Biofilms in the Food Environment (Institute of Food Technologists Series) [1 ed.] 0813820588, 9780813820583

Citation preview

Biofilms in the Food Environment

The IFT Press series reflects the mission of the Institute of Food Technologists— advancing the science and technology of food through the exchange of knowledge. Developed in partnership with Blackwell Publishing, IFT Press books serve as essential textbooks for academic programs and as leading edge handbooks for industrial application and reference. Crafted through rigorous peer review and meticulous research, IFT Press publications represent the latest, most significant resources available to food scientists and related agriculture professionals worldwide.

IFT Book Communications Committee Dennis R. Heldman Joseph H. Hotchkiss Ruth M. Patrick Terri D. Boylston Marianne H. Gillette William C. Haines Mark Barrett Jasmine Kuan Karen Banasiak

IFT Press Editorial Advisory Board Malcolm C. Bourne Fergus M. Clydesdale Dietrich Knorr Theodore P. Labuza Thomas J. Montville S. Suzanne Nielsen Martin R. Okos Michael W. Pariza Barbara J. Petersen David S. Reid Sam Saguy Herbert Stone Kenneth R. Swartzel

Biofilms in the Food Environment

EDITORS

Hans P. Blaschek r Hua H. Wang r Meredith E. Agle

Hans P. Blaschek, Ph.D. is Professor of Food Microbiology and Assistant Dean of the College of Agricultural, Consumer and Environmental Sciences, University of Illinois, Urbana-Champaign, Urbana, IL. Hua H. Wang, Ph.D. is Assistant Professor, Food Microbiology, in the Department of Food Science and Technology, The Ohio State University, Columbus, OH. Meredith E. Agle, Ph.D. is a Food Scientist in Bakery Research and Development at Rich Products, Buffalo, New York. Copyright  C Blackwell Publishing and the Institute of Food Technologists 2007 All rights reserved Blackwell Publishing Professional 2121 State Avenue, Ames, Iowa 50014, USA Orders: Office: Fax: Web site:

1-800-862-6657 1-515-292-0140 1-515-292-3348 www.blackwellprofessional.com

Blackwell Publishing Ltd 9600 Garsington Road, Oxford OX4 2DQ, UK Tel.: +44 (0)1865 776868 Blackwell Publishing Asia 550 Swanston Street, Carlton, Victoria 3053, Australia Tel.: +61 (0)3 8359 1011 Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee codes for users of the Transactional Reporting Service are ISBN-13: 978-0-8138-2058-3/2007. First edition, 2007 Library of Congress Cataloging-in-Publication Data Blaschek, Hans. Biofilms in the food environment/Hans Blaschek, Hua Wang, Meredith Agle. – 1st ed. p. cm. Includes index. ISBN-13: 978-0-8138-2058-3 (hardcopy) 1. Biofilms. 2. Food–Microbiology. 3. Food–Safety measures. I. Wang, Hua, 1965– II. Agle, Meredith. III. Title. QR100.8.B55B62 2007 579′ .17—dc22 2006025830 The last digit is the print number: 9 8 7 6 5 4 3 2 1

Titles in the IFT Press series r Accelerating New Food Product Design and Development (Jacqueline H.P. Beckley, J.C. Huang, Elizabeth J. Topp, M. Michele Foley, and Witoon Prinyawiwatkul) r Biofilms in the Food Environment (Hans P. Blaschek, Hua H. Wang, and Meredith E. Agle) r Food Carbohydrate Chemistry (Ronald E. Wrolstad) r Food Irradiation Research and Technology (Christopher H. Sommers and Xuetong Fan) r Foodborne Pathogens in the Food Processing Environment: Sources, Detection and Control (Sadhana Ravishankar and Vijay K. Juneja) r High Pressure Processing of Foods (Christopher J. Doona, C. Patrick Dunne, and Florence E. Feeherry) r Hydrocolloids in Food Processing (Thomas R. Laaman) r Microbiology and Technology of Fermented Foods (Robert W. Hutkins) r Multivariate and Probabilistic Analyses of Sensory Science Problems (Jean-Francois Meullenet, Rui Xiong, and Chris Findlay) r Nondestructive Testing of Food Quality (Joseph Irudayaraj and Christoph Reh) r Nonthermal Processing Technologies for Food (Howard Q. Zhang, Gustavo V. Barbosa-Canovas, V.M. Balasubramaniam, Editors; C. Patrick Dunne, Daniel F. Farkas, James T.C. Yuan, Associate Editors) r Packaging for Nonthermal Processing of Food (J. H. Han) r Preharvest and Postharvest Food Safety: Contemporary Issues and Future Directions (Ross C. Beier, Suresh D. Pillai, and Timothy D. Phillips, Editors; Richard L. Ziprin, Associate Editor) r Regulation of Functional Foods and Nutraceuticals: A Global Perspective (Clare M. Hasler) r Sensory and Consumer Research in Food Product Design and Development (Howard R. Moskowitz, Jacqueline H. Beckley, and Anna V.A. Resurreccion) r Thermal Processing of Foods: Control and Automation (K.P. Sandeep) r Water Activity in Foods: Fundamentals and Applications (Gustavo V. BarbosaCanovas, Anthony J. Fontana Jr., Shelly J. Schmidt, and Theodore P. Labuza)

CONTENTS

List of Contributors Preface

ix xiii

Chapter 1.

Biofilms in the Food Industry Meredith E. Agle

Chapter 2.

Shigella: Survival on Produce and Biofilm Formation Meredith E. Agle and Hans P. Blaschek

19

Chapter 3.

Biofilm Development by Listeria monocytogenes Scott E. Hanna and Hua H. Wang

47

Chapter 4.

Inactivation of Listeria monocytogenes Biofilms using Chemical Sanitizers and Heat 73 Revis A.N. Chmielewski and Joseph F. Frank

Chapter 5.

Mixed Culture Biofilms Michele Y. Manuzon and Hua H. Wang

Chapter 6.

Prokaryote Diversity of Epithelial Mucosal Biofilms in the Human Digestive Tract Denis O. Krause, H. Rex Gaskins, and Roderick I. Mackie

Chapter 7.

Beneficial Bacterial Biofilms Gregor Reid, Pirkka Kirjavainen, and Bryan Richardson

Chapter 8.

Applications of Biofilm Reactors for Production of Value-added Products by Microbial Fermentation Ali Demirci, Thunyarat Pongtharangkul, and Anthony L. Pometto III

Index

3

105

127

153

167

191 vii

LIST OF CONTRIBUTORS

Meredith E. Agle Rich Products, One Robert Rich Way, Buffalo, NY 14213, U.S.A. Chapter 1, Chapter 2 Hans P. Blaschek Department of Food Science and Human Nutrition, University of Illinois, Urbana-Champaign, 1207 W. Gregory Drive, 488 ASL, MC-630, Urbana, IL 61801, U.S.A. Chapter 2 Revis A. N. Chmielewski 313 Food Science Building, Department of Food Science, University of Georgia, Athens, GA 30602-7610, U.S.A. Chapter 4 Ali Demirci Department of Agricultural and Biological Engineering, 231 Agricultural Engineering Building, The Pennsylvania State University, University Park, PA 16802, U.S.A. Chapter 8 Joseph F. Frank Department of Food Science and Technology, University of Georgia, 211 Food Science Bldg., Athens, GA 30602-7610, U.S.A. Chapter 4 H. Rex Gaskins University of Illinois at Urbana-Champaign, Department of Animal Sciences, 1207 W. Gregory Drive, Urbana, IL 61801, U.S.A. Chapter 6

ix

x

List of Contributors

Scott E. Hanna 5117 Crestwood Hill, San Antonio, TX 78244, U.S.A. Chapter 3 Pirkka Kirjavainen Canadian R&D Centre for Probiotics, Lawson Health Research Institute, 268 Grosvenor Street, London, Ontario, N6A 4V2, Canada School of Public Health and Clinical Nutrition, University of Kuopio, Finland Chapter 7 Denis Krause 236 Animal Science Building, University of Manitoba, Winnipeg, MB, R3T 2N2, Canada Chapter 6 Roderick I. Mackie University of Illinois, 1207 W. Gregory Drive, Urbana, IL 61801, U.S.A. Chapter 6 Michele Y. Manuzon Department of Food Science and Technology, Ohio State University, 2015 Fyffe Ct., Columbus, OH 43210-1007, U.S.A. Chapter 5 Anthony L. Pometto III Department Food Science and Human Nutrition, 2312 Food Sciences Building, Iowa State University, Ames, IA 50011, U.S.A. Chapter 8 Thunyarat Pongtharangkul 249 Agricultural Engineering Building, The Pennsylvania State University, University Park, PA 16802, U.S.A. Chapter 8 Gregor Reid Lawson Health Research Institute, Room H214, 268 Grosvenor Street, London, Ontario, N6A 4V2, Canada Chapter 7

List of Contributors

xi

Bryan Richardson St. Joseph’s Health Care London, 268 Grosvenor Street, London, Ontario, N6A 4V2, Canada Chapter 7 Hua H. Wang Department of Food Science and Technology, Ohio State University, 2015 Fyffe Court, 219 Parker Food Science Bldg., Columbus, OH 43210-1007, U.S.A. Chapter 3, Chapter 5

PREFACE

This book examines biofilms produced by food-borne microorganisms, the risks associated with biofilms in the food chain, the beneficial applications of biofilms in the food environment, and approaches for biofilm removal to improve sanitation and safety in the food environment. Specifically, this book provides an introduction into the emerging and exciting field of biofilm research in the food environment, a summary of advanced knowledge in medical microbiology and engineering and its applicability to food biofilm research, and potential directions for biofilm intervention and industrial beneficial applications that may have direct impact on food safety and public health. This book is intended to serve as a comprehensive reference source for the food science community including industry scientists, university researchers, and regulatory agencies. Not only are general concepts regarding biofilms in the food environment covered herein, but also included are in-depth reviews on biofilm structures, the correlation between strain virulence and biofilm-forming abilities, cutting-edge technologies to investigate microbial compositions in ecosystems and cell-to-cell interactions, and updated findings on molecular attributes and mechanisms involved in biofilm development which might lead to targeted approaches for biofilm prevention and removal. The topics covered and approaches discussed are truly interdisciplinary in nature. Biofilm formation involving food-borne pathogens present on surfaces in the food environment and its correlation to pathogen persistence and food-borne illnesses were examined in Listeria monocytogenes and Shigella; results from various studies suggest that biofilm-related cells are more resistant to adverse environments, and stress responses may trigger biofilm formation. It is possible that stress responses and biofilm formation share some common metabolic pathways. Therefore, further characterization of molecular regulatory mechanisms involved in stress responses and biofilm formation may shed light on identification of new targets and development of new strategies for biofilm intervention. xiii

xiv

Preface

Biofilm intervention is a universal theme and this is an important area in food-related research as well. A comprehensive discussion on the types of physical treatments and chemical sanitizers, their mode of action on biofilm removal, and a summary of the effectiveness of various treatments will be very useful for industry scientists and academic researchers. While the formation of biofilms by pathogenic or spoilage microorganisms may have a negative impact on food safety and quality, biofilm formation can also have beneficial applications in the food environment. Attachment of beneficial microbes to the host intestinal tissues or gut microbiota can improve the overall health of the gut microenvironment. While microbial resistance to extreme environmental conditions is considered problematic in sanitation, such features have great application in fermentation where the production yield can be significantly improved by culture immobilization via biofilm formation and such biofilm-producing cultures are able to withstand high acid and low oxygen environment often associated with batch fermentation. The animal gastrointestinal ecosystem is considered one of the most complicated biofilms in nature. It is evident that food intake has a major impact on the ecosystem formulation but our knowledge in this area still remains at the infant stage. Due to the availability of the population genetic tools and its significance in public health, this area is inevitably becoming a research focus for microbiologists and food scientists in the coming years. We hope an overview of the human gut biofilms will help interested parties, particularly scientists new to the field to have a jump start in this fascinating research area. Biofilms in the food environment is still very much an emerging research area and the systems to be studied are complicated. In many cases, researchers are not just dealing with a pure bacterial culture, but rather consortia made up of a broad spectrum of organisms, i.e. food-borne pathogens, spoilage microbes, commensals, starters, and beneficial organisms. In addition, food processing and storage conditions, food ingredients, the host response, and immune system can all affect the behavior of these microorganisms. Therefore, a comprehensive knowledge of the food system, the microbiology, the host and environment, as well as the availability of cutting-edge research tools is a must for advancement in this field. We hope this book can serve as a reference source for applied and regulatory scientists and as well as academic researchers contemplating their future work. Hans P. Blaschek Hua H. Wang Meredith E. Agle

Biofilms in the Food Environment Edited by Hans P. Blaschek, Hua H. Wang, Meredith E. Agle Copyright © 2007 by Blackwell Publishing and the Institute of Food Technologists

Biofilms in the Food Environment

1

Biofilms in the Food Environment Edited by Hans P. Blaschek, Hua H. Wang, Meredith E. Agle Copyright © 2007 by Blackwell Publishing and the Institute of Food Technologists

Chapter 1 BIOFILMS IN THE FOOD INDUSTRY Meredith E. Agle

Introduction The first microbial biofilms were discovered on the surface of teeth by A. van Leeuwenhoek using primitive microscopes. The theory of biofilms was first described by J.W. Costerton in 1978. A biofilm is defined as a microbially derived sessile community which is characterized by cells that are irreversibly attached to a substratum, interface, or each other. The biofilm is irreversibly attached to the surface and rinsing cannot remove it. These cells are embedded in an extracellular polymeric matrix. Cells in a biofilm exhibit an altered growth and gene transcription compared to unattached cells (Donlan and Costerton 2002). Biofouling is the undesirable formation of a layer of microorganisms and their decomposition products on surfaces in contact with liquids. In the food industry this may lead to reduced heat transfer, increased resistance to flow, and corrosion. Biofilm formation can result in postprocessing contamination and cross contamination (Kumar and Anand 1998). Bacteria grow preferentially in the biofilm mode in industrial and natural systems. Bacteria have the ability to attach in turbulent conditions with Reynolds number greater than 5,000. High shear may serve to impinge bacteria on the surface (Donlan and Costerton 2002). Under conditions of higher flow and higher shear, cell clusters may be elongated and form streamers. Biofilms grown under high-shear conditions were smoother and denser than those grown under low-shear conditions (Stoodley and others 2002). Bacteria can colonize smooth as well as rough surfaces. Cells have been reported to attach more rapidly to hydrophobic surfaces— nonpolar surfaces such as plastic—rather than hydrophilic surfaces, such as glass or metal (Donlan 2002). Materials exposed to aqueous medium are conditioned by polymers in the medium increasing the rate and extent of attachment (Donlan 2002). Bacterial cells, organic molecules (such as 3

4

Biofilms in the Food Environment

proteins), and inorganic molecules can absorb to surfaces forming a conditioning film. These components can be transferred to the surface via turbulent fluid flow. This results in a higher concentration of nutrients at the surface when compared to the bulk fluid. The presence of the conditioning film alters the surface free energy, electrostatic charge, and hydrophobicity of the surface. A conditioning film is not, however, required for bacterial attachment (Kumar and Anand 1998). After a biofilm has been formed it is very viscoelastic and rubbery. Biofilms in low-shear environments have low tensile strength, while biofilms in high-sheer environments are very strong and resist mechanical breakage (Donlan and Costerton 2002). Confocal microscopy revealed that biofilms are not homogeneous monolayers of cells. Biofilms are heterogeneous and consist of microcolonies, which are the basic units of the biofilm. Biofilms are approximately 15% cells and 85% matrix by volume. The cells are enclosed in matrix which forms mushrooms and towers. Interspersed between these towers are water channels. These water channels can carry nutrients, dissolved oxygen, and antimicrobials to the cells in the microcolonies. The exchange of nutrients in the biofilm structure allows the biofilm to develop a high degree of thickness and complexity. Individual cells are maintained in optimal nutritional conditions in locations throughout the biofilm (Stoodley and others 2002). Measurements using microelectrodes reveal that the pH and the dissolved oxygen content of the biofilm are reduced near the substratum (Watnick and Kolter 2000). Biofilms can form as both single and multispecies communities, all of which share the same general organization (Donlan and Costerton 2002). Multispecies biofilms tend to be thicker than those of a single species. Microcolonies can break off the biofilm and serve as a seed to form new biofilms elsewhere. Cells that have shed may also revert to the planktonic mode. Upon attachment a variety of genes are up- and down-regulated in the cells (Donlan 2002). The formation of a biofilm can occur by one of three mechanisms: redistribution of attached cells by surface motility, binary division of attached cells, or the recruitment of cells from the bulk fluid to the developing biofilm. Biofilms can take over 10 days to reach structural maturity (Stoodley and others 2002). In Pseudomonas putida and Escherichia coli biofilms the cells in the center of the clusters decreased as clusters grew larger, but increased when carbon was supplied. This implies that the activity in the inside of the cluster may be limited by the amount of nutrients present. E. coli grow in the biofilm mode under conditions of nutrient availability. Other organisms grow preferentially in biofilms under conditions of nutrient deprivation. When nutrient deprivation occurs cells detach and return to the planktonic mode. Environmental factors

Biofilms in the Food Industry

5

such as temperature, osmolarity, pH, iron, and oxygen can also influence biofilm formation (O’Toole and others 2000). Biofilms form in a stepwise fashion. First, individual cells adhere to a surface by only a small amount of exopolysaccharide. This phase is reversible and cells may leave the surface and become planktonic again. As the biofilm grows the microcolonies and water channels form. Cells in the biofilm can alter their physiological state according to their niche. Long-range forces such as van der Waals, electrostatic, and hyrdrophobic are involved in reversible attachment. At this point, cell can still be removed by rinsing. After initial attachment the organism must maintain contact with the surface, attach irreversibly, and grow to form a biofilm. Irreversible attachment is mediated by short-range forces such as dipole– dipole, hydrogen, ionic, and covalent bonds as well as hydrophobic interactions. After irreversible attachment, rinsing will no longer remove the cells. Cells must be removed by scraping (Kumar and Anand 1998). The transition from weak to strong interactions with the surface is often mediated by the production of exopolymeric substances (EPS), which consists of a diverse array of biosynthetic polymers which may include substituted and unsubstituted polysaccharides, substituted and unsubstituted proteins, nucleic acids, and phospholipids. No population growth in the biofilm may be normal because cell division may be impeded by the surrounding exopolysaccharide. As biofilms mature, channels and pores are developed and the bacteria are redistributed away from the substratum. Acyl-homoserine lactone autoinducers have been detected in naturally occurring biofilms, implying that bacteria in biofilms may undergo densitydependent regulation. Cells in biofilms also have the ability to exchange genetic elements at an increased rate. This may allow for the acquisition of new genes for antibiotic resistance, virulence, and environmental survival (Watnick and Kolter 2000).

Antimicrobial Resistance Biofilms are more resistant to antimicrobials, such as antibiotics and disinfectants, than planktonic cells. Cells in biofilms may also exhibit increased resistance to UV light (O’Toole and others 2000). This increased resistance may be due to delayed penetration of the antimicrobial through the biofilm, altered rate of growth of cells in the biofilm, and other physiological changes that occur in the biofilm mode of growth. In order to kill cells in a biofilm the antimicrobial must penetrate the biofilm. The extracellular polymeric substance surrounding the cells may hinder diffusion. Ciprofloxacine required only 40 sec to treat a sterile surface, whereas

6

Biofilms in the Food Environment

21 min was needed to penetrate a Pseudomonas aeruginosa biofilm. A 2% solution of alginate, the polysaccharide found in Pseudomonas biofilms, inhibited the diffusion of several antibiotics. Additionally, the biofilm matrix may inactivate the antibiotic. Positively charged compounds such as aminoglycoside antibiotics may bind the negatively charged polymers of the biofilm, resulting in slower penetration (Stewart and Costerton 2001). Biofilm cells grow at a slower rate than planktonic cells and therefore take up antimicrobials more slowly. The slowest growing E. coli cells in a biofilm were more resistant to antibiotics. Older P. aeruginosa (10-dayold) biofilms were more resistant to antibiotics than younger (2-day-old) biofilms of the same organism (Donlan and Costerton 2002). The heterogeneous nature of biofilms that consist of cells representing a wide variety of different metabolic states allows cells to survive a metabolically directed attack (Costerton and others 1999). Certain antibiotics affect growing cells, such as penicillin which targets cell wall synthesis. Cells in biofilms that are not growing would be resistant to such an antibiotic. Conditions such as nutrient limitation and the build-up of toxic by-products favor the expression of stress-induced genes and the formation of biofilms (Donlan and Costerton 2002). Cells in a biofilm may develop a protected phenotype in response to growing on a surface, similar to spore formation (Costerton and others 1999). Cells in biofilms do not exhibit the familiar mechanisms for antibiotic resistance, such as efflux pumps, modifying enzymes, and target mutations (Stewart and Costerton 2001). Cells that detach from the biofilm do not exhibit the resistance of cells in the biofilm and quickly become susceptible to antibiotics. The resistance, therefore, is not acquired through mutations or mobile genetic elements (Stewart and Costerton 2001).

Multispecies Biofilms Biofilms in nature are generally multispecies. Mixed species biofilms are frequently thicker and more stable than biofilms consisting of a single species. Siebel and Characklis (1991) reported that P. aeruginosa and Klebsiella pneumoniae form biofilms of 30 and 15 µm, respectively, but form a 40-µm-thick biofilm together. Cells in multispecies biofilms distribute themselves based on the ability to survive in the various microenvironments in the biofilm and based on the symbiotic relationships among the groups of microorganisms. Organisms in multispecies biofilms are not randomly distributed, but are organized to meet the needs of each species (Watnick and Kolter 2000).

Biofilms in the Food Industry

7

Carpentier and Chassaing (2004) examined the ability of 29 microorganisms isolated from food processing environments to interact with Listeria monocytogenes. Sixteen strains decreased the ability of L. monocytogenes to form biofilms. Eleven strains had no effect and four strains increased the ability of L. monocytogenes to form biofilms. No correlation was observed between the amount of EPS produced by the different strains and their effect on the biofilm formation of L. monocytogenes, implying that not the quantity of EPS produced, but rather the type of EPS produced that is important for formation of L. monocytogenes biofilms in coculture. This work confirms that the resident microflora in a food processing environment has a strong effect on the presence of L. monocytogenes.

Disinfectants Bremer and others (2002) reported that pH-adjusted chlorine solutions were more effective in reducing the number of L. monocytogenes and Flavobacterium cells than unadjusted solutions. Cell death increased with increasing chlorine concentration and increased exposure time. Chlorine was more effective against cells on stainless steel compared to cells attached to conveyor belt material. This may be attributed to the inability of the chlorine to reach cells in the “pores” of the belt. Parkar and others (2004) examined the effectiveness of various cleaning regimes on the biofilms of the sporeforming thermophile, Bacillus flavothermus on stainless steel. Cleaning with caustic (2% NaOH, 75◦ C, 30 min, dH2 O rinse) followed by acid (1.8% HNO3 , 75◦ C, 30 min, dH2 O rinse) was the most effective caustic acid treatment for biofilm removal. This treatment killed all cells in the biofilm and removed most cells and polysaccharide from the stainless steel. Reducing concentrations of caustic or acid or reducing temperatures resulted in increased bacterial survival and increased detection of polysaccharide remaining on the surface of the stainless steel. Residue remaining on the surface after cleaning may serve as an attachment site for microorganisms or organic material, which may result in more rapid biofilm formation or product spoilage. The effectiveness of the cleaning process should be monitored not only by the number of cells remaining, but also by the presence of any cell residue on the cleaned surface. Langsrud and others (2003) isolated 14 strains of Serratia marcescens from disinfecting footbaths found in dairy plants. These strains were able to form biofilms on stainless steel. The strains were resistant to TEGO—an amphoteric disinfectant—and benzalkonium chloride. The strains, however, were sensitive to other disinfectants (peracetic acid and

8

Biofilms in the Food Environment

hypochlorite). This suggests that rotation of disinfectants may be useful in eliminating resistant microorganisms (Lansgrud and others 2003).

Food Processing Surfaces Arnold and others (2004) reported significantly fewer bacterial cells attached to electropolished stainless steel when compared to the control, untreated stainless steel surface. Electropolished samples exhibited decreased surface roughness. Both electropolished stainless steel samples and untreated controls were treated with a corrosive treatment to mimic processing conditions. All samples exhibited increased reddish-brown discoloration. The electropolished samples were less discolored than the controls and seemed to resist surface oxidation. After the corrosive treatment the control stainless steel surface was much smoother and exhibited reduced bacterial attachment. The electropolished stainless steel samples also exhibited reduced bacterial attachment after the corrosive treatment. Stainless steel is frequently used in the construction of food processing equipment. Regular mechanical or chemical cleaning can damage stainless steel surfaces. Microorganisms and organic material can gather in these sites and be protected from disinfectants. Boyd and others (2001) examined the surface of worn stainless steel. Four samples were examined: 316 grade, 316 grade abraded with 240 grit, 304 brushed grade, and 304 quartz paste abraded. Samples were treated with a 1% starch solution or with full fat milk powder to mimic food soil and then cleaned by spraying or brushing. The 316 abraded stainless steel sample exhibited unidirectional wear on the surface and the 304 abraded sampled exhibited bidirectional wear. Microscopic examination revealed that spray-cleaned samples retained soiling material. Greater amounts of soil absorption were observed on the damaged portion of the steel samples. Brush cleaning removed more soil than spray cleaning. The 304 brushed grade sample retained the most soil because this surface had the deepest grooves, followed by the 316 abraded with 240 grit and then the 304 quartz abraded sample. The retention of material was greater for surfaces with sharp deep scratches compared to surfaces with wider defects. Samples soiled with fat milk powder were spray cleaned or bushed cleaned, and then analyzed using ToF-SIMS (time of flight secondary ion mass spectrometry). This technique provided elemental, molecular, and polymer structure information by bombarding a sample with ions and then mass analyzing the secondary particles that are emitted. Spray cleaning left a larger amount of fatty acid material and removed more proteinaceous material when compared to samples cleaned by brushing (Boyd and others 2001).

Biofilms in the Food Industry

9

Moretro and others (2003) examined the ability of Staphylococcus isolated from food and food processing environments to form biofilms. Strains formed thicker biofilms when sodium chloride or glucose was added to the medium. Biofilm formation was examined on polystyrene, which is hydrophobic and may be used for food packages, and on stainless steel, which is hydrophlilic. Biofilm formation on polystyrene and stainless steel were correlated. The authors reported a higher prevalence in qac genes among biofilm forming strains of Staphylococcus. These genes encode efflux pumps which confer resistance to quaternary ammonium compounds. Borucki and others (2003) examined the ability of different strains of L. monocytogenes to form biofilms. Persistent strains of Listeria, those that were repeatedly isolated from bulk milk samples from the same dairy, were better biofilm formers than strains that were sporadically isolated from bulk milk samples. The authors demonstrated that L. monocytogenes does indeed produce EPS by staining with ruthenium red, a stain specific for carbohydrates. Midelet and Carpentier (2004) examined the ability of Pseudomonas fluorescens and Staphylococcus sciuri biofilms to transfer from stainless steel to a solid model food. These authors reported that the addition of calcium chloride to the liquid used to establish the biofilm led to increased surface coverage. This may be attributed to the calcium ions crosslinking between the anionic polysaccharides. S. sciuri was observed to have weaker attachment than P. fluorescens. Microcolonies were found to preferentially detach from the biofilm compared to single cells. When biofilms were treated with a chlorinated alkaline agent P. fluorescens cells detached more readily, but the attachment strength of S. sciuri increased. When biofilms were treated with a disinfectant containing glutaraldehyde and a quaternary ammonia, compound attachment strength and microcolony cohesion were increased due to the fixative action of glutaraldehyde (Midlet and Carpentier 2004).

Exopolysaccharides The main cement for cells in biofilms is a mixture of polysaccharides known as exopolysaccharides (EPS), which are secreted by cells in the biofilm. The types of EPS secreted vary from organism to organism. The majority of EPS are polyanionic owing to the presence of uronic acids or ketal-linked pyruvate. The primary configuration of the EPS is determined by composition. The secondary configuration often takes the form of aggregated helices. Exopolysaccharide is normally found in ordered

10

Biofilms in the Food Environment

conformations. The EPS are long chains with molecular masses of (0.5– 2.0) × 106 Da, which can associate in a variety of ways. Electrostatic forces and hydrogen bonds are the dominant forces that govern these interactions. The increased EPS production in biofilms may be a result of a stress response as in the case of colonic acid production in E. coli. The amount of EPS produced depends on the nutrients present. Synthesis of EPS is promoted by excess carbon sources with limiting nitrogen, potassium, and phosphorus. Bacterial mutants that are unable to produce EPS are unable to form biofilms. They may, however, be able to attach to surfaces. Exopolysaccharides allow for the binding of large amounts of water and contribute to mechanical stability of the biofilm, allowing it to withstand shear forces (Sutherland 2001). The EPS from several organisms have been characterized. Danese and others (2000) reported that the wcaF gene product was required for the production of colonic acid. Colonic acid was not required for initial attachment like the polysaccharides of Shewannella putrefaciens and Vibrio cholerae. It was, however, necessary for the establishment of the complex three-dimensional (3-D) structure of the E. coli biofilm. Mutants that could not produce colonic acid were still able to attach to abiotic surfaces (Danese and others 2000). Alginate is the primary component of P. aeruginosa biofilms. The algC gene involved in the production of alginate is transcribed at a higher rate (∼fourfold) in P. aeruginosa cells grown in biofilms compared to planktonic cells (Davies and others 1993). Genes in the intercellular adhesion locus (icaADBC) in Staphylococcus aureus and Staphylococcus epidermis encode genes involved in the synthesis of β-1-6-linked poly-Nacetylglucosamine referred to as PNAG. Staphylococcus strains deficient in PNAG production do not exhibit mushroom-like colonies and wide water channel-like strains, which produce larger amounts of PNAG.

Quorum Sensing Microbial biofilms provide a suitable environment for cell-to-cell signaling due to the large cell density. This quorum sensing occurs in a densitydependent manner via low-molecular-weight signaling compounds. The concentration of these compounds depends on population density. When a critical concentration is reached certain genes are turned on or off in the bacterial cells. There are several different molecules involved in cell-to-cell signaling. The most common autoinducer in Gram-negative microorganism is N-acyl-homoserine lactones (AHL). 2-Heptyl-3-hydroxy-4-quinolone (PQS) is involved in quorum sensing in P. aeruginosa. Amino acids and short posttranslationally modified peptides are used by Gram-positive

Biofilms in the Food Industry

11

microorganisms. Autoinducer-2 (AI-2) first discovered in Vibrio is produced by both Gram-positive and Gram-negative microorganisms (Van Houdt and others 2004). P. aeruginosa possesses two cell-to-cell signaling systems: lasR-lasI and rhlR-rhlI. The lasI gene product directs the synthesis of autoinducer N-(3-oxododecanoyl)-L-homoserine lactone. The lasR gene product requires a certain level of homoserine lactones to activate the transcription of virulence genes in this organism. The system directs the production of N-butryl homoserine lactone, which causes the transcription of virulence genes and RpoS. Davies and others (1998) examined biofilm production in knockout mutants, which were unable to produce either of the autoinducers described above. The biofilm produced by mutant cells was 80% thinner than the wild-type and the cells were densely packed. This phenotype was attributed to lasI gene. When the lasI gene product N-(3oxododecanoyl)-L-homoserine lactone was added to lasI mutant, biofilms that were similar to the wild-type were formed. These authors concluded that the quorum sensing molecule N-(3-oxododecanoyl)-L-homoserine lactone is required for biofilm formation. No significant difference was observed between the EPS of the mutant and wild-type P. aeruginosa cells (Davies and others 1998). Van Houdt and others (2004) isolated 68 strains of Gram-negative bacteria from a plant processing fresh vegetables. These strains were examined for their ability to form biofilms. Various degrees of biofilm-forming ability were observed and all strains were significantly better at forming biofilms than E. coli DH5α. The 68 strains were examined for their ability to produce AHL, PQS, or AI-2. No bacteria tested produced PQS, 26 isolates produced AI-2, and 5 isolates were positive for AHL production. These strains were identified as Vibrio diazotrophicus, Serratia plymuthica (2), and Panthoea agglomerans (2). The authors did not find a correlation between biofilm formation and the production of autoinducers (Van Houdt and others 2004).

Microscopic Examination of Biofilms Biofilms are often not very homogeneous, resulting in a specimen that is difficult to visualize. Thick biofilms may be problematic because the bacteria in lower layers cannot be observed or quantified and organisms in the upper layers may be lost if harsh fixation and staining techniques are used. Additional difficulties may occur depending on the surface on which the biofilm is located. Opaque and irregularly shaped surfaces require optics with a large depth of field. A variety of microscopic techniques such

12

Biofilms in the Food Environment

as scanning electron microscopy (SEM), environmental scanning electron microscopy (ESEM), atomic force microscopy (AFM), and confocal scanning laser microscopy (CSLM) can be used to image biofilms. The inventors of the ESEM wanted to observe liquid and hydrated samples. The operational pressure must be at least 4.6 torr at 0◦ C, as this is the minimum pressure needed to maintain liquid water. This is possible with multiple pressure-limiting apertures as well as environmental secondary electron detectors. In ESEM the use of multiple apertures permits smaller pressure differences, allowing for larger diameters at each aperture and maintaining a greater total pressure difference between the sample chamber and the column. By dividing the column into differential pressure zones separated by pressure-limiting apertures the electron gun can remain under a high vacuum and the sample chamber may contain a gas (Cameron and Donald 1994). The use of larger apertures does not place limits on the beam current. In ESEM water is the most common imaging gas and a separate vacuum pump permits fine control of its vapor pressure in the chamber. The electron beam emits primary electrons which strike the sample, resulting in the emission of secondary electrons. These secondary electrons collide with water molecules that serve as a cascade amplifier delivering the secondary electron signal to the positively biased gaseous secondary electron detector. The water molecules, which are positively charged due to the loss of electrons, are attracted to the specimen where a negative charge has been produced by the electron beam. This serves to suppress charging artifacts. The field emission gun produces a brighter filament image, or electron beam, than other sources. The accelerating voltage of the beam can be reduced to permit the imaging of fragile samples. Environmental SEM is a modification of conventional SEM. The specimen chamber can operate with up to 10 torr of vapor pressure, allowing for the examination of hydrated samples. Sample preparation such as fixation or staining is not required for ESEM. This technique allows intact biofilms to be examined in fully hydrated state at high magnifications. ESEM allows for the visualization of the biofilm in its naturally hydrated state. The electron beam, however, can cause damage to the sample. This damage occurs very quickly in ESEM samples. ESEM can also be used to image plant tissue in its hydrated state. Hamm and others (2002) examined alfalfa stems with ESEM as they were dehydrated. Conventional SEM allows for visualization of complex surface structures at very high magnifications. Samples are fixed with an aldehyde such as glutaraldehyde or formaldehyde, stained with a heavy metal stain, dehydrated using a graded ethanol or acetone series, and coated with a conductive material. Transmission electron microscopy (TEM) allows

Biofilms in the Food Industry

13

for the visualization of internal cross-sectional detail of the biofilm. TEM specimens are also fixed and stained as in SEM and are often embedded. Biological specimens such as biofilms tend to be sensitive to the harsh treatments required to prepare samples for SEM and TEM. The dehydration process may cause the biofilm to shrink, resulting in shrinking of components of the glycolax to 1% of their original volume (Surman and others 1996). The polymeric substances in the biofilm appear more fibrous rather than a thick gelatinous matrix when biofilms are dehydrated for SEM (Donlan and Costerton 2002). Care must be taken when interpreting these micrographs because of the presence of artifacts. Comparative visualization by other techniques is recommended. Both SEM and TEM offer high resolution and give information on spatial arrangement and cellular ultrastructure. Little and others (1991) examined biofilms with both ESEM and SEM. In the ESEM mode individual bacteria could not be distinguished in the hydrated biofilm, whereas in SEM, bacteria were observed as an individual monolayer. The authors reported that it was impossible to image individual cells in the monolayer in the ESEM mode. When a biofilm that had been examined using ESEM was treated with solvents much of the polymeric material was removed, revealing the bacteria as well as decreasing the surface area of the biofilm (Little and others 1991). In ESEM the biofilm was observed to have diatoms on the surface. These diatoms were removed with an acetone wash, demonstrating the negative effects that the harsh preparation protocol required for SEM may have on the sample (Little and others 1991). Atomic force microscopy (AFM) uses a sharp probe to map the contours of a sample. An AFM has a silicon nitride tip located on a flexible cantilever. The tip is scanned over a sample with a small repulsive force between the tip and the sample. Undulations in the surface topography of the sample result in the deflection of the cantilever. The undulations are detected by a laser located on the back of the cantilever, which is reflected onto a split photodetector. A feedback signal is then applied to the pizeoscanner, which is converted to a false color image that depicts the surface topography of the sample. AFM imaging in air results in very high resolution images of the surface of the biofilm. Imaging can also be carried out in liquid, but image quality is often poor. AFM also allows for the construction of 3-D images. Modulation contrast microscopy, which is a modification of bright-field microscopy, allows for noninvasive imaging of biofilms without the need for staining. The image has high-contrast resolution, a 3-D appearance, and does not contain the halos or artifacts that are often present in phasecontrast microscopy. This technique revealed a heterogeneous matrix

14

Biofilms in the Food Environment

with a diverse group of viable bacteria. Since a coverslip was used some compression of the biofilm occurred. The advantages of this technique are minimal sample preparation and capability to observe an intact, hydrated biofilm (Surman and others 1996). The differential interference contrast (DIC) and fluorescence microscope consists of a conventional light microscope with UV fluorescence and differential interference contrast through a mercury lamp. This system allows for the examination of biofilms without prior preparation and without a coverslip, so good topographical data are obtained without the compression of the biofilm by a coverslip. This technique allows for the measurement of the depth of the biofilm. The use of stains permitted differentiation between viable and nonviable cells (Surman and others 1996). Confocal scanning laser microscopy (CSLM) allows for optical sectioning and the construction of 3-D images. The optical sectioning capabilities are based on the confocal pinhole principle, which removes light that does not originate from the specimen plane in focus. Using several different fluorophores, a confocal microscope equipped with different detectors can produce multiple images simultaneously with the optical series. In this way spatial relationships of differential structures can be determined (Surman and others 1996). The resolution of a confocal microscope is 1.4 times greater than that of other optical microscopes (Carmichael and others 1999). CSLM can provide detailed information on location and viability of microorganisms without disturbing the physical location of the organism relative to the plant structure (Takeuchi and Frank 2001). Reisner and others (2003) used CSLM to examine the architecture of E. coli K-12 biofilms in continuous flow cell cultures over time. The presence of IncF plasmids induced biofilm formation similar to that of P. aeruginosa. Mature E. coli K-12 biofilms possessed 70–100-µm structures that extend into the liquid phase. Mattila and others (1997) used SEM, TEM, and confocal microscopy to observe seawater biofilms on stainless steel. Using microscopy, biofilms were observed forming in a stepwise fashion. First individual rods attached, next came the attachment of oval-shaped organisms, followed by spiral-shaped bacteria. Finally, a thin layer covered the surface. This layer was visible only in SEM when it was disturbed. It was not visible in TEM or using the light microscope. This demonstrates the advantage of using several microscopic imaging techniques. SEM also revealed the presence of mushroom-like microcolonies. TEM sections of the interior of the film showed small cells packed in an exopolymeric material and CSLM demonstrated the presence of polysaccharide in the matrix. Development of biofilm was also observed using CSLM, with total coverage of the stainless steel surface reaching 10–20%.

Biofilms in the Food Industry

15

Cookson and others (2002) used SEM to examine Shiga toxin-producing E. coli biofilms on glass and thermanox, which is a flexible polymer that is resistant to high temperatures and a variety of chemicals. Biofilms of curli knockout mutants were 2-D with limited fimbrial expression. E. coli O157:H7 biofilms were also 2-D with no visible fimbriae. Extracellular material indicative of dehydrated EPS was observed on E. coli O128:H2 microcolonies when the cells were grown at 25◦ C.

Summary Biofilms are the preferential mode of growth for many types of organisms. Biofilms can form on the surface of food processing equipment as well as on the surface of food such as meat or fresh produce. Cells in biofilms produce EPS, which enhances the structure of the biofilm. The composition of the EPS varies from organism to organism. Cells in biofilms are often more resistant to disinfectants. This is problematic for the food industry because cells of pathogenic or spoilage microorganisms may survive cleaning and disinfection and may detach and contaminate the food product.

References Arnold JW, Boothe DH, Suzuki O, Bailey GW. 2004. Multiple imaging techniques demonstrate the manipulation of surfaces to reduce bacterial contamination and corrosion. J Microsc 216:215–221. Borucki MK, Peppin JD, White D, Loge F, Call DR. 2003. Variation in biofilm formation among strains of Listeria monocytogenes. Appl Environ Microbiol 69(12):7336–7342. Boyd RD, Cole D, Rowe D, Verran J, Paul AJ, West RH. 2001. Cleanability of soiled stainless steel as studied by atomic force microscopy and time of flight secondary ion mass spectrometry. J Food Prot 64:87–93. Bremer PJ, Monk I, Butler R. 2002. Inactivation of Listeria monocytogenes/Flavobacterium spp. biofilms using chlorine: Impact of substrate, pH, time and concentration. Lett Appl Microbiol 35(4):321–325. Cameron RE, Donald AM. 1994. Minimizing sample evaporation in the environmental scanning electron microscope. J Microsc 173(3):227–237. Carmichael I, Harper IS, Coventry MJ, Taylor PWJ, Wan J, Hickey MW. 1999. Bacterial colonization and biofilm development on minimally processed vegetables. J Appl Microbiol Symp Suppl 85:45S–51S. Carpentier B, Chassaing D. 2004. Interactions in biofilms between Listeria monocytogenes and resident microorganisms from food industry premises. Int J Food Microbiol 97:111– 122. Cookson AL, Cooley WA, Woodward MJ. 2002. The role of type 1 and curli fimbriae of Shiga toxin-producing Escherichia coli in adherence to abiotic surfaces. Int J Med Microbiol 292(3–4):195–205.

16

Biofilms in the Food Environment

Costerton JW, Stewart PS, Greenberg EP. 1999. Bacterial biofilms: A common cause of persistent infections. Science 284(5418):1318–1322. Danese PN, Pratt LA, Kolter R. 2000. Exopolysaccharide production is required for development of Escherichia coli K-12 biofilm architecture. J Bacteriol 182(12): 3593–3596. Davies DG, Chakrabarty AM, Geesey GG. 1993. Exopolysaccharide production in biofilms: Substratum activation of alginate gene expression by Pseudomonas aeruginosa. Appl Environ Microbiol 59(4):1181–1186. Davies DG, Parsek MR, Pearson JP, Iglewski BH, Costerton JW, Greenberg EP. 1998. The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280:295–298. Donlan RM. 2002. Biofilms: Microbial life on surfaces. Emerg Infect Dis 8(9):881–890. Donlan RM, Costerton JW. 2002. Biofilms: Survival mechanisms of clinically relevant microorganisms. Clin Microbiol Rev 15(2):167–193. Hamm M, Debeire P, Monties B, Chabbert B. 2002. Changes in the cell wall network during the thermal dehydration of alfalfa stems. J Agric Food Chem 50(7):1897–1903. Kumar CG, Anand SK. 1998. Significance of microbial biofilms in the food industry: A review. Int J Food Microbiol 42:9–27. Lansgrud S, Moretro T, Sundheim G. 2003. Characterization of Serratia marcescens surviving in disinfecting footbaths. J Appl Microbiol 95:186–195. Little B, Wagner P, Ray R, Pope R, Scheetz R. 1991. Biofilms: An ESEM evaluation of artifacts introduced during SEM preparation. J Ind Microbiol 8:213–222. Mattila K, Carpen L, Hakkarainen T, Salkinoja-Salonen MS. 1997. Biofilm development during enoblement of stainless steel in Baltic sea water: A microscopic study. Int Biodeter Biodegrad 40(1):1–10. Midelet G, Carpentier B. 2004. Impact of cleaning and disinfection agents on biofilm structure and on microbial transfer to a solid model food. J Appl Microbiol 97:262– 270. Moretro T, Hermansen L, Holck AL, Sidhu MS, Rudi K, Langsrud S. 2003. Biofilm formation and the presence of the intercellular adhesion locus ica among staphylococci from food and food processing environments. Appl Environ Microbiol 69(9):5648– 5655. O’Toole G, Kaplan HB, Kolter R. 2000. Biofilm formation as microbial development. Annu Rev Microbiol 54:49–79. Parkar SG, Flint SH, Brooks JD. 2004. Evaluation of the effect of cleaning regimes on biofilm of thermophilic bacilli on stainless steel. J Appl Microbiol 96:110–116. Reisner A, Haagensen JA, Schembri MA, Zechner EL, Molin S. 2003. Development and maturation of Escherichia coli K-12 biofilms. Mol Microbiol 48(4):933–946. Siebel MA, Characklis WG. 1991. Observations of binary population biofilms. Biotechnol Bioerg 37:778–789. Stewart PS, Costerton JW. 2001. Antibiotic resistance of bacteria in biofilms. Lancet 358(9276):135–138. Stoodley P, Sauer K, Davies DG, Costerton JW. 2002. Biofilms as complex differentiated communities. Annu Rev Microbiol 56:187–209. Surman SB, Walker JT, Goddard DT, Morton LHG, Keevil CW, Weaver W, Skinner A, Hanson K, Caldwell D, Kurtz J. 1996. Comparison of microscope techniques for the examination of biofilms. J Microbiol Methods 25:57–70. Sutherland I. 2001. Biofilm exopolysaccharides: A strong and sticky framework. Microbiology 47(1):3–9.

Biofilms in the Food Industry

17

Takeuchi K, Frank JF. 2001. Expression of red-shifted green fluorescent protein by Escherichia coli O157:H7 as a marker for the detection of cells on fresh produce. J Food Prot 64(3):298–304. Van Houdt R, Aertsen A, Jansen A, Quintana AL, Michiels CW. 2004. Biofilm formation and cell-to-cell signaling in Gram-negative bacteria isolated from a food processing environment. J Appl Microbiol 96:177–184. Watnick P, Kolter R. 2000. Biofilm, city of microbes. J Bacteriol 182(10):2675–2679.

Biofilms in the Food Environment Edited by Hans P. Blaschek, Hua H. Wang, Meredith E. Agle Copyright © 2007 by Blackwell Publishing and the Institute of Food Technologists

Chapter 2 SHIGELLA: SURVIVAL ON PRODUCE AND BIOFILM FORMATION Meredith E. Agle and Hans P. Blaschek

Minimally Processed Produce Fresh-processed produce is a mushrooming industry. A 27% increase in fresh produce consumption had occurred from 1970 to 1993. In 1975 the average number of commodities offered for sale in the produce departments of U.S. supermarkets was 65. This skyrocketed to 340 items in 1995—a 432% increase. Fruit and vegetable production had risen from 85 billion pounds in 1970 to 136.8 billion pounds in 1994 (De Roever 1998) and the industry continues to grow today. Two factors have promoted this growth. First, consumers believe fresh produce is healthy and convenient. Secondly, fresh-processed produce provides a uniform product with less waste and less labor input for the food service industry (Hurst and Schuler 1992). There are two purposes in the minimal processing of fresh produce: to supply fresh produce in a convenient form while retaining nutritional value, and to extend the shelf life, allowing for the distribution of the product (Ahvenainen 1996). The popularity of minimally processed produce is due to the fact that it requires little labor for preparation and little waste is produced (Garg and others 1990). Minimally processed produce is marked by several features: the presence of cut surfaces or damaged plant tissue, the lack of sterility or microbial stability, the active metabolism of the plant tissue, and the confinement of the product (Nguyen-the and Carlin 1994). The minimal shelf life of these products should be at least 4–7 days, preferably up to 21 days (Ahvenainen 1996). The deterioration of minimally processed produce occurs for a number of reasons such as aging, biochemical changes, and microbial spoilage (Ahvenainen 1996). 19

20

Biofilms in the Food Environment

The greatest safety concern in the fresh-processed produce industry is microbial quality. CDC statistics from 1973 to 1992 show a doubling in the annual reporting of foodborne outbreaks (De Roever 1998). Organisms such as Salmonella, Shigella, Escherichia coli, and Campylobacter are often found in the intestinal tract of animals and are likely to contaminate raw vegetables by contact with feces, sewage, untreated irrigation water, or surface water. Increased recognition of fruits and vegetables as causative agents of foodborne illness along with better means of detecting pathogens may have resulted in increased reporting of foodborne outbreaks (Beuchat 1998). Increased reporting and improved diagnostics could both stimulate the observed increase in produce-related outbreaks, but one would expect to observe a rise in the total number of outbreaks, not just in produce-related outbreaks (De Roever 1998). Several factors may be responsible for the increase in produce-related outbreaks. Large and more centralized food production centers as well as a longer food chain allow for the proliferation of pathogens as well as an increase in their radius of distribution. The globalization of the food market allows consumers to be exposed to exotic microflora from foreign lands. The desire for convenience has caused an increase in the demand for minimally processed fruits, vegetables, and juices. This lack of thermal treatment allows for the survival of pathogens, which may cause illness. The large number of salad bars along with the increase in the amount of food consumed outside of the home increases the risk for food handling errors with fresh produce. Greater consumption of organic produce which is frequently fertilized with manure-containing pathogens such as E. coli O157:H7 may also be responsible for the increase in produce-related outbreaks (De Roever 1998). Additionally, the public may be more susceptible to foodborne disease because of the increased number of immunocompromised, elderly, and chronically ill (De Roever 1998). In the United States distribution patterns for fresh produce result in lots that are widely dispersed. Additionally, contamination of produce occurs sporadically and at low levels. Produce-related outbreaks, therefore, are often geographically diffuse and have low rates of attack. The high turnover rate, short shelf life, variable geographic origin, along with the complex network of growers, distributors, retailers who are often in different states or countries, make traceback difficult (De Roever 1998). Investigations of foodborne outbreaks involve three components. First, the epidemiological component identifies an association between a risk factor and becoming ill. Second, the environmental investigation identifies the circumstances that lead to the contamination of the food by the microorganisms. Third, analysis of patient specimens as well as food specimens

Shigella

21

allows for confirmation of the diagnosis as well as substantiation of the environmental findings. Investigations of foodborne outbreaks often do not provide information regarding all three components. Unlike meat and poultry outbreaks, fruit and vegetable outbreaks do not have certain characteristics that make them likely to be reported. They often do not cause an illness with symptoms serious enough to warrant medical attention. Organisms involved in produce-related outbreaks do not often have an established method of detection in the food and in clinical specimens. Fresh fruits and vegetables, unlike meat, do not have extended shelf lives, and are not frozen. Samples, therefore, are unavailable for testing. Foodborne outbreaks, for these reasons, are often underreported (De Roever 1998).

Ecology of Microbes on Produce The microflora present on market produce is representative of the organisms present at the time of harvest. This is also true for the presence of pathogens. Fresh produce primarily contains Gram-negative bacteria. The levels of microorganisms on plants in the field are highly variable. Microorganisms are often associated with the leaves and surfaces of fruits and vegetables and the inner tissue is considered sterile. The application of microorganisms to the surface of fresh produce often results in their internalization over time. Soil, irrigation water, animals, and farm workers can all serve as potential sources of microorganisms. Organisms such as Clostridium botulinum, C. perfringens, Bacillus cerus, and Listeria monocytogenes can be isolated from fecal-free soil and can be found on fruits and vegetables. Diseases associated with fresh fruits and vegetables are primarily transmitted via the fecal–oral route. Controlling fecal contamination is an important concern (De Roever 1998). Coliforms of the nonfecal variety can be found in the soil and associated with fruits and vegetables. The association of thermotolerant coliforms such as Klebsiella with produce limits the value of fecal coliforms as an indicator of fecal contamination. The ability of enteric bacteria to survive in soil depends on the type of soil, the initial inoculum, the ability of the soil to retain moisture, pH, nutrient availability, and the presence of microflora. Manure used as fertilizer or in irrigation water can contaminate fresh produce with harmful pathogens. Kudva and others (1998) reported that E. coli O157:H7 can survive for well over a year in nonaerated ovine manure exposed to environmental conditions. This organism can survive in aerated ovine and bovine manure for 4 months and 47 days respectively. Solomon and others (2002) reported that E. coli O157:H7 in manure used to fertilize soil or in

22

Biofilms in the Food Environment

irrigation water was able to enter the roots of mature lettuce plants and be transported to locations within edible portions of the plant. No direct contact between the leaves and the source of contamination is required for E. coli to be integrated into the lettuce tissue. A major source of fecal contamination is irrigation water. Irrigation water may be contaminated through direct introduction of sewage or ground water runoff. The use of irrigation water containing large numbers of microorganisms results in increased frequency of pathogen detection. The greatest amount of contamination is observed with leafy vegetables, which provide a large surface area for attachment. High humidity also favors survival and spread of microorganisms. Animals are a source of pathogenic microorganisms that may contaminate produce. Farm workers may also harbor enteric pathogens that can contaminate produce (De Roever 1998). Dust, insects, domestic and wild animals, harvesting equipment, transport containers, and processing equipment can all serve to transmit pathogens. Fruit flies exposed to apple juice contaminated with E. coli tested positive for the presence of E. coli. Fruit flies carrying E. coli were able to transfer the organisms to wounds on apples. Fruit flies were also capable of transferring organisms from apple wounds contaminated with E. coli to wounds on apples that were not previously contaminated (Janisiewicz and others 1999). The storage temperature of minimally processed produce determines the respiration rate, and therefore the gaseous atmosphere surrounding the produce. Temperature also determines the rate of senescence of minimally processed produce. Both factors can affect the microorganisms present (Nguyen-the and Carlin 1994). The minimum temperature for growth of most mesophilic enteric pathogens is 8–10◦ C. Unrefrigerated products often spoil before pathogen outgrowth is sufficient to cause illness. A number of pathogens such as L. monocytogenes and Yersinia enterocolitica can survive and grow under refrigeration conditions (De Roever 1998). The growth, survival, and inactivation of microorganisms on fresh produce is dependent on several factors including the characteristics of the microorganisms present, the physiological state of the plant tissue, the environmental characteristics (pH, water activity, etc.), and the effect of processing on the microflora or metabolism of the plant. The major determinants of pathogen growth on fresh produce are pH and storage temperature. The colonization of the surface of fresh produce by microorganisms is a naturally occurring phenomenon. Most vegetables have a pH greater than 4.5 and are able to support the growth of pathogens, emphasizing the importance of storage temperature. Due to the high levels of nutrients

Shigella

23

and moisture, as well as a near-neutral pH, produce can support a wide range of microorganisms. Spinach and lettuce, for example, contain ∼8% carbohydrates, 2% protein, and more than 88% moisture and a pH range of 5.1–6.8. These conditions are suitable for bacterial growth (Carmichael and others 1999). Fruits are often more acidic and do not support the growth of pathogens. Yeasts and molds can grow in lower pH conditions than most bacteria. Spoilage of fruits is often caused by yeasts and molds. Some yeasts and molds produce alkaline products during metabolism that can reduce the acidity and raise the pH of the product, allowing for the survival or growth of harmful pathogens (Beuchat 2002). Fruits and vegetables affected by soft rot also provide more suitable conditions for the survival of pathogens. Beuchat (1998) reported that the presence of pathogenic microorganisms is caused by exposure to environmental factors rather than the surface topography. However, produce with deep crevices and highly textured surfaces may be more likely to harbor soil containing large numbers of microorganisms. This may explain why large numbers of microorganisms are often found on the surface of leafy produce when compared to produce with smooth surfaces. Pathogens on the surface of fruits and vegetables that are peeled, such as bananas and oranges, are less of a concern because the peel is not consumed. Care, however, must be taken when peeling not to contaminate the inner flesh (Beuchat 1998). Each fruit and vegetable has a unique combination of optimal conditions such as growing conditions, harvesting and cooling practices, storage conditions, as well as unique physical characteristics and composition. For example, berries, which are very delicate and perishable, are not washed after harvesting. Tomatoes are picked green and flushed from totes with water through a flume into the packing house. Apples have a smoother more delicate skin than citrus. Factors such as these must be taken into consideration when determining microbiological hazards (De Roever 1998). Minimal processing of vegetables involves washing, trimming, peeling, slicing, and sanitizing. The goal is to minimize handling while maintaining freshness, quality, and maximal shelf life (Carmichael and others 1999). The act of cutting allows enzymes and substrates to join, resulting in discoloration. Cutting also allows for the release of nutrient rich fluids, which allow for microbial growth (Hurst and Schuler 1992). Shredding or slicing may be a major source of contamination in fruit and vegetable processing plants. Passing a knife through a contaminated surface results in contamination of the newly cut surface (De Roever 1998). The large number of cut surfaces along with high humidity provides excellent conditions for the growth of microorganisms (Carmichael and others 1999).

24

Biofilms in the Food Environment

Contamination of produce may also occur in food service environments. There is a great deal of direct hand contact with the produce and a heat or chemical step is not normally taken to inactivate the organisms. Poor personal hygiene practices of food service workers may also lead to contamination. Cross contamination with other food products such as uncooked meat may also be a problem (De Roever 1998). Steps can be taken to minimize the contamination of fresh produce. Untreated manure should not be used as fertilizer. Good worker hygiene should be practiced both on the farm and in the field. Quality water should be used for irrigation, washing, and production of ice. Proper temperatures should be maintained during processing, storage, and transport of produce to minimize growth of pathogens. Pectinolytic strains of Pseudomonas cause soft rot in minimally processed leafy vegetables. Raising the temperature and the carbon dioxide concentration causes lactic acid bacteria to predominate (Ahvenainen 1996). Nguyen-the and Carlin (1994) reported that mesophilic plate counts for minimally processed produce samples ranged between 103 and 109 CFU/g. Lactic acid bacteria counts were as high as 109 CFU/g. Gram-negative rods such as Pseudomonas, Erwinia, and Enterobacter are normally prevalent, with pseudomonads comprising over 50% of the population. Ten to 20% of mesophilic organisms isolated from lettuce were reported to be pectinolytic. Large numbers of pectinolytic pseudomonads were reported on shredded carrots and shredded chicory samples (Nguyen-the and Carlin 1994). Organisms that are found on minimally processed produce are also found on the plants in the field (Nguyen-the and Carlin 1994). Garg and others (1990) examined the microflora of minimally processed produce. The inner leaves of lettuce and cabbage contained 104 CFU/g. Removing the outer leaves significantly reduced microbial loads. Counts were higher after shredding. The factory’s goal was to maintain 300 mg/L chlorine in the wash, but levels fluctuated due to the large amount of organic material in the water. Many of the processed vegetables contained large numbers of psychrotrophs. None of the vegetables tested yielded high numbers of lactic acid bacteria, which was indicative of temperature abuse. Over 80% of the lactic acid bacteria isolated were leuconostocs. Spinach contained the most spores, 3.1 × 103 spores/g. Gram-negative rods were the predominant organisms on spinach, cauliflower, and carrots. Most species were members of the genus Pseudomonas. King and others (1991) examined the microbial quality of lettuce. The total counts and yeast and mold counts varied depending on the degree of outer leaf removal and the amount of soil present on the surface of the lettuce. Bacterial counts were always greater than yeast and mold counts.

Shigella

25

The bacteria count decreased from outer leaves successively from outer to inner leaves. The initial counts of cored lettuce were not significantly different from that of salad mix. This demonstrates that processing (cutting, chlorine rinse, centrifuging, and cooling) did not alter microbial counts. Of organisms isolated, 97.3% were Gram-negative rods, and 56.7% were Pseudomonas, Serratia, and Erwinia. Each comprised 8.1% of the bacterial population. The isolation of mold was very infrequent and these organisms probably do not play a part of the normal or spoilage microflora of lettuce. Abdul-Raouf and others (1993) examined the ability of E. coli O157:H7 to survive on salad vegetables. The organisms grew on shredded lettuce at 12 and 21◦ C and decreased significantly over the 14-day test period at 5◦ C. Psychrotroph populations increased on samples stored at 5 and 12◦ C over time as did the populations of mesophiles on lettuce samples stored at 21◦ C. The pH of the lettuce samples decreased over time. The largest pH drop was in lettuce samples stored at 21◦ C. As was observed with lettuce, E. coli declined on cucumber slices stored at 5◦ C, whereas increases in the E. coli population were observed at 12 and 21◦ C. The psychrotroph population on cucumbers at 5◦ C increased over time. The mesophilic population of cucumbers stored at 21◦ C also increased over the initial 3-day period. The rate in pH drop of the cucumber samples was proportionate to the increase in storage temperature. E. coli on shredded carrots stored at 5◦ C decreased significantly over time, but were still detectable after 14 days. At 12◦ C E. coli populations declined for the first 3 days and did not increase when the organisms were initially present in small numbers. In the large inoculum 12◦ C and 21◦ C samples, E. coli grew in shredded carrots over time. Psychrotrophs on carrots at 5 and 12◦ C and mesophiles on carrots at 21◦ C increased over time. The pH of carrots decreased over time. The pH of inoculated carrots decreased at a more rapid rate than that of the uninoculated carrots. The drop in pH of the produce samples may have had an effect of the viability of E. coli O157:H7. The pH drop is attributed to the fermentative capabilities of E. coli.

Cleaning and Sanitizing Fresh produce can be heavily contaminated after harvest and organisms can rapidly multiply during transport in the warm, humid conditions. Removing outer leaves can greatly reduce the microbial load. Washing reduces the number of microorganisms present, but does not completely remove them. Water washes can successfully reduce populations of surfaces microorganism from fresh fruits and vegetables. Not all vegetables, however, can withstand the stresses associated with washing. Even if

26

Biofilms in the Food Environment

vegetables are washed with sterile water, complete elimination of microorganisms will not occur because viable organisms remain in the tissue (Beuchat 1992). Washing produce can effectively remove soil from fruits and vegetables, but cannot completely remove microorganisms. Fresh produce may contain as many as 106 CFU/g microorganisms after harvesting. Washing in water can result in a 1–2 order-of-magnitude reduction in the initial microbial load (Beuchat 1995). Inadequate disinfection of wash water can shorten the shelf life of minimally produce by increasing the microbial load (Seymour 1999). Twenty percent of processors surveyed used only water rinses for disinfecting minimally processed produce. Washing reduces cellular components released during slicing or peeling. These components may serve as source of nutrients for microorganisms. The temperature of the wash water is also important. If the temperature of the water is less than that of the produce, the pressure differential will result in uptake of the bacteria by the produce. Infiltration is dependent upon time, temperature, and pressure. It occurs when the water pressure on the surface of the produce overcomes the internal gas pressure and the hydrophobic nature of the produce surface. The addition of detergents to the wash water results in increased internalization by reducing the surface tension of the water at the air–water interface with damaged plant cells, which lead into the plant tissue. Cells that have infiltrated the tissue may survive and grow. These cells may be difficult to reach with sanitizing solutions (Beuchat 2002). Zhuang and others (1995) examined the effects of temperature differentials on the uptake of Salmonella by tomatoes. A significantly higher number of cells were taken up by the tomato core tissue when tomatoes at 25◦ C were dipped into cells suspensions at 10◦ C, compared to the number of cells taken up when tomatoes were dipped into cell suspensions at 25◦ C or 37◦ C. It may be beneficial for packing houses to maintain wash tanks at temperatures higher than that of the incoming tomatoes. Chlorine is widely used for treatment of wastewater, drinking water, and in the food industry for disinfecting equipment. The Food and Drug Administration (FDA) permits the use of sodium hypochlorite, calcium hypochlorite, and gaseous chlorine as disinfectants in wash, spray, and flume water for treatment of fresh produce (Seymour 1999). Title 21 of the Code of Federal Regulations (CFR) specifies that a maximum level of 0.2% can be used in wash water. This concentration is not normally used for disinfection, but is used for lye peeling of fruits and vegetables. Between 50 and 200 ppm is required to destroy bacteria and fungi in packing houses. High levels of chlorine are often needed to satisfy the chlorine demand of large recirculated wash water systems (Hurst and

Shigella

27

Schuler 1992). Several theories exist to explain the effectiveness of chlorine as an antimicrobial; however its mode of action remains undetermined. Chlorine may combine with cellular membrane proteins forming N-chloro compounds, which interfere with metabolism. Additionally, necessary enzymes may be oxidized by chlorine (Beuchat 1992). When hypochlorites or chlorine is added to water the following reactions occur: Cl2 + H2 O → HOCl + H+ + Cl Ca(OCl)2 + H2 O → Ca + H2 O + 2 OCl− Ca(OCl)2 + 2 H2 O → Ca(OH)2 + 2 HOCl HOCl → H+ + OCl− Free available chlorine refers to elemental chlorine (Cl2 ), hypochlorous acid (HOCl), and the hypochlorite ion (OCl− ). The dissociation of HOCl is pH dependent. The HOCl and OCl− equilibrium is maintained even when HOCl is utilized for its antimicrobial activity. Hypochlorous acid is referred to as free chlorine. When free chlorine reacts with organic matter combined chlorine compounds are formed. Free and combined chlorine together are measured as total chlorine (Seymour 1999). The solution pH has a significant effect on the behavior of chlorine in water. When chlorine compounds are added to water chlorine gas, hypochlorous acid, and hypochlorite ions are generated in proportions determined by the pH of the solution. Lethality is determined by the amount of the HOCl. Hypochlorite ions are relatively inactive and chlorine gas will rapidly dissipate. A decrease in pH shifts the equilibrium toward HOCl. At a pH of 6 and 8 the concentrations of HOCl are 98 and 32%, respectively. It is recommended that disinfection of produce with chlorine should occur in conditions with a pH less than 7.5 (Beuchat 1998). Toxic chlorine gas is formed at a pH lower than 4. The equilibrium favors HOCl as temperature decreases at a fixed pH. After the available chlorine has been combined, additional chlorine must be added to produce more free chlorine to maintain the disinfecting capacity of the system. Certain types of fruits and vegetables with large organic loads require the addition of more chlorine to the wash system to maintain chlorine levels suitable for disinfection. Exposure time plays an important role in the efficacy of chlorine as a disinfectant. Quick dips are not nearly as effective as longer exposures. Most of the disinfection is thought to occur within the first few minutes of treatment. Microorganisms may be protected from chlorine by surface structures of the plant, such as stoma or cracks and crevices as well as biofilms. Longer treatment times may not increase microbial

28

Biofilms in the Food Environment

death. Contact times vary according to the type of washing system used. Extended exposure to chlorine may result in bleaching and off flavors in the finished product (Seymour 1999). The efficacy of decontamination using chlorine is highly dependent upon the product disinfected. When dipped into 300 ppm chlorine, significant reductions in microbial load were observed with lettuce, whereas no decrease was observed with carrots or red cabbage (Nguyen-the and Carlin 1994). Variable results are often observed with chlorine. This may be attributed to several factors: hypochlorite may not fully wet the hydrophobic surface of the waxy cuticle of vegetables, and cells may also be in a biofilm that serves to protect against disinfectants. Additionally, contact with host tissue may inactivate sanitizers such as chlorine (Nguyen-the and Carlin 1994). Chlorine may be more effective in inactivating cells in water washes used during processing in order to prevent cross contamination (Nguyen-the and Carlin 1994). Failure to maintain adequate amounts of chlorine in wash water may lead to increased numbers of microorganisms on produce (Beuchat 1992). Produce should be rinsed following treatment with chlorine to reduce the concentration of chlorine and to improve the organoleptic properties of the produce (Ahvenainen 1996). LeChevallier and others (1985) reported that coliform bacteria have similar susceptibility to chlorine as enterotoxigenic E. coli with greater than 90% injury observed with 0.25–0.5 mg chlorine/L. Salmonella typhimurium, Y. enterocolitica, and Shigella spp. are significantly more resistant requiring between 0.9 and 1.5 mg chlorine/L to cause injury. Chlorine injury in E. coli and Salmonella decreased the ability of the organisms to attach to Henle cells. The authors suggest that chlorine may have damaged the fimbriae which are required for attachment. In an examination of the current industry practices for fruit and vegetable decontamination, Seymour (1999) reported 80% of those surveyed used a disinfectant in the wash water, with chlorine being the most widely used disinfectant for minimally processed fruits and vegetables. Of those surveyed, 67% of respondents maintain chlorine levels of 50–200 ppm. This does not accurately measure the system’s disinfecting capacity, as it does not measure combined chlorine. Eighty-nine percent of those who use chlorine used a dip, while 11% used a spray. Eighty percent agitate the produce while washing to ensure adequate surface contact. More than half of those surveyed (60%) reported using washing temperatures of less than 5◦ C. This is desirable as it inhibits enzymatic reactions as well as microbial growth (Seymour 1999). The effectiveness of chlorine decreases as temperature increases. The solubility of chlorine is highest at 4◦ C (Beuchat 1998). Forty percent of processors recirculate wash water. Only 23% of

Shigella

29

those surveyed used a final rinse after disinfection. Only 8% of producers who test wash water for total viable counts do so more than once a month. Ninety-five percent of processors surveyed conducted microbial tests on finished products. Fifty-three percent of processors test once a week and 20% test once a day. Ninety-two percent have limits for microbial counts. Eighty-six percent of those surveyed clean their wash systems once a day and 98% carried out tests to verify sanitation (Seymour 1999). Beuchat and others (1998) evaluated the efficacy of a spray chlorine treatment for the removal of Salmonella, E. coli O157:H7, L. monocytogenes, yeasts, molds, and total aerobic microorganisms from apples, tomatoes, and lettuce. Treatment with chlorine yielded additional reductions of 0.35–2.30 orders of magnitude in pathogen populations on the surface of fruits and vegetables. Chlorine concentrations of 2,000 ppm were generally more effective than 200 ppm. Reductions in microorganisms occurred within 1 min of the application of chlorine. Zhuang and others (1995) examined the survival of Salmonella montevideo on tomato surfaces and on tomato scar tissue under conditions that would be observed in a production environment. Significant increases were observed in Salmonella on tomatoes stored at 20◦ C for 7 days and on tomatoes stored at 30◦ C for 1 day. No changes in Salmonella levels occurred on tomatoes stored at 10◦ C for 18 days. This shows the potential for survival of Salmonella during transport, storage, ripening, and consumption. A significant reduction of Salmonella on the surface was achieved by dipping tomatoes in a solution containing 60 ppm chlorine. An additional reduction was observed using a 110 ppm chlorine dip. No additional reduction was observed using a 320 ppm chlorine dip. Chlorine was less effective in killing organisms in the core than on the surface. Chlorine concentrations of 110 and 320 ppm resulted in significant reductions in Salmonella on core tissue. Salmonella populations in chopped tomatoes remained constant at 5◦ C, but increased after 96 and 22 h for samples stored at 20 and 30◦ C, respectively. pH increases were observed in chopped tomato samples over time. Organic acids act by reducing the intracellular pH of bacterial cells by ionizing the undissociated acid. Acids affect the cell’s ability to maintain pH, as well as inhibiting substrate transport and metabolic pathways. Bacteria that cause foodborne illness cannot grow in conditions with a pH less than 4 (Beuchat 1998). Washes containing organic acids have been successfully used to disinfect beef, lamb, and poultry carcasses (Beuchat 1998). Organic acids such as lactic, acetic, citric, and propionic at levels of 300–500 mg/mL yielded reductions, in total counts, similar to those achieved by water. Peracetic acid yielded a 2-order-of-magnitude reduction

30

Biofilms in the Food Environment

in total counts and fecal coliforms on salads. Oxytetracycline (50 ppm) yielded a threefold greater reduction in total counts on a fresh vegetable mix than water. A combination of chlorine and the surfactant Tween 80% was more effective than Tween or chlorine alone in reducing the microbial load of a salad mix (Nguyen-the and Carlin 1994). Zhang and Farber (1996) reported that a 1% lactic acid solution combined with a 100 ppm chlorine solution was more effective against L. monocytogenes on lettuce than either alone. Irradiation at doses less than 2 kGy is usually more effective at killing microorganisms on fresh produce than chemical disinfection, often yielding a 3–4-log reduction in total counts (Nguyen-the and Carlin 1994). Higher doses of irradiation are required for inactivating viruses (Beuchat 1998). Carmichael and others (1999) reported that lettuce entering a commercial process had a microbial load of 105 CFU/g, primarily consisting of pseudomonads. Washing and sanitizing resulted in a 100-fold reduction in the microbial load of the lettuce.

Foodborne Outbreaks Involving Shigella In March of 1999 several people became ill after dining at a Chicago area restaurant. Of all the ill patrons identified, four tested positive for Shigella boydii 18. Ill individuals were defined as those who had a stool culture that was positive for S. boydii 18 or who reported acute onset of diarrhea and fever within 72 h of dining at the restaurant in question. The eight ill individuals all consumed the bean salad, which was determined to be the causative agent in this outbreak. No leftover food samples were obtained for testing. Several items that were prepared in the same way as the original were tested. These “check-up” samples had high plate and coliform counts, but Shigella was not isolated from the samples. Thirty-three employees of the restaurant were tested and determined to be negative for Shigella and Salmonella. In August of 1998 two restaurant associated outbreaks of Shigella sonnei occurred in Minnesota. Chopped parsley was determined to be the vehicle of transmission. Two hundred ten people were affected. At the same time in California 9 people were affected by S. sonnei after consuming foods sprinkled with chopped parsley. Twenty-seven food handlers tested negative for Shigella. In Massachusetts, 6 people became ill with shigellosis after consuming chicken sandwiches and coleslaw containing chopped parsley. Again restaurant employees tested negative for Shigella. In Canada, 35 people became ill after consuming a smoked salmon and parsley dish that contained fresh parsley. Once again S. sonnei

Shigella

31

was determined to be the causative agent and food handlers tested negative. In addition to these outbreaks, four additional outbreaks of S. sonnei occurred, affecting 218 people who had consumed uncooked parsley. All isolates from the outbreaks had the same pulsed-field gel electrophoresis pattern. The parsley in these outbreaks was traced back to a farm in Mexico. At this farm water supplies used to wash produce and produce ice were not chlorinated and susceptible to contamination. Because water in the hydrocooler was recirculated, microorganisms in the water or from the parsley may have survived in the absence of chlorine and contaminated many cases of parsley. Workers and villagers did not drink this water and consumed bottled water or water from other sources. Workers had limited hygiene education and limited sanitary facilities were available. Parsley was washed by most food handlers before it was served. Chopped parsley was allowed to remain at room temperature until it was served (Crowe 1999). In 1994 a multistate outbreak of Shigella flexneri 6A was traced back to the consumption of scallions. This outbreak was detected because of a sevenfold increase in the reported number of S. flexneri 6A cases in the state of Illinois. Sixteen cases of Shigella were confirmed. The scallions were traced back to five U.S. and at least one Mexican farm. Several potential sources of contamination were found during the growth, harvest, and shipping of the scallions. Additionally, precautions to prevent the outgrowth of Shigella were not taken. S. flexneri 6A is rarely found in the United States, and is quite common in Mexico. Contamination of the scallions may have occurred during harvest or packaging in Mexico (De Roever 1998). In May and June of 1994 an increase in the number of cases of S. sonnei was observed in several European countries including Norway. Iceberg lettuce imported from Spain was determined to be the causative agent. Shigella was not isolated from lettuce samples, but large numbers of fecal coliforms (up to 80,000 CFU/g) were isolated, implying heavy fecal contamination. In Norway, 110 people were affected in this outbreak. The authors estimate 1,650 days of illness, 19 admissions to the hospital, 76 days of hospital stay, and 550 days lost from work. Twenty-eight cases of S. sonnei attributed to this outbreak occurred in England (Frost and others 1995). In 1986, 347 people in two Texas counties contracted shigellosis caused by S. sonnei. Individuals became ill after consuming shredded lettuce from three area restaurants. Restaurants which received lettuce that was not shredded were not involved in the outbreak. This implies that the lettuce was not contaminated in the field or during transport to the plant, and was most likely contaminated during shredding. One of the workers

32

Biofilms in the Food Environment

who prepared the lettuce had loose stools and cramps while at work. Specimens obtained later from the worker did not test positive for Shigella. The ill worker fed heads of washed lettuce into the shredder and the shredded lettuce was then packed into bags without preservatives or chemicals. Washing of machinery etc. did not occur until the shredding for the day was complete. Temperature of the lettuce shredding area was 14◦ C. After shredding, the lettuce remained in the shredding area for as long as 6 h and was then taken to restaurants where it was stored at 4◦ C until it was served. The 8,800 boxes of lettuce involved in the outbreak were harvested from one field. No human feces was present in the field or in its drainage ditch. Other areas to which lettuce from the harvest was shipped did not report any S. sonnei outbreaks (Davis and others 1988). In 1983 contaminated lettuce was the source of two outbreaks of S. sonnei at two universities located about 100 km apart in Texas. A total of 140 students were affected. Tossed salad was associated with illness at both schools. Food handlers were tested and found to be negative. Both universities received lettuce shipments from one company that purchased produce from several states. The lettuce was not traced back to a specific farm.

Survivability on Fresh Produce Davis and others (1988) examined the ability of an outbreak strain of S. sonnei to survive on lettuce. S. sonnei survived on lettuce at a constant level for 3 days at 5◦ C. A 1-log decrease was observed after 7 days. The outbreak strains survived, but did not grow between 5 and 15◦ C and grew well at 22◦ C. Escartin and others (1989) reported that Shigella grew on inoculated sliced papaya after storage at room temperature for 2 h and on jicama after 4–6 h. S. flexneri on watermelon increased from 2.79 to 4.49 log CFU/cube after 6 h at room temperature. The surface pH of the papaya and the jicama were 5.69 and 5.97, respectively. Rafii and others (1995) isolated 17 species of bacteria from packaged carrots, radishes, broccoli, cauliflower, lettuce, and celery. The amount of organisms varied with each sample and E. coli was among the organisms detected, but Shigella was not. The ability of S. flexneri to survive in phosphate-buffered saline was monitored. After 1 month at refrigeration temperatures the level of S. flexneri was reduced from the initial level of 109 to 107 , and remained at that level for 2 months. S. flexneri survived for several days at levels of 105 –106 CFU/g, on both sterile (sterilized using ethylene oxide) and nonsterile vegetables at both ambient and refrigeration temperatures.

Shigella

33

Rafii and Lunsford (1997) examined the ability of S. flexneri to survive in commercially prepared salads including coleslaw, crab salad, carrot salad, cabbage salad, and potato salad. Twenty-eight different species of bacteria were isolated from the salads, with populations ranging from 6.0 × 101 to 1.75 × 105 CFU/g. No pathogens were isolated from the salads. Cabbage, onion, and green pepper were also examined. Cabbage contained 1.6 × 102 CFU/g, while no organisms were detected on the pepper or onion. At 4◦ C S. flexneri survived in all four salads for at least 11 days and on the vegetables for at least 12 days. The initial amount of S. flexneri in coleslaw was 1.18 × 105 . Levels decreased to 2.16 × 104 CFU/g after 13 days. The initial level of S. flexneri in crab salad was 1.09 × 106 CFU/g, which decreased to 2.10 × 105 CFU/g by day 8 and remained at that level until day 20. For carrot salad the initial level of S. flexneri was 4.30 × 106 CFU/g, which decreased to 3.52 × 105 CFU/g after 3 days and 4.2 × 102 CFU/g after 10 days. S. flexneri was inoculated to a level of 1.32 × 106 CFU/g in potato salad, which decreased to a level of 8.50 × 102 on day 1l. S. flexneri was not killed by the low pH or the normal microflora of the salads. After inoculation the levels of S. flexneri were 5.25 × 106 , 5.10 × 106 , and 3.45 × 106 CFU/g for pepper, onion, and cabbage respectively. The levels declined to 2.20 × 104 , 2.10 × 105 , and 9.5 × 103 CFU/g after 12 days. S. flexneri was detected on cabbage at 4◦ C even after 26 days (Rafii and Lunsford 1997). Satchell and others (1990) examined the ability of S. sonnei to survive in shredded cabbage. S. sonnei in vacuum-packed cabbage initially increased, but showed a 4-order-of-magnitude decrease after day 2. In modified atmosphere and normal atmosphere sample levels of inoculated cabbage, the S. sonnei levels increased and remained high for 1–3 days and decreased thereafter. Aerobic plate counts (APCs) remained high throughout for aerobic and modified atmosphere (30% nitrogen, 70% carbon dioxide) samples. Samples stored at refrigeration temperature supported the survival of S. sonnei through 7 days with a relatively constant level of microorganisms. The type of package did not have any affect on the survival of S. sonnei. Over the 7-day period the pH of the cabbage samples stored at room temperature decreased from pH 5 to 4 for the modified atmosphere and vacuum-packed samples, and from pH 6 to 4 for the aerobically packaged samples. The decrease in pH of these samples may have contributed to the decrease in the level of Shigella. The pH of the refrigerated samples remained at a constant pH of about 6. Wu and others (2000) examined the ability of S. sonnei isolated from the 1998 parsley outbreak described above to survive on parsley. S. sonnei was suspended to an optical density of 0.5 in phosphate-buffered saline containing 5% horse serum as an organic soil. Parsley was inoculated by

34

Biofilms in the Food Environment

submerging in this suspension for 1 min with constant agitation. Parsley was allowed to dry for 1 h at room temperature in a laminar flow hood. Samples were stored at 4 or 21◦ C and 10-g samples were removed, diluted with 0.1% peptone water, and stomached for 2 min. Samples were enumerated using MacConkey agar supplemented with 20 µg/mL tetracycline. Shigella on chopped parsley grew to 9.20 and 6.32 log CFU/g from initial inoculums of 6.48 and 3.49 log CFU/g within 2 days at 21◦ C. Whole parsley supported less than 1 log of growth after day 1 and a decline in the S. sonnei population was observed after 2 days. S. sonnei grew rapidly on chopped parsley stored at 21◦ C for 24 h from an initial level of 2.72 to a final level of 6.53 log CFU/g. No lag phase was observed. This rapid growth may be attributed to the release of nutrients from the parsley cells upon cutting. S. sonnei levels on both whole and chopped parsley stored at 4◦ C declined over a 14-day period. Wu and others (2000) observed a greater than 6.1-order-of-magnitude reduction when parsley inoculated with S. sonnei was treated with vinegar containing 5.2% acetic acid for 5 min. Vinegar containing 7.6% acetic acid reduced the initial load of log 7.07 S. sonnei CFU/g to an undetectable level. Vinegar containing 2.6% acetic acid yielded a 3.3-order-of-magnitude reduction. Parsley treated with the higher acetic acid vinegar was discolored and had a strong vinegar odor. Treatment of parsley with 150 ppm chlorine resulted in a 6-order-of-magnitude reduction of S. sonnei. Treatment with 250 ppm chlorine yielded a reduction of S. sonnei from log 7.28 CFU/g to undetectable levels.

Parsley Parsley is an herb that is commonly used in the preparation of commercial and homemade foods. Often it is added after cooking, and therefore not subject to a heat treatment prior to consumption (Kaferstein 1976). Johannessen and others (2002) examined produce samples in Norway for the presence of thermotolerant coliform bacteria, E. coli O157:H7, Salmonella, L. monocytogenes, Staphylococcus, and Y. enterocolitica. No E. coli O157:H7 or Salmonella was detected. E. coli was isolated from a cilantro sample and two samples of parsley. Johannessen and others (2002) suggested that the E. coli may have originated from fecally contaminated water, soil, or improper handling of the product. In a study of domestic produce in the United States, 1 of 64 parsley samples tested was contaminated with Shigella (FDA 2001). Abu-Ghazaleh (2001) examined the ability of E. coli strains isolated from raw sewage and final effluent of wastewater treatment plants to survive

Shigella

35

on parsley stored at 28◦ C for 10 days. Strains resistant and sensitive to antibiotics were tested. Resistant strains were found to survive better with a mean survival of 11.2% compared to 6.9% survival of sensitive strains. Park and Sanders (1992) purchased 1,564 vegetable samples from supermarkets and outdoor farmers’ markets. Samples were examined for the presence of Campylobacter. These organisms were detected in 1 out of 42 (2.4%) parsley samples obtained from farmers’ market and none of the 65 summer or 70 winter parsley samples obtained from supermarkets. The authors attributed the presence of this organism to the use of untreated farm water for cleaning produce, soil-containing untreated sewage, fecal contamination from animals, or infected farmers who transmit the organism. An additional 76 parsley samples were obtained from local farmers’ markets. Two samples (2.6%) tested positive for Campylobacter. After decontamination by washing three times with tap water no samples tested positive for Campylobacter. Garcia-Villanova Ruiz and others (1987) examined 23 parsley samples from supermarkets and outdoor markets in Granada. Of the parsley samples, all had greater than 106 CFU/100 g aerobic microorganisms. Ten samples had between 109 and 1010 CFU/100 g. Twenty-two of the parsley samples had coliform counts greater than 103 CFU/100 g and 13 samples had greater than 103 CFU/100 g E. coli. One of the 23 parsley samples contained S. typhimurium. Garcia-Villanova Ruiz and others (1987) reported that microbial counts of the produce samples analyzed were higher during the summer than in the winter. This may be attributed to higher temperatures and the use of contaminated irrigation water during the summer. Rosas and others (1984) examined parsley which had been irrigated with wastewater. Using MPN calculations parsley contained total plate count of 4.3 × 104 CFU/100 g and fecal coliforms of 7 × 103 CFU/100 g. More than 90% of the organisms for both total and fecal counts were found in the roots. The authors attribute this to the contact of the roots with contaminated soil. When parsley was washed with tap water for 30 sec, 88% of the total number of organisms were removed, whereas 55% of the coliforms were removed. Abdelnoor and others (1983) examined the microbial microflora of fresh produce in Lebanon before and after water washing. Twenty-five percent of washed parsley samples contained E. coli. Of the unwashed parsley samples, 7.7% contained Staphylococcus spp. Unwashed parsley samples contained greater than 105 cells/g. Kaferstein (1976) examined the microflora of parsley. Salmonella and coagulase-positive staphylococci were not isolated from any samples. Fresh parsley from retail shops was heavily contaminated and E. coli was isolated from all 11 samples tested. Rinsing with water had little effect

36

Biofilms in the Food Environment

and fecal contamination (E. coli) was detected in two-thirds of the rinsed samples. Blanching greatly reduced the microbial load. Freezing resulted in a 1-log reduction in the number of microorganisms present, with 50% of the sample showing fecal contamination. Aseptically harvested samples of parsley had 2-order-of-magnitude fewer total aerobic microorganisms and Enterobacteriaceae than parsley samples obtained from retail markets. The aseptically harvested samples did not differ from the retail samples in yeast, mold, and clostridiacounts. Only 5% of the aseptically harvested samples were positive for E. coli, where commercially available parsley was frequently contaminated with this organism (74% of samples), implying that the contamination may be of human origin. Similar results were observed when the presence of Streptococcous was examined.

Cilantro Cilantro (Coriandrum sativuum) is one of the most widely used fresh herbs. It is featured in the cuisines of China, Southeast Asia, India, and Central and South America. It is commonly found in salsas and in poultry and seafood dishes. It is also known as coriander and Chinese parsley (Potter 1996). In March of 1999 an outbreak of Salmonella Thompson occurred. Illness was associated with eating fresh uncooked cilantro or eating a salsa containing fresh cilantro. No farm investigations were carried out due to the lack of records. It was impossible to determine if there was a common grower who supplied the restaurants involved in the outbreak. All restaurants reported washing and chopping the cilantro. Whole and chopped cilantro and the salsa were stored under refrigerated conditions. Campbell and others (2001) examined the ability of Salmonella to survive on cilantro and in fresh salsa containing cilantro. At room temperature a log increase was observed in the Salmonella population on cilantro. A 3-order-of-magnitude increase was observed on chopped cilantro. The organisms grew faster on chopped cilantro due to the release of nutrients from broken plant tissue. Refrigeration can impede the growth of Salmonella on cilantro for 3 days and in fresh salsa for 1 day. Even though salsa has a low pH it will support the growth of Salmonella. A 300-fold increase was observed in the Salmonella population of salsa stored at room temperature after 1 day (Campbell and others 2001). Brandl and Mandrell (2002) examined the ability of Salmonella serovar Thompson to survive on cilantro plants. On cilantro Salmonella grew rapidly at elevated temperatures. Populations of Pantoea agglomerans and Pseudomonas chloroaphis, organisms that are commonly found on

Shigella

37

the surface of plants, were 10 times greater than that of Salmonella serovar Thompson on cilantro plants incubated at 22◦ C. At 30◦ C Salmonella serovar Thompson reached significantly higher populations and represented a greater portion of the total portion of microorganisms on the surface of the cilantro. Higher temperature conditions may have allowed Salmonella serovar Thompson to utilize a greater share of the available nutrients, therefore increasing the competitive fitness of the organism. Salmonella serovar Thompson may not have achieved populations as high as the other organisms, because it may be unable to use the wide variety of nutrients on the surface of the cilantro leaf. This organism may be better adapted to metabolize carbon and nitrogen found in the human gut. Salmonella serovar Thompson was able to survive in dry conditions such as 60 or 50% relative humidity for several days and was able to grow to maximal levels when high humidity conditions were restored. Several serovars of Salmonella including S. enterica serovar Derby, S. enterica serovar Newport, S. enterica serovar Enteritidis, and S. enterica serovar Thompson exhibited similar colonization trends on cilantro. Cilantro plants were inoculated with a GFP Salmonella serovar Thompson and observed using confocal laser scanning microscopy. The organisms were observed on leaf veins and adjacent areas as well as the base of the leaf where the petiole joins the leaf. This may be attributed to the waxy trichnome of the cilantro leaf surface as well as the tendency of surface water to accumulate in the depressions of leaf veins. When the GFP Salmonella together with a DsRed P. agglomerans were used to inoculate the surface of cilantro leaves the organisms were detected in large mixed aggregates. Salmonella also formed mixed colonies with the native microflora. Salmonella serovar Thompson was found to infect damaged tissue and reach higher cell densities than on healthy cilantro tissue. Lesions may provide access to the nutritious inner leaf tissue as well as protection from environmental stresses. This clearly demonstrates that human pathogens can colonize plants in the field prior to harvest (Brandl and Mandrell 2002). In a survey of domestic fresh produce conducted by the FDA 1 of 62 cilantro samples tested was contaminated with Shigella (FDA 2001). One of the cilantro samples was also contaminated with Salmonella.

Acid Tolerance Stationary phase E. coli K-12 and S. flexneri can survive for several hours at low pH (2–3) (Small and others 1994). This may contribute to the low infective dose associated with shigellosis. S. flexneri cultures grown initially at an external pH range of 5–8 were reported to be 100% acid resistant,

38

Biofilms in the Food Environment

surviving at pH 2.5 for 2 h. Exponential phase cultures grown at pH 5 were also acid resistant; however, survival decreased 10–100-fold as the pH of the growth medium was raised to pH 8. Lin and others (1995) reported the minimum growth pH for S. flexneri to be pH 4.8, which was greater than that required for both E. coli (pH 4.4) and S. typhimurium (pH 3.0). S. flexneri and E. coli, however, both survived exposure to low pH levels better than S. typhimurium. Small and others (1994) reported a 1.3 kb fragment cloned from S. flexneri was found to confer acid resistance on E. coli HB101 and S. flexneri Tn10. This fragment was identified as homolog of rpoS, the growth-phase-dependent sigma factor δ 38 (Small and others 1994). RpoS is a component of the RNA polymerase that directs the polymerase to promoters that are poorly recognized by the housekeeping sigma factor σ 70 (Waterman and Small 1996). RpoS is a major regulator of late-log and stationary phase growth. Waterman and Small (1996) used transposon (TnlacZ and TnphoA) mutagenesis to create acid-sensitive S. flexneri mutants. Mutations were observed in the hdeA gene in two mutants and in an open reading frame (ORF) downstream of the gadB gene. This ORF encodes a protein that has homology to several inner membrane amino acid antiporters. The fusions, positively regulated by RpoS, were induced upon entry into late-exponential phase growth. Zaika (2001) examined the ability of S. flexneri 5348 to survive in brain heart infusion broth adjusted to pH 2–5 with HCl at various temperatures. Survival increased as pH increased at all temperatures. At pH 2 and 19◦ C S. flexneri was undetectable after 30 h. S. flexneri was present at 2 log CFU/mL after 8 days at pH 3 and after 23 days at pH 4. At 12◦ C a 5-order-of-magnitude decrease in CFU/mL was observed at pH 3 after 13 days. A 1.1-log decrease in the S. flexneri population was observed after 58 days at pH 5. At pH 5 growth occurred at 19, 28, and 37◦ C. Fehlhaber (1981) examined the ability of strains of S. flexneri and S. sonnei to survive in nutrient broth at pH 3–4.5 at room temperature. Fifteen strains of S. flexneri survived for 30 min at pH 3. All strains survived for 4 h at pH 4.5, two strains survived for 2 days. This author used mid-exponential phase cells, which have been reported to be less acid tolerant than cells in the stationary phase (Fehlhaber 1981). Bagamboula and others (2002) examined the behavior of multiple strains of S. sonnei and S. flexneri in media at different pH from 3.25 to 5 and in different fruit products. At pH 5 all Shigella strains tested grew to maximal levels within 6 h. The minimum pH for growth of S. flexneri was 4.75. S. sonnei was able to grow at low pH 4.5. At pH 4.25 a great deal of variability in the ability of strains to survive was observed. At pH 4.0 and 3.75 all strains were recovered after 24 h, but populations were greatly reduced. The authors reported that laboratory strains survived better in

Shigella

39

acidic conditions than clinical isolates. This is contrary to what was expected as clinical isolates have been exposed to the acidic conditions of the stomach. Shigella decreased rapidly at 22◦ C in apple juice (pH 3.3– 3.4) and tomato juice (pH 3.9–4.1). Some strains were detected for as long as 14 days in tomato juice and 8 days in apple juice. The survival of low numbers of Shigella on strawberries and fresh fruit salad was examined. S. flexneri was not detected on strawberries stored at 4◦ C for 4 or 48 h. It was, however, detected in all samples of fresh fruit salad at 4◦ C for 4 or 48 h (Bagamboula and others 2002). Tetteh and Beuchat (2001) examined the effects of organic acids on the survival and growth of unadapted, acid-adapted, and acid-shocked S. flexneri. At the same pH, propionic acid was the most inhibitory, followed by acetic and lactic acids. Acid-adapted cells were found to be more resistant to acid environments than were acid-shocked or unadapted cells.

Produce Wash Consumers realize that fruits and vegetables should be washed prior to consumption. Ninety-five percent of consumers recognize the need for thorough washing of produce, but most use only water for this purpose. Of those who wash produce, 5% use a household cleaner such as dishwashing liquid. These liquids may be problematic because they provide large volumes of persistent suds, which is difficult to remove. Additionally, many components of these products may be undesirable, as they may not be completely removed from the food product. Ingredients in a product used for fruits and vegetables washes should have generally recognized as safe (GRAS) status as described in the Code of Federal Regulations (CFR). Any product making a bactericidal claim must be registered and approved by the Environmental Protection Agency (EPA). A variety of sanitizers have been examined for their ability to kill pathogens. Since there is no standardized method for the evaluation of produce sanitizers it is difficult to compare results between laboratories. In September of 1997 the EPA created a scientific advisory panel to discuss the development a standardized method to evaluate produce sanitizers. The sanitizers should be effective against five strains of E. coli O157:H7, L. monocytogenes, and Salmonella, and a reasonable performance would be described as a 2-log reduction in the number of pathogens present (Harris and others 2001). Beuchat and others (2001a) discussed several factors involved in the development of methods to examine the efficacy of sanitizer on microorganisms on fresh produce. In order to compare results among laboratories,

40

Biofilms in the Food Environment

well-characterized reference strains should be used. A cocktail of five strains in equal concentrations should be used as the inoculum, because if the strains exhibit different degrees of susceptibility the most robust strain will prevail. Cells used for the inoculum should be in the stationary phase, because cells in this growth phase are less susceptible to environmental stresses. Fruits and vegetables can be contaminated at anytime from the field to the consumer by a wide variety sources such as dust, rain water, irrigation water, soil, sewage, feces, contact surfaces, and workers. Pathogens are likely to be found in organic material. In order to simulate these conditions inocula should be prepared with a 5% solution of horse serum. Higher levels of inocula should be used for disinfection and retrieval studies whereas lower levels should be used in challenge studies. Special considerations must be made depending on the type of fruit or vegetable tested. Homogenizing samples in order to enumerate microorganisms may release components that could be lethal to the organisms to be enumerated. Rubbing may be an effective way to remove microorganism from certain fruits and vegetables such as apples. Agitation or sonnication may be more appropriate for leafy vegetables. Removal of microorganisms may be complicated by factors such as cutting or bruising or treatment with wax or oil. Inoculation by dipping or spraying with a cell suspension may be appropriate especially if the contamination may have occurred in a commercial immersion process. A problem associated with dipping is the actual number of organisms applied or adhering to the produce is not known. Spotting 10 or 50 µL of an inoculum of known cell density on the surface of produce may be more effective and would represent contamination from a point such as soil, equipment, or a worker’s hand. A negative temperature differential (warm produce and cold inoculum) can result in uptake of the organisms by the plant tissue. The inoculated produce should be dried for a specific amount of time prior to disinfection. Procter and Gamble (2000) developed several produce washes containing GRAS ingredients, which could be used to remove unwanted deposits such as wax or soil from produce. These washes also exhibited bactericidal properties. These products contain a variety of ingredients such as potassium hydroxide, ethanol, glycerin, oleic acid, sodium bicarbonate, phosphoric acid, citric acid, and essence. Procter and Gamble tested the efficacy of commercial FitTM produce wash on bacterial suspensions and on produce inoculated with several human pathogens. The results can be seen in Tables 2.1 and 2.2. For the in vitro suspension testing a modification of AOAC 960.09 was used. Commercial FitTM was tested at a concentration of 5 g/L. Staphylococcus aureus, L. monocytogenes, E. coli, Pseudomonas aeruginosa, Pseudomonas cepacia, and Salmonella choleraesuis were tested. Reductions of at least 6 orders of magnitude

41

Shigella Table 2.1.

Efficacy of Commercial FitTM Produce Wash on Bacterial Suspensions

Bacteria Staphylococcus aureus (ATCC 6538) Listeria monocytogenes (ATCC 19117) Escherichia coli (ATCC 11229) Pseudomonas aeruginosa (ATCC 15442) Pseudomonas cepacia (ATCC 25416) Salmonella choleraesuis (ATCC 10708)

Log Reduction Using FitTM (5 g/L) for 1 min

Log Reduction Using FitTM (5 g/L) for 5 mina

>5 log10

>6 log10

>6 log10

>6 log10

>6 log10

>7 log10

>6 log10

>6 log10

>6 log10

>6 log10

>4 log10

>7 log10

Source: Tables modified from data obtained from Pettigrew (2000). a FitTM recommended usage is to soak processed produce for 5 min.

were observed for all organisms after 1- or 5-min treatments with commercial FitTM produce wash, except for S. choleraesuis which exhibited a 4-order-of-magnitude decrease after 1 min of treatment. The efficacy of commercial FitTM (5 g/L), water or chlorine (200 ppm) after a 5-min treatment of Salmonella on tomatoes, and E. coli O157:H7 or S. aureus on lettuce, broccoli, and tomatoes was examined. The results are listed in Table 2.2.

Produce

Efficacy of Commercial FitTM Produce Wash Against Human Pathogens Log Reduction Using FitTM (5 g/L)

Salmonella sp. Tomato 4.17 Escherichia coli O157:H7 Lettuce 1.43 Tomato 2.49 Broccoli 1.62 Staphylococcus aureus Lettuce 2.08 Tomato 2.57 Broccoli 1.81

Log Reduction Using FitTM (5 g/L) vs. Water

Log Reduction HOCl (200 ppm)

Log Reduction HOCl (200 ppm) vs. Water

2.99a

5.18

4.00

0.89a 1.74a 0.82a

1.79 2.11 1.55

1.25 1.35 0.75

0.91a 1.70a 1.16a

2.09 2.48 1.72

0.92 1.60 1.08

Source: Tables modified from data obtained from Pettigrew (2000). a Difference between FitTM and Water Means are significant at the 95% confidence interval.

42

Biofilms in the Food Environment

Table 2.2. Commercial FitTM was significantly better than water in killing pathogenic microorganisms on produce. The efficacy of commercial FitTM on produce was dependent on the type of produce tested. Commercial FitTM was effective against Gram-negative and Gram-positive organisms. Harris and others (2001) developed a method of assessing the efficacy of produce sanitizers to be recommended to the EPA. The method simulated home washing of produce. Tomatoes were used as the model vegetable and the procedure involved spot inoculation with a multistrain cocktail, a short drying period, exposure to the sanitizer, a water rinse, and peptone water wash. A 2-order-of-magnitude greater reduction of Salmonella on tomatoes was observed following treatment with alkaline FitTM produce wash versus inoculated tomatoes treated with water or with Dey and Engley (D/E) neutralizer broth. Between 2.82 and 4.08 log CFU/tomato was recovered from tomatoes treated with FitTM produce wash vs. 8.15– 8.60 log CFU/tomato recovered from tomatoes treated with D/E broth. D/E broth exhibited almost identical results to water. Results were consistent across a private contract laboratory, an industry laboratory, and an academic laboratory. Beuchat and others (2001b) examined the ability of FitTM produce wash and chlorine to kill Salmonella and E. coli O157:H7 on alfalfa sprouts. Treating the seeds may be a more effective means of reducing the large numbers of pathogens on sprouts. When seeds were treated with 200 ppm of chlorine for 30 min, a 1.9-log reduction was observed in Salmonella. Salmonella was reduced by 2.3 orders of magnitude when treated with 20,000 ppm chlorine or FitTM produce wash for 15 or 30 min. Treatment with 20,000 ppm chlorine or FitTM reduced the germination percentage of the seeds. E. coli was not detected in the D/E broth used to neutralize samples treated with FitTM . It was, however, detected in the D/E broth upon enrichment. Treatment with 20,000 ppm chlorine yielded a 2-orderof-magnitude reduction of E. coli. Alfalfa seeds are often scarified to promote rapid and uniform germination. This may result in cells lodging on the seed surface. Using a different inoculation procedure and seeds from a different supplier Salmonella and E. coli were used to inoculate sprout seeds. Salmonella was reduced by 0.2, 2.5, and 1.7 orders of magnitude when seeds were treated with 200 ppm chlorine, 20,000 ppm chlorine, or 20,000 ppm FitTM produce wash, respectively. Similar reductions in E. coli were observed for 20,000 ppm chlorine and for FitTM produce wash, both of which were greater than 200 ppm chlorine. The presence of the additional organic load in the second inoculation procedure may have lessened the effectiveness of the 200 ppm chlorine and the FitTM treatments, but not the 20,000 ppm chlorine treatment. Beuchat and others (2001a,b)

Shigella

43

concluded that differences in the seeds obtained from two suppliers and in the organic loads of the inoculum altered the efficacy of the sanitizers. FitTM and 20,000 ppm chlorine caused similar reductions in E. coli and Salmonella on alfalfa seeds. Takeuchi and Frank (2001) compared the efficacy of an alkaline prototype produce wash on E. coli O157:H7 on lettuce. The produce wash yielded a 0.7–1.1 log CFU/cm2 reduction, which was greater than the reduction observed with a baking soda-salt solution.

References Abdelnoor AM, Batshoun R, Roumani BM. 1983. The bacterial flora of fruits and vegetables in Lebanon and the effect of washing on the bacterial content. Zentralbl Bakteriol Hyg Abt. 177(3–4):342–349. Abdul-Raouf UM, Beuchat LR, Ammar MS. 1993. Survival and growth of Escherichia coli O157:H7 on salad vegetables. Appl Environ Microbiol 59(7):1999–2006. Abu-Ghazaleh BM. 2001. Fecal coliforms of wastewater treatment plants: Antibiotic resistance, survival on surfaces and inhibition by sodium chloride and ascorbic acid. New Microbiol 24(4):379–387. Ahvenainen R. 1996. New approaches in improving the shelf life of minimally processed fruit and vegetables. Trends Food Sci Technol 7:179–187. Bagamboula CF, Uyttendaele M, Debevere J. 2002. Acid tolerance of Shigella sonnei and Shigella flexneri. J Appl Microbiol 93(3):479–486. Beuchat LR. 1992. Surface disinfection of raw produce. Dairy Food Environ Sanit 12(1):6–9. Beuchat LR. 1995. Pathogenic microorganisms associated with fresh produce. J Food Prot 59(2):204–216. Beuchat LR. 1998. Surface decontamination of fruits and vegetables eaten raw: A review. Food Safety Issues, Food Safety Unit, WHO, Geneva, Switzerland. WHO/FSE/98.2. Beuchat LR. 2002. Ecological factors influencing survival and growth of human pathogens on raw fruits and vegetables. Microbes Infect 4(4):413–423. Beuchat LR, Farber JM, Garrett EH, Harris LJ, Parish ME, Suslow TV, Busta FF. 2001a. Standardization of a method to determine the efficacy of sanitizers in inactivating human pathogenic microorganisms on raw fruits and vegetables. J Food Prot 64(7): 1079–1084. Beuchat LR, Harris LJ, Ward TE, Kajs TM. 2001b. Development of a proposed standard method for assessing the efficacy of fresh produce sanitizers. J Food Prot 64(8):1103– 1109. Beuchat LR, Nail BV, Adler BB, Clavero MRS. 1998. Efficacy of spray application of chlorinated water in killing pathogenic bacteria on raw apples, tomatoes, and lettuce. J Food Prot 61(10):1305–1311. Brandl MT, Mandrell RE. 2002. Fitness of Salmonella enterica serovar Thompson in the cilantro phyllosphere. Appl Environ Microbiol 68(7):3614–3621. Campbell JV, Mohle-Boetani J, Reporter R, Abbott S, Farrar J, Brandl M, Mandrell R, Werner SB. 2001. An outbreak of Salmonella serotype Thompson associated with fresh cilantro. J Infect Dis 183(6):984–987.

44

Biofilms in the Food Environment

Carmichael I, Harper IS, Coventry MJ, Taylor PWJ, Wan J, Hickey MW. 1999. Bacterial colonization and biofilm development on minimally processed vegetables. J Appl Microbiol Symp Suppl 85: 45S–51S. Crowe L. 1999. From the Centers for Disease Control and Prevention. Outbreaks of Shigella sonnei infection associated with eating fresh parsley—United States and Canada, July– August 1998. JAMA 281(19):1785–1787. Davis H, Taylor JP, Perdue JN, Stelma GN Jr, Humphreys JM Jr, Rowntree R, Greene KD. 1988. A shigellosis outbreak traced to commercially distributed shredded lettuce. Am J Epidemiol 128(6):1312–1321. De Roever C. 1998. Microbiological safety evaluations and recommendations on fresh produce. Food Control 9(6):321–347. Escartin EF, Ayala AC, Lozano JS. 1989. Survival and growth of Salmonella and Shigella on sliced fresh fruit. J Food Prot 52(7):471–472. Fehlhaber K. 1981. Untersuchungen uberlebensmittelhygienisch bedeutsame Eigenschaften von Shigellen. Arch Vet Med Leipzig 35:955–964. Food and Drug Administration. 2001. Survey of Domestic Fresh Produce: Interim Results (July 31, 2001). Frost JA, McEvoy MB, Bentley CA, Anderson Y, Rowe B. 1995. An outbreak of Shigella sonnei infection associated with consumption of iceberg lettuce. Emerg Infect Dis 1(1): 26–29. Garcia-Villanova RB, Vargas RG, Garcia-Villanova R. 1987. Contamination on fresh vegetables during cultivation and marketing. Int J Food Microbiol 4:285–291. Garg N, Churey JJ, Splittstoesser DF. 1990. Effect of processing conditions on the microflora of fresh-cut vegetables. J Food Prot 53(8):701–703. Harris LJ, Beuchat LR, Kajs TM, Ward TE, Taylor CH. 2001. Efficacy and reproducibility of a produce wash in killing Salmonella on the surface of tomatoes assessed with a proposed standard method for produce sanitizers. J Food Prot 64(10):1477–1482. Hurst WC, Schuler GA. 1992. Fresh produce processing: an industry perspective. J Food Prot 55(10):824–827. Janisiewicz WJ, Conway WS, Brown MW, Sapers GM, Fratamico P, Buchanan RL. 1999. Fate of Escherichia coli O157:H7 on fresh-cut apple tissue and its potential for transmission by fruit flies. Appl Environ Microbiol 65(1):1–5. Johannessen GS, Loncarevic S, Kruse H. 2002. Bacteriological analysis of fresh produce in Norway. Int J Food Microbiol 77(3):199–204. Kaferstein FK. 1976. The microflora of parsley. J Milk Food Technol 39(12):837–840. King AD Jr, Magnuson JA, Torok T, Goodman N. 1991. Microbial flora and storage quality of partially processed lettuce. J Food Sci 56(2):459–461. Kudva IT, Blanch K, Hovde CJ. 1998. Analysis of Escherichia coli O157:H7 survival in ovine or bovine manure and manure slurry. Appl Environ Microbiol 64(9):3166–3174. LeChevallier MW, Singh A, Schiemann DA, McFeters GA. 1985. Changes in virulence of waterborne enteropathogens with chlorine injury. Appl Environ Microbiol 50(2):412– 419. Nguyen-the C, Carlin F. 1994. The microbiology of minimally processed fresh fruits and vegetables. Crit Rev Food Sci Nutr 34(4):371–401. Park CE, Sanders GW. 1992. Occurrence of thermotolerant campylobacters in fresh vegetables sold at farmers’ outdoor markets and supermarkets. Can J Microbiol 38(4):313–316. Pettigrew CA. 2000. Antimicrobial efficacy of Fit powder. Personal communication, Cincinnati, 21 August 2000. Potter TL. 1996. Essential oil composition of cilantro. J Agric Food Chem 44(7):1824–1826.

Shigella

45

Procter and Gamble. 2000. Fresher better tasting produce made possible by innovation from the Procter and Gamble Company and T.G.I. Friday’s restaurants, Dallas, 18 April 2000. Rafii F, Holland MA, Hill WE, Cerniglia CE. 1995. Survival of Shigella flexneri on vegetables and detection by polymerase chain reaction. J Food Prot 58(7):727–732. Rafii F, Lunsford P. 1997. Survival and detection of Shigella flexneri in vegetables and commercially prepared salads. J AOAC Int 80(6):1191–1197. Rosas I, Baez A, Coutino M. 1984. Bacteriological quality of crops irrigated with wastewater in the Xochimilco plots, Mexico City, Mexico. Appl Environ Microbiol 47(5):1074–1079. Satchell FB, Stehphenson P, Andrews WH, Estela L, Allen G. 1990. The survival of Shigella sonnei in shredded cabbage. J Food Prot 53(7):558–562. Seymour IJ. 1999. Review of current industry practice on fruit and vegetable decontamination. Review No. 14. Campden & Chorleywood Food Research Association, Gloucestershire, U.K. Small P, Blankenhorn D, Welty D, Zinser E, Slonczewski JL. 1994. Acid and base resistance in Escherichia coli and Shigella flexneri: Role of rpoS and growth pH. J Bacteriol 176(6):1729–1737. Solomon EB, Yaron S, Matthews KR. 2002. Transmission of Escherichia coli O157:H7 from contaminated manure and irrigation water to lettuce plant tissue and its subsequent internalization. Appl Environ Microbiol 68(1):397–400. Takeuchi K, Frank JF. 2001. Direct microscopic observation of lettuce leaf decontamination with a prototype fruit and vegetable washing solution and 1% NaCl–NaHCO3 . J Food Prot 64(8):1235–1239. Tetteh GL, Beuchat LR. 2001. Sensitivity of acid-adapted and acid-shocked Shigella flexneri to reduced pH achieved with acetic, lactic, and propionic acids. J Food Prot 64(7):975– 981. Waterman SR, Small PLC. 1996. Identification of ss-dependent genes associated with the stationary-phase acid-resistance phenotype of Shigella flexneri. Mol Microbiol 21(5):925–940. Wu FM, Doyle MP, Beuchat LR, Wells JG, Mintz ED, Swaminathan B. 2000. Fate of Shigella sonnei on parsley and methods of disinfection. J Food Prot 63(5):568–572. Zaika LL. 2001. The effect of temperature and low pH on survival of Shigella flexneri in broth. J Food Prot 64(8):1162–1165. Zhang S, Farber JM. 1996. The effects of various disinfectants against Listeria monocytogenes on fresh-cut vegetables. Food Microbiol 13:311–321. Zhuang RY, Beuchat LR, Angulo FJ. 1995. Fate of Salmonella montevideo on and in raw tomatoes as affected by temperature and treatment with chlorine. Appl Environ Microbiol 61(6):2127–2131.

Biofilms in the Food Environment Edited by Hans P. Blaschek, Hua H. Wang, Meredith E. Agle Copyright © 2007 by Blackwell Publishing and the Institute of Food Technologists

Chapter 3 BIOFILM DEVELOPMENT BY LISTERIA MONOCYTOGENES Scott E. Hanna and Hua H. Wang

Introduction Listeria monocytogenes is one of the most important food-borne pathogens, capable of causing severe illness in susceptible individuals, particularly the young, old, pregnant, and immunocompromised. Listeriosis has a mortality rate of 20–30%, among the highest in food-borne diseases. L. monocytogenes is widespread in the environment. It can grow at temperatures ranging from 1 to 45◦ C, at acidity levels from pH 4 to 9, and at salt concentrations up to 10% (Yousef and Carlstrom 2003). These wide ranges allow it to flourish in conditions that inhibit the growth of most bacteria. The psychrotrophic nature of L. monocytogenes makes it especially dangerous in ready-to-eat foods that are stored at refrigeration temperatures, since even small numbers can multiply during storage. Many listeriosis cases are linked to secondary contamination of L. monocytogenes during or after food processing. L. monocytogenes is capable of surviving various stress conditions that are commonly found in foods and food processing environment. It tends to maintain itself in niches in the processing environment, particularly in hard to clean areas such as drains, rollers on conveyor belts, and worn or cracked rubber seals around doors (Tompkin 2002). An individual strain—known as a persistent strain—can sometimes be repeatedly isolated from the same facility over a period of months or years, although sporadic contamination also occurs (Kathariou 2002). The formation of biofilms involving L. monocytogenes is believed to be a main reason for such persistence. Once bacteria form biofilms, they become more resistant to cleaning and sanitation treatment, and cells detaching from the biofilm can further turn into the source of persistent contamination. 47

48

Biofilms in the Food Environment

Although improving sanitation has been the main drive for L. monocytogenes biofilm studies, recent advancement in understanding microbial virulence and biofilm forming mechanisms inspires the investigation of L. monocytogenes biofilm at the molecular level. Molecular attributes essential for L. monocytogenes strains to form biofilms on abiotic surfaces may also be involved in L. monocytogenes pathogenicity, from host tissue attachment to invasion. Many identified L. monocytogenes virulence factors essential for the bacteria attaching to and invading host tissues and ultimately causing disease, such as internalin, listeriolysin O (LLO), and ActA, are surface proteins that might also have a role in L. monocytogenes attachment to abiotic surfaces. Therefore a comprehensive understanding of the L. monocytogenes biofilm development mechanism is essential for designing effective strategies for proper disease prevention and treatment.

Single Culture L. monocytogenes Biofilm Much of the knowledge on L. monocytogenes biofilm has been gained from studying the behavior of the organism in monoculture. Results from such studies provide key insights into the unique features of L. monocytogenes biofilm formation, as well as biofilm characteristics shared with other bacterial species. Examining the contributions of environmental factors and strain specificities on L. monocytogenes biofilm formation can facilitate the identification of key biofilm attributes and stress-responsive elements involved in protecting microorganisms in adverse environment, and thus enable the development of targeted antagonistic strategies to minimize L. monocytogenes contamination.

L. monocytogenes Biofilm Development Relatively little is known about the events involved in L. monocytogenes biofilm development, maturation, and detachment. Marsh and others (2003) monitored the development of L. monocytogenes biofilms up to 72 h. Distinctive L. monocytogenes biofilm structures, from initiation to well-developed three-dimensional “honeycomb” networks, were captured by scanning electron microscopy (SEM). Chavant and others (2002) examined L. monocytogenes biofilm development over a longer period of time using SEM and observed similar biofilm developmental stages.

Biofilm Development by Listeria monocytogenes

49

Initiation Early adherence of L. monocytogenes cells to the underlying surface is the initiating event in biofilm formation. Takhistov and George (2004) observed rapid adherence (within 3–5 sec) of a few L. monocytogenes cells to random areas on an aluminium surface. This adherence was initially sparse and uniform but became more clustered over time, eventually leading to the formation of microcolonies. This initial adhesion contains both reversibly and irreversibly bound cells, with the number of irreversibly bound cells increasing over time (Beresford and others 2001). Vatanyoopaisarn and others (1999) reported that the presence of flagella is important for early L. monocytogenes adherence. This group observed that a nonflagellated mutant attached to stainless steel at a 10-fold reduced rate than the corresponding wild type at 22◦ C during the first 4 h; however, after 6–24 h of incubation there was no significant difference. At 37◦ C, a temperature at which L. monocytogenes does not produce flagella, there was no difference in attachment between the two strains. Meylheuc and others (2001), however, saw no difference in the attachment of cells at 20 and 37◦ C to stainless steel or polytetrafluoroethylene (PTFE) at 2 h. Chae and Schraft (2000) also noted the lack of correlation between the amount of initial adhesion and the amount of biofilm a particular strain produced on glass after 24 h of growth. Initial attachment is dependent on the interaction between the cell wall and the underlying surface. Studies evaluating the surface physiochemistry of L. monocytogenes as it relates to biofilm formation have evaluated hydrophilic interactions between these two components. Briandet and others (1999) found that glucose and lactic acid lowered the hydrophilicity of L. monocytogenes and increased its attachment to stainless steel. While Meylheuc and others (2001) did not observe a difference in attachment between flagellated and nonflagellated L. monocytogenes, they did note that at 20◦ C the cells were more electronegative, possibly due to the extra COO− groups present on the flagella. Smoot and Pierson (1998) also found no correlation between cell hydrophobicity and attachment. Their study suggests that proteins play a role in cell adherence to both stainless steel and Buna-N rubber, since L. monocytogenes attachment on both these surfaces was reduced 99.9% in the presence of trypsin.

Development, Maturation, and Detachment Relatively little is known about the events involved in L. monocytogenes biofilm development and detachment. Takhistov and George (2004) observed that following the initial attachment of L. monocytogenes, new

50

Biofilms in the Food Environment

cells preferentially attach to the extracellular polymeric substances (EPS) produced by the initial adherent cells. They also noted that daughter cells did not detach from the parent cells but instead formed microcolonies, growing toward areas of higher nutrients and fewer cells. The number and size of these colonies increased with time, eventually forming intercolony bridges. They describe this appearance as a “bacterial web,” similar to the net-like and honeycomb patterns that were reported by Marsh and others (2003). As the biofilm ages, the EPS begins to decrease. Takhistov and George (2004) attribute this to the use of the EPS as an energy source. As the EPS is used up, it becomes easier for cells to detach; eventually the EPS becomes weakened to the point that the entire biofilm can detach from the surface. In this study, maximum surface population of L. monocytogenes was reached at 90 h of incubation with detachment of cells occurring after 120 h. It was also observed that dead cells did not detach from the surface.

Structural Characteristics and EPS Production The structure of L. monocytogenes biofilm is different from the “classic” mushroom-shaped growth observed with many other species. Chae and Schraft (2000) described a two-layer biofilm structure by L. monocytogenes. The biofilms were cultivated on a glass surface with static nutrient supply, and confocal scanning laser microscopy (CSLM) was used to reveal the three-dimensional structural features. It was found that L. monocytogenes biofilm architecture consisted of upper and lower layers containing more than 105 cells/cm2 and an area between the layers with less than 105 cells/cm2 . The upper and lower layers were approximately 4.54 µm and 5.24 µm thick, respectively, with 2.13 µm between them. The authors acknowledged that the biofilm structure might be different under flow conditions. Using SEM, Marsh and others (2003) observed the development of a “honeycomb” biofilm structure on stainless steel coupons cultured in static conditions by two out of the three virulent L. monocytogenes strains examined (Figure 3.1). The two strains that formed this complex biofilm structure also exhibited a net-like pattern during early attachment to the surface, which can be readily observed by wide-field fluorescence microscopy (WFM). The third strain, which produced only a sparse, random adherence of cells, did not exhibit this early pattern (Figure 3.2). The EPS produced by L. monocytogenes is quite different from those found in Pseudomonas spp. and Staphylococcus spp., and its production is affected by experimental conditions. Mafu and others (1990a) observed extracellular material (ECM) production on a variety of surfaces in as little as 1 h. This ECM had a stringy, fibrillar appearance, similar to the EPS

Biofilm Development by Listeria monocytogenes

51

Figure 3.1. The “honeycomb” biofilm structure by L. monocytogenes strain Scott A. (From Marsh and others 2003.)

seen by Marsh and others (2003) that appeared to anchor the cells in place. Ronner and Wong (1993) observed ECM production on both stainless steel and Buna-N rubber within 2 days at room temperature, which is consistent with the report by Herald and Zottola (1988) showing that L. monocytogenes produced extracellular fibrils on stainless steel at 21◦ C but not at 10 or 35◦ C. Under continuous flow conditions, Sasahara and Zottola (1993) did not observe EPS formation by L. monocytogenes on a glass surface. Similar results were seen on condensate-forming stainless steel, a more stationary setting (Hassan and others 2004). In both these studies L. monocytogenes could readily use the EPS produced by other bacteria species to form biofilm. It may be worth noting that among these studies, incubating the biofilms in a fixed growth medium—with or without agitation—induced EPS production by L. monocytogenes, while incubating under flow conditions or in a condensate did not. Finally, there appears to be a difference among L. monocytogenes strains with respect to EPS production. Borucki and others (2003) evaluated 80 different L. monocytogenes strains for biofilm-forming capabilities using a microtiter plate assay, and found that while all the strains produced extracellular polysaccharides, the highest biofilm formers produced noticeably more EPS.

52

Biofilms in the Food Environment

(a)

(b)

(c)

(d)

Figure 3.2. WFM assessment of complex structure development by L. monocytogenes Scott A grown in TSB on plastic chamber slides for 3–72 h, according to patterns observed after sample processing. (a) 3 h; (b) 6 h; (c) 24 h; (d) 72 h. At 72 h the net-like pattern is not distinct. Multiple layers may be dense, network may be dissociated, or structure may have peeled from the surface. Pictures of (a) and (b) were captured using a 10× objective lens and the bars represent 100 µm; pictures of (c) and (d) were captured using a 40× objective lens and the bars represent 25 µm. (From Marsh and others 2003.)

Strain Variability L. monocytogenes strains are often grouped according to serotype, phylogenetic lineage, or source. Two different antigens are used to determine the serotype, somatic (1–4) and flagellar (a–d). The 13 existing serotypes are further grouped into lineages, with division I consisting of serotypes 4b, 1/2b, and 3b, and division II containing serotypes 1/2a, 1/2c, 3a, and 3c (Kathariou 2002). The serotypes most commonly implicated in human listeriosis are 4b, 1/2b, and 1/2a. The source of the bacteria—whether from humans, food, animals, or the environment—can also be used to segregate one strain from another. It is of particular interest whether group-specific features, such as cell surface antigens or virulence factors, contribute to biofilm formation on abiotic surfaces. Since some strains of L. monocytogenes can persist in a processing environment for months, or even years, a serious concern is whether such strains may have enhanced biofilmforming capabilities and possibly be more virulent compared to other,

Biofilm Development by Listeria monocytogenes

53

more sporadically isolated strains. Therefore revealing the potential relationship among strain persistence, biofilm formation, and virulence could facilitate identifying molecular attributes or regulatory pathways involved in these events and be important for disease prevention and treatment. Norwood and Gilmour (1999) evaluated 111 strains of L. monocytogenes for their biofilm-forming capabilities on stainless steel. They tested 35 persistent strains, which were isolated repeatedly from the same environment over a period of months, and 24 sporadic strains, which were isolated only once during this period; the remainder were from stock cultures. The CFU/cm2 for each strain was evaluated after 24 h of incubation in dilute (1:15) tryptic soy broth (TSB) at 25◦ C. The persistent strains had a higher mean CFU/cm2 than did the sporadic strains ( p = 0.041); however, there was substantial overlap between these two categories. When the serotypes of the known strains were compared, the mean of the 1/2c strains was significantly higher than that of the 4b and 1/2a strains; also, the 4b strains’ mean was higher than that of the 1/2a strains. There was no significant difference in adherence when comparing the source of the strains (meat, dairy, or clinical). Chae and Schraft (2000) compared 13 strains of L. monocytogenes, evaluating their adhesion to and ability to form biofilm on glass slides. All the strains formed biofilm within 24 h following the initial adhesion assay, but the amount formed did not necessarily correspond to the initial attachment numbers. There was no trend among the different serotypes or sources for either adherence or biofilm formation, and the initial inoculum CFU/mL also seemed to play no role in the adherence or biofilm growth. By comparing 3 persistent and 14 sporadic L. monocytogenes strains, Lunden and others (2000) showed that the persistent strains had higher adherence at 1 and 2 h of contact time with stainless steel. Coupons were incubated during this time at 25◦ C in TSB with shaking, with the persistent strains showing a 2.7- to 4.6-fold higher amount of adherent cells. After 72 h of incubation, however, some of the nonpersistent strains had a higher CFU/cm2 than the persistent strains. As with Norwood and Gilmour (1999), this group found the 1/2c serotype to be the most adherent; two of the three persistent strains were of this serotype. Kalmokoff and others (2001) evaluated 36 strains of L. monocytogenes, as well as some other bacterial species, for adhesion and biofilm formation on stainless steel in brain-heart infusion. They found no correlation between source or serotype and 2-h adhesion capabilities. Noteworthy is the fact that under these experimental conditions the group found that there was little difference in the amount of adhesion among the L. monocytogenes strains, and overall they adhered poorly compared to other bacteria. After 72 h of incubation, there was a wide variability among the strains

54

Biofilms in the Food Environment

in the numbers of cells attached to the biofilm, but still no clear trend separating one serotype or source from another. Borucki and others (2003), evaluating 80 strains grown in modified Welshimer’s broth (MWB) using a microtiter plate assay, reported a statistically significant difference in biofilm formation between phylogenetic lineages of L. monocytogenes. This group found that division II strains, which are less commonly associated with food-borne outbreaks, were better biofilm formers. Although the 3a, 1/2c, and 1/2a serotypes had the highest amounts of biofilm, the difference compared to other individual serotypes was not statistically significant. They also found a significantly higher amount of biofilm from persistent strains than sporadic ones. These results support the theory that persistent strains have enhanced biofilmforming capabilities but do not support a consistent relationship between this enhanced biofilm formation and disease incidence. Marsh and others (2003) compared biofilm formation of 3 L. monocytogenes outbreak strains (Scott A, V7, and F2365), and found that the strains varied in biofilm-forming capabilities, which is in agreement with the finding by Borucki and others (2003). However, another comparison of biofilm formation, this time of 31 L. monocytogenes strains, found no difference between persistent and sporadic isolates (Djordjevic and others 2002). In this study, the strains were incubated in MWB with glucose at 32◦ C for 20 and 40 h using a microtiter plate assay. While there was no difference between persistent and sporadic strains in terms of biofilm formation, in this study division I strains produced significantly more biofilm than other strains. While a significant amount of work has been conducted, so far these studies fail to provide overwhelming evidence for a correlation between L. monocytogenes strain, serotype, or source and the ability to form biofilm. Overall, it seems that serotypes less commonly implicated in disease (such as 1/2c) may be better biofilm formers, but the opposite has also been shown. Persistent strains appear to form more biofilm than sporadic strains in most of the studies, but not in every experiment. Differing growth conditions, surfaces, and media may play a role in the various results, since all these environmental conditions can affect L. monocytogenes biofilm formation.

Environmental Factors Affecting L. monocytogenes Biofilm Formation Numerous environmental and stress factors can affect the formation of biofilm by L. monocytogenes, including those that would normally be found in a food processing or storage environment. Temperature, nutrient

Biofilm Development by Listeria monocytogenes

55

levels, and pH have all been shown to affect L. monocytogenes biofilm production. Also, the surface to which the bacteria attach plays a key role. Surface L. monocytogenes has been shown to adhere to a wide variety of surfaces commonly found in food processing environments, including stainless steel, rubber, polymers and plastics, and glass. Beresford and others (2001) evaluated a number of these surfaces by immersing coupons in liquid culture of the L. monocytogenes strain 10403S and determining the number of attached cells. No significant difference in total attached cells at either initial adherence or 2-h adherence was noted for any of the surfaces tested. When the numbers of strongly adhering cells present at the initial adherence were compared to the numbers present after the 2-h incubation, using a wash step to remove loosely adhering cells, a significant increase in strongly attached cells was observed on all the surfaces except polypropylene. Stainless steel, a material commonly found in food processing facilities, has been shown to be one of the best surfaces for L. monocytogenes attachment. Smoot and Pierson (1998) found that the strain Scott A adhered much more rapidly to stainless steel than to Buna-N rubber; in fact, the attachment to the former surface was too rapid to quantify. Schwab and others (2005) also observed that cells immediately attached to stainless steel, even before incubation. Meylheuc and others (2001) found strain LO28 adhered better to stainless steel than to PTFE. Buna-N rubber has been shown to have inhibitory effects on L. monocytogenes biofilm growth, even after a simulated CIP process (Ronner and Wong 1993; Helke and Wong 1994). Blackman and Frank (1996) found that L. monocytogenes biofilm growth on stainless steel, Teflon, nylon, and polyester floor sealant varied with respect to temperature and growth media. With TSB as the growth media, biofilm production was the best among the surfaces on the floor sealant at both 21◦ C and 10◦ C, but in a chemically defined minimal media the sealant did not support any biofilm growth. In all cases, L. monocytogenes formed less biofilm on nylon than on stainless steel or Teflon. Nutrients So far, results from most studies indicate that L. monocytogenes prefers forming a biofilm under relatively high nutrient conditions, unlike many other bacterial species. Takhistov and George (2004) noted that L. monocytogenes biofilm grows toward areas of higher nutrient density. Stepanovic and others (2004) showed that the biofilm quantities produced

56

Biofilms in the Food Environment

by L. monocytogenes strains on plastic were highest in brain-heart infusion and lowest in dilute (1:20) TSB, with nondilute TSB and meat broth falling between these; as a comparison Salmonella species observed in the same study formed much more biofilm in the dilute media. Helke and Wong (1994) showed enhanced survival of L. monocytogenes biofilm in the presence of milk soils than in peptone buffered saline on stainless steel. This is consistent with the finding by Blackman and Frank (1996) that L. monocytogenes in TSB produced much more biofilm than in chemically defined minimal media, despite reaching similar numbers of planktonic cells in both media. The effects of several specific nutrients on L. monocytogenes biofilm formation have also been studied. Kim and Frank (1995) examined the effects of PO4 , amino acids, tryptone, and various carbohydrates in MWB on L. monocytogenes biofilm formation. Biofilm production decreased if the PO4 level was either increased or decreased. A reduction in amino acid levels produced a corresponding decrease in biofilm formation during the first 7 days of incubation, but after 12 days the amount of biofilm was the same regardless of amino acid concentration. The substitution of tryptone for individual amino acids also enhanced biofilm formation during the first 7 days, but again biofilm levels at day 12 were also the same. Among the carbon sources studied, L. monocytogenes in mannose and trehalose produced significantly greater biofilm amounts than glucose, while it produced less in glucosamine. It is worth noting that the contribution of nutrients to biofilm formation may differ from their roles in attachment, reinforcing the distinction between attachment and biofilm growth. For instance Kim and Frank (1994) reported that the 4-h attachment of L. monocytogenes was inhibited by tryptone, while the reduction of PO4 or use of different carbon sources had no effect. Attachment was also reduced in increased ammonia, decreased iron, and the presence of soytone. L. monocytogenes strains also vary in their nutrient requirements for optimal biofilm formation. Moltz and Martin (2005) studied eight strains and found that while two of the strains formed the most biofilm in TSB, the other six produced the most in MWB. Such findings may partly explain the differences in strain-to-strain variability in biofilm formation by L. monocytogenes among different studies with differing experimental conditions. Temperature One of the most studied conditions for L. monocytogenes biofilm formation is temperature. Since this species can grow in a wide range of

Biofilm Development by Listeria monocytogenes

57

temperatures, including many of those found in a food processing or food storage environment, such work may provide insight into the persistence of L. monocytogenes. Unfortunately, the results of these studies are not always consistent. Djordjevic and others (2002) noted an increase in L. monocytogenes biofilm formation at 32◦ C over 20◦ C, with the biofilm levels formed at 20◦ C never reaching the levels at 32◦ C. Duffy and Sheridan (1997), using meat cultures, also observed increased adhesion of L. monocytogenes at 30◦ C compared to both 25◦ C and 37◦ C; however, the total viable count, including the background microflora, was the highest at 25◦ C. In contrast, Herald and Zottola (1988) found that attachment of L. monocytogenes cells was better at 21◦ C than 35◦ C or 10◦ C. Moltz and Martin (2005) also showed that while cell numbers were higher at 20◦ C than 37◦ C after 2 h, after 20 h the 37◦ C biofilm was more extensive. As a psychrotroph, L. monocytogenes has been shown to produce biofilm at refrigeration temperatures as well. Norwood and Gilmour (2001) found that the strains Scott A and FM876 could adhere to stainless steel at 4, 18, and 30◦ C in both single culture and in mixed culture with other species, although the adherence was better at the two higher temperatures. Mafu and others (1990a) observed attachment of L. monocytogenes to a variety of surfaces at room temperature within 20 min and at refrigeration temperatures within 60 min. Bremer and others (2001) noted that L. monocytogenes biofilm survival was enhanced at 4◦ C compared to 15◦ C and −20◦ C in a monoculture, but showed least survival at 4◦ C in a mixed culture with Flavobacterium spp. Moltz and Martin (2005) also found L. monocytogenes capable of producing biofilm at 4◦ C, although in lesser amounts than at 20◦ C or 37◦ C. Helke and Wong (1994) found improved survival of biofilm at 6◦ C over 25◦ C, possibly due to decreased drying at this temperature; they also noted that the biofilm survival was improved at higher temperatures with a higher relative humidity.

Acidity The pH level can also affect L. monocytogenes adherence. Herald and Zottola (1988) saw an increase in attachment to stainless steel at pH 8 compared to pH 5; they also observed that the EPS fibrils produced by the cells at pH 8 were absent at pH 5. Smoot and Pierson (1998) found a lower rate of attachment to Buna-N rubber from cells grown at pH 5.5 than from cells grown at pH 7 or 8.5; however, cells grown at pH 7 showed decreased attachment when placed in a pH 9 environment compared to similar cells placed in neutral and acidic conditions. Conversely, Stopforth and

58

Biofilms in the Food Environment

others (2002) observed no significant difference in attachment and biofilm formation between acid-adapted and nonadapted L. monocytogenes. Overall, results from these studies suggest that environmental factors have significant effects on L. monocytogenes biofilm formation, and the optimal biofilm conditions vary among strains. A better understanding of environmental conditions on L. monocytogenes biofilm formation and identification of biofilm attributes at the molecular level will eventually lead to the identification of essential elements and characterization of the common microbial network involved in microbial defensive mechanisms such as stress responses and biofilm formation.

Molecular Attributes In contrast to the relatively well-established understanding on L. monocytogenes pathogenicity and virulence factors, little is known about molecular attributes and regulatory pathways involved in L. monocytogenes biofilm formation. By comparing the protein expression patterns of planktonic and biofilm cultures using two-dimensional gel elctrophoresis, Tremoulet and others (2002) reported that 22 proteins were upregulated in biofilm and 9 were downregulated. However, few of these could be identified using existing databases. The upregulated proteins included 30S ribosomal proteins, enzymes involved in carbon metabolism, oxidative stress proteins, and regulators of quorum sensing. The only identified downregulated protein was flagellin. Helloin and others (2003) also compared the proteomes of L. monocytogenes in planktonic and biofilm cultures using two-dimensional gel electrophoresis, including samples with and without a carbon source. In biofilm under carbon starvation, 5 of the 14 upregulated proteins were identified, including amino acid and nucleic acid metabolism regulators as well as superoxide dismutase. Nine of the 26 upregulated proteins in biofilm grown with glucose were identified; they included regulators of growth, protein synthesis and repair, and stress response. Two of these proteins were also upregulated in planktonic cells undergoing carbon starvation, suggesting that biofilm formation and response to starvation in L. monocytogenes may be linked. Taylor and others (2002) observed that relA and hpt L. monocytogenes mutants did not grow as well as the wild type after attachment to polystyrene. Both these genes are important for mounting a stringent response during amino acid starvation. Also, relA was transcribed at an increased level in the wild type following attachment to the surface. This study suggests that the ability to mount a response to starvation is important in L. monocytogenes biofilm growth. To examine the potential role of

Biofilm Development by Listeria monocytogenes

59

stress-responsive regulatory network on biofilm formation, Schwab and others (2005) compared the attachment of a L. monocytogenes general stress response sigma factor σ B mutant and its wild-type strain to stainless steel. After cultivating at 24◦ C for 24 h, the numbers of cells attached to the surfaces for both strains were similar; however, the numbers of both attached and planktonic sigB cells were significantly less than the wild type after 48 and 72 h, possibly due to differences in long-term survival. When the incubation temperature was raised to 37◦ C, the wild type attached in significantly greater numbers than at the lower temperature; however, the sigB cell numbers that attached were unchanged. They also observed an increase in attachment for both strains following the addition of salt to the media, suggesting that σ B -independent stress proteins may play a role in attachment. The use of new technologies to analyze gene expression should provide further insights into the cellular processes involved in L. monocytogenes biofilm formation. Quantitative real-time reverse-transcriptase polymerase chain reaction is a highly sensitive and precise way to measure gene transcription, and the development of microarrays can increase analysis throughput. The availability of L. monocytogenes genome sequences, particularly information from comparative genomics of Listeria species and serotypes, will significantly facilitate identification of virulence factors and biofilm determinants (Doumith and others 2004; Nelson and others 2004). Knowledge of the precise mechanisms involved in biofilm formation can lead to novel strategies for its control, and may also aid in the understanding of how L. monocytogenes biofilm production differs from that of other bacteria.

Mixed Culture Biofilms L. monocytogenes is a relatively poor biofilm former when compared to other species of bacteria. Kalmokoff and others (2001) evaluated the 72-h biofilms produced on stainless steel under flow conditions at room temperature by 36 strains of L. monocytogenes and compared them to a variety of Gram-positive and -negative species. While Salmonella enteriditis and Enterococcus faecium formed extensive microcolonies, and Escherichia coli and Pseudomonas aeruginosa produced extensive biofilms, L. monocytogenes showed only sparse attachment. In fact, under the test conditions used only 1 of the 36 L. monocytogenes strains formed a rudimentary biofilm. The other 35 strains showed only sparse attachment with few clusters of cells. Stepanovic and others (2004) also reported that L. monocytogenes produced much less biofilm than Salmonella species

60

Biofilms in the Food Environment

under static media conditions, although all of the 48 strains tested did form biofilm. However, monoculture biofilms are rarely found in natural environment. Microorganisms can become part of a microbial ecosystem by acting as primary surface colonizers, or as later biofilm partners through establishing interaction with other organisms (Kolenbrander 2000). As noted earlier, L. monocytogenes is capable of integrating itself into EPS and biofilm formed by other bacteria (Sasahara and Zottola 1993; Hassan and others 2004), and such mixed culture biofilms are likely the dominant forms of L. monocytogenes persistent in the natural environment. To reveal the contribution of other abundant microbes in the food system on L. monocytogenes survival and persistence, the interactions between L. monocytogenes and several food-borne bacteria, including the general microflora from food processing facilities, have been investigated. Pseudomonas spp. are common spoilage organisms, particularly at refrigeration temperatures, and are widely distributed in foods (Jay 2003). Hassan and others (2004) examined L. monocytogenes and Pseudomonas putida in forming a biofilm in condensate on stainless steel at 12◦ C and compared this with a L. monocytogenes monoculture. The P. putida, isolated from a food processing plant, was allowed to form a biofilm on stainless steel coupons in dilute TSB for 48 h. After this, a cocktail of five environmental strains of L. monocytogenes was introduced to the preexisting biofilm and incubated for an additional 48 h, then compared to the L. monocytogenes monoculture biofilm. L. monocytogenes attached to the surface in significantly greater numbers (>3-log difference) with the P. putida biofilm than without. Despite a lack of nutrients both organisms survived in the condensate for 35 days, although no growth was evident; the higher initial attachment of L. monocytogenes in the mixed culture also led to higher cell numbers at 35 days than in monoculture. When protein was added along with the condensate, L. monocytogenes survival was enhanced in both the mixed and monocultures. Both species could also readily detach from the surface while in the biofilm state, allowing for potential contamination in a food processing environment. A mixed culture of L. monocytogenes with a different Pseudomonad, Pseudomonas fragi, has also been studied (Sasahara and Zottola 1993). In this case, biofilm was formed on glass coverslips under either a continuous flow system or agitation in beakers using TSB with 0.6% yeast extract (TSBYE) as growth medium. In monoculture, L. monocytogenes attached only sparingly in the agitation vessel and not at all in the continuous flow conditions. It adhered better in mixed culture, forming microcolonies around the preattached P. fragi cells. It appeared there was a substance, possibly

Biofilm Development by Listeria monocytogenes

61

the P. fragi EPS, that attracted and entrapped the L. monocytogenes cells into the P. fragi biofilm. Schwab and others (2005) also combined L. monocytogenes and Pseudomonas fluorescens cultures to form biofilm on stainless steel coupons. While the cell numbers of the wild-type L. monocytogenes and the sigB mutant strain in planktonic culture were not significantly different in mixed culture compared to monoculture, both attached to the stainless steel surface in significantly greater numbers in the mixed cultures. The P. fluorescens appeared to promote the attachment of L. monocytogenes cells by entrapping them in its biofilm. Bremer and others (2001) combined seven L. monocytogenes strains with two species of Flavobacterium, isolated from a seafood plant, and compared biofilm formation on stainless steel at different temperatures. This was done under flow conditions, after the initial biofilm formation, to remove unattached cells. Sparse growth of single cells or short chains of cells were visible in the L. monocytogenes monoculture, while the mixed culture showed extensive, multilayered biofilm. In the presence of L. monocytogenes, however, the Flavobacterium rapidly died off, going from >108 cells/cm2 initially to no cells by day 10. The L. monocytogenes numbers also decreased, but the decline was much less in the mixed culture than in the monoculture. This was attributed to the EPS produced by the “primary colonizer” Flavobacterium that likely protected L. monocytogenes cells from desiccation. Not all bacteria enhance L. monocytogenes biofilm, however. Leriche and Carpentier (2000) inoculated L. monocytogenes to 1-day-old Staphylococcus sciuri biofilm and compared the biofilm formation by L. monocytogenes monoculture. Staph. sciuri inhibited L. monocytogenes growth in the biofilm, since L. monocytogenes exhibited a higher proportion of the cells in mixed culture in the planktonic phase than it did in the biofilm. The results suggested that the extracellular substances produced by Staph. sciuri biofilms are involved in the decreased adhesion of L. monocytogenes in the biofilm. This property was evident in all three media tested, TSB-YE with glucose (TSB-YEg), dilute TSB-YEg, and whey. Leriche and others (1999) also studied L. monocytogenes biofilm growth in association with a nisin-producing strain of Lactococcus lactis. In this study, L. monocytogenes either was combined with Lc. lactis and inoculated onto stainless steel coupons or was inoculated onto a preexisting Lc. lactis biofilm. The antilisterial activity of Lc. lactis in the simultaneous inoculation began within the first 6 h of incubation, reducing the L. monocytogenes numbers to undetectable levels when a 106 CFU/mL L. monocytogenes inoculum was used. When 108 CFU/mL L. monocytogenes was used for inoculation, the cell numbers were reduced approximately

62

Biofilms in the Food Environment

2 logs and then stabilized for the duration of the experiment (48 h). Similar results were observed when L. monocytogenes was inoculated onto a preexisting Lc. lactis biofilm. Planktonic L. monocytogenes cells were more sensitive to the nisin than sessile cells, demonstrating the enhanced resistance of biofilm cells toward adverse environments. Also, replacement of the media with fresh TSB-YE at 22 or 24 h in an attempt to further stimulate nisin production had no effect on L. monocytogenes numbers, suggesting that the surviving cells had become resistant to the bacteriocin. Norwood and Gilmour (2000) combined L. monocytogenes with P. fragi and Staphylococcus xylosus in a mixed culture biofilm. After achieving a steady state for the three organisms in a chemostat, the mixed culture was inoculated into a constant-depth film fermenter to maintain a consistent thickness of the biofilm on the stainless steel surface. They used dilute TSB (2 g/L) supplied at 30 mL/h at 21◦ C, and allowed the biofilm to grow for 28 days. While in the chemostat, L. monocytogenes numbers fell sharply from day 3 to day 6, at which time steady-state levels of the organisms were achieved with L. monocytogenes, making up only 1.8% of the total population. This percentage was unchanged during the incubation in the constant-depth film fermenter , with P. fragi becoming the dominant organism in the biofilm. They also observed that the biofilm cells continued to produce EPS up until day 17. Other studies have examined L. monocytogenes biofilm formation in conjunction with naturally occurring background microflora. Stopforth and others (2002) used an inoculum from meat decontamination washings with a L. monocytogenes strain isolated from meat to study the interaction in a biofilm cultivated on stainless steel coupons in TSB-YE. The background microflora, which was 2.0 × 102 CFU/mL in the initial inoculum, had achieved attachment numbers of 1.3 × 106 CFU/cm2 by day 2 and increased to 2.5 × 106 CFU/cm2 by day 4; these numbers then remained largely unchanged through day 14. The L. monocytogenes, on the other hand, attached in far fewer numbers (1.6 × 105 –3.2 × 105 CFU/cm2 by day 2) and decreased in number to 1.0 × 104 –3.4 × 104 CFU/cm2 by day 4; this number also remained largely unchanged through day 14. It appears that the natural microflora restricted the growth of L. monocytogenes in these settings. Jeong and Frank (1994) evaluated L. monocytogenes biofilm growth in the presence of eight organisms isolated from meat and dairy processing environments. This was done by inoculating a mixture of L. monocytogenes culture and test organism culture on stainless steel coupons in dilute TSB at 10◦ C and comparing the results with a L. monocytogenes monoculture. Initial attachment (during the first 5 days) of L. monocytogenes varied depending on the competitive culture; but none of the species enhanced

Biofilm Development by Listeria monocytogenes

63

L. monocytogenes attachment. Two isolates, a Flavobacterium species and a Pseudomonas species, allowed L. monocytogenes cell numbers to reach the levels of the monoculture in 1% TSB. Another two isolates, a different Pseudomonas and a Streptococcus species, allowed over a 100fold increase in L. monocytogenes growth in the biofilm, but both had substantially inhibited its initial attachment such that the final numbers never equaled those in monoculture. A Bacillus species was overall the most inhibitory to L. monocytogenes biofilm growth. In all cases, however, L. monocytogenes was able to grow in the mixed culture biofilm, although the numbers often did not increase until after day 9. Also, L. monocytogenes was able to survive throughout the 25 days of the study in all the cultures. In another study, Carpentier and Chassaing (2004) examined the effects of 29 individual species of bacteria isolated from various food processing facilities on L. monocytogenes biofilm formation. These resident species were allowed to attach to stainless steel coupons in TSB for 3 h before L. monocytogenes culture was inoculated. The resulting biofilms were then compared to L. monocytogenes monoculture biofilm. Sixteen of the strains had a negative effect on the L. monocytogenes biofilm growth, with 3 (P. fluorescens, a Bacillus species, and an unidentified Gram-positive rod) causing a 3-log reduction in the L. monocytogenes count. Only 4 strains (Kocuria varians, Staphylococcus capitis, Stenotrophomonas maltophilia, and Comamonas testosteroni) enhanced L. monocytogenes numbers, with a 0.5–1.0-log increase over the monoculture. The other 11 strains had no significant effect on L. monocytogenes growth. There was no link between EPS production by a given strain and its effect on the L. monocytogenes count, suggesting that perhaps the nature of the EPS produced, rather than the quantity, is the key factor. Overall, results from these studies suggest that the specific resident microflora in a food processing facility play an important role in determining the likelihood of L. monocytogenes establishment and becoming persistent in the environment. While some bacteria seem to enhance L. monocytogenes biofilm growth and others exhibit antagonistic effects, it is worth noting that except in the presence of nisin-producing bacteria (Leriche and others 1999), L. monocytogenes cells were all capable of survival in mixed culture biofilms in these studies, including up to 35 days in a nutrient-free condensate (Hassan and others 2004). Furthermore, although L. monocytogenes may be a minority in the initial ecosystem, it has the potential to outcompete other dominating organisms in foods in stress conditions such as refrigeration. Therefore the above studies highlight the challenge L. monocytogenes can pose once it becomes established in a food processing environment.

64

Biofilms in the Food Environment

Prevention and Control of L. monocytogenes Biofilm Improving the overall sanitation and preventing L. monocytogenes biofilm from establishing itself in a food processing environment, therefore, is critical to minimize its contamination incidence from the food supply. However, L. monocytogenes is notorious for maintaining itself in inaccessible, hard to reach places, particularly in biofilms. Numerous studies have therefore been conducted to find ways to inhibit L. monocytogenes adhesion to surfaces and to evaluate the effectiveness of sanitizers on L. monocytogenes biofilm once it becomes established. One approach is to use competitive bacteria to exclude L. monocytogenes. As was previously discussed, nisin-producing bacteria can reduce or even eliminate L. monocytogenes biofilms (Leriche and others 1999). Similar results were obtained when several microbial isolates from commercial food processing facility drains were tested against L. monocytogenes (Zhao and others 2004). In this study, 24 strains were assessed for antilisterial activities on tryptic soy agar (TSA). These strains were combined individually with L. monocytogenes to form a biofilm on stainless steel coupons in TSB at various temperatures. Nine of the 24 isolates produced substantial inhibition to L. monocytogenes growth (4–5log reduction compared to controls) and were identified belonging to Enterococcus durans, Lc. lactis subsp. lactis, and Lactobacillus plantarum. Particularly, one each of the E. durans and Lc. lactis strains were highly inhibitory to L. monocytogenes due to the production of enterocin and nisin, respectively. The Lc. lactis strain even retained its antilisterial activity at 4◦ C, although it did not grow at this temperature. The authors concluded that these 2 strains might be well suited for use as competitive exclusion microorganisms to prevent L. monocytogenes biofilm formation. Adsorption of nisin directly onto a surface has also been shown to reduce L. monocytogenes colonization (Bower and others 1995). In this study, nisin was adsorbed onto both hydrophilic and hydrophobic silica surfaces and the silica placed in L. monocytogenes culture. The surfaces were examined at 0, 4, 8, and 12 h of incubation for viable cells using iodonitrotetrazolium, which forms red crystals when exposed to cellular respiration. While 95% of the cells remained viable in nisin-free control samples, less than 20% of the adhered cells had visible crystals on the nisinadsorbed surfaces. There were also fewer total adhered cells on the nisintreated surfaces, and the cells that did attach failed to grow or reproduce by 12 h. Meylheuc and others (2001) found similar effects when using P. fluorescens biosurfactants. Pretreatment of stainless steel with purified,

Biofilm Development by Listeria monocytogenes

65

anionic P. fluorescens biosurfactant caused a 92% decrease in culturable L. monocytogenes when compared to nontreated surfaces. This decrease was not observed when PTFE was used as the biofilm surface, indicating that the surface characteristics of PTFE were not as dramatically changed by the biosurfactants as was stainless steel. Al-Makhlafi and others (1994) studied the effects of the milk proteins β-lactoglobulin, α-lactalbumin, β-casein, and bovine serum albumin (BSA) adsorbed onto silica surfaces on L. monocytogenes adhesion. On hydrophobic silica, all the proteins inhibited L. monocytogenes attachment to varying degrees compared to the bare surface, while on hydrophilic surfaces all the proteins except BSA enhanced attachment compared to the bare surface. On both types of silica surfaces, BSA showed the most profound inhibition, well below that of either bare surface. Later work (Al-Makhlafi and others 1995) showed that the order of adsorption of the proteins is important, since the addition of BSA to the surface after β-lactoglobulin, rather than before or concurrently, did not produce this inhibition. Many of the commonly used sanitizing agents in the food processing industry have been studied for their effects on surfaces contaminated with L. monocytogenes. Mafu and others (1990b) exposed stainless steel, glass, polypropylene, and nitrile rubber to L. monocytogenes culture for 15 min, and then treated them with various concentrations of sodium hypochlorite,two different iodophores, and a quaternary ammonia compound (QAC). They found that higher concentrations were needed to inactivate the L. monocytogenes on the porous polypropylene and rubber surfaces, and that higher concentrations of hypochlorite and QAC were needed than either of the iodophores. Also, higher concentrations were needed at 4◦ C compared to 20◦ C. While the short incubation time in this study likely did not result in a true biofilm, it does give some indication of how attached cells respond to sanitizers. Ronner and Wong (1993) also observed that L. monocytogenes biofilm grown on the porous Buna-N rubber was more resistant to sanitizers than biofilm grown on stainless steel, and attributed this at least in part to the slower growth on the rubber. They used QAC, iodine, chlorine, and an anionic acid to treat 2-day biofilm and planktonic cultures, with greatest effect on the planktonic culture (7–8-log reduction) and the least effect in biofilm on the rubber (1–2-log reduction), with biofilm on stainless steel being intermediate (3–5-log reduction). They also noted that the chlorine and the anionic acid generally removed EPS better than the other sanitizers. Chlorine has also been evaluated in studies on L. monocytogenes mixed culture biofilms. Bremer and others (2002) used different concentrations

66

Biofilms in the Food Environment

in conjunction with differing surfaces, contact time, and pH to inactivate a L. monocytogenes and Flavobacterium species mixed culture biofilm. They noted improved effectiveness on a stainless steel surface over a conveyor belt material, likely due to the cells growing within the weave of the belt. Adjusting the pH to 6.5 also improved the efficacy of the chlorine. Norwood and Glimour (2000) found that in a 28-day biofilm of L. monocytogenes, P. fragi, and Staph. xylosus, 1,000 ppm chlorine for 20 min was needed to produce a statistically significant reduction in cell numbers. In contrast, 10 ppm for 30 sec inactivated all cells in planktonic culture. The effectiveness of chlorine, peracetic acid, and peroctanoic acid on a L. monocytogenes and Pseudomonas species mixed-culture biofilm was evaluated (Fatemi and Frank 1999). The 4-h attachment and 48-h biofilm samples grown on stainless steel, in both skim milk and TSB, were used. The peracid sanitizers were much more effective in the presence of skim milk than was the chlorine, which has little activity in the presence of organic material. Further, peroctanoic acid was more effective than peracetic acid under these conditions. Oh and Marshal (1996) used monolaurin and acetic acids, both separately and in combination, to treat L. monocytogenes biofilm. They found that either compound alone did not effectively inactivate the biofilm, but that a combination of both acids was able to completely inactivate a 1-day old 105 CFU/cm2 biofilm. However, after 7 days of growth, the efficiency of the combined reagents decreased once the L. monocytogenes population reaches 106 CFU/cm2 or higher. As the biofilm matured, it became resistant to these sanitizing agents. Stopforth and others (2002) found similar results using the beef decontamination washings. The effectiveness of the sanitizers used—sodium hypochlorite, QAC, and peracetic acid—varied with the age of the biofilm. In general, the L. monocytogenes biofilm was more resistant to the sanitizers at day 7 than it was at day 2; however, by day 14 it had become sensitized to them. In contrast, the biofilm from the overall microflora increased in resistance to the sanitizers throughout the 14 days of the study. Overall, they found that peracetic acid took the shortest time to inactivate the biofilms and was more effective in killing attached cells than planktonic ones, although the biofilm cells tended to be more resistant to the sanitizers than suspended cells. Chavant and others (2004) also compared how planktonic and sessile cells respond to different sanitizers, in this case using a L. monocytogenes monoculture in stationary and exponential planktonic growth as well as in 6-h, 1-day, and 7-day biofilms on stainless steel. They also observed that while QAC inactivated 98% or more of cells in most cases,

Biofilm Development by Listeria monocytogenes

67

after 7 days of biofilm growth this mortality rate dropped to around 40%. Acetic acid (pH 5) had little effect under any conditions, although this improved slightly with the addition of Na2 SO4 ; monolauric acid was also only partially effective. The use of NaOH to raise the pH to 12, with or without Na2 SO4 , was completely effective, except in the very early stages of biofilm development (6-h growth) where it produced a mortality rate of less than 80%. Arizcun and others (1998) demonstrated the effectiveness of a high pH on inactivating L. monocytogenes biofilm. They tested 16 decontamination protocols on a 3-day biofilm, using different sanitizers and temperatures. The most effective protocols applied 63◦ C heat for 30 min, lowering the cell count by more than 5.5 log to 0.22 CFU/cm2 ; however, such treatment is impractical in many food processing settings. The use of NaOH with a pH 10.5 followed or proceeded by acetic acid at pH 5.4, each at 55◦ C for as little as 5 min, provided similar results. Treatments at 20◦ C, or at 55◦ C without the use of NaOH, had little effect. This enhanced antibacterial effect of NaOH may be due to its ability to dissolve EPS, destroying its ability to protect the cells within. A recent study looked at the efficacy of two cleaning and sanitizing combinations on L. monocytogenes biofilm, rather than focusing on only the sanitizing step (Somers and Wong 2004). Combination A contained a chlorinated alkaline, low phosphate detergent and a dual peracid sanitizer; combination B contained a solvated alkaline cleaner and hypochlorite sanitizer. The researchers used a variety of surfaces to grow the L. monocytogenes biofilm, compared the efficacy of the protocols on two types of stainless steel, three conveyor belt materials, two rubber surfaces, a brick wall, and flooring, and found that biofilms were more resistant on the brick and conveyor materials. They also found that while both detergents reduced biofilm cells and material, the hypochlorite sanitizer was more effective at further reducing biofilm numbers than was the peracid sanitizer. While the use of competitive exclusion organisms and the adsorption of proteins to a food processing surface may hold promise for the future, currently cleaning and sanitation are the main methods of biofilm prevention and control. The studies cited highlight the importance of routine enforcement of such activities, since L. monocytogenes has been shown to become more resistant to sanitizers over time. The use of heat and a combination of acid and alkaline cleaners and sanitizers appear to be effective approaches of reducing established biofilms. Attention should also be paid to the surface that is being cleaned, since porous surfaces seem to enhance the resistance of L. monocytogenes to sanitizers.

68

Biofilms in the Food Environment

References Al-Makhlafi H, Nasir A, McGuire J, Daeschel MA. 1994. Influence of preadsorbed milk proteins on adhesion of Listeria monocytogenes to hydrophobic and hydrophilic silica surfaces. Appl Environ Microbiol 60(10):3560–3565. Al-Makhlafi H, Nasir A, McGuire J, Daeschel MA. 1995. Adhesion of Listeria monocytogenes to silica surfaces after sequential and competitive adsorption of bovine serum albumin and β-lactoglobulin. Appl Environ Microbiol 61(5):2013–2015. Arizcun C, Vasseur C, Labadie J. 1998. Effect of several decontamination procedures on Listeria monocytogenes growing in biofilms. J Food Prot 61(6):731–734. Beresford MR, Andrew PW, Shama G. 2001. Listeria monocytogenes adheres to many materials found in food-processing environments. J Appl Microbiol 90(6):1000– 1005. Blackman IC, Frank JF. 1996. Growth of Listeria monocytogenes as a biofilm on various food processing surfaces. J Food Prot 59(8):827–831. Borucki MK, Peppin JD, White D, Loge F, Call DR. 2003. Variation in biofilm formation among strains of Listeria monocytogenes. Appl Environ Microbiol 69(12):7336–7342. Bower CK, McGuire J, Daeschel MA. 1995. Suppression of Listeria monocytogenes colonization following adsorption of nisin onto silica surfaces. Appl Environ Microbiol 61(3):992–997. Bremer PJ, Monk I, Butler R. 2002. Inactivation of Listeria monocytogenes/Flavobacterium spp. biofilms using chlorine: Impact of substrate, pH, time and concentration. Lett Appl Microbiol 35(4):321–325. Bremer PJ, Monk I, Osborne CM. 2001. Survival of Listeria monocytogenes attached to stainless steel surfaces in the presence or absence of Flavobacterium spp. J Food Prot 64(9):1369–1376. Briandet RT, Meylheuc C, Maher MN, Bellon-Fontaine. 1999. Listeria monocytogenes Scott A: Cell surface charge, hydrophobicity, and electron donor and acceptor characteristics under different environmental growth conditions. Appl Environ Microbiol 65(12):5328– 5333. Carpentier B, Chassaing D. 2004. Interactions in biofilms between Listeria monocytogenes and resident microorganisms from food industry premises. Int J Food Microbiol 97(2):111–122. Chae MS, Schraft H. 2000. Comparative evaluation of adhesion and biofilm formation of different Listeria monocytogenes strains. Int J Food Microbiol 62(1–2): 103–111. Chavant P, Gaillard-Martinie B, Hebraud M. 2004. Antimicrobial effects of sanitizers against planktonic and sessile Listeria monocytogenes cells according to the growth phase. FEMS Microbiol Lett 236(2):241–248. Chavant P, Martinie B, Meylheuc T, Bellon-Fontaine MN, Hebraud M. 2002. Listeria monocytogenes LO28: Surface physicochemical properties and ability to form biofilms at different temperatures and growth phases. Appl Environ Microbiol 68(2):728– 737. Djordjevic D, Wiedmann M, McLandsborough LA. 2002. Microtiter plate assay for assessment of Listeria monocytogenes biofilm formation. Appl Environ Microbiol 68(6):2950– 2958. Doumith M, Cazalet C, Simoes N, Frangeul L, Jacquet C, Kunst F, Martin P, Cossart P, Glaser P, Buchrieser C. 2004. New aspects regarding evolution and virulence of Listeria monocytogenes revealed by comparative genomics and DNA arrays. Infect Immun 72(2):1072– 1083.

Biofilm Development by Listeria monocytogenes

69

Duffy G, Sheridan JJ. 1997. The effect of temperature, pH and medium in a surface adhesion immunofluorescence technique for detection of Listeria monocytogenes. J Appl Microbiol 83(1):95–101. Dussurget O, Pizarro-Cerda J, Cossart P. 2004. Molecular determinants of Listeria monocytogenes virulence. Annu Rev Microbiol 58:587–610. Fatemi P, Frank JF. 1999. Inactivation of Listeria monocytogenes/Pseudomonas biofilms by peracid sanitizers. J Food Prot 62(7):761–765. Hassan AN, Birt DM, Frank JF. 2004. Behavior of Listeria monocytogenes in a Pseudomonas putida biofilm on a condensate-forming surface. J Food Prot 67(2):322–327. Helke DM, Wong ACL. 1994. Survival and growth characteristics of Listeria monocytogenes and Salmonella typhimurium on stainless steel and buna-n rubber. J Food Prot 57(11):963–968. Helloin E, Jansch L, Phan-Thanh L. 2003. Carbon starvation of Listeria monocytogenes in planktonic state and in biofilm: A proteomic study. Proteomics 3(10):2052–2064. Herald PJ, Zottola EA. 1988. Attachment of Listeria monocytogenes to stainless steel surfaces at various temperatures and pH values. J Food Sci 53(5):1549–1552. Jay JM. 2003. Taxonomy, role and significance of microorganisms in food. In Modern Food Microbiology, 6th Edition. Aspen Publishers, Gaithersburg, MD, p. 23. Jeong DK, Frank JF. 1994. Growth of Listeria monocytogenes at 10◦ C in biofilms with microorganisms isolated from meat and dairy processing environments. J Food Prot 57(7):576–586. Kalmokoff ML, Austin JW, Wan, XD, Sanders G, Banerjee S, Farber JM. 2001. Adsorption, attachment and biofilm formation among isolates of Listeria monocytogenes using model conditions. J Appl Microbiol 91(4):725–734. Kathariou S. 2002. Listeria monocytogenes virulence and pathogenicity, a food safety perspective. J Food Prot 65(11):1811–1829. Kim KY, Frank JF. 1994. Effect of growth nutrients on attachment of Listeria monocytogenes to stainless steel. J Food Prot 57(8):720–724. Kim KY, Frank JF. 1995. Effect of growth nutrients on biofilm formation by Listeria monocytogenes on stainless steel. J Food Prot 58(1):24–28. Kolenbrander PE. 2000. Oral microbial communities: Biofilms, interactions, and genetic systems. Annu Rev Microbiol 54:413–437. Leriche V, Carpentier B. 2000. Limitation of adhesion and growth of Listeria monocytogenes on stainless steel surfaces by Staphylococcus sciuri biofilms. J Appl Microbiol 88(4):594– 605. Leriche V, Chassaing D, Carpentier B. 1999. Behaviour of Listeria monocytogenes in an artificially made biofilm of a nisin-producing strain of Lactococcus lactis. Int J Food Microbiol 51(2–3):169–182. Lunden JM, Miettinen MK, Autio TJ, Korkeala HJ. 2000. Persistent Listeria monocytogenes strains show enhanced adherence to food contact surfaces after short contact times. J Food Prot 63(9):1204–1207. Mafu AA, Roy D, Goulet J, Magny P. 1990a. Attachment of Listeria monocytogenes to stainless steel, glass, polypropylene, and rubber surfaces after short contact times. J Food Prot 53(9):742–746. Mafu AA, Roy D, Goulet J, Savoie L, Roy R. 1990b. Efficacy of sanitizing agents for destroying Listeria monocytogenes on contaminated surfaces. J Dairy Sci 73(12): 3428–3432. Marsh EJ, Luo H, Wang H. 2003. A three-tiered approach to differentiate Listeria monocytogenes biofilm-forming abilities. FEMS Microbiol Lett 228(2):1969–1973.

70

Biofilms in the Food Environment

Meylheuc T, vanOss CJ, Bellon-Fontaine MN. 2001. Adsorption of biosurfactant on solid surfaces and consequences regarding the bioadhesion of Listeria monocytogenes LO28. J Appl Microbiol 91(5):822–832. Moltz AG, Martin SE. 2005. Formation of biofilms by Listeria monocytogenes under various growth conditions. J Food Prot 68(1):92–97. Nelson KE, Fouts DE, Mongodin EF, Ravel J, DeBoy RT, Kolonay JF, Rasko DA, Angiuoli SV, Gill SR, Paulsen IT, Peterson J, White O, Nelson WC, Nierman W, Beanan MJ, Brinkac LM, Daugherty SC, Dodson RJ, Durkin AS, Madupu R, Haft DH, Selengut J, Van Aken S, Khouri H, Fedorova N, Forberger H, Tran B, Kathariou S, Wonderling LD, Uhlich GA, Bayles DO, Luchansky JB, Fraser CM. 2004. Whole genome comparisons of serotype 4b and 1/2a strains of the food-borne pathogen Listeria monocytogenes reveal new insights into the core genome components of this species. Nucleic Acids Res 32(8): 2386–2395. Norwood DE, Gilmour A. 1999. Adherence of Listeria monocytogenes strains to stainless steel coupons. J Appl Microbiol 86(4):576–582. Norwood DE, Gilmour A. 2000. The growth and resistance to sodium hypochlorite of Listeria monocytogenes in a steady-state multispecies biofilm. J Appl Microbiol 88(3):512– 520. Norwood DE, Gilmour A. 2001. The differential adherence capabilities of two Listeria monocytogenes strains in monoculture and multispecies biofilms as a function of temperature. Lett Appl Microbiol 33(4):320–324. Oh DH, Marshall DL. 1996. Monolaurin and acetic acid inactivation of Listeria monocytogenes attached to stainless steel. J Food Prot 59(3):249–252. Ronner AB, Wong ACL. 1993. Biofilm development and sanitizer inactivation of Listeria monocytogenes and Salmonella typhimurium on stainless steel and Buna-n rubber. J Food Prot 56(9):750–758. Sasahara K, Zottola E. 1993. Biofilm formation by Listeria monocytogenes utilizes a primary colonizing microorganism in flowing systems. J Food Prot 56(12):1022–1028. Schwab U, Hu Y, Wiedmann M, Boor K. 2005. Alternative sigma factor σ B is not essential for Listeria monocytogenes surface attachment. J Food Prot 68(2):311–317. Smoot LM, Pierson MD. 1998. Influence of environmental stress on the kinetics and strength of attachment of Listeria monocytogenes Scott A to Buna-N rubber and stainless steel. J Food Prot 61(10):1286–1292. Somers EB, Wong, AC. 2004. Efficacy of two cleaning and sanitizing combinations on Listeria monocytogenes biofilms formed at low temperature on a variety of materials in the presence of ready-to-eat meat residue. J Food Prot 67(10):2218–2229. Stepanovic S, Cirkovic I, Ranin L, Svabic-Vlahovic M. 2004. Biofilm formation by Salmonella spp. and Listeria monocytogenes on plastic surface. Lett Appl Microbiol 38(5): 428–432. Stopforth JD, Samelis J, Sofos JN, Kendall PA, and Smith GC. 2002. Biofilm formation by acid-adapted and non-adapted Listeria monocytogenes in fresh beef decontamination washings and its subsequent inactivation with sanitizers. J Food Prot 65(11):1717–1727. Takhistov P, George B. 2004. Linearized kinetic model of Listeria monocytogenes biofilm growth. Bioprocess Biosyst Eng 26(4):259–270. Taylor CA, Beresford M, Epton HAS, Sigee DC, Shama G, Andrew PW, Roberts IS. 2002. Listeria monocytogenes relA and hpt mutants are impaired in surface-attached growth and virulence. J Bacteriol 184(3):621–628. Tompkin RB. 2002. Control of Listeria monocytogenes in the food-processing environment. J Food Prot 65(4):709–725.

Biofilm Development by Listeria monocytogenes

71

Tremoulet F, Duche O, Namane A, Martinie B, The European Listeria Genome Consortium Labadie JC. 2002. Comparison of protein patterns of Listeria monocytogenes grown in biofilm or in planktonic mode by proteomic analysis. FEMS Microbiol Lett 210(1):25–31. Vatanyoopaisarn S, Nazli A, Dodd CER, Reed CED, Waites WM. 2000. Effect of flagella on initial attachment of Listeria monocytogenes to stainless steel. Appl Environ Microbiol 66(2):860–863. Yousef AE, Carlstrom C. 2003. Listeria monocytogenes. In Food Microbiology. John Wiley & Sons, Inc., Hoboken, NJ, pp. 138–139. Zhao T, Doyle MP, Zhao P. 2004. Control of Listeria monocytogenes in a biofilm by competitive-exclusion microorganisms. Appl Environ Microbiol 70(7):3996–4003.

Biofilms in the Food Environment Edited by Hans P. Blaschek, Hua H. Wang, Meredith E. Agle Copyright © 2007 by Blackwell Publishing and the Institute of Food Technologists

Chapter 4 INACTIVATION OF LISTERIA MONOCYTOGENES BIOFILMS USING CHEMICAL SANITIZERS AND HEAT Revis A.N. Chmielewski and Joseph F. Frank

Introduction Control of Listeria monocytogenes is a challenge for many food processing plants, especially those producing ready-to-eat products that are exposed to the environment after heat processing. L. monocytogenes can attach to and grow on environmental and food contact surfaces, forming biofilms. Listeria cells that survive the cleaning/sanitizing process may be attached to or growing on surfaces. Listeria can form biofilms at points in the system that are not adequately exposed to cleaning chemicals, either because of poor equipment design or maintenance or because the surface is not targeted for cleaning. Biofilms containing Listeria, usually in combination with other microflora, contain extracellular polymers (EPS) that act as a biological glue anchoring the cells to the surface and protecting them from removal and inactivation. Chemical cleaners must be able to dissolve this EPS for cells to be removed. If the EPS is not dissolved then chemical sanitizers must penetrate residual EPS to inactivate underlying cells. The presence of food residues, along with biofilm EPS, makes effective cleaning/sanitizing more difficult. Microorganisms within biofilm are more resistant to both heat and biocides than are suspended cells (LeChevallier and others 1988; Frank and Koffi 1990; Lee and Frank 1991). The mechanism of increased heat resistance of Listeria in biofilms is not known. The underlying cause of many Listeria control failures is lack of effective cleaning. Ineffective cleaning allows growth of biofilms, whereas 73

74

Biofilms in the Food Environment

effective cleaning removes protective organic matter from the surface, allowing sanitizing chemicals to work. Food soils are often complex blends of carbohydrate, protein, and fats along with other ingredients (salts, spices, etc.) that may have been exposed to heat or acids. Interactions between food components can make cleaning more difficult. For example, studies of milk soil removal showed a linear reduction of fat residues with increased temperature of the detergent, but this effect stopped at 65◦ C in the presence of protein soil (Dunsmore 1981). If food soils are not completely removed, they accumulate over time, entrap microorganisms, and lead to biofilms that cannot be eliminated by chemical sanitizer application. Sanitation is most effective in inactivating attached or biofilm cells when applied after thorough cleaning (Dunsmore and Thomson 1981; Krysinski and others 1992; Frank and others 2003). Because of the importance of the cleaning step as a prerequisite to effective biofilm control, this chapter will provide background information on cleaning effectiveness and then discuss chemical and heat sanitation as means to control Listeria in biofilms left behind by ineffective cleaning.

Cleaning and Biofilm Control Cleaning is the primary means of biofilm control in food plants. Environmental surfaces that are likely to harbor biofilms (floor drains, equipment mounting brackets, the undersides of equipment) are those that are difficult to clean. Soft, moderately worn surfaces, such as gaskets and conveyer belts, and surfaces of corroded metal may be difficult or not possible to clean, and therefore become microbial growth niches. Cleaning Compounds Cleaning compounds are designed to remove specific soils such as protein, fats, carbohydrate, and mineral deposits. Cleaning compounds widely used in the food industry were not specifically designed to remove biofilm polymers. Cleaners function to remove soil by decreasing surface tension of water so that soil can be dislodged or loosened; suspending soil particle in an emulsion by allowing the cleaning compound to surround the soil to form a micelle; and by preventing resuspension of the soil. The factors affecting cleaning efficacy include contact time; physical force (laminar or turbulent flow); concentration of the cleaning compound; temperature of the cleaning fluid; water chemistry; cleanup workers’ skill, type of cleanup equipment used, composition of the soil; and type of surface.

Inactivation of Listeria monocytogenes Biofilms

75

If the cleaning solution does not contact the soiled surface for the appropriate amount of time, at the appropriate temperature, or with the appropriate amount of physical force, then effective cleaning will not occur. Cleaning compounds contain chelators (sequestrants) that bind minerals and decrease water hardness. Cleaning compounds also contain agents that emulsify, saponify, and disperse lipids; hydrolyze protein; and reduce surface tension. Various types of cleaners and their effectiveness in biofilm removal are presented in Table 4.1. Alkali cleaners are generally effective for food soil and biofilm removal. Alkali cleaners applied under static conditions (foam or gel, no high pressure or scrubbing) are able to remove 99% fat and 93% protein from stainless steel surfaces within 30 min of application, and remove 7 log CFU/50 cm2 of Listeria biofilm cells within 10 min (Frank and others 2003). A neutral cleaning agent applied under similar conditions was effective at removing fat (99% in 30 min) but not as effective at removing protein (77% in 30 min) from stainless steel (Table 4.1). The neutral cleaning agent was also effective at removing biofilms from stainless steel but not in the presence of protein (Frank and others 2003). The cleaning method employed and microflora population may also influence cleaning ability. Gibson and others (1999) found that alkali and neutral cleaners were not as effective as acid cleaners for the removal of Staphylococcus and Pseudomonas biofilms. The alkali and neutral cleaners were able to remove 3–5-log biofilm cells whereas the acid cleaner had a 6-log reduction. The acid cleaner may have removed the biofilm by mineral solubilization. Wirtanen and others (1996) demonstrated that alkali cleaner with EDTA was the most effective CIP (clean in place) treatment to remove and inactivate Bacillus biofilm (Wirtanen and others 1996). Chelating agents are considered important in the removal of biofilm by destabilizing the polysaccharide matrix. Chen and Stewart (2000) observed the detachment of biofilm cells after treatment with chelating agents. Cleaning that removes biofilm cells, but leaves EPS residual, may not be sufficiently effective for food industry applications, since the residual EPS serves as an attractant for soil and bacteria. Antoniou and Frank (2005) found that commercial alkali cleaning compound applied at cold or room temperature without scrubbing or other physical force was effective at removing biofilm EPS, whereas hot 1.28% alkali (no additives) did not completely remove the EPS when applied with agitation. The best explanation for this discrepancy is the presence of chelating agents in the commercial cleaner. They also reported that the bacterial cells within the biofilm were more easily removed from the surface by alkali treatment than were the biofilm EPS.

Hot alkali cleaner, 2%, 85◦ C 2% cold alkali cleaner

Alkali cleaners

Alkali detergent (1.6–10%), Alkali peroxide (3%), detergent + ClO2 , anionic detergent (1.6%)

5% alkali cleaner, pH 11.6, 20-min exposure

Hot alkali cleaner, 1.2–6.0%, 68–70◦ C

Concentration >95% reduction in 4-day-old biofilm for both hot and cold water CIP cleaning on [PTFE (Teflon); Buna-N and Vitron rubber] gasket materials, and 70–85% biofilm reduction in cold CIP and 80–95% reduction in hot CIP for EPDM rubber gaskets 1.2% alkali was ineffective in removal of Pseudomonas biofilms. However, concentration of 2.0–6.0% was effective in removal of biofilms from stainless steel In laboratory tests, there was a 3–5-log reduction of Staphylococcus aureus and Pseudomonas aeruginosa biofilms on stainless steel In field tests, there was an approximately 1-log reduction in microbial load On stainless steel surfaces there was an approximately 4-log cells/cm2 reduction while on polyester/polyurethane surfaces there was a 4 log units Reduction of L. monocytogenes and Listeria innocua was by >6 log units in the presence of tryptic soy broth and 6-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil 0.5% of EO water 4-log reduction of E. coli O157:H7 in a and 2% suspension. When combined inoculum level is 1 × anode/cathode 105 ,there is only a 2-log water at OPR >848 mV reduction 10 ppm, ORP >8-log reduction of 1123 mV, L. monocytogenes and pH 2.5 E. coli in suspension. 3-log reduction of B. cereus in vegetative cells after 30 sec of treatment and 8-log reduction in L. monocytogenes, E. coli, and B. cereus and a 3-log reduction in spores after 120 sec of treatment 7-log reduction of L. monocytogenes and E. coli 1.5–2-log reduction of L. monocytogenes, E. coli respectively on lettuce 4-log reduction of Salmonella after 7 days storage at 4◦ C 1-log reduction of L. monocytogenes and E. coli O157:H7 respectively on raw salmon 2-log reduction of L. monocytogenes, E. coli O157:H7, and Salmonella on eggs, fruits, and vegetables

Kim and others (2000)

2 ppm (mg/L), ORP 915 mV, pH 2.6 150 ppm, ice

20–50 ppm, ORP 795 mV, pH 2.6 70–90 ppm, ORP 1150 mV, pH 2.6, 64 min

18– 64 ppm, ORP 1083–1150 mV, pH 2.6, 15–64 min

Ozone

Ozone

85

14.3 ppm (mg/L), Reduction of E. coli 3 min O157:H7 attached to alfalfa seeds and sprouts by 1 log CFU/g Ozone + UV 6-log reduction of E. coli in poultry processing water 0.065–1.0 µg/mL, 0.5–6-log reduction of 0.5 min E. coli 4 ppm, 3 min 7.47-log reduction of L. monocytogenes 0.198–0.188 ppm, 3-log reduction of 1–5 min Salmonella and 4-log reduction of L. monocytogenes

Park and others (2004) Koseki and others (2004)

Fabrizio and others (2002) Ozer and Demirci (2005)

Sharma and Demirci (2003), Stan and Daeschel (2003), Bialka and others (2004), Koseki and others (2004), Wang, and others (2004) Singh and others (2003)

Diaz and Law (1997)

Khadre and others (2001) Robbins and others (2005) Restaino and others (1995)

(continued )

86

Biofilms in the Food Environment

Table 4.2. Summary of Research on the Effectiveness of Chemical Sanitizers for Inactivating Biofilm Cells (Continued ) Concentration Ozone + UV

Iodine

Peroxygen Peroxygen

Effectiveness

Ozone (0.77 ppm) 6-log reduction of E. coli in + UV (4,500 poultry processing water µW/cm2 ) 25 ppm, 10-min 4-log CFU/cm2 reduction of L. monocytogenes exposure biofilms on stainless steel surfaces 25 ppm, 10-min 2-log cells/cm2 reduction of exposure L. monocytogenes on stainless steel and 1-log reduction on polyester surfaces but 6-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil

Rossoni and Gaylarde (2000)

180 ppm peracetic acid for 10 min (tray test method) 80–160 ppm, 5 min

Peracetic acid: acetic acid (8%), hydrogen peroxide (27.5%), and peroxyacetic acid (5.8%) Peracetic acid + octanoic acid: acetic acid (24%), hydrogen peroxide (5–20%), peroxyacetic acid (1–5%), and octanoic acid (1–5% Surfactant sanitizers Cationics: auaternary ammonium

2.0 mL/L, 10 min

Makela and others (1991) Fatemi and Frank (1999)

Frank and others (2003)

1.3 mL/L, 10 min

>5-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil

Frank and others (2003)

100, 200, 400, and 800 ppm, 30-sec exposure

2–3-log CFU/cm2 reduction of L. monocytogenes biofilms on stainless steel after 30-sec exposure and 4–5 log after 20 min

Frank and Koffi (1990)

(continued )

88

Biofilms in the Food Environment

Table 4.2. Summary of Research on the Effectiveness of Chemical Sanitizers for Inactivating Biofilm Cells (Continued )

Quaternary ammonium

Concentration

Effectiveness

Reference

200 ppm, 1-min exposure

3-log CFU/cm2 reduction of L. monocytogenes biofilms on stainless steel 6-log reduction of attached L. monocytogenes cells in the presence of milk soil >4-log CFU/mL reduction of Listeria biofilm on stainless steel Reduction of attached lactic acid bacteria was by >4 log units or 99.99% 4–5-log reduction of attached L. monocytogenes attached to chitin 2–5-log reduction of attached Staph. aureus to various surfaces >5-log reduction of L. monocytogenes biofilm on stainless steel in the presence of soil 4-log cells/cm2 reduction of L. monocytogenes on stainless steel but 848 mV must be maintained.

Inactivation of Listeria monocytogenes Biofilms

93

Storage of EO water beyond 95 h resulted in a decrease in ORP (Stevenson and others 2004). In water or cell suspension test, acidic EO water at 10 ppm, ORP 1123 mV, pH 2.5 was effective in inactivating L. monocytogenes and E. coli in suspension by >8 log units and Bacillus cereus vegetative cells by 3 log units after 30 sec of treatment. There was 8-log reduction in L. monocytogenes, E. coli, and B. cereus and a 3-log reduction in spores after 120 sec of treatment (Kim and others 2000). Ice generated from acidic EO water at 150 ppm residual chlorine reduced L. monocytogenes and E. coli on lettuce by 1.5 and 2 log units, respectively. However, ice with 240 ppm residual chlorine caused significant damage to the lettuce leaves (Koseki and others 2004). Immersion of broiler poultry in EO water (pH 2.6, 20–50 ppm chlorine, and ORP 795 mV) for 45 min was able to reduce Salmonella attached to poultry skin by 4 log units after 7 days of storage at 4◦ C (Fabrizio and others 2002). The immersion of raw salmon in EO water showed about a 1-log reduction of L. monocytogenes and E .coli O157:H7 after 64 min of treatment (Ozer and Demirci 2005). The disinfection of eggs, fruits, and vegetables with acidic EO water showed minimal inactivation of 2-log reduction of L. monocytogenes, E .coli O157:H7, and Salmonella (Sharma and Demirci 2003; Singh and others 2003; Stan and Daeschel 2003; Bialka and others 2004; Koseki and others 2004; Wang and others 2004). The studies summarized in Table 4.3 demonstrate that EO water as a disinfectant shows marginal result in the inactivation of microorganisms attached to foods. Acidic EO water seemed to be more effective in inactivating microorganisms than alkali EO water (Fabrizio and others 2002). Sequential use of alkali then acidic EO water does not seem to be more effective than EO water alone (Stevenson and others 2004). However, direct application of EO water (pH 2.6, ORP 1160 mV, 56 mg/L residual chlorine, for 300 sec) to biofilm on stainless steel successfully reduced Listeria cells in the biofilm by about 9 log units (Kim and others 2001). One of the primary disadvantages of EO water is that it requires fairly pure water to get an effective ORP for microbial inactivation (Stevenson and others 2004). Ozone Ozone is an effective antimicrobial for water treatment systems. An attractive aspect of ozone is that it decomposes leaving no chemical residue. The efficacy of ozone as a sanitizer in the food industry is mixed. Khadre and others (2001) summarized ozone inactivation studies on spoilage and pathogenic microflora in foods. He reported that Gram positives and negatives exposed in 0.066–1.0 µg/mL ozone for 0.5–10 min produced

94

Biofilms in the Food Environment

0.5–6-log CFU/mL reduction. Ozonation studies of water and water in the presence of organic matter showed that ozone treatment of 0.198 ppm for 1–5 min gave a >3-log reduction for Salmonella and a 4-log reduction for L. monocytogenes (Restaino and others 1995) and ozone treatment of 0.2–1.8 ppm, pH 5.9 for 30 sec gave a 0.7–7-log reduction for L. monocytogenes (Kim and Yousef 2000). Ozone treatment of alfalfa sprouts show a 4-log reduction of L. monocytogenes biofilm and a 3-log reduction of Salmonella typhimurium biofilm on stainless steel, but a 6-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil with application of 2.0 mL/L of peracetic acid solution. Similarly, 1.3 mL/L of peroctanoic/peracetic acid solution exposed for 10 min produced a >5-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil (Frank, and others, 2003). At cold temperature (4◦ C), an effective cleaning regiment in removing Listeria biofilm in the presence of fat and protein soil involved cleaning with alkali (10 min) followed by sanitizing with 1.3 mL/L of peracetic/peroctanoic acid solution exposed for 10 min. This produced a 5.3-log reduction of L. monocytogenes biofilm (Frank, and others, 2003). Acid and Anionic Sanitizers Lactic and acetic acids are used directly as sanitizing agents or in sanitizer formulations. Arizcun and others (1998) observed that sequential treatment with acetic acid and lactic acid at 55◦ C reduced Listeria biofilm by about 4 log units, and sequential treatment with acetic acid and sodium hydroxide at 55◦ C reduced Listeria biofilm by 5.7 log units with a 10-min contact time. Ammor and others (2004) demonstrated that a sanitizing solution of monolaurin (0.075%) with acetic acid

96

Biofilms in the Food Environment

(pH 5.4) reduced pathogens and spoilage organisms such as Staphylococcus, Pseudomonas, and Listeria by 3–4 log units while allowing the maintenance of desirable microflora in the meat fermentation environment. Heat (65◦ C) plus monolaurin reduced Listeria biofilm by >6 log units (Oh and Marshall, 1995). Anionic acid sanitizer at 200 ppm with 2-min exposure produced a 3–4-log reduction of L. monocytogenes biofilms on stainless steel surfaces (Ronner and Wong, 1993). Acid anionic sanitizer at 0.4% with 10-min exposure produced a 4-log reduction of L. monocytogenes on stainless steel and on polyester surfaces but only a 1-log reduction on polyester/polyurethane conveyer belt material (Krysinski and others 1992). Orthophosphoric acid anionic sanitizer at 200 and 400 ppm reduced L. monocytogenes biofilm on stainless steel by 2–3 log units after 30-sec exposure and by about 4 log CFU/cm2 after 20 min (Frank and Koffi, 1990). Fatty Acid Sanitizers Medium chain fatty acids can have useful bactericidal activity. A 10-min exposure of 0.04% C8 –C10 solution reduced L. monocytogenes on stainless steel by 4 log units, by 2 log units on polyester surfaces, and by 6-log reduction of Listeria in the presence of milk serum using the carrier test method for the efficacy study of QAC. Other Sanitizers Dunsmore and others (1980) demonstrated that in a CIP system with milk soil, Pseud. aeroginosa, M. luteus, and Strep. faecalis were resistant to 200 mg/L chlorhexidene sanitizer showing 3 log or less reduction but E. coli and Enter. aerogenes were more sensitive, exhibiting a >5-log reduction (Dunsmore and others 1980). Best and others (1990) demonstrated using the carrier test method that 4% chlorheximide was effective in inactivating L. monocytogenes in the presence of 2% milk soil, and a study of dental water systems showed 0.2% chlorheximide inactivated cells of a mixed culture biofilm but removed only 31% of the mixed cells from dental silicone and polyurethane tubing (Walker and others 2003). The efficacy of trisodium phosphate in the inactivation of microbial biofilm was mixed, showing a 1–3-log CFU/cm2 reduction of L. monocytogenes biofilm on stainless steel and a 3-log reduction of Campylobacter and Salm. typhimurium biofilms (Somers and others 1994). Research on QAC (and other sanitizers), before 1990, primarily used suspension or carrier test methodology. The carrier test evaluates the sanitizer on a microbial suspension dried onto a surface. Microorganisms in biofilms are generally many times more resistant to sanitizers than indicated by the carrier test, because the carrier test does not allow formation of protective EPS.

Hot Water Sanitation It may be appropriate to use heat to inactivate biofilm when chemical sanitizing is not effective, and cooking of organic residues is not a concern.

98

Biofilms in the Food Environment

Also, some equipment is designed to be sanitized by hot water or steam. When chemical sanitizers are not effective at controlling Listeria, hot water sanitation is used on specific pieces of equipment in CIP or COP (clean out of place) cleaning. However, if food residues, especially those containing protein, cook onto a surface, they become much more difficult to remove, and ordinary cleaning processes are not likely to work. In addition, biofilm EPS will not be removed from a surface by hot water sanitation. Since surfaces with food or biofilm residues are attractive to microorganisms, unclean surfaces sanitized using hot water may become rapidly recontaminated when processing proceeds. Little is known about this potential problem. Hot water sanitation is successfully used in dairy, brewery, winery, and other food processing plants for specific applications. Hot water sanitation does not leave chemical residues or have a significant waste disposal cost (Agius and others 2004). Effective hot water sanitation requires proper maintenance of water chemistry especially water hardness, iron, manganese, nitrate/nitrite, hydrogen sulfide, and pH. It requires safety training of personnel. The recommended temperature for hot water sanitation is between 68 and 85◦ C depending on whether the application is CIP, COP, or recirculation (Stanfield 2003). In CIP systems, 77◦ C for 5 min is generally recommended (Stanfield 2003) since temperatures above 77◦ C may cause cavitation (rattling of pipes). Data from hot water sanitation studies show that heat treatments (70–82◦ C, 5-min exposure) can reduce L. monocytogenes biofilm by about 4 log units (Frank and Koffi 1990; ; Lee and Frank 1991; Krysinski and others 1992) but may leave surviving cells. Wirtanen found that hot water (90◦ C for 15 min) and super hot water (125◦ C for 30 min) rinses were effective at removing 3-day Bacillus biofilms (Wirtanen and others 1996). Since there has been little research on heat treatments required to inactivate L. monocytogenes biofilms (Table 4.3), we initiated a project to develop a predictive model for heat inactivation of L. monocytogenes in biofilms.

Modeling Heat Inactivation of Biofilms Most microbial heat inactivation predictive models use decimal reduction time (D-values) to model death kinetics. D-values are widely used owing to the simplicity of the concept. The major assumption behind D-value models, however, is that microbial death follows a log-linear reduction with time of heating. Models based on D-values must often ignore nonlinear portions of the curve called shoulders and tails. More sophisticated methods of analyzing nonlinear data may use the modified Gompertz model,

Inactivation of Listeria monocytogenes Biofilms

99

which includes the shoulder, linear reduction, and tail portions of an inactivation curve in the predictive equation. Heat inactivation of cells within a biofilm exhibits tailing because the cells are clumped within the polysaccharide matrix. Removing the cells from the surface and breaking up the clumps reduces the heat resistance of the cells and therefore would not provide an accurate model (Frank and Koffi 1990). In addition, the most useful part of heat inactivation is at the end of the curve where there are very low numbers, because these are the survivors that can affect product safety and shelf life if the heat treatment is inadequate. Fraction negative data overcome these concerns, because data are obtained from enrichment culture of the heated system, allowing the determination of low numbers of survivors. This data-gathering method, while less precise than plate count techniques, avoids the errors associated with cell clumping and does not require the detachment of surviving cells from the surface. Fraction negative modeling is based on a binomial response where a fraction of samples in each treatment show either growth or no growth. This approach provides for the determination of heat treatments based upon complete inactivation of the target population, thus eliminating tailing effects. In addition, fraction negative modeling does not assume log-linear inactivation. Since L. monocytogenes often occurs in food plants in mixed species biofilms with food residues, we developed models to predict the heat inactivation of L. monocytogenes in monoculture biofilms (strains Scott A and 3990) and in biofilms with competing bacteria (a Pseudomonas sp. and Pantoea agglomerans) formed on stainless steel and Buna-N rubber in the presence of food-derived soil. Biofilms were produced on stainless steel coupons and rubber disks using 1/10 diluted tryptic soy broth with incubation for 48 h at 25◦ C. Duplicate biofilm samples were heat-treated for 1, 3, 5, and 15 min at 70, 72, 75, 77, and 80◦ C and tested for survivors using enrichment culture. The experiment was repeated six times. Predictive models for each surface were developed using logistic regression analysis of the fraction negative data: 1. Prediction model for L. monocytogenes biofilms on stainless steel. ⎡ ⎤ −0.4706∗ Scott A ⎦ Ln[P/(1 − P )] = 18.0527 + ⎣ −0.5316∗ LM 3990 −0.0616∗ multispecies 0.2299∗ TEMP − 0.1108∗ TIME 2. Validation model of stainless steel: Ln[P/(1 − P )] = 35.9399 − 0.4655∗ TEMP − 0.1892∗ TIME

100

Biofilms in the Food Environment

3. Prediction model for L. monocytogenes biofilms on Buna-N rubber: ⎤ ⎡ ∗  Scott A 0.4092 ∗   0.3718 Soil Ln P/ (P − 1) = 19.8811+ + ⎣ 0.1245∗ LM 3990 ⎦ 0.000∗ Soil −0.5337∗ Mixed −0.2981∗ TEMP − 0.1122∗ TIME 4. Validation model of Buna-N rubber: Ln[P/(P − 1)] = 21.1155 − 0.3175∗ TEMP − 0.1317∗ TIME Each predictive equation estimates the probability of inactivation of L. monocytogenes in monoculture (Scott A and 3990) and in multispecies biofilms after heat treatment under various time–temperature conditions in the presence of soil on the respective surface. Examples of predictions based on these models are presented in Table 4.4. The predictive models were validated using a five-strain cocktail of L. monocytogenes in the presence of food soil (Chmielewski and Frank 2004). The validation equations indicate that the model predictions are conservative. The predictive models demonstrate that hot water sanitation of stainless steel and rubber surfaces can be effective in inactivating L. monocytogenes in a biofilm on stainless steel if time and temperature are Table 4.4. Probability of Complete Inactivation of L. monocytogenes Biofilm Cells Subject To Various Heat Treatments as Estimated by Fraction Negative Models for Various Surface, Microflora, and Soil Conditions Culture

Surface

Soil

Scott A 3990 Multispecies Scott A 3990 Multispecies Scott A 3990 Multispecies Scott A 3990 Multispecies Scott A 3990 Multispecies

Stainless steel Stainless steel Stainless steel Stainless steel Stainless steel Stainless steel Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber Buna-N rubber

Food soil Food soil Food soil Food soil Food soil Food soil Food soil Food soil Food soil Food soil Food soil Food soil No soil No soil No soil

Temperature (◦ C)

Time (min)

Inactivation probability (%)

80 80 80 76 76 76 80 80 80 76 76 76 76 76 76

2.5 11.7 6.3 11.0 >15 15 9