaciens population. Consequently, it can be assumed that any decrease in the 91 number of S. putrefaciens organisms exposed to ozonated or chlorinated d-d water would be due to the presence of dissolved ozone or chlorine in the d-d water and not due to the d-d water. 4.3 Biofilm - Control Figures 15 and 16 illustrates the effects of distilled-deionized water at 10° ± 0.5°C and 0.5° ± 0.5°C on biofilm covered SS chips. These figures show a similar trend as that observed with planktonic suspensions of S. putrefaciens exposed to similar testing conditions. Basically, d-d water appeared to have little or no effect on the S. putrefaciens population. Figures 15 and 16 show small variation among the S. putrefaciens biofilm population throughout the 3 min test. The biofilm populations on the SS chips retrieved throughout the 3 min testing period did not vary greatly from the initial populations. The initial populations were determined by averaging the biofilm population of 3 of the 10 SS chips used in the sanitizer tests. In Figure 15 the initial CFU/cm 2 on TSAc prior to exposure to 10° ± 0.5°C d-d water was 1.1 x 107 CFU/cm 2 . The average number of CFU/cm 2 from these SS chips during the 3 min test was 9.9 x 106 CFU/cm 2 ± 9.3 x 105 CFU/cm 2 . Hence, there was an average of 9.4% variation among the biofilm population from the SS chips exposed to 10° ± 0.5°C d-d water. On the other hand, the starting population on TSAc+0.5% NaCl was 8.6 x 106 CFU/cm 2 . The average population during the 3 min test was 7.9 x 106 CFU/cm 2 ± 6.5 x 105 CFU/cm 2 . Thus, in TSAc + 0.5% NaCl there was a 8.1% variation among the biofilm population from the SS phips retrieved 92 1.00E+08 1.00E+07 i : 1 .OOE+06 1.00E+05 + E L L 1 .OOE+04 o cn o 1.00E+03 + 1.00E+02 + 1 .OOE+01 + 1.00E+00 30 60 1 90 Time (s) -fi--•—TSAc + no NaCl -®—TSAc + 0.5% NaCl -A— TSAc + 2.0% NaCl 120 150 180 Fig. 15. Effect of distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 93 1.00E+08 1 .OOE+07 1.00E+06 + 1.00E+05 + E o L L 1.00E+04 o CD O 1.00E+03 + 1.00E+02 + 1.00E+01 + 1.00E+00 4 30 60 90 Time (s) -•—TSAc + no NaCl - O — T S A c + 0.5% NaCl -A—TSAc+ 2.0% NaCl 120 150 180 Fig. 16. Effect of distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 94 throughout the 3 min. The starting S. putrefaciens population on TSAc + 2.0% NaCl prior to exposure to 10° ± 0.5°C d-d water was 9.8 x 105 CFU/cm 2 . The average CFU/cm 2 during the 3 min test was 9.2 x lO 5 CFU/cm 2 ± 5.4 x 104 CFU/cm 2 . This indicated that on TSAc + 2.0% NaCl, there was 5.9% variation among the biofilm populations from the sampled SS chips. Figure 16 illustrates the effects of biofilm covered SS chips exposed to d-d water at 0.5° ± 0.5°C. As noted earlier, the results were similar to those of biofilms exposed to 10° ± 0.5°C d-d water. The initial population in TSAc prior to exposure to 0.5° ± 0.5°C d-d water was 1.2 x 107 CFU/cm 2 The average population during the 3 min test was 1.1 x 107 CFU/cm 2 ± 5.3 x 105 CFU/cm 2 . Consequently, on TSAc, there was a 4.7% variation among the biofilm population. On TSAc + 0.5% NaCl, the initial population was 9.0 x 106 CFU/cm 2 . The average population during the test was 8.5 x 106 CFU/cm 2 ± 5.7 x 105 CFU/cm 2 . On TSAc + 0.5% NaCl, there was a 6.7% variation in the biofilm population during the 3 min test. Finally, in TSAc + 2.0% NaCl, the initial population was 5.4 x 105 CFU/cm 2 . The average biofilm population during the test was 5.4 x 105 CFU/cm 2 ± 4.0 x 104 CFU/cm 2 . This corresponds to a 7.5% variation among the biofilm population from the SS chips retrieved every 30 sec throughout the 3 min test. Once again, it was also observed that the organisms did not grow as well on TSAc + 0.5% NaCl and TSAc + 2.0% NaCl media as compared to the TSAc. Of the organisms exposed tp 10° ± 0.5°C d-d water 20.7% did not grow on TSAc + 0.5% NaCl and 90.9% did not grow on TSAc + 2.0% NaCl. Similarly, 23.0% and 95.4% of the organisms exposed to 0.5° ± 0.5°C d-d water did not grow on TSAc + 0.5% NaCl and TSAc + 2.0% NaCl, respectively. 95 These results suggest that d-d at either 10° ± 0.5°C or 0.5° ± 0.5°C had minimal or no biocidal effect on S. putrefaciens biofilms. However, it is interesting to note that the biofilms appeared to have provided some kind of protection to cells which enabled them to grow better when plated on media containing 2.0% NaCl. For example, prior to exposure to 10° ± 0.5°C d-d water, only 5.4 x 103 CFU/mL of the starting planktonic S. putrefaciens population grew on TSAc + 2.0% NaCl; that is, approximately 0.5% of the initial population. On the other hand, 9.9 x 105 CFU/cm 2 of the starting S. putrefaciens biofilm population grew on TSAc + 2.0% NaCl; that is, approximately 9.0% of the initial population. This difference was maintained throughout the 3 min test. The S. putrefaciens planktonic and biofilm populations exposed to d-d water and plated on TSAc, TSAc + 0.5% and 2.0% NaCl were incubated for 48 hr at 25°C. Results from sanitizer tests involving d-d water at 10° ± 0.5°C and 0.5° ± 0.5°C, suggest that d-d water had minimal or no effect on S. putrefaciens biofilms. Consequently, decreases in biofilm populations exposed to either ozonatecj or chlorinated d-d water would be due to the effects of ozone or chlorine on the microorganisms and not due to d-d water. Temperature, 10° ± 0.5°C and 0.5° ± 0.5°C, also appeared to have minimal effects on the survival of S. putrefaciens exposed to d-d water. 4.4 Planktonic Suspensions - Ozone The effect of 0.25 ppm ozonated d-d water at 10° ± 0.5°C and 0.5° ± 0.5°C on S. putrefaciens suspensions are illustrated on Figures 17 and 18. Both figures show similar patterns of cell inactivation. 96 Fig. 17. Effect of ozonated distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests) 97 1 .OOE+07 1.00E+06§ 1.00E+05 + 1.00E+04 + Z> Ll_ o O 1.00E+03 + 1.00E+02 + 1 .OOE+01 + 1.00E+00 -H TSAc + no NaCl -A TSAc + 0.5% NaCl -H TSAc + 2.0% NaCl • - - Ozone concentration + 0.05 - X -30 60 90 Time (s) 120 150 180 Fig. 18. Effect of ozonated distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests) 98 S. putrefaciens exposed to 0.25 ppm ozonated d-d water at 10° ± 0.5°C exhibited a sharp decrease in CFU/mL after only 30 sec exposure to ozone. This sharp decrease in CFU/mL was observed with all three media. On TSAc plates, the starting S. putrefaciens population prior to exposure to ozone was 1.1 x 106 CFU/mL. After 30 sec, only 4.1 x 103 CFU/mL could be detected. This is about a 23/5 log decrease in the starting number of S. putrefaciens organisms. During this time, ozone decreased from an initial concentration of 0.25 ppm to 0.04 ppm. There was no detectable ozone after 30 sec. S. putrefaciens population decreased slightly after 30 sec. After 3 min, 1.8 x 103 CFU/mL were detected; a total 24/s log decrease in the number of starting organisms. Thus, after the first 30 sec, the S. putrefaciens population remained relatively stable. Similarly, the starting population on TSAc + 0.5% NaCl prior to ozone exposure was 8.2 x 105 CFU/mL. Again, there was a sharp decrease in the number of surviving organisms after 30 sec. After 30 sec, only 3.1 x 103 CFU/mL were detected. This is about a 23/s log decrease in the starting population. At 3 min, 1.3 x 103 CFU/mL were detected; approximately a 24/s log decrease from the initial population. The starting population on TSAc + 2.0% NaCl media was 2.2 x 104 CFU/mL. After 30 sec, only 6.2 x 101 CFU/mL were detected. This was approximately a 27/io log decrease. During the remainder of the test, the S. putrefaciens surviving population remained stable. Figure 18 shows a similar survival pattern for organisms exposed to 0.25 ppm ozonated d-d water held at 0.5° ± 0.5°C as those exposed to ozonated d-d water at 10° ± 0.5°C. On TSAc the starting population prior to exposure to ozone was 1.1 x 106 CFU/mL. After 30 sec, the original population decreased to 2.3 x 103 CFU/mL; 99 approximately a 24/s log decrease. After 3 min, 1.8 x 103 CFU/mL were detected. Thus, the population remained fairly constant after the first 30 sec. Ozone concentration decreased from 0.25 ppm to 0.03 ppm in only 30 sec. By 60 sec, there was no detectable ozone in the system. The starting population on TSAc + 0.5% NaCl was 8.8 x 105 CFU/mL. After 30sec, it decreased to 1.8 x 103 CFU/mL; about a 24/s log decrease. Once again, the surviving number of organisms remained constant during the remaining 2V2 min of the test run. The starting population on TSAc + 2.0% NaCl was 4.0 x 103 CFU/mL. After 30 sec, it decreased to 7.7 x 10° CFU/mL; a 24/5 log decrease. There was no apparent change in number of surviving organisms during the remainder of the test. It appeared that the inactivation of S. putrefaciens population by 0.25 ppm ozonated water took place almost immediately after the organisms and ozone came into contact. Once the ozone was depleted, the population stabilized. From the inactivation patterns observed on Figures 17 and 18, it appeared that ozone had an all or none effect on the planktonic organisms since injury could not be detected from the inactivation curves for TSAc media containing 0.5% and 2.0 % NaCl. However, it was noted that ozonated cells grew at a slower rate than those which were not ozonated; that is, as compared to the starting population at time zero which were not ozonated. Colonies that had been exposed to ozone and later plated on TSAc and TSAc + 0.5% NaCl were smaller than their non-ozonated counterparts (starting population at time zero). They were incubated for at least 12 hr longer than planktonic cells exposed to d-d water. Similarly, organisms growing on the 2.0% NaCl media grew at a s lower rate than organisms treated with ozone and plated on TSAc or TSAc + 0.5% NaCl. The colonies"' 100 on the TSAc + 2.0% NaCl were smaller than their non-ozonated counterparts grown on TSAc +2.0% NaCl (starting population at time zero). Consequently, they were incubated for at least 24 hr longer than planktonic cells treated with d-d water. Therefore, even though no injury could be detected by examining the graphical data, ozone appears to have affected the surviving organisms in some manner. Thus, the reported all or none effect does not seen to be true in this case. Temperature, 10° ± 0.5°C and 0.5° ± 0.5°C, did not appear to have influenced the results in any substantial manner. Both treatments gave very close results. 4.5 Biofilm - Ozone The effects of 0.25 ppm ozonated d-d water on S. putrefaciens biofilm are illustrated in Figures 19 and 20. Both of these figures show a similar pattern of ozone consumption and microbial inactivation. There was a sharp decrease in the number of surviving organisms after 30 sec exposure to 0.25 ppm ozonated d-d water followed by a slow decline over the remainder of the 3 min test. This pattern of microbial survival was similar on all enumeration media. Likewise, ozone concentration declined gradually but it still remained quite high after 3 min. In Figure 19, the average starting biofilm population prior to exposure to ozonated water held at 10° ± 0.5°C was 1.3 x 107 CFU/cm 2 when enumerated with TSAc. After 30 sec, it decreased to 2.2 x 105 CFU/cm 2 , about al 4/s log decrease in the number of starting organisms. The number of surviving organisms decreased slowly over the remaining 2xli min. After 3 min, only 4.2 x 104 CFU/cm 2 were detected. Thus, the population had been 101 1.00E+08 1.00E+07 + 0.25 + 0.20 -* ± 0.15 -TSAc + no NaCl -TSAc + 0.5% NaCl -TSAc + 2.0% NaCl - Ozone concentration 1.00E+01 + 1.00E+00 + 0.10 E Q_ CL c o ra c 0) o c o o 0 c o N o + 0.05 to.oo 30 60 90 120 150 180 Time (s) Fig. 19. Effect of ozonated distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 102 1.00E+02 + 1.00E+01 + 1.00E+00 - H — T S A c + no NaCl -A TSAc + 0.5% NaCl -H TSAc + 2.0% NaCl • - - Ozone concentration - f -+ 0.05 30 60 90 Time (s) 120 150 180 Fig. 20. Effect of ozonated distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 103 decreased by about 27/io logs. On TSAc+0.5% NaCl, the starting population was 1.0 x 107 CFU/cm 2 . After 30 sec, the population was reduced to 1.5 x 105 CFU/cm 2 ; a l 4/s log decrease. During the remaining 2xli min, the surviving population gradually decreased to 3.4 x 104 CFU/cm 2 ; close to a 27/io log decrease. The starting populatipn on TSAc + 2.0% NaCl was 4.5 x 105 CFU/cm 2 . After 30 sec, it decreased to 5.2 x 103 CFU/cm 2 ; almost a 2 log decrease. After 3 min, the population was 8.6 x 102 CFU/cm 2 ; a 24/s log decrease from the initial population. On the other hand, ozone concentration declined slowly from an initial concentration of 0.25 ppm to 0.17 ppm after three min. Biofilm exposed to 0.25 ppm ozonated d-d water at 0.5° ± 0.5°C, showed the same pattern of microbial inactivation. On TSAc, the starting population prior to exposure to ozonated water was 1.4 x 107 CFU/cm 2 . After 30 sec, it decreased to 6.1 x 105 CFU/cm 2 . This constituted about a IV2 log decrease from the initial population. After the remaining 2V2 min the surviving population decreased to 1.2 x 105 CFU/cm 2 ; that is, about a 2Vio log decrease. On TSAc + 0.5% NaCl, the starting population was 1.0 x 107 CFU/cm 2 . After 30 sec, it decreased to 4.6 x 105 CFU/cm 2 ; about a VI2 log decrease. The population gradually decreased to 7.2 x 104 CFU/cm 2 after 3 min exposure to ozone. This was a 23/io log decrease from the initial number of organisms. Finally, on TSAc + 2.0% NaCl, the starting population of 3.3 x 105 CFU/cm 2 decreased to 4.4 x 103 CFU/cm 2 after 30 sec; approximately a l 4/s log decrease. At the end of the 3 min test, the population had gradually decreased to 1.9 x 103 CFU/cm 2 ; a 22/5 log decrease from the initial population. During the 3 min test, ozone concentration also decreased gradually from a starting concentration of 0.25 ppm to 0.19 ppm. 104 S. putrefaciens counts on TSAc + 0.5% NaCl and TSAc + 2.0% NaCl revealed no injury as a result of ozonation. There was no need for longer incubation of the inoculated plates as in the case of the planktonic cells exposed to ozone because the incubation times for the ozonated biofilm cells were similar to those of untreated biofilm cells. Thus, no assumptions of injury could be made based on extra recovery time. In this case, the data agrees with the reported all or none effect of ozone on microorganisms. Once again, the test temperatures, 10° ± 0.5°C and 0.5° ± 0.5°C, appeared to have minimal effects on the efficiency of ozone as a sanitizer. The sanitizer test data illustrates different ozone microbial inactivation patterns for planktonic and biofilms cells. In Figures 17 and 18, the inactivation of planktonic cells by ozone took place in just 30 sec. Most of the ozone was consumed within this 30 sec. The rapid consumption of ozone was most likely due to the increased surface area available for its interaction with the organisms and their extracellular polymers. At the end of the 3 min test, there was no ozone in the system and there were a considerable number of surviving organisms. On the other hand, in Figures 19 and 20 there was a gradual decrease in the number of surviving organisms and ozone concentration. Although at the end of the test there was a large number of surviving organisms in the biofilm, there was also a substantial concentration of ozone in the system. The data also show that a considerable proportion of the organisms in the biofilms were killed during the first 30 sec. After the initial 30 sec exposure, there was a gradual decrease in the number of surviving organisms. This suggests, that ozone first attacked the organisms present on the surface of the biofilm on the SS chips. However, it 105 was less effective in inactivating the organisms embedded in the underlying matrix of the biofilm. It is possible that as ozone came in contact with the surface of the biofilm and made its way deeper into the biofilm matrix, its efficiency was diminished by its interaction with the organic matter making up the biofilm. Thus, the first layers of the biofilm may have acted as a barrier towards further penetration of ozone into the biofilm, therefore, decreasing its efficiency as a sanitizing agent for S. putrefaciens biofilms as compared to planktonic cells. 4.6 Planktonic Suspensions - Chlorine Figures 21 and 22 illustrate the effect of 1.0 ppm chlorinated d-d water on S. putrefaciens suspensions. Both figures show similar patterns of chlorine inactivation of S. putrefaciens organisms. In Figure 21, the 10° ± 0.5°C chlorine in the chlorinated d-d water was almost depleted after only 60 sec. At the end of the 3 min test, there was no detectable chlorine in the system. On the other hand, the S. putrefaciens population suffered a sharp decrease during the first 30 sec exposure to chlorine. The population only decreased slightly during the remaining 272 min of the test. This is due to the fact that there was basically no chlorine left in the system to interact with the organisms. The pattern of chlorine inactivation of planktonic S. putrefaciens cells was similar in all three enumeration media. On TSAc, the starting population prior to exposure to 1.0 ppm chlorine was 1.2 x 106 CFU/mL. After 30 sec, the population decreased to 1.9 x 105 CFU/mL; this was a 4 / 5 of a log decrease. After 3 min, there were only 1.4 x 105 106 1.00E+07 1.00E+06 1.00E+05 + .1.00E+04 + E o o '1.00E+03 + 1.00E+02 + 1.00E+01 + 1.00E+00 Time (s) Fig. 21. Effect of chlorinated distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests) 107 1.00E+07 + 1.00 1.00E+06 1.00E+05 + ,1.00E+04 + E l_L O cn o J1.00E+03 + 1.00E+02 + 1.00E+01 + 1.00E+00 " • E 1 1 1 1 *~ u ' )" 1 • . « —i i X ' T — 1 HH TSAc + no NaCl -A TSAc+ 0.5% NaCl -X—TSAc+ 2.0% NaCl • - - Chlorine concentration -+• 0.80 E CL CL C + 0.60 B CO c CD O c o o CD c + 0.40 O O + 0.20 0.00 30 60 90 Time (s) 120 150 180 Fig. 22. Effect of chlorinated distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens suspension. Bars represent standard deviations (derived from three replicate tests) 108 CFU/mL. This was only a 9 / i 0 of a log decrease in the initial population. On TSAc + 0.5% NaCl, the initial population was 1.1 x 106 CFU/mL. After 30 sec exposure to 1.0 ppm chlorinated water, the population decreased to 1.4 x 105 CFU/mL which corresponded to a 9 / 1 0 of a log decrease. After 3 min, the population decreased slightly to 9.3 x 104 CFU/mL, which was about lVio log decrease from the initial population. Finally, the initial population on TSAc + 2.0% NaCl was 6.1 x 103 CFU/mL. The population decreased to 5.8 x 102 CFU/mL; that is, 1 log during the first 30 sec of exposure to 1.0 ppm chlorine. After 3 min, the population decreased slightly to 4.2 x 102 CFU/mL; a l 3/io log decrease from the initial population. S. putrefaciens planktonic cells exposed to 1.0 ppm chlorinated d-d water held at 0.5° ± 0.5°C experienced similar decreases in S. putrefaciens population and chlorine concentration as S. putrefaciens planktonic cells exposed to 1.0 ppm chlorinated d-d water held at 10° ± 0.5°C. Chlorine concentration decreased from 1.1 ppm to 0.03 ppm in only 60 sec. After 60 sec, there was no detectable chlorine in the system. On TSAc, the starting population of 1.3 x 106 CFU/mL decreased to 1.8 x 105 CFU/mL during the first 30 sec exposure to chlorine. This was close to 9/io of a log decrease from the initial population. After 3 min the population was 1.6 x 105 CFU/mL, also approximately 9 / i 0 of a log decrease in the initial population. On TSAc + 0.5% NaCl, the initial population of 1.1 x 106 CFU/mL decreased to 1.3 x 105 CFU/mL after 30 sec of exposure to chlorine. This was about 9 / i 0 of a log decrease from the number of starting organisms. By the end of the 3 min test, this population was decreased to 9.5 x 104 CFU/cm 2; that is, lVio log decrease from the initial population . The starting population on the TSAc + 2.0% NaCl 109 decreased from 6.2 x 103 CFU/mL to 9.3 x 102 CFU/mL during the first 30 sec of exposure to chlorine. This corresponds to a bit more than 8 / 1 0 of a log decrease from the initial number of organisms. This population further decreased to 7.5 x 102 CFU/mL during the remaining 2V2 min of the test. Thus, the initial population decreased by close to 9/ioof a log. Examination of the inactivation data revealed no microbial injury. However, planktonic cells exposed to chlorine and grown on 2.0% NaCl required longer incubation times. After 48 hr incubation at 25°C, these colonies were smaller than their non-chlorinated counterparts grown on TSAc + 2.0% NaCl (starting population at time zero). Thus, they were incubated for an extra 24 hr prior to enumeration. This suggested that the surviving organisms may have been affected by the chlorine. The temperatures employed in these tests, 10° ± 0.5°C and 0.5° ± 0.5°C, did not appear to have affected the efficiency of chlorine as a sanitizer. Chlorine inactivation of planktonic cells appeared to follow a similar inactivation trend as ozone inactivation of planktonic cells. Most of the killing occurred in the first 30 sec. Also, most of the chlorine was probably gone in the first 30 sec. Unfortunately, chlorine concentration could not be measured every 30 sec but only every min due to the large amount of sample (100 mL) required to measure chlorine concentration. This rapid decrease in both organisms and chlorine concentration is again probably due to the increased surface area available for interaction between the organisms and chlorine. However, ozone appears to be much more effective in inactivating planktonic cells than chlorine. These planktonic tests have shown that ozonated d-d water at 0.25 ppm was > 110 70 times more effective in inactivating Shewanella putrefaciens than 1.0 ppm chlorinated d-d water. 4.7 Biofilm - Chlorine The effects of 1.0 ppm chlorinated d-d water held at 10° ± 0.5°C and 0.5° ± 0.5°C are illustrated in Figures 23 and 24. Both of these figures show similar patterns of S. putrefaciens inactivation and chlorine consumption. In biofilms exposed to 1.0 ppm chlorinated d-d water at 10° ± 0.5°C, there was a gradual decrease in chlorine consumption from 1.0 ppm to 0.8 ppm after 3 min. On TSAc, the initial population suffered a sharp decrease during }he first 30 sec, from 1.1 x 107 CFU/cm 2 to 4.4 x 105 CFU/cm 2 . This was a Vh log decrease in the initial number of organisms. After 90 sec, the population decreased to 2.9 x 104 CFU/cm 2 . This corresponded to a 27/io log decrease in the initial number of organisms. After 120 sec, the S. putrefaciens population was 3.1 x 103 CFU/cm 2 ; a 37/io log decrease from the initial population. Finally, after 3 min only 8.4 x 102 CFU/cm 2 remained. This corresponded to a full 4 log decrease in the initial population after 3 min of exposure to chlorine. The same general results were observed for S. putrefaciens plated on TSAc + 0.5% NaCl. The initial population of 8.8 x 106 CFU/cm 2 decreased to 2.5 x 105 CFU/cm 2 after 30 sec of exposure to chlorine; that was a l 7/io log decrease. After 60 sec, it dropped to 3.4 x 104 CFU/cm 2 ; that was a 23/s log decrease. After 120 sec it further dropped to 2.0 x 103 CFU/cm 2 ; a 3 4A log decrease from initial number. Finally, at the end of the 3 min exposure to chlorine the initial population was decreased to 4.6 x 102 CFU/cm 2 ; which was a 4 log decrease. The number of & putrefaciens that could grow on TSAc + 2.0% NaCl decreased from 2.3 x I l l 1.00E+08 Fig. 23. Effect of chlorinated distilled-deionized water (10°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 112 1.00E+08 1.00E+07 1.00E+00 + 1 .OOE-01 + 1.00 + 0.80 + 0.60 + 0.40 E CL Q_ C o ••—» CO l_ c CD O c o o 0) c o sz o + 0.20 0.00 180 Fig. 24. Effect of chlorinated distilled-deionized water (0.5°C ± 0.5°C) on Shewanella putrefaciens biofilm. Bars represent standard deviations (derived from three replicate tests) 113 105 CFU/cm 2 to 2.7 x 101 CFU/cm 2 in just 30 sec. This was almost a 4 log decrease. At the end of the 3 min test the initial population decreased to 1.2 x 10° CFU/cm 2 ; a 5 V2 log decrease in the initial population. This suggests that part of the surviving population was injured because a larger apparent kill was observed by plating on TSAc + 2.0% NaCl than on TSAc. The biocidal effects of 1.0 ppm chlorinated d-d water at 0.5° ± 0.5°C on S. putrefaciens biofilm were very similar to those observed at 10 °± 0.5°C. Chlorine concentration decreased gradually from 1.0 ppm to 0.7 ppm in 3 min. At the same time, S. putrefaciens population on TSAc decreased from 1.1 x 107 CFU/cm 2 to 4.4 x 105 CFU/cm 2 ; a V/5 log decrease in just 30 sec. The population further decreased to 2.4 x 103 CFU/cm 2 after 120 sec, which was a 34/s log decrease from the initial number of organisms. After 3 min exposure to chlorine, the population decreased to 1.3 x 103 CFU/cm 2 ; a 4 log decrease. The initial population that grew on TSAc + 0.5% NaCl was decreased from 8.4 x 106 CFU/cm 2 to 2.3 x 105 CFU/cm 2 in 30 sec, which was a l 7/io log decrease in the initial population. The population continued to decrease gradually. At 120 sec, the population had decreased to 1.3 x 103 CFU/cm 2 or in other words by 3 4 /5 logs. At the end of the 3 min test, the population decreased to 7.2 x 102 CFU/cm 2 or 4 logs. S. putrefaciens biofilm exposed to chlorine and plated on TSAc + 2.0% NaCl decreased rapidly from an initial population of 2.7 x 105 CFU/cm 2 to 3.3 x 10° CFU/cm 2 in just 30 sec. This decrease was equivalent to close to a 5 log reduction in the initial population. After 3 min exposure to chlorine, the population decreased to 1.1 x 10° CFU/cm 2 , which was a 53/s log decrease from the initial population. It is worth noting 114 that in some of the replicate tests at 10° ± 0.5°C and 0.5° ± 0.5°C in which organisms were exposed to chlorine followed by plating on TSAc + 2.0% NaCl, there were no surviving organisms. If these sanitizing tests had proceeded for a few more minutes longer, there would probably have been no growth observed on TSAc + 2.0% NaCl. A l l the organisms would have been either killed or injured. Sanitizer test data also showed that chlorine had inflicted injury upon the surviving organisms as can be observed from the survival curve for S. putrefaciens plated on TSAc + 2.0% NaCl. Furthermore, the cells exposed to 1.0 ppm chlorinated d-d water and grown on TSAc + 2.0% NaCl were smaller than their counterparts which were not exposed to chlorine and grown on TSAc + 2.0% NaCl (starting population at time zero). Thus, they were incubated for an additional 24 hr at 25°C prior to their enumeration. Temperature (10° ± 0.5°C and 0.5° ± 0.5°C) did not appear to have affected the efficiency of chlorine as a sanitizer. Chlorine inactivation of biofilms differed considerably from that of planktonic cells. In biofilms, there was a gradual yet considerable decrease in the number of organisms with a relatively small consumption of available chlorine in the system. Because of the smaller surface area, chlorine was not subjected to increased inactivation by the organic matter as it was in a planktonic system. Thus, chlorine was more effective in inactivating cells present in biofilms than it was in inactivating planktonic cells. Chlorine was also more effective than ozone in inactivating biofilm cells at either 10° ± 0.5°C or 0.5° ± 0.5°C given the concentration of each sanitizer used in these tests. Like ozone, chlorine also caused a substantial decrease in the number of organisms during the first 30 sec. However, chlorine seemed to be more effective in penetrating through the 115 biofilm and thus inactivating more organisms as demonstrated by the greater rate of inactivation during the three min tests. It can be seen from Figures 19, 20, 23 and 24, that chlorine inactivation curves were much steeper than those of ozone. This increased ability to kill a greater number of organisms may be due to the fact that chlorine was not as easily inactivated by the organic matter making up the biofilm as was ozone. However, it must be pointed out that the concentration of chlorine was four times greater than that of ozone. Comparison of results obtained in these ozone and chlorine tests to information found in published studies is difficult. Difficulties arise from the lack of studies comparing ozone and chlorine sanitizing efficacy on both planktonic and biofilm cells. Most studies focus on ozone or chlorine and rarely compare their sanitizing efficacy on biofilms. In addition, most of these studies employed different test organisms, procedures and materials. For example, it is difficult to compare the studies performed by Bott (1991) and Restaino et al. (1995) to this study due to the fact that these studies tested ozone efficacy as a sanitizer in very different environments. Boty (1991) tested ozone efficacy in removing Pseudomonas fluorescens biofilms in a complex apparatus that simulated cooling water systems. However, given the differences between these studies, they do agree on the efficacy of ozone in inactivating planktonic pells. The few studies that have compared the efficacy of ozone and chlorine in planktonic cells agree that ozone is a stronger sanitizer than chlorine. Domingue et al. (1988) found that ozone was a stronger biocide than chlorine. Ozone decreased L. pneumophda planktonic population by more than 2 logs during a 5 min exposure to 0.10 to 0.30 ppm of ozone/mL 116 (Domingue et al., 1988). Chlorine also decreased the L. pneumophila population by more than 2 logs but after exposure to 0.30 and 0.40 ppm free chlorine/mL for 30 and 5 min, respectively. Greene et al. (1993) compared ozonated and chlorinated water efficacy against biofilms of milk spoilage bacteria. The biofilms were grown on stainless steel plates. Greene et al. (1993) found that both ozone at 0.5 ppm for 10 min and 100 ppm Antibac B® (active ingredient, sodium dichloro-s-triazinetrione) for 2 min decreased A. faecalis and P. fluorescens biofilm populations by more than 2 logs. In the present study, 0.25 ppm ozonated d-d water at 10° ± 0.5°C decreased the initial S. putrefaciens biofilm population by about 2 7 / 1 0 logs at the end of a 3 min test. On the other hand, 1.0 ppm chlorinated water at 10° ± 0.5°C decreased the initial S. putrefaciens biofilm population by 4 logs at the end of a 3 min test. In summary, the inactivation data from all the treatments indicated that ozone was a better sanitizing agent than chlorine for the inactivation of planktonic cells. However, chlorine appeared to be a better choice against biofilms; given the concentrations employed in these tests. Perhaps, the test data for the two sanitizing agents was an indication of the different microbial inactivation mechanisms for ozone and chlorine. Different inactivation mechanisms could affect their relative sanitation effectiveness in practical situations. The organisms also underwent considerable changes in their resistance to chlorine and ozone when they went from one state to another; that is, from the planktonic to the biofilm form. 117 4.8 Statistical Analysis Anova - General Linear Model and Bonferroni Pairwise Comparisons were used to evaluate the sanitation data for 5. putrefaciens cells inoculated on TSAc agar. However, planktonic and biofilm data were evaluated separately. It was not possible to statistically compare the results arising from planktonic tests with those of biofilm tests since both were measuring microbial inactivation in different systems; that is, planktonic data was expressed as CFU/mL form while the biofilm data was expressed as CFU/cm 2 form. The results from the Anova tests are shown in Tables 9, 9a, 10 and 10a. The Anova for planktonic data, shown in Table 9, indicates that there was a significant difference among the factors. In order to determine which factors were different, the data were subjected to the Bonferroni Pairwise Comparisons (Table 9a). The analysis revealed that there was a significant difference between the sanitizers (p < 0.05). Sanitizerl (distilled-deionized water) was significantly different (p < 0.05) from sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm). However, there was no significant difference (p > 0.05) between sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm). There was also no significant difference (p > 0.05) between temperatures, 10° ± 0.5°C and 0.5° ± 0.5°C. Anova also indicated no significant difference (p > 0.05) between sanitizer*temperature interactions. But, Bonferroni Pairwise Comparisons showed that although there was no significant difference between sanitizerl *temperaturel (distilled-deionized water * 10° ± 0.5°C) and sanitizerl*temperature2 (distilled-deionized water * 0.5° ± 0.5°C), both were significantly different (p < 0.05) from sanitizer2*temperaturel (ozone at 0.25 ppm * 10° CO cb CO c o CO CD > CD o 1— LO o CO LO LO 00 o o d V co LO CO LO o d d T- a> a> o o o o X CO oi X X X co o c o n t co co r: CM CO CM CN ^ ~ ~ T " o o o o o X X X X X CM (D N N CO CM CO CO CO CO •^ J- CO CO CN LO O 00 N S t i n t t n ^ CM I— "co CN 2 Q. E 0} <D CN N cl) N E c o CD 0 CD b CO I— CO LU o < «-co £ ^ 1 O T3 sz <» l i cn 0) CO ? C TJ o = 45 !2 CD T3 3 » 1 1 N . N § 1 •o -6 "c TJ T3 CD CO c T3 0) CO c o N O E I Q. E Q . 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E E E E CD CD CD CD I— I— I— I— * * * * CM CM CO CO CD CD CD CD N N N N c c c E (0 (0 (0 CO CO CO CO CO T-CMCOTflOCON-OOCDO CO 122 ± 0.5°C), sanitizer2*temperature2 (ozone at 0.25 ppm * 0.5° ± 0.5°C), sanitizer3*temperaturel (chlorine at 1.0 ppm * 10° ± 0.5°C) and sanitizer3*temperature2 (chlorine at 1.0 ppm * 0.5° ± 0.5°C). Results from Anova of the planktonic data confirmed the expected difference between sanitizers. Ozone and chlorine are well known biocidal agents and as expected they were found to have a significantly (p < 0.05) greater biocidal activity than distilled-deionized water. Surprisingly; however, there was no significant difference (p > 0.05) between ozone and chlorine in terms of their biocidal effect on S. putrefaciens. However, sanitiation data clearly showed that there was a substantial difference. Ozonated distilled-deionized water at 0.25 ppm inflicted > 70 times the number of deaths as did 1.0 ppm chlorinated distilled-deionized water. Upon closer examination of the statistical data obtained through minitab, it was noted that perhaps the inclusion of distilled-deionized water as a sanitizer (control) may have skewed the results of the analysis. It may have caused the obvious difference between ozone and chlorine to become non-significant. The statistical data also indicated that the temperatures employed in these tests did not significantly (p > 0.05) affect microbial inactivation of planktonic cells by the sanitizers. Examination of sanitizer*temperature interactions, only emphasized the significant difference ( p < 0.05) between distilled-deionized water and both ozone and chlorine. Once again, no significant differences (p > 0.05) were observed between sanitizer*temperature interactions involving ozone and chlorine. Similar results were observed, when tyofilm data was analyzed (Table 10 and 10a). As expected there was a significant difference (p < 0.05) between the sanitizers. 123 Sanitizerl (distilled-deionized water) was significantly (p < 0.05) different from sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm) However, there was no significant (p > 0.05) difference between sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm). There was also no significant difference (p > 0.05) between temperatures; that is, 10° ± 0.5°C and 0.5° ± 0.5°C. As with planktonic data, Anova of biofilm data indicated that there was no significant (p > 0.05) difference among sanitizer*temperature interactions. But, Bonferroni Pairwise Comparisons revealed that although there was no significant difference (p > 0.05) between sanitizerl *temperaturel (distilled-deionized water * 10° ± 0.5°C) and sanitizerl*temperature2 (distilled-deionized water * 0.5° ± 0.5°C); both were significantly different from sanitizer2*temperaturel (ozone at 0.25 ppm * 10° ± 0.5°C), sanitizer2*temperature2 (ozone at 0.25 ppm * 0.5° ± 0.5°C), sanitizer3*temperaturel (chlorine at 1.0 ppm * 10° ± 0.5°C) and sanitizer3*temperature2 (chlorine at 1.0 ppm * 0.5° ± 0.5°C). However, there was no significant difference (p > 0.05) between sanitizer*temperature interactions involving ozone and chlorine. Thus, the results from sanitizer*temperature interactions only re-enforces the significant difference between distilled-deionized water and ozone or chlorine as sanitizers. Anova of biofilm data indicated that temperature did not play a significant role in the inactivation of S. putrefaciens. However, just as with the planktonic data, sanitizerl (distilled-deionized water) was significantly (p < 0.05) different from sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm). This indicates a change in lethality due to the use of sanitizer; that is, no sanitizer vs. sanitizer. But, it was also found that 124 sanitizer2 (ozone at 0.25 ppm) was not significantly (p > 0.05) different from sanitizer3 (chlorine at 1.0 ppm). As with the planktonic data, sanitation data for S. putrefaciens biofilm exposed to 0.25 ppm ozonated distilled-deionized water and 1.0 ppm chlorinated distilled-deionized water, also showed obvious differences in their biocidal efficiency. These differences are clearly illustrated in Figures 19, 20, 23 and 24. Again, the inclusion of sanitizerl (distilled-deionized water) as a control may have skewed the results making the differences between ozone and chlorine non-significant. Thus, it is difficult to statistically compare results from the microbial sanitation tests because of the great differences in surviving microbial populations. Therefore, it was decided to determine i f there were any significant differences between ozone and chlorine by performing Anova and Bonferroni Pairwise Comparisons using only data for ozone and chlorine as sanitizers for both planktonic suspensions and biofilms based on results for TSAc data. The results from the analysis of the planktonic and biofilm data are illustrated in Tables 11, 11a, 12 and 12a. Table 11 and 11a show that there was a significant (p < 0.05) difference between the sanitizers. Sanitizer2 (ozone at 0.25 ppm) was significantly (p < 0.05) different from sanitizer3 (chlorine at 1.0 ppm)r On the other hand, temperature appeared to have no significant (p > 0.05) effect on the inactivation of S. putrefaciens cells. Table 11 also indicated no significant (p > 0.05) differences among sanitizer*temperature interactions. 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The results from the sanitizer*temperature interactions help to confirm the differences among the sanitizers. Tables 12 and 12a show the results of the analysis of the biofilm data. As in the case of the planktonic cells subjected to ozone and chlorine, the biofilm data gave similar results. It was found that there was a significant (p < 0.05) difference between sanitizer2 (ozone at 0.25 ppm) and sanitizer3 (chlorine at 1.0 ppm) in their inactivation of biofilm cells. However, there was also a significant (p < 0.05) difference between the two test temperatures, 10° ± 0.5°C and 0.5° ± 0.5°C. In addition, it was found that sanitizer2*temperaturel (ozone at 0.25 ppm * 10° ± 0.5°C), was significantly (p < 0.05) different from sanitizer2*tempeature2 (ozone at 0.25 ppm * 0.5° ± 0.5°C), sanitizer3*temperaturel (chlorine at 1.0 ppm * 10° ± 0.5°C), and sanitizer3* temperature2 (chlorine at 1.0 ppm * 0.5° ± 0.5°C). Sanitizer2*temperature2 (ozone at 0.25 ppm * 0.5° ± 0.5°C) was also significantly (p < 0.05) different from sanitizer3*temperaturel (chlorine at 1.0 ppm * 10° ± 0.5°C) and santizer3*temperature2 (chlorine at 1.0 ppm * 0.5° ± 0.5°C). The results form Tables 11, 11a, 12 and 12a confirm the biocidal differences observed between ozone and chlorine. Figures 17, 18, 21, 22 clearly show that ozone was a more effective sanitizer than chlorine for the inactivation of planktonic 130 suspensions. However, chlorine appears to be better suited than ozone for the inactivation of biofilm cells given the concentration of each sanitizer employed in this study. Anova also indicates that temperature did have an influence in the inactivation of biofilm cells; that is, ozonated water at 10° ± 0.5°C appeared to have a greater biocidal effect on biofilm cells than 0.5° ± 0.5°C. In many instances an increase in temperature is accompanied by an increase in the biocidal effectiveness of sanitizers against microorganisms. However, the same effect of temperature was not observed for ozonated planktonic suspensions nor in chlorinated planktonic suspensions and biofilms. In some instances, microbial inactivation data may not be suitable for statistical analysis. These series of sanitizer tests may be one of them. In this case, the great differences between the number of surviving organisnsms treated with the control from those treated with ozone and chlorine may have caused the statistical data to be skewed and thus not to confirm the observed differences between ozone and chlorine. Only the removal of the control, sanitizerl (distilled-deionized water), allowed the observed differences between ozone and chlorine to come to light. In addition, using data derived from one point in a sanitation curve, in this case results at the 3 min mark, may leave out important significant differences. For example, statistical analysis may conclude that there were not significant differences among treatments. However, patterns of cell inactivation over time may be of significant importance when comparing efficiency of different sanitizers. 131 5. CONCLUSION AND RECOMMENDATIONS Chlorine and ozone are well known for their biocidal properties and thus useful as sanitizers in the food industry. The sanitation tests performed on S. putrefaciens indicated that both chlorine and ozone are good biocidal agents. However, the data also suggested that they may be better suited for different sanitation tasks. S. putrefaciens suspensions exposed to chlorine and ozone experienced a sharp decrease in the number of organisms after only 30 sec exposure. However, 0.25 ppm ozone was more than 70 times more effective in inactivating S. putrefaciens than distilled-deionized water containing 1.0 ppm available chlorine. S. putrefaciens suspension at 10° ± 0.5°C exposed to 0.25 ppm ozonated distilled-deionized water experienced a 2 3 /5 log decrease from the initial number of organisms after only 30 sec. Under the same conditions, 1.0 ppm chlorinated distilled-deionized water decreased the S. putrefaciens population by only 4 / 5 of a log. Most of the inactivation by either chlorine or ozone took place within the initial 30 sec. After the initial decline in the planktonic population and simultaneous consumption of both chlorine and ozone, the surviving population remained stable. The rapid consumption of chlorine and ozone was probably due to the increased surface area available for interaction between the sanitizer and the organisms plus their extracellular components (slime). On the other hand, biofilms exposed to chlorine and ozone experienced a gradual decrease in the number of surviving organisms. Chlorine and ozone concentrations also declined but gradually. However, at the end of the 3 min test, chlorine and ozone concentrations remained quite high. Biofilm sanitation data indicated that chlorinated distilled-deionized water at 1.0 ppm was more effective against biofilms 132 than 0.25 ppm ozonated distilled-deionized water under the conditions used in the study. Biofilms exposed to 0.25 ppm ozonated distilled-deionized water experienced a sharp decrease in their initial population after only 30 sec exposure to ozone. After this initial drop in the population, the population continued to decrease but at a substantially slower rate. In comparison, biofilms exposed to 1.0 ppm chlorinated distilled-deionized water suffered a continuous decline in the S. putrefaciens population. Rate of chlorine inactivation of S. putrefaciens was greater than that of ozone. This was clearly illustrated by the steeper inactivation curves of biofilms treated with 1.0 ppm chlorinated distilled-deionized water. These results suggested that ozone at 0.25 ppm was less effective in inactivating organisms embedded in a biofilm matrix. The data suggested that during the first 30 sec, ozone first attacked the surface layers of the biofilm and thus inactivated a large number of organisms. However, ozone was not as effective in inactivating cells embedded in the underlying layers of the biofilm matrix. Because ozone is such a powerful oxidizing agent and very reactive with organic matter, it is likely that as ozone came in contact with the surface of the biofilm and as it made its way through the biofilm matrix it reacted with the organic matter in its path and thus lost some of its potential effect on the S. putrefaciens cells. The organic layers making up the biofilm matrix probably acted as a protective barrier against the killing effect of ozone, decreasing its sanitizing efficiency against S1. putrefaciens cells. On the other hand, chlorine at 1.0 ppm, was more effective in penetrating the biofilm matrix and inactivating S. putrefaciens cells. This greater biocidal efficiency against biofilms may be due to the fact that chlorine is not as easily inactivated by organic matter as is ozone. Thus, 133 although ozone is a stronger oxidizing agent than chlorine and it is a more powerful sanitizer against S. putrefaciens suspensions than chlorine, chlorine at 1.0 ppm appeared to be better suited for use as a sanitizer against biofilms than 0.25 ppm ozone. It would be of considerable interest to determine the sanitizing effect of 1.0 ppm ozonated distilled-deionized water on microbial biofilms compared to 1.0 ppm chlorinated distilled-deionized water. For example, if ozone concentration was increased to 1.0 ppm, would the sanitation pattern be similar to that observed at 0.25 ppm?. Given the advantages of using ozone as a substitute for current sanitizers, the sanitation data obtained in this study and gathered by other researchers is promising. It must be noted that after 3 min exposure of biofilm to ozonated distilled-deionized water at 10° ± 0.5°C, the initial S. putrefaciens biofilm population of 1.3 x 107 CFU/cm 2 was decreased to 4.2 x 104 CFU/cm 2 . On the other hand, the S. putrefaciens biofilm population exposed to 1.0 ppm chlorinated distilled-deionized water at 10° ± 0.5°C decreased from 1.1 x 107 CFU/cm 2 to 8.4 x 102 CFU/cm 2. These differences are not great when considering that chlorine concentration was 4 times higher than ozone. If ozone is used as part of an effective cleaning and sanitation procedure; it can be as effective as other sanitizers in minimizing bacterial contamination of food from microbial biofilms. Its use as a sanitizer may require some adjustments. Plant operations may need longer cleaning and sanitation procedures than those used with other sanitizers in order to compensate for some of ozone drawbacks such as its high reactivity with organic matter. Thus, there is a lot of work that still needs to be done. For example, it is important to test sanitizer efficiency against both planktonic microbial suspensions and biofilms to 134 determine the suitability of sanitizers to deal with different physical states of spoilage or pathogenic organisms of concern for the food industry. Unfortunately, most of the data available on sanitizers involves planktonic suspensions. It is also important to perform tests that take into consideration that biofilms are complex systems involving in many cases more than one microbial population and its extracellular material in addition to food based soil as well. Just as a specific sanitizer may be more effective against planktonic suspensions than biofilms; it may also be more effective against one organism of the possible two or three making up a biofilm. Future tests could perhaps determine the efficiency of chlorine and ozone in not only inactivating cells embedded in the biofilm but also in helping to remove the biofilm from the SS chips. There is also the need for more adequate determination of cell injury. In this study, it was noted that ozonated cells required longer incubation times than non-ozonated cells when plated on TSAc containing sodium chloride. Longer incubation times do indicate some degree of stress or injury; hence, the need to be able to determine this "injury" in a specific and definitive way. The longer incubation times observed in this study contradict the often quoted all or none effect of ozone on microorganisms. The sanitation tests also revealed that the indigo trisulfonate method is not suitable to measure ozone concentration in suspensions containing large number of microorganisms. Some microorganisms; e.g. S. putrefaciens, adsorbed the indigo trisulfonate dye. Adsorption of the indigo trisulfonate dye by microorganisms will lead to erroneous ozone concentrations measurements and thus the possibility of inadequate ozonation of microbial suspensions. Selvakone (1997) also demonstrated that E. coli and 135 S. aureus adsorbed indigo trisulfonate. Although, the procedure used in this study to measure ozone concentration in microbial suspensions was practical; it was far from ideal. Thus, there is a need to find a method that works as well as the indigo trisulfonate method but that does not interact with microorganisms. 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Food Technology. 48(7): 107-114. 147 7 APPENDIX 7.1 Problems Determining Ozone Concentrations in S. putrefaciens Suspensions The objective of this thesis was to evaluate the effectiveness pf ozone as a sanitizing agent on suspensions of S. putrefaciens and S. putrefaciens biofilms grown on SS chips. For comparison purposes ozone effectiveness was evaluated against chlorine (sodium hypochlorite) and no sanitizer (distilled-deionized water). During preliminary work with ozononated suspensions of S. putrefaciens; problems were encpuntered when attempting to determine residual ozone concentrations. The ozone readings as determined by the indigo trisulfonate method gave inconsistent results. The following sections describes some of the preliminary tests and problems encountered determining ozone concentration using the indigo trisulfonate method. 7.2 Preliminary Ozone Sanitizing Tests - Table 13 7.2.1 Preparation of Shewanella putrefaciens Planktonic Inoculum S. putrefaciens (A. T. C. C. No 8071, Rockville, MD) cultures frozen at -80°C in 1 mL of 50% glycerol solution were activated by consecutive transfers into TSB and TSA as described by Prahst (1994). The frozen vials were thawed and 1 mL was transferred into a test tube (Length: 150 mm, ID: 15 mm) holding 5 mL of TSB broth. These cultures were incubated for 24 hr at 10°C in a rotatory-type shaking water bath agitated at 100 rpm. One loop of these cultures was subsequently streaked onto TSA plates which were incubated at 25°C for 48 hr or longer until distinct individual colonies were 148 obtained. A colony from a TSA plate was then transferred to two test tubes (Length: 150 mm, DD: 15 mm) containing 5 mL of TSB and incubated at 10°C for 24 hr in the rotatory-type shaking water bath at 100 rpm. One mL of each of these suspensions was then transferred to another set of test tubes containing 5 mL of TSB and incubated again at 10°C for 24 hr in the rotatory-type shaking water bath (Orbit Shaker Bath Model No 3543, Lab-Line Instruments, Inc., Melrose Park, EL.) at 100 rpm. This step was repeated one more time. After these series of inoculations and incubations, an individual 5 mL culture was transferred to each of two 250 mL erlenmeyer flasks containing 100 mL of TSB. These consecutive transfers were required to give the bacterial culture enough time to grow and adapt to the new incubation temperatures. The 100 mL microbial suspensions destined for experimental trials at 10° ± 0.5°C, were incubated for approximately 48 hr at 10°C in a rotatory-type shaking water bath maintained at 120 rpm. The two flasks containing 100 mL of the microbial suspension were then transferred to a sterile centrifuge bottle and pelleted by centrifugation at 9000 g for 20 min at 10°C. The resulting supernatant was discarded while the bacterial pellet was re-suspended in 200 mL sterile, distilled-deionized water and centrifuged one more time at 9000 g for 20 min at 10°C. Once again the supernatant was discarded, the bacterial pellet was re-suspended in 27 mL of sterile distilled-deionized water. This 27 mL bacterial suspension was then used as the inoculum for planktonic sanitizer tests. 149 7.2.2 Addition of Shewanella putrefaciens Planktonic Inoculiim to 0.5 ppm Ozonated d-d Water at 10° ± 0.5°C In order to determine the initial S. putrefaciens population in each 27 mL inoculum, three 1 mL samples were transferred to three test tubes containing 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI). These samples were serially diluted and the number of organisms enumerated using the drop plate method. The remaining 24 mL inoculum was then added to a 500 mL erlenmeyer flask containing 500 mL of 0.5 ppm ozonated distilled-deionized water at 10° ± 0.5°C. The suspension was thoroughly mixed with the aid of a 38 mm magnetic stirrer set at 650 rpm using a Corning Laboratory Stirrer/Hot Plate, Model PC-320. The bacteria were thoroughly dispersed through the water in less than 20 sec. 7.2.3 Enumeration of Surviving Shewanella putrefaciens cells Exposed to 0.5 ppm Ozonated d-d Water at 10° ± 0.5°C One mL samples from the 500 mL test suspension were retrieved every 30 sec during a 3 min period. Each 1 mL sample was diluted with 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI) containing 15 mg/L sodium sulfite (Sigma Chemical Company, St. Louis, MO). Anhydrous sodium sulfite was used to neutralize ozone residuals (Domingue et al., 1988). These diluted samples were then serially diluted and the surviving organisms enumerated using the drop plate method. 150 7.2.4 Determination of Ozone Residual Concentration At the same time 1 mL samples were retrieved to determine the number of surviving S. putrefaciens organisms exposed to 0.5 ppm ozonated d-d water, 1 mL samples were removed to measured ozone concentration. These samples were retrieved every 30 sec for 3 min. Each 1 mL sample was carefully added to 1 mL of indigo trisulfonate solution. Once the sample and the indigo trisulfonate solution had reacted, 8 mL of ozone demand free water was added. These 10 mL suspensions were then vortexed and their absorbance taken at 600 nm. One mL of indigo trisulfonate solution diluted with 9 mL of ozone demand free water was used as the reference. 7.2.5 Results from Preliminary Ozone Sanitizing Tests - Table 13 The results from the sanitation test described in the previous sections are illustrated in Table 13. Table 13 shows that after the addition of the S. putrefaciens inoculum to the 500 mL of 0.5 ppm ozonated d-d water at 10° ± 0.5°C, the starting population of 5.7 x 106 CFU/mL was decreased by three logs in just 3 min. After this initial 3 log decrease in the initial population, the surviving population repiained stable. This suggests that either the surviving organisms were resistant to residual ozone remaining in the system or that ozone had been completely consumed. Logic indicates that the latter was the most likely reason. Thus the absorbance values for ozone concentration should have been close to zero. However, this was not the case. As can be seen from Table 13, absorbance values of 0.010 to 0.013 were obtained throughout the 3 min testing period. If any ozone remained in the system, negative absorbance values Table 13. Effect of ozonated distilled-deionized water (10°C) Shewanella putrefaciens3 suspension. Determination of [ozone] Time Absorbance [ozone] CFU/mL (s) @ 600 nm (ppm) 0 -0.021 0.500 0 . 0 x 1 0 ° 30 0.013 ? 1.0x 103 60 0.011 ? 3.0 x10 3 90 0.011 ? 1.0x 103 120 0.011 ? 4.0 x 103 150 0.011 ? 1.0x 103 180 0.010 ? 2.0 x 103 a. Once the S. putrefaciens inoculum was added to the ozonated suspension, there would have been 5.7 x 106 CFU/mL. 152 would have been recorded. If there was no ozone, absorbance values should have been zero or very close to zero. There should not be positive absorbance values. 7.3 Preliminary Ozone Sanitizing Tests - Trouble-shooting To determine why ozone concentrations could not be measured accurately with the indigo trisulfonate method several tests were performed. Examples of these tests are illustrated in Tables 14 and 15. 7.3.1. Preliminary Ozone Sanitizing Tests - Table 14 7.3.2 Preparation of Shewanella putrefaciens Planktonic Inoculum S. putrefaciens (A. T. C. C. No 8071, Rockville, MD) cultures frozen at -80°C in 1 mL of 50% glycerol solution were activated by consecutive transfers into TSB and TSA as described by Prahst (1994). The frozen vials were thawed and 1 mL transferred into a test tube Length: 150 mm, ED: 15 mm) holding 5 mL of TSB broth. These^  cultures were incubated for 24 hr at 10°C in a rotatory-type shaking water bath agitated at 100 rpm. One loop of these cultures was subsequently streaked onto TSA plates which were incubated at 25°C for 48 hr or longer until distinct individual colonies were obtained. A colony from a TSA plate was then transferred to two test tubes (Length: 150 mm, EO: 15 mm) containing 5 mL of TSB and incubated at 10°C for 24 hr in the rotatory-type shaking water bath at 100 rpm. One mL of each of these suspensions was then 153 transferred to another set of test tubes containing 5 mL of TSB and incubated again at 10°C for 24 hr in the rotatory-type shaking water bath (Orbit Shaker Bath Model No 3543, Lab-Line Instruments, Inc., Melrose Park, IL.) at 100 rpm. This step was repeated one more time. After these series of inoculations and incubations, an individual 5 mL culture was transferred to each of two 250 mL erlenmeyer flasks containing 100 mL of TSB. These consecutive transfers were required to give the bacterial culture enough time to grow and adapt to the new incubation temperatures. The 100 mL microbial suspensions destined for experimental trials were incubated for approximately 48 hr at 10°C in a rotatory-type shaking water bath maintained at 120 rpm. The two flasks containing 100 mL of the microbial suspension were then transferred to a sterile centrifuge bottle and pelleted by centrifugation at 9000 g for 20 min at 10°C. The resulting supernatant was discarded while the bacterial pellet was re-suspended in 200 mL sterile, distilled-deionized water and centrifuged one more time at 9000 g for 20 min at 10°C. Once again the supernatant was discarded, the bacterial pellet was re-suspended in 27 mL of sterile distilled-deionized water. This 27 mL bacterial suspension was then used as the inoculum for planktonic sanitation tests. 7.3.3 Addition of the Shewanella putrefaciens Planktonic Inocujum to d-d WateratlO°±0.5°C In order to determine the initial S. putrefaciens population in each 27 mL inoculum, three 1 mL samples were transferred to three test tubes containing 9 mL of 154 0.1% peptone water (Difco Laboratories, Detroit, MI). These samples were serially diluted and the number of organisms enumerated using the drop plate method. The remaining 24 mL inoculum was then added to a 500 mL erjenmeyer flask containing 500 mL of distilled-deionized water at 10° ± 0.5°C. The suspension was thoroughly mixed with the aid of a 38 mm magnetic stirrer set at 650 rpm using a Coming Laboratory Stirrer/Hot Plate, Model PC-320. The bacteria were thoroughly dispersed through the water in less than 20 sec. 7.3.4 Enumeration of Surviving Shewanella putrefaciens cells Exposed to d-d Water at 10°±0.5°C One mL samples from the 500 mL test suspension were retrieved every 30 sec during a 3 min period. Each 1 mL sample was diluted with 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI) containing 15 mg/L sodium sulfite (Sigma Chemical Company, St. Louis, MO). These diluted samples were then serially diluted and the surviving organisms enumerated using the drop plate method. 7.3.5 Absorbance Values @ 600nm At the same time 1 mL samples were retrieved to determine the number of surviving S. putrefaciens organisms exposed to d-d water, 1 mL samples were removed to take absorbance values. These samples were retrieved every 30 sec for 3 min. Each 1 mL sample was carefully added to 1 mL of indigo trisulfonate solution. Once the sample and the indigo trisulfonate solution had reacted, 8 mL of ozone demand free water was 155 added. These 10 mL suspensions were then vortexed and their absorbance taken at 600 nm. One mL of indigo trisulfonate solution diluted with 9 mL of ozone demand free water was used as the reference. Another 1 mL sample was also taken after 30 sec exposure to d-d water (A*). This 1 mL sample was added to 9 mL of ozone demand free water and its absorbance taken at 600 nm using as a standard d-d water in the reference cell of the spectrophotometer. 7.3.6 Results from Preliminary Ozone Sanitizing Test - Table 14 Results from this test are shown in Table 14. The data in Table 14, shows that the 1 mL samples from the 500 mL microbial suspension added to the 1 mL indigo trisulfonate dye and 8 mL ozone demand free water gave positive absorbance values. These values range from 0.012 to 0.013. Because the 500 mL microbial suspension was not ozonated, these absorbance values should have been zero or fairly close to zero. Some interference was causing these erroneous readings. During the test, an additional sample (A**) was retrieved after 30 sec. This 1 mL sample was diluted with just 9 mL of ozone demand free water. The absorbance value of this sample using distilled-deionized water as the standard in the reference cell of the spectrophotometer was 0.097. Thus, the interference and resulting positive absorbance values recorded when measuring ozone concentration with the indigo trisulfonate method were the result of the light scattering caused by the suspended bacterial cells. The suspended cells gave the impression of a decreased ozone concentration. The larger the number of organisms, the greater the interference and corresponding positive absorbance value. 156 Table 14. Effect of distilled-deionized water (10°C) inoculated with Shewanella putrefaciens on indigo trisulfonate dye used to measure ozone concentration. Time (sec) Absorbance @ 600 nm of microbial suspension — ?m " CFU/mL 0 0.000 0.0x10 0 a 30b 0.013 2.1 x 107 60 0.012 2.0 x10 7 90 0.012 2.2 x 107 120 0.013 1.7x 107 150 0.012 1.9x 107 180 0.013 2.0 x 107 a. Once the S. putrefaciens inoculum was added to the ozonated water, there would have been 2.0 x 107 CFU/mL. b. The absorbance value of an additional 1 mL sample taken also at 30 s and diluted with just 9 mL of ozone demand free water was 0.097. pzone demand free water was used as the standard. 157 7.4 Preliminary Ozone Sanitizing Tests - Table 15 Results from the previous tests lead to attempts to remove the bacterial interference. A l l of these attempts did not work. For example, attempts were made to remove the cells by first taking the samples from the 500 mL of ozonated d-d water with special syringes fitted with 0.45 p. filter units (Millipore Products Division, Bedford, MA) . The filters would remove suspended organisms from the 1 mL samples prior to their addition to the indigo trisulfonate dye. Results showed that the filters did remove the organisms but they also interacted with the ozone present in the suspensions. Tests performed with just 0.5 ppm ozonated d-d water and no organisms confirmed that the filters totally consumed all the ozone present in the ozonated samples. Thus, it was impossible to measure ozone concentrations in this way. However, filters could be used to remove the bacterial cells after the sample had reacted with the indigo trisulfonate dye. The filters could be used to remove the cells from the 10 mL suspension consisting of lmL samples of the test S. putrefaciens suspension, lmL indigo trisulfonate solution and 8 mL ozone demand free water. The following sections describe how this test was performed and its results. 7.4.1 Preparation of Shewanella putrefaciens Planktonic Inoculum The planktonic inocula were prepared as described in section 7.2.1. 158 7.4.2 Addition of the Planktonic Inoculum to 0.5 ppm Ozonated d-d Water at 10° ± 0.5°C In order to determine the initial S. putrefaciens population in the 27 mL inoculum, three 1 mL samples were transferred to three test tubes containing 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI). These samples were serially diluted and then enumerated using the drop plate method. The 24 mL inocula was then added to a 500 mL erlenmeyer flask containing 500 mL of 0.5 ppm ozonated distilled-deionized water at 10° ± 0.5°C. The 500 mL suspension was thoroughly mixed with the aid of a 38 mm magnetic stirrer set at 650 rpm using a Corning Laboratory Stirrer/Hot Plate, Model PC-320. The bacteria were thoroughly dispersed through the water in less than 20 sec. 7.4.3 Enumeration of Surviving Shewanella putrefaciens Exposed to 0.5 ppm Ozonated d-d Water at 10° ± 0.5°C One mL samples from the 500 mL test suspension were retrieved every 30 sec during a 3 min period. Each 1 mL sample was diluted with 9 mL of 0.1% peptone water (Difco Laboratories, Detroit, MI) containing 15 mg/L sodium sulfite (Sigma Chemical Company, St. Louis, MO). Anhydrous sodium sulfite was used to neutralize ozone residuals (Domingue et al., 1988). These diluted samples were then serially diluted and the surviving organisms enumerated using the drop plate method. 159 7.4.4 Determination of Ozone Residual Concentration At the same time 1 mL samples were retrieved to determine the number of surviving S. putrefaciens organisms exposed to 0.5 ppm ozonated d-d water, 1 mL samples were removed to measured ozone concentration. These samples were retrieved every 30 sec for 3 min. Each 1 mL sample was carefully added to 1 mL of indigo trisulfonate solution. Once the sample and the indigo trisulfonate solution had reacted, 8 mL of ozone demand free water was added. These 10 mL suspensions were then vortexed and filtered using a syringe fitted with a 0.45pm filter (Millipore Products Division, Bedford, MA) . The filter would remove the suspended microorganisms but not the indigo trisulfonate in solution. The absorbance of the filtered solution was then measured at 600 nm. One mL of indigo trisulfonate solution diluted with 9 mL of ozone demand free water was used as the standard in the reference cell of the spectrophotometer. 7.4.5 Results from Preliminary Ozone Sanitizing Test - Table 15 Results from this test are shown in Table 15. The data from Table 15 indicates that the filter could remove some of the interference caused by the microorganisms. Table 15 shows that at time zero; that is, prior to the addition of the bacterial inoculum to the ozonated solutions, the ozone concentration was approximately 0.5 ppm. However, the absorbance values for samples taken after 30 sec corresponded to ozone concentrations higher than the initial 0.5 ppm. In fact, calculated ozone concentrations 160 Table 15. Effect of ozonated distilled-deionized water (10°C) on Shewanella putrefaciens suspension. Determination of TOzone] Time Absorbance [Ozone] CFU/mL (sec) @ 600 nm (ppm) 0 -0.021 0.50 0.0x10 0 a 30b -0.160 3.81 1.8 x 107 60b -0.127 3.02 1.2x 107 90b -0.119 2.83 1.2x 107 120b -0.119 2.83 1.2x 107 150b -0.118 2.81 1.0x 107 180b -0.118 2.81 1.6 x 107 a. Once the S. putrefaciens inoculum was added to the ozonated suspension, there would have been 4.3 x 107 CFU/mL. b. Samples taken at these time intervals were filtered prior to taking their absorbance value @ 600 nm. The removal of the microorganisms by the filter also removed the indigo trisulfonate adsorbed by the S. putrefaciens cells. This created a large decrease in the absorbance values making it appear as though there was more ozone in the system than at time zero. ' 161 were as high as 3.81 ppm. These values were wrong. Something had happened during the filtering of the samples. At this point, it was decided to centrifuge the 10 mL suspensions made of 1 mL of the ozonated microbial samples, 1 mL indigo trisulfonate dye and 8 mL ozone demand free d-d water. It was observed that after these samples were centrifuged for 20 min at 9000 g at 10°C, a blue pellet formed at the bottom of the centrifuge tubes. This indicated that not only did the presence of the bacteria cause spectrophotometric interference but the indigo trisulfonate used to measure ozone concentration was adsorbing to the 5*. putrefaciens cells causing additional interference. This would explain why in some circumstances ozone concentrations were higher than expected. The suspended organisms with the attached indigo trisulfonate dye would give the appearance of a lower ozone concentration than was actually present. When the cells were rernoved with the filters, they also removed some of the indigo trisulfonate dye. This resulted in an over estimation of the ozone concentration in the system. In summary, through a series of tests, it was observed that it was pot possible to directly determine ozone concentration in ozonated microbial suspensions consisting of large number of microorganisms. This was due to the spectrophotometric interference caused by the presence of the microbial cells as well as their adsorption of the indigo trisulfonate dye. Therefore, an indirect method of measuring ozone concentration in which the interfering factors were removed was necessary. It was also necessary to standardize both the initial microbial populations as well as the ratio of indigo trisulfonate dye to 162 ozonated microbial suspension samples in order to get a reasonable working range for both microbial populations and absorbance values. Thus the set up described in section 3.2.10 of Materials and Methods was employed to achieve this goal. It js important to note that the experimental set-up for the sanitation tests described in the Materials and Methods is quite different from those employed during the preliminary tests used to resolve the absorbance interferences cause by the S. putrefaciens cells. For example, in the preliminary tests, microbial population were enumerated using the drop plate method while in the sanitizer tests described in Material and Methods, the hydrophobic grid-membrane method was employed. The hydrophobic grid-membrane method proved to be more suitable for enumeration of both high and low number of surviving organisms from sanitizer tests. In addition, 1000 mL of 0.25 ppm ozonated d-d water were employed for sanitizer tests. Microbial inocula were standardized so that essentially the same number of organisms were obtained in each inoculum use for sanitizer tests. The amount of sample retrieved for ozone concentration readings was increased from 1 to 5 mL, while the indigo trisulfonate dye was correspondingly increased from 1 to 3 mL. This change of sample to indigo trisulfonate ratio provided a better working range of absorbance values for the determination of ozone concentration. 7.5 Variability Among Inoculated Stainless Steel Chips Before sanitizer tests were done, other tests were performed to determine the variability in the number of organisms present in the biofilms on SS chips. For this purpose, S. putrefaciens were grown on two sets of 15 SS chips. One set of 15 SS chips 163 was inoculated with S. putrefaciens derived from the same 1 mL frozen starter culture. The other set of 15 SS chips were inoculated with S. putrefaciens using 15 different 1 mL frozen starter cultures. The preparation of the biofilm inocula and the actual inoculation of the SS chips was performed as described in section 3.2.5 of Materials and Methods. It was found that SS chips inoculated from the same inoculum, had an average of 1.16 x 107 CFU/cm 2 with a standard deviation of ±1.34 x 106 CFU/cm 2 . Percentage wise, this is a 11.63% variability among SS chips inoculated from the same starter culture. SS chips inoculated by different sets of S. putrefaciens starter cultures, had an average of 1.17 x 107 CFU/cm 2 and a standard deviation of ±1.66 x 106 CFU/cm 2 . In other words, there was a 14.21% variability among chips inoculated from different starter cultures. The variability observed among SS chips inoculated from the same starter culture could be due to slight differences in the surface of the SS chips that may have promoted attachment of more or fewer cells to the surface of the chips. The variability among chips inoculated from different starter cultures may also be partly due to slight differences in the surfaces of the SS chips as well as differences due to some cells being in different stages of their growth cycle and slight differences inherent to the starter inocula. It is worth noting that during the sanitizer tests, SS chips that had biofilms with very high number of CFU/cm 2, were excluded from analysis. Only data from replicates with similar initial CFU/cm 2 were pooled together for analysis. 

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