Biofilms in the Food Environment (Institute of Food Technologists Series)

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

Storgards, and others (1999b)

>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 <2-log reduction

Krysinski and others (1992)

Gibson and others (1999)

Antoniou and Frank (2005)

Reference

Effectiveness

Summary of Research on the Effectiveness of Cleaning Agents in Removing Biofilms

Types

Table 4.1.

Alkali with EDTA

Short CIP: 1% alkali, 3.5–4.6 EDTA Long CIP: 1% alkali, 2% EDTA 70◦ C, 7–20 min

Mild alkali cleaner (foam)

Sodium hydroxide (10%), tallow(bishydroxyethyl)amine oxide (5%), diethylene glycol methyl ether (3%), propylene glycol monomethyl ether (3%), and dipropylene glycol methyl ether (3%) Strong alkali cleaner (foam)

Bredholt and others (1999), Antoniou and Frank (2005),

Strong alkali foam cleaner was effective in reducing 95% of biofilm cells (Pseudomas, Bacillus, and L. monocytogenes) and reduce Listeria and other mixed cultures by at least 2 log units Mild alkali cleaner was less effective, with about 50% biofilm removal and about 1–2-log cfu/cm2 reduction in the presence of soil Short CIP cleaning was not effective in reducing Bacillus biofilm cells, showing 10–30% remaining Long CIP was effective in reducing 95% of biofilm cells

(continued )

Wirtanen and others (1996)

Frank and others (2003)

99% of fat, 93% of protein, and 7-log cfu/cm2 biofilm cells were removed from stainless steel surfaces

Concentration

2.5% acidic cleaner, pH 1.7, 20-min exposure Enzyme–detergent blend

Acid cleaner

Enzyme-based cleaners

Chlorinated alkali

Chlorinated cleaner

In laboratory tests 3–5-log cfu/cm2 reduction of Staph. aureus and Pseud. aeruginosa On stainless steel and polyester surfaces there was a 4-log cells/cm2 reduction while on polyester/polyurethane surfaces there was a <1-log reduction On stainless steel surfaces there was a 4-log cells/cm2 reduction while on polyester/polyurethane surfaces there was a <1-log reduction In laboratory tests 6-log reduction of Staph. aureus and 4–5-log reduction of Pseud. aeruginosa On stainless steel surfaces there were an approximately 4-log cells/cm2 reduction while on polyester/polyurethane surfaces there was a <2-log reduction

Effectiveness

Summary of Research on the Effectiveness of Cleaning Agents in Removing Biofilms (Continued )

Other cleaner formulations Neutral cleaner 1%, pH 8.3, 20-min exposure Anionic cleaner Anionic detergent (1.6%)

Types

Table 4.1.

Gibson and others (1999) Krysinski and others (1992)

Krysinski and others (1992)

Frank and others (2003) Krysinski and others (1992)

Reference

Inactivation of Listeria monocytogenes Biofilms

79

Cleaning Agent Concentration A sufficient concentration of cleaning agent must be applied to achieve a soil- and biofilm-free surface. However, increasing concentrations do not lead to increased soil removal if a saturation point has been attained ( Jackson 1985). Sodium hydroxide (5 N) inactivated or removed 93% of Listeria biofilm cells (Chavant and others 2004), whereas in a model system, 2.5% NaOH (66◦ C) was more effective at removing Pseudomonas biofilm EPS than the 1.28% recommended by the Model Pasteurized Milk Ordinance (Antoniou and Frank 2005). Likewise, an increase in temperature of the alkali cleaners increased the removal of biofilm. Method of Cleaning In some systems, high flow velocity or turbulent force is required for soil removal. Dunsmore and others (1981) showed that turbulent force was necessary for detergent cleaning and sanitation to be effective in removing dairy soil. Storgards and others (1999a) found that there were higher levels of biofilm remaining on rubber surfaces after low-velocity (0.8 m/sec) CIP cleaning compared to high-velocity (2.0 m/sec ) cleaning. High-pressure wash is effective in removal of microbial biofilms but creates aerosols and disperses microorganisms (Gibson and others 1999). Mechanical scrubbing is also effective at removing biofilms. Static application is the application of chemical cleaners without the use of physical force. Examples include the low-pressure application of foam or gel (viscous liquid) cleaners. Foam and gel cleaners contain emollients and stabilizers that keep the cleaning agent in contact with the surface and reduce dripping. In general, foam cleaners tend to drip more over time compared to gel cleaners. The effectiveness of foam cleaners was demonstrated in a study by Bredholt and others (1999) who observed greater than 95% decrease in biofilms in the presence of food soil. The efficacy of gel alkali cleaner was demonstrated by Frank and others (2003) who showed removal of 7 log CFU of Listeria biofilm cells within 10 min (Table 4.3). Ineffective use of these cleaning agents is usually the result of the agent not contacting the soiled surface for sufficient time. This is why gels or viscous liquids that do not drip from vertical surfaces are preferred in some applications to foam cleaners. Temperature and Contact Time The recommended temperature for detergent cleaning of food contact surfaces ranges from 40 to 90◦ C (Jackson 1985). There is a linear effect on

80

Biofilms in the Food Environment

cleaner effectiveness with increasing heat except for protein soil, which coagulates at 60◦ C (Dunsmore 1981). The removal of Listeria biofilm in the presence of soil increases with increasing time (Frank and others 2003). The CIP cleaning study of Wirtanen and others (1996) demonstrated that CIP treatment using alkali cleaners with EDTA at 70◦ C for 20 min was more effective at removing Bacillus biofilm than were shorter treatments. Surface Type The ease of biofilm removal depends to some extent on the type of surface being cleaned. For example, stainless steel surfaces are easier to clean than polyester/polyurethane ones (Holah and Thorne 1990; Krysinski and others 1992). When selecting surfaces such as stainless steel, surface roughness measurements that include an indicator of surface defects (microscopic scratches, pits, and burrs) were observed to be a better indicator of cleanability than the surface finish designation (Frank and Chmielewski 2001). Studies by Storgards and others (1999b) showed that repeated cleaning of rubber surfaces in hot alkali CIP cleaner caused cracks and deterioration and that subsequent biofilm formation became more difficult to remove. Various cleaning studies (Dunsmore and others 1981; Holah and Thorne 1990) support the conclusion that abraded surfaces are more difficult to clean and sanitize. Krysinski and others (1992) concluded that alkali, anionic, enzyme, alkali quaternary ammonium, and detergent plus chlorine dioxide cleaners effectively removed L. monocytogenes biofilm on stainless steel, while for polyester/polyurethane surfaces, chlorinated alkali and alkali peroxide were more effective than the other detergents and auxiliary cleaners. The Role of Water in Cleaning Water is used as a prerinse for removing large soil particles, for wetting of soil and softening, for suspension and transporting cleaning compound, for suspension and transport of soil to be removed, and for rinsing of cleaning compound and transport of sanitizer to the cleaned area. Water may be soft (free of minerals) or hard (containing minerals such as calcium, magnesium ions, and sodium bicarbonate). Hard water interferes with the chelating action of cleaning compounds, thereby limiting the performance of the cleaning compound. Although no research has been reported on the importance of water in cleaning biofilms, the expectation is that hard water would reduce the solubilization of minerals from the biofilm matrix, making it more difficult to destabilize and dissolve the EPS matrix.

Inactivation of Listeria monocytogenes Biofilms

81

Sanitizing Agents and Biofilm Inactivation Chemical sanitizers are approved based on their ability to reduce populations of Escherichia coli and Salmonella by 5 log units in 30 sec. Regulatory approval for food industry use does not require that the chemical be evaluated for inactivation of biofilm cells. Sanitizing chemicals do not inactivate microorganisms by attacking a specific molecular site, but have nonspecific activity, most commonly from general oxidation or disruption of the cell membrane and denaturation of cytoplasmic proteins, or reducing the internal pH of the cell. The effectiveness of a sanitizing chemical depends on exposure time, temperature, concentration, pH, presence of organic matter, water chemistry, and bacterial load. Some desirable properties in a sanitizer include having broad spectrum of activity: activity in the presence of organic matter, surfactant properties, lack of toxicity, good solubility in water, and ease of use. Chemical sanitizers can be categorized as oxidative (halogen, peroxides, ozone) acid, alkali, and phenolic. Research studies on the effectiveness of chemical sanitizing agents for inactivating biofilms are summarized in Table 4.2. Halogens Halogens include chlorine or chlorine-releasing agents. The recommended application is usually 200 ppm for 2-min exposure on nonporous surfaces and 800 ppm for porous surfaces. Chlorine can reduce numbers of Listeria within biofilms if the organic load is low, but some survivors are often found. To have an effective chlorine treatment the concentration, exposure time, and pH of the solution must be controlled. Ronner and Wong (1993) observed that chlorine and anionic sanitizers were more effective in inactivating L. monocytogenes and Salmonella biofilm cells than were iodine and quaternary ammonia compound. They also observed survival of L. monocytogenes biofilm after exposure to 200 ppm chlorine for 1 min. Lee and Frank (1991) demonstrated that an exposure time of 5 min at 200 ppm was required to inactivate Listeria biofilm by approximately 5 log units, and Bremer and others (2002) reported that a chlorine concentration of 200 ppm with exposure for greater than 2 min was required to obtain >4-log reduction of Listeria biofilm. Surface type can influence the efficacy of chlorine against biofilms (Ronner and Wong 1993). In a study by Ronner and Wang (1993), stainless steel surfaces were sanitized more effectively than were Buna-N, Vitron rubber, and PVC/polyester surfaces. Bal’a and others (1999) sanitized stainless steel surfaces with 75 ppm chlorine solutions for 1 min, demonstrating a 4-log CFU/surface reduction of Aeromonas hydrophila biofilm. Rossoni and Gaylarde (2000)

82

Biofilms in the Food Environment

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

Halogens Chlorine

Concentration

Effectiveness

Reference

100 ppm, 2-min exposure

4-log CFU/cm2 reduction of L. monocytogenes biofilms on stainless steel surfaces 1–2-log reduction on Buna-N rubber >4-logCFU/mL reduction of Listeria biofilm on stainless steel surfaces Reduction of Listeria biofilm was by greater than 99% (2 log units) on stainless steel surfaces Reduction of Listeria biofilm was by 92–99% (1–2 log units) on PVC/polyester conveyer belt material Reduction of dairy biofilms was by 0.6 log unit after 10-min exposure and >4 log units after 2-h exposure Reduction of atttached lactic acid bacteria was >4 log units Reduction of L. monocytogenes and Listeria innocua was by >6 log units in the presence of tryptic soy broth and <2-log reduction in the presence of milk Reduction of <2-log of L. monocytogenes attached to cabbage and lettuce

Ronner and Wong (1993)

200 ppm for greater than 2 min

200–600 ppm for 2–20 min with and without pH adjustment (6.5)

200 ppm for 10 min and 2–24 h

170–200 ppm for 10 min (tray test method) 60 ppm, 1 min

200 ppm, 10 min

Mustapha and Liewen (1989)

Bremer and others (2002)

Dufour and others (2004)

Makela and others (1991) Best and others (1990)

Zhang and Farber (1996)

83

Inactivation of Listeria monocytogenes Biofilms Table 4.2.

(Continued ) Concentration

Effectiveness

Reference

200 ppm, 10 sec

Reduction of 4-log of L. monocytogenes attached to brussel sprouts 4-log CFU/surface reduction of A. hydrophila biofilm on stainless steel 4-log CFU/cm2 reduction of Salmonella biofilm formed on cement and plastic surfaces 2-log CFU/mm2 reduction of E. coli, Pseud. fluorescens, and Staph. aureus attached to stainless steel 6-log CFU/cm2 reduction of L. monocytogenes and Psuedomonas attached to stainless steel 3–5-log reduction of L. monocytogenes on chitin after 5-min exposure at 100 ppm 2-log reduction of Staph. aureus on glass 4-log cells/cm2 reduction of L. monocytogenes on stainless steel and polyester surfaces but <1-log reduction on polyester/polyurethane conveyer belt material 2–3-log CFU/spotted site reduction of attached L. monocytogenes to apple pulp skin 3.5–6.5-log reduction of attached L. monocytogenes to apple pulp skin

Brackett (1987)

75 ppm, 1 min

100 ppm for 10 min

100–200 ppm, 10 min

80–160 ppm, 5 min

100–800 ppm, 2–20 min

Chlorine dioxide

300 ppm, 5-min exposure 5 ppm, 10 min

4 ppm, 10-min exposure

4 ppm, 30-min exposure

Bal’a and others (1999)

Joseph and others (2001)

Rossoni and Gaylarde (2000)

Fatemi and (1999)

Frank

McCarthy (1992)

Luppens and others (2002) Krysinski and others (1992)

Du and others (2002)

(continued )

84

Biofilms in the Food Environment

Table 4.2. Summary of Research on the Effectiveness of Chemical Sanitizers for Inactivating Biofilm Cells (Continued ) Concentration 5 ppm, 10 min

Sodium hypochlorite

Acidified sodium hypochlorite

Electrolyzed water

Effectiveness

Reduction of L. monocytogenes attached to freshly cut vegetables by 1 log CFU/g 0.30 ppm for 20 h Reduction of E. coli at 4 and 20◦ C attached to apples by approximately 5 log units 25 ppm (mg/L), Reduction of E. coli 3 min O157:H7 attached to alfalfa seeds and sprouts by 1 log CFU/g 200–500 ppm, Reduction of E. coli 3–10 min O157:H7 attached to alfalfa seeds and sprouts by >2 log CFU/g 200 ppm, 10 min >4-log reduction of L. monocytogenes biofilm on stainless steel in the presence of fat and protein soil 150 ppm, 10 min >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 <1-log reduction in Bacillus spores

Reference Zhang and Farber (1996)

Sapers and others (2003)

Singh and others (2003)

Taormina and Beuchat (1999)

Frank and others (2003)

Frank and others (2003)

Stevenson and others (2004)

Kim and others (2000)

Inactivation of Listeria monocytogenes Biofilms Table 4.2.

(Continued ) Concentration

Effectiveness

Reference

56 ppm, ORP 1160 mV, pH 2.6

>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 <1-log reduction on polyester/polyurethane conveyer belt material Iodophor 80 ppm <1-log reduction of attached L. monocytogenes on (0.008%) (carrier stainless steel surfaces in test method) the presence of milk Iodine, 25 ppm 5-log reduction of Pseud. aeroginosa, E. coli, and M. luteus, Strep. faecalis, and Enter. aerogenes in milk soil Iodine, 25 ppm 3-log reduction of attached L. monocytogenes attached to chitin

Reference Diaz and Law (1997) Ronner and Wong (1993)

Krysinski and others (1992)

Best and others (1990)

Dunsmore and others (1980)

McCarthy (1992)

4-log cells/cm2 reduction of Krysinski et al. (1992) L. monocytogenes on stainless steel and polyester surfaces but <1-log reduction on polyester/polyurethane conveyer belt material Knowles and 1.2 ppm peracetic >3-, 2.9-, 0.8-, and 1.2-log Roller (2001) reduction of acid plus 160 L. monocytogenes, Salm. ppm hydrogen typhimurium, Staph. peroxide aureus, Sacch. cerevisiae respectively attached to stainless steel 105 ppm, 10-min exposure

Inactivation of Listeria monocytogenes Biofilms Table 4.2.

87

(Continued ) Concentration

Effectiveness

Reference

250–1,000 ppm, 10 min

1 log or less CFU/mm2 reduction of E. coli, Pseud. fluorescens, and Staph. aureus attached to stainless steel Reduction of atttached lactic acid bacteria was by >4 log units or 99.99% 7-log CFU/cm2 reduction of L. monocytogenes and Psuedomonas attached to stainless steel >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 <2-log reduction on polyester surfaces and polyester/polyurethane conveyer belt material 4-log cells/cm2 reduction of L. monocytogenes on stainless steel and polyester surfaces but <2-log reduction on polyester/polyurethane conveyer belt material

Ronner and Wong (1993)

100 ppm, 1-min exposure

200 ppm for at least 1 min

140–230 ppm for 10 min (tray test method) 100 ppm, 10 min

200 ppm, 35 sec

200 ppm, 10 min

Neutral QAC, pH 200 ppm, 10-min exposure 6.4 Acidic QAC, pH 2.4

Acidic quats + ClO2

200 ppm quats + 5 ppm

Best and others (1990)

Mustapha and Liewen (1989) Makela and others (1991) McCarthy (1992)

Frank and Chmielewski (1997) Frank, and others (2003)

Krysinski and others (1992)

Krysinski and others (1992)

Inactivation of Listeria monocytogenes Biofilms Table 4.2.

89

(Continued ) Concentration

Acids and anionics Anionic acid 200 ppm, 2-min exposure

Effectiveness

3–4-log CFU/cm2 reduction of L. monocytogenes biofilms on stainless steel Acid anionics 0.4%, 10-min 4-log cells/cm2 reduction exposure of L. monocytogenes on stainless steel and on polyester surfaces but only 1-log reduction on polyester/polyurethane conveyer belt material Orthophosphoric 200 and 400 ppm, 2–3-log CFU/cm2 acid 30-sec to reduction of 20-min L. monocytogenes exposure biofilms on stainless steelafter 30-sec exposure and about 4-log reduction after 20 min Acids Acetic acid (pH 4-log CFU/cm2 reduction 5.4, 76.7 mM) of L. monocytogenes and lactic acid on glass slides (pH 5.4, 84.4 mM) at 55◦ C 5.7-log CFU/cm2 Acetic acid (pH 5.4, 76.7 mM) reduction of and NaOH (pH L. monocytogenes on 10.5, 100 mM) glass slides Reduce Pseudomonas, Acid and nonionics Acetic acid + Listeria, and monlaurin, pH Staphylococcus 5.4 biofilms by >3 log CFU/mL while maintaining the desirable microflora with <1-log reduction of Lactobacillus Nonionic: glyceryl 75 ppm Less than 50% reduction monolaurate in L. monocytogenes biofilm cells

Reference Ronner and Wong (1993)

Krysinski, and others (1992)

Frank and Koffi (1990)

Arizcun and others (1998)

Ammor and others (2004)

Chavant and others (2004)

(continued )

90

Biofilms in the Food Environment

Table 4.2. Summary of Research on the Effectiveness of Chemical Sanitizers for Inactivating Biofilm Cells (Continued ) Concentration Other fatty acids: 0.13%, 10-min C8 –C10 exposure

Others Chlorheximide: chlorhexidine gluconate

4%, 10-min exposure

Chlorheximide

200 mg/L in a simulated CIP system

Chlorheximide

0.2%, 16-h contact time

Trisodium phosphate

8%, 10–30 min

Effectiveness

Reference

4-log cells/cm2 reduction Krysinski and others of L. monocytogenes (1992) on stainless steel, 2-log reduction on polyester surfaces, and <1-log reduction on polyester/polyurethane conveyer belt material >6-log (CFU/mL) reduction of L. monocytogenes on stainless steel in the presence of 2% milk (carrier test method) 3 log or less reduction of Pseud. aeroginosa, M. luteus, and Strep. faecalis in the presence of milk soil and 5-log reduction of E. coli and Enter. aerogenes >5-log reduction in viable cells (Legionella pneumophila, Mycobacterium spp., Candida, and Pseud. aeroginosa) from a 14-day biofilm formed on silicone and polyurethane tubing but only a 31% removal of biofilm materials 1–3-log CFU/cm2 reduction of L. monocytogenes biofilm on stainless steel and 3-log reduction of Campylobacter and Salm. typhimurium biofilms

Best and others (1990)

Dunsmore and others (1980)

Walker and others (2003)

Somers and others (1994)

Inactivation of Listeria monocytogenes Biofilms

91

obtained an approximately 2-log CFU/mm2 reduction of E. coli, Pesudomonas fluorescens, and Staph. aureus attached to stainless steel using 100–200 ppm chlorine solution. McCarthy (1992) showed a 3–5-log reduction of L. monocytogenes on chitin after 5-min exposure at 100 ppm. Studies of attached Listeria in the presence of milk-based soil indicated an inactivation of about 2-log reduction (Best 1990, p. 324), whereas a 6-log reduction of Listeria in a mixed culture with Pseudomonas biofilm was obtained by Fatemi and Frank (1999); a 4-log reduction on brussel sprouts by Brackett (1987); and a <2-log reduction in cabbage and lettuce by Zhang and Farber (1996). Salmonella biofilms formed on cement and plastic surfaces were reduced by about a 4 log CFU/surface upon the exposure to 100 ppm chlorine for 10 min (Joseph and others 2001). In general, studies summarized in Table 4.3 indicate that to inactivate biofilm cells, chlorine concentration greater than 200 ppm with contact time of at least 5 min is required, but that 5-log reduction in biofilm cell numbers cannot be expected.

Chlorine dioxide Chlorine dioxide (ClO2 ) is a broad spectrum disinfectant which, in some applications, is more effective than chlorine. It has low toxicity, is nonflammable, and is stable in solution (Marriott 1999; Dychdala 2001). Chlorine dioxide retains activity in the presence of organic load and at high pH (Troller 1983). In food sanitation, stabilized chlorine dioxide (i.e., with anthium dioxide added) is applied as a gas, foam, or liquid. It is applied at 100 ppm as a no-rinse sanitizer, at 3–5 ppm as processing water, and at 1–5 ppm as foam (Marriott 1999). Chlorine dioxide at 7 ppm is successfully used in brewery CIP systems as an alternative to hot water sanitation. In this process, however, alkali cleaners must be replaced with acid cleaners since ClO2 reacts with the alkali. One advantage of chlorine dioxide is that it does not produce toxic by-products that are produced by chlorine, such as trihalomethane, haloacetic acid, or chlorophenols (Troller 1983; Dychdala 2001; Agius and others 2004). In studies testing the efficacy of cleaners and sanitizers on food contact surfaces (Krysinski and others 1992), application of 5 ppm chlorine dioxide for 10 min produced a 4-log cells/cm2 reduction of L. monocytogenes on stainless steel and polyester surfaces but <1-log reduction on polyester/polyurethane conveyer belt material. Sanitation studies of fruits and vegetables show that chlorine dioxide at 5 ppm for 10 min reduced L. monocytogenes attached to freshly cut vegetables by 1 log CFU/g (Zhang and Farber 1996). ClO2 used at 4 ppm for at least 10 min inactivated 2–4 logunits of L. monocytogenes attached to the pulp skin of apples. An exposure time of 30 min at 4 ppm

92

Biofilms in the Food Environment

Table 4.3. Summary of Research on the Heat Inactivation of Attached or L. monocytogenes Biofilm

Adherent L. monocytogenes cells, 1 day Adherent L. monocytogenes microcolonies L. monocytogenes biofilm cells Adherent cells, 7 days

Temperature (◦ C)

Time (min)

Effectiveness (log reduction)

82

10

4

Krysinski and others (1992)

70

5

4

Frank and Koffi (1990)

68

3

65

5

1.3

Reference

Lee and Frank (1991) Oh and Marshall 1995)

was required to obtain a 4–6-log reduction of L. monocytogenes on apple pulp skin (Du and others 2002). Saper and others (2003) showed that ClO2 used at 7.5 mg or about 0.3 ppm reduced E. coli attached to apples by about 5 log units. Taormina and Beuchat (1999) found that 200 ppm acidified chlorine dioxide inactivated >2 log units of E. coli O157:H7 attached to alfalfa sprouts. These studies indicate that chlorine dioxide can rapidly and effectively inactivate microorganisms in the presence of organic matter. The studies summarized in Table 4.3 suggest that chlorine dioxide is an effective sanitizer in inactivating attached microorganisms but little information is available on its effectiveness against biofilms. Electrolyzed Water Electrolyzed (EO) water is produced in two forms by an application of an electrical current across a cathode and an anode in a divided chamber containing pure water and sodium chloride: the anode produces acidic or oxidizing water (pH <2.5) and the cathode produces alkali or reducing water (pH >11) (Park and others 2004; Stevenson and others 2004). Studies showed that acidic EO water characterized by residual chlorine of 2 mg/L combined with an oxidation–reduction potential (ORP) of >841 mV is effective in inactivating L. monocytogenes and E. coli O157:H7 in suspension. The effects of water source, dilution, storage, inoculum levels, and organic load on the efficacy of EO water were studied by Stevenson and others (2004). The conclusion of that study was that there was a minimum concentration of 0.5% of EO water and 2% combined anode/cathode water to inactivate 4 log units of E. coli O157:H7 in suspension and that to maintain bactericidal activity an ORP of >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 <1-log reduction in Bacillus spores. In acidic EO water generated by an ROX unit at 56 ppm chlorine, ORP 1160 mV, and pH of 2.6, there was a >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 <1-log reduction of E. coli O157:H7 even after sequential treatments over 3 days (Singh and others 2003). The ozone treatment/UV treatment (UV intensity of 4,500 μW/cm2 and ozone dosage of 0.77 ppm/min) of poultry processing water resulted in a 6-log reduction of E. coli (Diaz and Law 1997). The sanitation of stainless steel with ozone (4 ppm, 3 min) resulted in the inactivation of 7 log CFU of biofilm cells (Robbins and others 2005). Iodine Iodophor at 25 ppm shows mixed results in its effectiveness against microbial biofilm. Ronner and Wong (1993) demonstrated a >4-log reduction of L. monocytogenes biofilm and a 3-log reduction of Salmonella typhimurium biofilm on stainless steel, but a <1-log reduction of both L. monocytogenes and Salm. typhimurium biofilms on rubber surfaces resulting from treatment with 25 ppm iodophor, while other studies obtained an approximate 2-log reduction of L. monocytogenes biofilm on stainless steel and 1-log or less reduction on polyester and polyester/polyurethane (Krysinski and others 1992). Iodine at 80 ppm (0.008%) was not effective in the carrier test on stainless steel in the presence of milk soil, as there was <1-log reduction of attached L. monocytogenes cells (Best and others 1990). Dunsmore and others (1980) demonstrated that in a CIP system with milk soil, iodine efficacy varied depending on the type of microorganism. Pseud. aeroginosa, E. coli, and Micrococcus luteus were more resistant to iodophore sanitizer than were Streptococcus faecalis and Enterobacteri aerogenes. Pseud. aeroginosa exhibited a <3-log reduction when exposed to 12.5 mg/L iodine, while E. coli and M. luteus exhibited a >5-log reduction for the same sanitizer exposure. However at 25 mg/L, populations of all five bacterial species were reduced by 5 log units (Dunsmore and others 1980). McCarthy (1992) demonstrated a 3-log reduction of L. monocytogenes attached to chitin after treatment with 100 ppm iodophore for 5 min. Peroxygen Compounds Peroxygen compounds are oxidizing agents that are often effective against biofilms. Hydrogen peroxide (H2 O2 ) is used as an antiseptic at 3%, as a sterilant for aseptic packages at 25 ppm, and as a surface sanitizer at

Inactivation of Listeria monocytogenes Biofilms

95

125–250 ppm. Peracetic acid sanitizer is an equilibrium mixture of hydrogen peroxide, acetic acid, and peroxyacetic acid. Peroctanoic acid-based sanitizer is an equilibrium mixture of hydrogen peroxide, acetic acid, peroxyacetic acid, octanoic acid, and peroxyoctanoic acid. Krysinski and others (1992) found that 25 ppm peracetic acid sanitizer reduced L. monocytogenes on stainless steel and polyester by 4 log CFU/cm2 but <1-log reduction was observed on polyester/polyurethane conveyer belt material. A solution containing 1.2 ppm peracetic acid plus 160 ppm hydrogen peroxide effectively reduced L. monocytogenes, Salm. typhimurium, Staph. aureus, and Saccharomyces cerevisiae attached to stainless steel surfaces by >3, 2.9, 0.8, and 1.2 log CFU/surface respectively (Knowles and Roller 2001). Rossoni and Gaylarde (2000) used 250–1000 ppm peracetic acid solution with a 10-min exposure time and showed a 1 log or less CFU/mm2 reduction of E. coli, Pseud. fluorescens, and Staph. aureus attached to stainless steel. Makela and others (1991) used the tray test method to determine the efficacy of 180 ppm peracetic acid after 10-min exposure on attached lactic acid bacteria to stainless steel. The test showed a >4-log reduction in the attached population. Fatemi and Frank (1999) used 80–160 ppm of peracetic/peroctanoic acid solution for 5 min and obtained a 6–7-log CFU/cm2 reduction of L. monocytogenes and Pseudomonas biofilms on stainless steel in the presence of milk soil. Frank and others (2003) obtained 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 <1 log unit on polyester/polyurethane conveyer belt material. Surfactant Sanitizers Surfactant or surface active ingredients are ampholytic compounds that accumulate at the surface interface of immiscible liquids and lower the surface tension. There are three categories of surfactants: anionics, cationics, and nonionics. Quaternary ammonia compounds (QAC) are cationic surfactants that denature proteins and disrupt the microbial cell membrane. Common QAC include alkyl dimethylethyl benzyl ammonium chloride, betaine, alkyldimethylamine, pyridium quaternary, cepacol, and ethylpyridium chloride. QAC are commonly used sanitizers in the food industry. Manufacturers generally recommend application of 200 ppm QAC for 2 min on food contact surfaces. Frank and Koffi (1990) observed that a 20-min exposure to 200 ppm QAC was required to reduce L. monocytogenes biofilm by 4–5 log units. Ronner and Wong (1993) observed that 200 ppm QAC at 1-min exposure was effective in reducing L. monocytogenes biofilms on stainless steel surfaces by 3 log units. QAC at 100–200 ppm for 10-min exposure inactivated L. monocytogenes attached to chitin by 4–5 log units (McCarthy 1992). Frank and others (1997) observed that QAC at 200 ppm for 35 sec was effective in reducing attached Staph. aureus by 2–5 log CFU/cm2 for

Inactivation of Listeria monocytogenes Biofilms

97

various smooth and abraded surfaces. Acidic QAC at 200 ppm, pH 2.4 was effective on stainless steel with at least 4-log reduction of L. monocytogenes biofilm, while on polyester and polyester/polyurethane there was about a 2-log reduction (Krysinski and others 1992). Neutral QAC at 200 ppm, pH 6.4 was effective on stainless steel with at least a 4-log reduction of L. monocytogenes biofilm, while on polyester and polyester/polyurethane there was about a 2-log and 1-log reduction respectively (Krysinski and others 1992). Chlorinated alkali cleaning followed by application of QAC is effective in the removal of Listeria biofilm from both stainless steel and conveyer belt material (Krysinski and others 1992). QAC are most effective at temperatures greater than 37◦ C but they are also effective at cold temperatures for long exposure (Gelinas and others 1984). Best and others (1990) observed a >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 <1 <1 >15 <1 <1 <1 <1 <1

75 75 75 75 75 75 90 90 90 90 90 90 90 90 90

Inactivation of Listeria monocytogenes Biofilms

101

controlled. The models provide processors with a risk management tool that provides predicted probabilities of L. monocytogenes inactivation and allows a choice of three different heat resistance assumptions (inactivation based on strain Scott A, strain 3990, or mixed microflora). An important challenge in the practical application of these models is that it may be difficult to identify and verify cold spots within complex equipment. Sites within equipment that are not reached by the flow of cleaning solutions may not also be reached by the flow of hot water. In addition, soft parts such as gaskets and air pockets may insulate or interfere with heat transfer to the extent that unexpected cold spots develop. Users of the heat inactivation models may assume that if they exceed heat treatments employed to develop the model then the risk of Listeria survival will be less but extrapolations of the models will not provide accurate risk estimates. Acknowledgments The authors’ research was supported by state and Hatch funds provided to the Georgia Agricultural Experiment Station. Additional funding was provided by the Center for Food Safety and contributions from the food industry. References Agius G, Burkeen S, Mynatt J. 2004. Benefits of using chlorine dioxide as an alternative to hot water sanitation. Master Brewer Assoc Americas Tech Q 41:42–44. Ammor S, Chevallier I, Laguet A, Labadie J, Talon R, Dufour E. 2004. Investigation of the selective bactericidal effect of several decontaminating solutions on bacterial biofilms including useful, spoilage and/or pathogenic bacteria. Food Microbiol. 21:11–17. Antoniou K, Frank JF. 2005. Removal of Pseudomonas putida biofilm and associated extracellular polymeric substances from stainless steel using alkali cleaning. J Food Prot 68:179–183. Arizcun C, Vasseur C, Labadie J. 1998. Effect of several decontamination procedure on Listeria monocytogenes growing in biofilms. J Food Prot 61:731–734. Bal’a MFA, Jamilah ID, Marshall DL. 1999. Moderate heat or chlorine destroys Aeromonas hydrophila biofilms on stainless steel. Dairy Food Envron. Sanit 19:29–34. Best M, Kennedy ME, Coates F. 1990. Efficacy of a variety of disinfectants against Listeria spp. Appl Environ Microbiol 56:377–380. Bialka KL, Demirci A, Knabel SJ, Patterson PH, Puri VM. 2004. Efficacy of electrolyzed oxidizing water for the microbial safety and quality of eggs. Poult Sci 83:2071–2078. Brackett RE. 1987. Antimicrobial effect of chlorine on Listeria monocytogenes. J Food Prot 50:999–1003. Bredholt S, Maukonen J, Kajanpaa K, Alanko T, Olofson U, Husmark U, Sjoberg AM. 1999. Microbial methods for assessment of cleaning and disinfection of food-processing surfaces cleaned in a low-pressure system. Eur Food Res Technol 209:145–152.

102

Biofilms in the Food Environment

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 35:321–325. 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 236:241–248. Chen X, Stewart PS. 2000. Biofilm removal caused by chemical treatments. Water Res 34:4229–4233. Chmielewski RA, Frank JF. 2004. A predictive model for heat inactivation of Listeria monocytogenes biofilm on stainless steel. J Food Prot 67:2712–2718. Diaz ME, Law SE. 1997. Ultraviolet photon enhanced ozonation for microbiological safety in poultry processing water. Paper presented at ASAE Meeting, St. Joseph, MI, 10–14 August 2004. Paper No. 976054. Du J, Han Y, Linton RH. 2002. Inactivation by chlorine dioxide gas of Listeria monocytogenes spotted onto different apple surfaces. Food Microbiol 19:481–490. Dufour M, Simmonds RS, Bremer PJ. 2004. Development of a laboratory scale clean-in-place system to test the effectiveness pf “natural” antimicrobials against dairy biofilms. J Food Prot 67:1438–1443. Dunsmore DG. 1981. Bacteriology control of food equipment surfaces by cleaning systems. I. Detergent effects. J Food Prot 44:15–20. Dunsmore DG, Thomson MA. 1981. Bacteriological control of food equipment surfaces by cleaning systems. II. Sanitizer effects. J. Food Prot 44: 21–27. Dunsmore DG, Thomson MA, Murray G. 1981. Bacteriological control of food equipment surfaces by cleaning systems. III. Complementary cleaning. J Food Prot 44:100– 108. Dunsmore DG, Westwood DA, Jay DB, Embling M. 1980. Simulator technique for assessing the bacteriological control of food equipment surfaces by cleaning systems. J Food Prot 43:850–855. Dychdala GR. 2001. Chlorine and chlorine compounds. In Disinfection, Sterilization, and Preservation, 5th Edition (GR Dychdala, Editor). Lippinscott Williams & Wilkins, Philadelphia, p. 1481. Fabrizio KA, Sharma RR, Demirci A, Cutter C. 2002. Comparison of electrolyzed oxidizing water with various antimicrobial interventions to reduce Salmonella species on poultry. Poul Sci 81:1598–1605. Fatemi P, Frank JF. 1999. Inactivation of Listeria monocytogenes/Pseudomonas biofilms by peracid sanitizers. J Food Prot 62:761–765. Frank J, Chmielewski R. 2001. Influence of surface finish on the cleanability on stainless steel. J Food Prot 68:1178–1182. Frank J, Koffi R. 1990. Surface-adherence growth of Listeria monocytogenes is associated with increased resistance to surfactant sanitizers and heat. J Food Prot 53:550–554. Frank JF, Chmielewski RA. 1997. Effectiveness of sanitation with quaternary ammonium compound or chlorine on stainless steel and other domestic food-preparation surfaces. J Food Prot 60:1–6. Frank JF, Ehlers J, Wicker L. 2003. Removal of Listeria monocytogenes and poutry soilcontaining biofilms using chemical cleaning and sanitizing agents under static conditions. Food Prot Trends 23:654–663. Gelinas P, Goulet J, Tastayre GM, Picard GA. 1984. Effect of temperature and contact time on the activity of eight disinfectants—a classification. J Food Prot 47:841–847. Gibson H, Taylor JH, Hall KE, Holah JT. 1999. Effectiveness of cleaning techniques used

Inactivation of Listeria monocytogenes Biofilms

103

in the food industry in terms of the removal of bacterial biofilms. J Appl Microbiol 87:41–48. Holah JT, Thorne RH. 1990. Cleanability in relation to bacterial retention on unused and abraded domestic sink materials. J Appl Bacteriol 69:599–608. Jackson AT. 1985. Cleaning of food processing plant. In Developments in Food Preservation-3 (AT Jackson, Editor) Elsevier Science Publications, New York, pp. 95– 125. Joseph B, Otta SK, Karunasagar I, Karunasagar I. 2001. Biofilm formation by Salmonella spp. on food contact surfaces and their sensitivity to sanitizers. Int J Food Microbiol 64:367–372. Khadre MA, Yousef AE, Kim J.-G. 2001. Microbiological aspects of ozone applications in foods: A review. J Food Sci 66:1242–1252. Kim C, Hung YC, Brackett RE. 2000. Efficacy of electrolyzed oxidizing (EO) and chemically modified water on different types of foodborne pathogens. Int J Food Microbiol 61:199– 207. Kim C, Hung YC, Brackett RE, Frank JF. 2001. Inactivation of Listeria monocytogenes biofilms by electrolyzed oxidizing water. J Food Process Preserv 25:91–100. Kim JG, Yousef AE. 2000. Inactivation kinetics of foodborne spoilage and pahtogenic bacteria by ozone. J Food Sci 65:521–528. Knowles J, Roller S. 2001. Efficacy of chitosan, carvacol, and a hydrogen peroxide based biocide against foodborne microorganisms in suspension and adhered to stainless steel. J Food Prot 64:1542–1548. Koseki S, Yoshida K, Isobe S, Itoh K. 2004. Efficacy of acidic electrolyzed water ice for pathogen control on lettuce. J Food Prot 67:2544–2549. Krysinski EP, Brown LJ, Marchisello TJ. 1992. Effect of cleaners and sanitizers on Listeria monocytogenes attached to product contact surfaces. J Food Prot 55:246–251. LeChevallier MW, Cawthon CD, Ramon RG. 1988. Inactivation of biofilm bacteria. Appl Environ Microbiol 54:2492–2499. Lee S-H, Frank J. 1991. Inactivation of surface adherent Listeria monocytogenes hypochlorite and heat. J Food Prot 54:4–6. Luppens S, Reij M, van der Heijden RWL, Rombouts F, Abee T. 2002. Development of a standard test to assess the resistance of Staphylococcus aureus biofilm cells to disinfectants. Appl Environ Microbiol 68:4194–4200. Makela PM, Korkeala HJ, Sand EK. 1991. Effectiveness of commercial germicide products against the ropy slime-producing lactic acid bacteria. J Food Prot 54:632–636. Marriott NG. 1999. Principles of Food Sanitation, 4th Edition. Aspen, Gaithersburg, MD, p. 364. McCarthy SA. 1992. Attachment of Listeria monocytogenes to chitin and resistance to biocides. Food Technol 46:84–88. Mustapha A, Liewen MB. 1989. Destruction of Listeria moncytogenes by sodium hypochlorite and quaternary ammonium sanitizers. J Food Prot 52:306–311. Oh D-H, Marshall DL. 1995. Destruction of Listeria monocytogenes biofilms on stainless steel using monolaurin and heat. J Food Prot 57:251–255. Ozer NP, Demirci A. 2005. Electrolyzed oxidizing water treatment for decontamination of raw salmon inoculated with Escherichia coli O157:H7 and Listeria monocytogenes Scott A and response surface modeling. J Food Eng 21:559–566. Park H, Hung Y-C, Chung D. 2004. Effect of chlorine and pH on efficacy of electrolyzed water for inactivating Escherichia coli O157:H7 and Listeria monocytogenes. Int J Food Microbiol 91:13–18.

104

Biofilms in the Food Environment

Restaino L, Frampton EW, Hemphill JB, Palnikar P. 1995. Efficacy of ozonated water against various food-related microorganisms. Appl Environ Microbiol 61:3471–3475. Robbins JB, Fisher CW, Moltz AG, Martin SE. 2005. Elimination of Listeria monocytogenes biofilms by ozone, chlorine, and hydrogen peroxide. J Food Prot 86:494–498. Ronner A, Wong A. 1993. Biofilm development and sanitizer inactivation of Listeria monocytogenes and Salmonella typhimurium on stainless steel and buna-n rubber. J Food Prot 56:750–758. Rossoni EM, Gaylarde CC. 2000. Comparison of sodium hypochlorite and peracetic acid as sanitising agents for stainless steel food processing surfaces using epifluorescence microscopy. J Food Microbiol 61:81–85. Sapers GM, Walker PN, Sites JE, Annous BA, Eblen DR. 2003. Vapor-phase decontaminatation of apples inoculated with Escherichia coli. J Food Sci 68:1003–1007. Sharma RR, Demirci A. 2003. Treatment of Escherichia coli O157:H7 inoculated alfalfa seeds and sprouts with electrolyzed oxidizing water. Int J Food Microbiol 86:231–237. Singh N, Singh RK, Bhunia AK. 2003. Sequential disinfection of Escherichia coli O157:H7 inoculated alfalfa seeds before and during sprouting using aqueous chlorine dioxide, ozonated water, and thyme essential oil. Lebensm Wiss U Technol 36:235–243. Somers EB, Schoeni JL, Wong A. 1994. Effect of trisodium phosphate on biofilm and planktonic cells of Campylobacter jejuni, Escherichia coli O157:H7, Listeria monocytogenes and Salmonella typhimurium. Int J Food Microbiol 22:269–276. Stan SD, Daeschel MA. 2003. Reduction of Salmonella enterica on alfalfa seeds with acidic electrolyzed oxidizing water and enhanced uptake of acidic electrolyzed oxidizing water into seeds by gas exchange. J Food Prot 66:2017–2022. Stanfield P. 2003. Cleaning and sanitizing a food plant. In Food Plant Sanitation (P Stanfield, Editor).Marcel Dekker, New York, p. 101. Stevenson SM, Cook SR, Bach SJ, McAllister TA. 2004. Effects of water source, dilution, storage and bacterial and fecal loads on the efficacy of electrolyzed oxidizing water for the control of Escherichia coli O157:H7. J Food Prot 67:1377–1383. Storgards E, Simola H, Sjoberg A-M, Wirtanen G. 1999a. Hygiene of gasket material used in food processing equipment. Part 1: New materials. Trans Inst Chem Eng 77:137–145. Storgards E, Simola H, Sjoberg A-M, Wirtanen G. 1999b. Hygiene of gasket material used in food processing equipment. Part 2: Aged materials. Trans Inst Chem Eng 77:146–155. Taormina PJ, Beuchat LR. 1999. Comparison of chemical treatments to eliminate enterohemorrhagic Escherichia coli O157:H7 on alfalfa seeds. J Food Prot 62:318–324. Troller JA. 1983. Sanitation in Food Processing. Academic Press, New York, p. 456. Walker JT, Bradshaw DJ, Fulford MR, Marsh PD. 2003. Microbiological evaluation of a range of disinfectant products to control mixed-species biofilm contamination of a laboratory model of a dental unit water system. Appl Environ Microbiol 69:3327–3332. Wang H, Feng H, Luo Y. 2004. Microbial reduction and storage quality of fresh-cut cilantro washed with acidic electrolyzed water and aqueous ozone. Food Res Int 37:949–956. Wirtanen G, Husmark U, Matilla-Sandholm T. 1996. Microbial evaluation of the biotransfer potential from surfaces with Bacillus biofilms after rinsing and cleaning procedures in closed food-processing systems. J Food Prot 59:727–733. Zhang S, Farber JM. 1996. The effects of various disinfectants against Listeria monocytogenes on fresh cut vegetables. Food Microbiol 13:311–321.

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 5 MIXED CULTURE BIOFILMS Michele Y. Manuzon and Hua H. Wang

Introduction Biofilms are probably the most prevalent form of microbial life in both natural and processing environments. Costerton and others (1995) defined biofilms as “matrix-enclosed bacterial populations adherent to each other and/or to surfaces or interfaces, and includes microbial aggregates and floccules and also adherent populations within the pore spaces of porous media.” While most of the knowledge on bacterial biofilms has been gained from studying the behavior of microorganisms in monoculture, such monospecies biofilms are rarely found in natural environment and industrial settings (Percival and others 2000; Sutherland 2001). Most ecosystems consist of heterogeneous microorganisms associated with each other, although certain species or genera might be predominant in these biofilms. Monospecies biofilms of Pseudomonas aeruginosa or Staphylococcus aureus may be found in clinical settings (Percival and others 2000), although the polymicrobial nature of many infectious biofilms is now being recognized (Brogden 2002). In natural ecosystems, certain microorganisms may act as primary colonizers attaching to the surfaces, while others can become biofilm partners by establishing interactions with existing organisms (Kolenblander 2000). Not only does the coexistence of different microorganisms in the environment lead to the formation of mixed culture biofilms, but such ecosystems also provide an ideal environment for microbial residents to coordinate metabolic activities and share genetic elements encoding beneficial traits with each other. Mixed culture biofilms therefore play a significant and active role in microbial evolution. The various intraspecies and interspecies interactions among microorganisms add to the overall complexity and diversity of natural biofilms. Therapeutic treatments and biofilm removal are in fact much more challenging in such mixed species ecosystems compared 105

106

Biofilms in the Food Environment

to monoculture biofilms. Accordingly, a comprehensive understanding of mixed culture biofilms including mechanisms involved in development, intracellular communication, and gene exchange is essential for targeted strategies to control this problem in natural, food processing, and clinical settings. This chapter will introduce current methodologies to examine the microbial composition of these ecosystems, current knowledge on mixed culture biofilm-forming mechanisms, and risks associated with these microbial consortia. Emerging strategies for the development of new sanitation ingredients for thorough cleaning will be discussed briefly. Many of these studies were conducted using organisms of medical significance, but the overall approach and knowledge could also be applied for biofilm studies and problem solving in the food environment. Analyses of Microbial Diversity in Biofilms Biofilms in nature are composed of different microorganisms and might include species of bacteria, archaea, yeasts, molds, algae, or protozoans. Analysis of the microbial composition of these mixed ecosystems requires identification of the different species present and their relative abundance in the community. However, conventional methods relying on pure culture techniques are very tedious and time-consuming, particularly when dealing with slow-growing microorganisms (von Wintzingerode and others 2002). Moreover, many microorganisms do not grow very well under laboratory conditions. In fact, several studies have shown that less than 1% of the total microbial population from environmental samples could be recovered using standard cultivation methods (Amann and others 1995). Wimpenny and others (2000) estimated that approximately 99.9% of microorganisms in natural biofilms could not be cultivated on standard microbiological media. Thus, previous studies on the identification of microorganisms based on traditional culturing procedures resulted in a skewed assessment of the actual diversity in microbial communities in what has been referred to as the ‘the great plate count anomaly” (Staley and Konopka 1985). Population analyses based on culture-dependent techniques usually underestimate the actual microbial heterogeneity. Methods that would not rely on the growth of pure cultures are therefore needed for more accurate assessment of microbial diversity in natural biofilm communities. DNA Sequencing With the development of molecular biology tools, it is now possible to identify microorganisms without the need to grow them in pure cultures.

Mixed Culture Biofilms

107

One of the approaches is to analyze the ribosomal RNA (rRNA) and intergenic transcribed spacer (ITS) sequences to determine the identity of microorganisms. Woese (1987) first recognized the importance of the rRNA in establishing phylogenetic relationships among living organisms, with the sequence homologies among different organisms directly correlated to their degree of relatedness. Bacteria commonly have three rRNAs: the 5S, 16S, and 23S (Olsen and others 1986). In eukaryotes, rRNAs occur in several class sizes as well: large subunit (25S–28S), small subunit (18S), and 5S. The genes encoding these eukaryotic rRNAs occur as tandem repeats and may be present in as high as 100–200 copies (Kurtzman and Blanz 1998). Several universal primers for amplification of these 18S rRNA genes have already been developed and applied successfully for the detection and identification of various yeasts and molds (Kappe and others 1996; Wan et al., 2006). Eukaryotic organisms generally have a fourth rRNA, the 5.8S with approximately 160 nucleotides, which exhibits high homology with the 5 end of the bacterial 23S rRNA (Olsen and others 1986). In prokaryotes, the sequences of the genes encoding the small ribosomal subunit RNA (16S rRNA) are often used for determining bacterial taxonomic classifications and evolutionary relationships (Weisburg and others 1991) since these genes have both highly conserved as well as moderately variable regions (Woese 1987). The 16S rRNA was first sequenced in 1978 from Escherichia coli (Brimacombe 1978). It is approximately 1500 base pairs (bp) in length, and has an intricate looped secondary structure with approximately 45 separate helices, and has multiple copies in the chromosome for most organisms. Today, the most extensively reported genes in the databases are 16S rRNA sequences (von Wintzingerode and others 2002). In many cases, the identity of an unknown bacterial strain (genus, and sometimes species) can be determined by comparing the homology of its 16S rRNA gene sequence to the sequences deposited in the databases. The 16S rRNA sequence has been an important parameter for bacterial strain identification. Assessing the 16S rRNA sequence pool has proven to be a very useful tool in ecosystem population analyses as well. The general procedures include extracting the total DNA from either the clinical or environmental samples, and using one or several pair(s) of universal primers to amplify the 16S rRNA gene by polymerase chain reaction (PCR). The PCR products are then cloned and the inserted DNA sequences from the clone libraries are determined. Even if some organisms cannot be recovered from the ecosystem by conventional culturing method, their DNA can be extracted and amplified. Thus their unique DNA sequences in the clone libraries represent their presence in the ecosystem. Assuming that the amount of PCR amplicons is correlated to the numbers of the templates in

108

Biofilms in the Food Environment

the original sample, the population diversity of the ecosystem can be calculated based on the frequencies of genus- or species-specific sequences in the libraries. However, dominant amplification often happens in many cases, thereby affecting the accuracy of the population assessment. Thus, approaches to minimize such incidences, such as comparing data from multiple clone libraries, cross-referencing with other direct population diversity estimation methods, and even applying special treatments to limit the amplification of dominant species in PCR in certain clone libraries, are important for a more comprehensive assessment of the ecosystem composition, particularly in revealing the presence of minor populations. Kroes and others (1999) successfully implemented such practices in their study to determine microbial population diversity within the human subgingival crevice. They found that greater diversity of bacterial 16S rDNA sequence types (phylotypes) was obtained by direct PCR amplification of the sample than through the use of culturing techniques. Ward (2002) also reported that most of the 16S rRNA sequences obtained directly from environmental samples did not match any sequences deposited in the GenBank. Aside from the analysis of 16S rRNA sequences, ribosomal intergenic spacer analysis (RISA) has also been used to study microbial community structure and composition. A pair of primers derived from 16S and 23S rRNA gene sequences are used in RISA. For instance, Larue and others (2005) reported the application of RISA to examine the microbial diversity in the digestive tract of herbivores. The results showed that microorganisms present in the planktonic populations of ruminal microorganisms differ from those associated with or adherent to plant biomass. While sequence analysis of rRNA genes is very useful for microbial identification in mixed communities, it does not provide insights into the organization and functionality of these ecosystems. Denaturing Gradient Gel Electrophoresis Muyzer and others (1993) first described the use of PCR-amplified V3 variable region of 16S rRNA genes and denaturing gradient gel electrophoresis (DGGE) for the analysis of genetic diversity in complex microbial communities. The method was used to analyze the composition of microbial mats from Wadden Sea sediment and bacterial biofilms obtained from wastewater treatment facilities. DGGE analysis of the different mixed populations showed the presence of 5–10 distinguishable bands in the separation patterns. Subsequent hybridization steps using group-specific nucleotides enabled the identification of specific microorganisms in the population. Using template DNA from known isolates, they were able to show the

Mixed Culture Biofilms

109

sensitivity of the method in detecting microorganisms representing only 1% of the total population. The developed technique could be used for the identification of different species as well as for determining their relative abundance in the community, thereby providing a novel method for both qualitative and quantitative analyses of genetic diversity in complex microbial populations. Lyautey and others (2005) also used DGGE to assess the bacterial diversity in a river epilithic biofilm. Molecular Probes The application of molecular probes is another way of assessing microbial ecosystems, and has the potential for in situ ecosystem functionality analysis. Basically, organism-specific oligonucleotide probes are first developed targeting rRNA or other signature genes. These probes can then be used to detect and possibly identify constituent microorganisms and to profile the community structure when combined with PCR amplification of target gene sequences using DNA extracted from the community population (Fletcher 1999). Such genus- or species-specific probes can further be used in fluorescent in situ hybridization or microarray to assess the spatial arrangement of cells or other functionalities of the ecosystem. Microarray The DNA microarray platform enables the analyses of multiple genes, and even complete microbial genome(s) simultaneously, and is becoming a popular tool in determining the phylogenetic and functional relationships among different microorganisms (Murray and others 2001). Rudi and others (2002) developed a 16S-rRNA-gene-based array method for the detection and enumeration of microorganisms. Various fluorescently labeled genus- or species-specific 16S rRNA gene probes were spotted on the array. DNA or RNA extracts from the ecosystem were hybridized to the array and the positive signals indicated the presence of the particular types of organisms. They compared the developed microarray method with both cultivation-dependent assays (enrichment and plating) and cultivationindependent assays based on direct fluorescence microscopy and scanning electron microscopy using ready-to-eat vegetables packed in modified atmospheres. The DNA-based array method gave a more accurate description of the microbial communities. Murray and others (2001) constructed a DNA microarray to assess the degree of phylogenetic and functional relatedness among strains belonging to the genus Shewanella, using 192 genes from the genome of Shewanella oneidensis MR-1. In another study, Peplies and others (2003) optimized the conditions for DNA microarray for

110

Biofilms in the Food Environment

the specific detection of microorganisms based on 16S rRNA genes using six environmental strains and 20 oligonucleotide probes. The improved method using directed application of capture oligonucleotides resulted in less false-positive and false-negative results. Fluorescent In Situ Hybridization During the past several years, microbiologists have developed techniques that allow the study of microorganisms in vivo and in situ (Costerton and others 1987). Microscopic and digital image analysis techniques, particularly confocal laser scanning coupled with fluorescent probe(s), are widely used in studying biofilm structure and function (Nancharaiaha and others 2005). In order to study the spatial arrangement of bacteria in a mixed culture biofilm, techniques that rely on specific microbial cell markers and maintain the natural ultrastructure of the biofilm are required. Confocal laser scanning microscopy (CLSM) is the method of choice for studying bacterial biofilms, mainly because of its noninvasive, three-dimensional illustration of the cells in the biofilm matrix and computational reconstruction of mature biofilms without distortion (Thurnheer and others 2004). Examination of microbial biofilms by CLSM may be accomplished using a wide variety of specific fluorescent probes as well as nonspecific fluorescent compounds (Costerton and others 1995). Fluorescent in situ hybridization (FISH) enables the detection of specific nucleic acid sequences in eukaryotic and prokaryotic cells through the binding of fluorescently labeled oligonucleotide probes to their complementary target sequences (Amann and others 1995). However, a major drawback for FISH is that only a small number of probes can be used in a single hybridization experiment, limiting its application in microbial heterogeneity analysis. For ecosystem composition analyses, multiple probes are often needed to achieve a high level of phylogenetic resolution (Peplies and others 2003) or to minimize false-positive or false-negative results in identification of selected target organisms by individual probes (Amann and others 1995). One of the critical challenges in using this method is the development of experimental conditions that would make the cell walls of the Gram-positive bacteria permeable to the probes without the concomitant loss of signal from Gram-negative bacteria that are simultaneously labeled with oligonucleotide probes. Preferably, several consecutive hybridization procedures could be performed without affecting the biofilm structure (Thurnheer and others 2004). The use of CSLM and FISH enables the study of microbial diversity in natural ecosystems without the selective bias of cultivation, extraction, or amplification (Davey and O’Toole 2000). In a recent study, Laramee

Mixed Culture Biofilms

111

and others (2000) studied the distribution of diclofop-methyl-degrading bacteria in a mixed biofilm. Oligonucleotide probes specific for bacteriaharboring genes for the degradation of related chlorinated aromatic compounds were successfully used for detecting selected isolates from the mixed culture biofilm using FISH. Another popular application of FISH is for the monitoring of gene transfer in mixed ecosystems. Christensen and others (1998) studied the conjugal transfer of green-fluorescent protein (gfp)-tagged version of the TOL plasmid in mixed culture biofilm of Pseudomonas putida RI, Acinetobacter sp. strain C6, and an unknown isolate D8 involved in benzyl alcohol degradation. Induction of the gfp gene enabled the detection of cells receiving the plasmid. Denaturing High-Performance Liquid Chromatography Barlaan and others (2005) reported the application of denaturing highperformance liquid chromatography (DHPLC) of amplified 16S rDNA sequences in microbial diversity assessment. They established the optimum conditions for microbial analysis using marine bacterial samples. PCRDHPLC analysis of microbial composition showed profiles with distinct peaks, representing the different microorganisms present and their relative abundance. Subsequent fraction collection and DNA sequencing of the profile peaks allowed the identification of the different species present in the sample. This technology may find application in characterizing microbial communities in nature. The different methods developed for studying biofilm structure and composition have their own advantages and limitations. The suitability of a particular procedure is largely determined by the purpose of the study and the level of sensitivity and resolution required, which often depends on the mass and complexity of the biofilms and on the availability of equipment and expertise (Fletcher 1999). In order to get a comprehensive picture of the ecosystem, it is necessary to combine the full cycle rRNA approach with other techniques to study the functional and physiological properties of the detected microorganisms (Kroes and others 1999; Wagner and others 2002).

Biofilm Ecosystem Development The ability of an organism to form biofilms in nature or at least integrate into an existing biofilm is essential to its survival in natural ecosystems or even within its eukaryotic host. Being part of a biofilm community confers

112

Biofilms in the Food Environment

several advantages to a microbial cell, including increased resistance to antimicrobial agents, as well as the capacity to withstand highly variable or adverse environmental conditions. All microorganisms, including those incapable of forming biofilms under laboratory conditions, could possibly form or become part of a biofilm ecosystem in nature. The ability of microorganisms to form biofilms is likewise affected by the presence of other organisms. For instance, Moretro and Langsrud (2004) concluded that some bacteria enhanced the formation of biofilms by Listeria monocytogenes, whereas others have inhibitory effects. Filoche and others (2004) also reported that the presence of Actinomyces naeslundii and Actinomyces gerencseriae promoted biofilm formation by Lactobacillus rhamnosus and Lactobacillus plantarum. Although both lactobacilli showed poor monoculture biofilm formation, they both significantly increased in numbers in the biofilms when they were cocultured with Actinomyces species. Streptococcus mutans promoted biofilm formation by lactobacilli to a significantly lesser extent, while Veillonella parvula had no effect. Coaggregation and Aggregation The ability of microorganisms to aggregate in liquid culture is correlated to their abilities to form biofilms. Autoaggregation and clumping have also been used to describe similar phenotypes. Cell aggregates in liquid cultures share the same features as biofilms, including the presence of an extracellular matrix and the presence of distinct chemical gradients within the aggregate. Coaggregation also enables cells to withstand highly variable conditions, which would otherwise adversely affect nonaggregated cells (Rickard and others 2003). Cell aggregates or flocs are often found in wastewater treatment systems as well as many other natural settings. This aggregative process may occur between similar or different microorganisms (for reviews, see Kolenblander and others 2005; Parsek and Greenberg 2005). When this occurs between genetically distinct microorganisms, it is referred to as coaggregation. Rickard and others (2003) defined coaggregation as “a process by which genetically distinct bacteria become attached to one another via specific molecules.” It was initially used to describe interactions among dental plaque bacteria (Kolenbrander 1988; Kolenblander 1997; Kolenblander and others 1995). Since then, coaggregation has been observed amongst bacteria isolated from various biofilm ecosystems and has been recognized as an important mechanism in the development of complex mixed-culture biofilms. Typically, coaggregation formed between pairs of bacteria is highly specific and is usually mediated by the interaction between a protein adhesin on

Mixed Culture Biofilms

113

one cell surface and a compatible receptor on the other. For instance, Sharma and others (2005) showed that coaggregation of Fusobacterium nucleatum and Tannerella forsythia is necessary for the formation of mixed-species biofilms. BspA protein, a T. forsythia surface adhesin, is involved in coaggregation of this organism with F. nucleatum, along with other adhesins. Both SspA and SspB are important cell surface adhesins found in autoaggregation or coaggregation involving Streptococcus gordonii (Kolenbrander 1989; Egland and others 2001). Ghigo (2001) showed that surface adhesin (pilin)-mediated cell aggregation is a critical factor in biofilm formation in E. coli. Reisner and others (2003) also demonstrated that the E. coli adhesins causing cell aggregation are involved in biofilm development. The staphylococcal clumping factors ClfA and ClfB are fibrinogen-binding proteins, and ClfA mediates staphylococcal adherence to host extracellular matrix components as well as abiotic surfaces (Vaudaux, and others 1995; Foster and Hook 1998). Certain surface components such as the staphylococcal biofilm-accumulation-associated protein (AAP) have greater impact on biofilm formation than other adhesins. Staphylococcal strains expressing AAP produce significantly larger amounts of biofilm than strains without this antigen (Hussain and others 1997). In Lactococcus lactis, the clumping protein CluA can also considerably facilitate biofilm development. Biofilms formed by lactococcal strains with induced overexpression of the clumping protein CluA is seven to eight times thicker than those without this protein (Luo and others 2005). Quorum Sensing The term quorum sensing refers to the ability of a microorganism to perceive and respond to microbial population density, usually through the generation and response to diffusible signal molecules. The involvement of quorum sensing in Gram-negative bacterial biofilm formation was well illustrated in the model organism P. aeruginosa (Davies and others 1998). Two cell-to-cell signaling systems, lasR–lasI and rhlR–rhlI, were identified in P. aeruginosa. The lasI gene product directs the synthesis of a diffusible extracellular signal, N-(3-oxododecanoyl)-L-homoserine lactone (3OC12 -HSL). The signal regulates the expression of lasR, and the product of which is a transcriptional regulator involved in activating a number of virulence genes, including lasI, and the rhlR–rhlI system. The rhlI product directs the synthesis of a second diffusible signal, N-buytrylL-homoserine lactone. The rhlR gene product is regulated by the signal and is required for the activation of virulence genes and expression of the stationary-phase σ factor, RpoS. The mutant strain defective in luxI gene

114

Biofilms in the Food Environment

forms only thin and uniform biofilm lacking three-dimensional structure. Meanwhile, exogenous 3OC12 -HSL can complement the phenotype mutation, suggesting the involvement of the quorum sensing in its biofilm development. Li and others (2002) illustrated the role of quorum sensing in Gram-positive bacteria biofilm formation. In this case, the signal is a peptide encoded by the comC gene. A two-component regulatory system (ComDE) composed of a sensor histidine kinase and a response regulator is involved in sensing the secreted ComC signal from the environment. Both the comD or comE mutants have altered biofilm architecture. The comC mutant is also defective in biofilm formation but the phenotype can be complemented by exogenous synthesized signal peptide. Since then, the role of quorum sensing in biofilm formation has been identified in many other bacteria and various signals have been characterized. However, signals themselves are not directly involved in biofilm development. It seems that the signals trigger the bacterial regulatory machinery leading to a cascade of reactions. The most likely notion is that among these reactions, the expression of key biofilm attributes, such as the cell surface adhesins, is enhanced. These surface adhesins directly contribute to the biofilm development. Conjugation The role of conjugation in biofilm development was first illustrated in the Gram-negative bacterium E. coli. Ghigo (2001) reported that natural conjugative plasmids induced bacterial biofilm development. He observed that biofilm formation was stimulated when cells harboring large conjugative plasmids were derepressed for transfer. He further demonstrated that the pilin encoding traA gene in plasmid F was essential for this stimulation. These findings suggested that conjugative pili might act as cell adhesin interconnecting the cells and stabilizing the biofilm structure in hydrodynamic biofilm systems. Plasmid-encoded factors have also been shown to enhance biofilm formation capabilities in other E. coli strains. Reisner and others (2003) further showed that the defective biofilm development by mutants malfunctioning in flagella, type 1 fimbriae, curli, and Ag43 synthesis could be complemented in trans by the function of F plasmid. In addition, F plasmid complemented the biofilm maturation defective strain DH5α which is unable to secrete the antoinducer-2 due to a frameshift mutation in luxS. Modification of the adherence factor can affect the shape and structure of the formed biofilm. These data indicate that cell–cell adherence is a key maturation and shaping factor. Because these surface adhesion factors are plasmid-encoded and are subject to conjugal gene transfer, dissemination of these plasmids among the bacterial

Mixed Culture Biofilms

115

population in fact enhances biofilm development and the stability of the biofilm structure. The role of conjugation in enhancing biofilm development was recently demonstrated in the Gram-positive bacterium L. lactis as well. Luo and others (2005) found that conjugation facilitates lactococcal biofilm development by enhancing the expression of CluA, which is a plasmid-encoded biofilm attribute. Furthermore, this enhanced biofilm-forming trait can be disseminated among lactococcal population very effectively via subsequent conjugation. A striking feature about lactococcal CluA protein is that it shares significant homology with more than 10 streptococcal aggregation factors or adhesins, many of which are chromosomally encoded. The biological significance of incorporating such adhesin into a conjugative element in lactococci itself is worth of a discussion. Likely it provided the microorganism with certain survival advantage during evolution, such as enhanced expression of the adhesin due to high copy number as well as additional uncharacterized mechanism triggered by conjugation, facilitated biofilm or aggregate formation to better withstand adverse environments, and accelerated mixed culture ecosystem development that enables further beneficial metabolic coordination and gene exchange. Quorum sensing contributes to biofilm development likely by affecting the expression of cell surface adhesins only in individual cells carrying the sensing system. Conjugation seems to be another mechanism bacteria have adapted to enhance biofilm development in a population, independent from quorum sensing. Not only do the cells carrying the plasmid express more cell surface adhesins which facilitates the biofilm or aggregation development, but the plasmid can further be transferred to the adjacent cells therefore shortly after many cells in the ecosystem will acquire and express the biofilm attribute(s). Therefore this could be a mechanism microorganisms have adapted for effective expansion of the ecosystem. A tightly bound ecosystem further provides an ideal environment for residents to share beneficial traits and coordinate metabolic activities.

Intercellular Communication Within Biofilm Communities Intercellular communication was originally perceived as a feature specific to eukaryotic organisms. However, there have been increasing reports on the presence of cell-to-cell signaling among different bacteria in recent years. The highly structured, coordinated microbial ecosystem could be regarded as protocellular organisms, since their biological processes are regulated for the benefit of the entire community (McNab and

116

Biofilms in the Food Environment

Lamont 2003). Microbial communication among constituent microorganisms through diffusible signals likely plays an important role in the coordination of metabolic and other biological activities within the ecosystem. Cooperative behavioral patterns in microorganisms have been observed for quite a long time. For instance, autoinduction of luminescence in Vibrio fisheri and Vibrio harveyi was described in the early 1970s. The swarming motility of various organisms is also exhibited as a multicellular behavior (for review, see Fuqua and others 1994, 2001). The identification of quorum sensing signals and the characterization of regulatory pathways involved revealed that microbial intercellular communication through the signals plays a key role in these coordinated activities. Quorum sensing is also involved in several other biological processes such as microbial biofilm formation, regulation of virulence gene expression, microbial cell competence development, acid tolerance, and bacteriocin production. Three distinct types of chemical compounds, including acylhomoserine lactones (AHLs) in Gram-negative bacteria; modified or unmodified peptides in Gram-positive bacteria; and the autoinducer-2 in both Gram-negative and Gram-positive bacteria, commonly serve as communication signals (for reviews, see Sturme and others 2002; Cvitkovitch and others 2003; Federle and Bassler 2003; McNab and Lamont 2003; Parsek and Greenberg 2005). Most cell-to-cell signaling systems elucidated so far involve intraspecies communication. Studies on communication among different microorganisms particularly those growing in mixed species biofilms are very limited. However, investigations on the genetic organizations of various organisms showed the likelihood of such interspecies communications. For instance, the genes encoding enzymes involved in the production of quorum sensing signals were missing in several microorganisms but many of them carry the receptor or sensing genes, possibly enabling them to engage in multispecies communication. Watnick and Kolter (2000) hypothesized that signaling systems in these multispecies biofilms would be considerably different from those observed in monoculture biofilms, and would play a significant role in determining the diversity and distribution of bacteria in these mixed culture ecosystems. Egland and coworkers (2004) provided evidence of intercellular signaling between the human oral microorganisms S. gordonii and Veillonella atypica growing in a dual-species biofilm. V. atypica was found to produce a diffusible signal that resulted in the increased expression of the α-amylase gene amyB of the other organism. S. gordonii is capable of producing lactic acid from the fermentation of either stored or exogenous carbohydrate polymers. Since V. atypica prefers lactic acid as carbon and energy source, it would largely benefit from the increased expression of the amyB of S. gordonii. The nature of the signal compound, however, is not yet known. Neither autoinducer-2 nor

Mixed Culture Biofilms

117

AHLs were detected in V. atypica cultures, suggesting that either the signaling molecule is different from known QS signals or the same signaling molecule was not detected by the bioassays used in the study. Riedel and others (2001) studied intercellular signaling in dual-species biofilms of P. aeruginosa and Burkholderia cepacia in artificial flow chambers and alginate beads in mouse lung tissues. Both of these organisms are known to utilize N-acylhomoserine lactone (AHL) as quorum sensing signals for the expression of virulence genes and for biofilm formation. Intercellular communication was found to be unidirectional in both model systems. B. cepacia was capable of responding to signals produced by P. aeruginosa, but not vice versa. Biofilms exhibit differential gene expression, as revealed by various transcriptome and proteome analyses of biofilms at different ages (Beloin and Ghigo 2005). In recent years, there has been a shift in research direction from the simple search for genes involved in the initial adhesive steps in biofilm formation to the search for late and more complex biofilm functions. This is an emerging and challenging area. More studies are required to fully understand the role of interspecies communication in biofilm developmental processes from adhesion to detachment. This knowledge could be used for the development of technologies to control biofilms of clinical or industrial importance. Safety and Public Health Risks Associated with Mixed Culture Biofilms Environmental Persistence As mentioned earlier, not all microorganisms are capable of forming biofilms in monoculture. However, almost all microorganisms can find at least a proper partner organism and become a resident of a mixed culture ecosystem. This may pose a significant health concern particularly when dealing with pathogenic microorganisms in both clinical and food processing environments. Any unsanitary conditions favoring microbial biofilm formation may increase the likelihood of pathogens being incorporated into existing biofilms, which may exhibit increased resistance to further sanitation or antibiotic treatments. Increased Resistance to Antimicrobial and Disinfecting Agents Biofilms provide a good example of how the collective strengths are greater than the sum of the individual components (Jenkinson and LappinScott 2001). One feature of biofilms that could be attributed to their multicellular nature is their markedly high resistance toward the action

118

Biofilms in the Food Environment

of biocidal agents. The increased resistance of biofilms to antibiotics and sanitizing agents has been well-documented. Microorganisms growing in a biofilm exhibit significantly less susceptibility to antibiotics and other sanitizing agents compared to their planktonic counterparts (Somers and others 2001; Donlan 2002; Petersen and others 2004; Bjarnsholt and others 2005). They also have increased resistance toward host immune system defense mechanisms (Donlan 2002; Petersen and others 2004), including the phagocytic action of polymorphonuclear leukocytes (PMNs) (Bjarnsholt and others 2005). Microbial cells in the outermost layers of a biofilm are more susceptible to the action of biocidal agents, whereas cells in the deeper layers could survive such treatments (Suchett-Kaye and others 1996). At present, there is still no clear understanding regarding how much of the characteristics of a particular biofilm are regulated by the developmental processes of its constituent organisms, and how much are controlled by its immediate environment (Kolter 2005). The increased resistance exhibited by biofilms toward biocidal agents may be attributed to greater cell density and physical exclusion of the antibiotic. The individual cells in the biofilm may also undergo physiological changes that confer greater resistance to the action of these inhibitory agents. Several other possibilities include the induction of the general stress response, which is an rpoS-dependent process in Gram-negative bacteria; increased expression of multiple drug resistance pumps; induction of quorum sensing systems; and differences in outer membrane protein profiles (Mah and O’Toole 2001). Brooun and others (2000) showed evidence that multidrug resistance pumps were important in conferring antibiotic resistance to P. aeruginosa biofilms against the antibiotic ofloxacin, using strains overexpressing and lacking the MexABOprM pump. Resistance of P. aeruginosa biofilms to ofloxacin was found to be dependent on the expression of MexAB-OprM but only in the low concentration range. Stewart (2001) cited the utilization of oxygen by outer layer of biofilm cells as another possible mechanism for their protective nature. Since some antibiotics need aerobic conditions for effective action, the depletion of oxygen protects the bacteria in the deeper layers of the biofilm. This cannot be accomplished by a planktonic cell, since its respiratory activity alone is inadequate to deplete oxygen in its immediate surrounding. Bjarnsholt and others (2005) reported that P. aeruginosa with functional quorum sensing systems was less susceptible to treatment with tobramycin and hydrogen peroxide, as well as against the phagocytic action of PMNs. On the other hand, cells in which the quorum sensing system was blocked either by mutation or by the presence of QS inhibitory drugs were sensitive to the bactericidal treatment and were readily phagocytosed by PMNs. The use of quorum sensing inhibitor drugs in combination

Mixed Culture Biofilms

119

with antibiotic treatment, as well as the action of PMNs, could possibly be used to eradicate biofilm-forming microorganisms and prevent the development of chronic infections. Horizontal Gene Transfer Horizontal gene transfer events occur in many natural ecosystems and are likely to be more efficient in biofilms than in suspended cells (Molin and Tolker-Nielsen, 2003). Several studies have shown that horizontal gene transfer occurs among microbial species in a biofilm by conjugation or transformation (Cvitkovitch and others 2003). Such horizontal gene transfer events presumably confer selective advantage by allowing microorganisms to share beneficial traits, such as antibiotic resistance genes, among the population effectively. In an earlier study, Kuramitsu and Trapa (1984) examined gene exchange among transformable strains of Streptococcus mutans, Streptococcus sanguis, and Streptococcus milleri in mixed cultures. They noted that both chromosomal and plasmid-encoded antibiotic resistance genes were readily transferred from S. mutans GS-5 to S. milleri NCTC 10707 or S. sanguis Challis during mixed growth. Gene transfer from the two organisms to strain GS-5 was not observed, but transfer from S. sanguis to S. milleri was possible. Recent studies have shown that natural transformation is another important transfer mechanism for gene exchange between streptococci in nature (Steinmoen and others 2002). Wang and others (2002) demonstrated that transformation of S. gordonii Challis occurred in both liquid and biofilm cultures, when grown with purified pKMR4PE plasmid with erythromycin resistance marker or when cocultivated with Treponema denticola with pKMR4PE. Biofilm provides an ideal environment for gene transmission. Moreover, quorum sensing signals in the biofilm can further facilitate cell competence development and the subsequent gene transfer. Li and others (2001) reported that transformation rate of S. mutans growing in biofilms was 10- to 600-fold higher than that of planktonic cells. Although it is anticipated that biofilm might have facilitated horizontal gene transfer, Luo and others (2005) showed that this contribution might be minimal. It is more likely that intrinsic mechanism(s) involved in high-frequency conjugal gene transfer also have a role in enhanced biofilm formation. Mixed Culture Biofilms and Cell-to-Cell Communications in the Food Environment Emerging evidences show that mixed culture biofilms and cell-to-cell communication may play important roles in affecting food safety and quality. Jay and others (2003) suggested that the slime layer that forms on the

120

Biofilms in the Food Environment

surface of refrigerated fresh meats during spoilage is a biofilm. Likewise, the tackiness associated with ground meats is presumably a mixed-culture biofilm. They further hypothesized that the quorum sensing is associated with the spoilage and biofilm formation in this type of food sample stored at low temperatures. In a related study, Bruhn and others (2004) demonstrated the presence of AHLs in commercial samples of vacuum-packed meat. Ninety-six AHL-producing bacteria were isolated from the samples, 92 of which were found to be members of the family Enterobacteriaceae. Hafnia alvei was found to be the most predominant AHL-producing microorganism. The authors suggested that H. alvei promotes changes in the phenotypes of other microorganisms in the food product, resulting in spoilage of the sample. In an earlier study, Flodgaard and others (2003) showed that food isolates belonging to family Enterobacteriaceae produced the same type and concentrations of AHLs when grown under food-relevant conditions and in culture media at higher temperatures. The AHLs produced in foods were also found to be relatively stable. Likewise, Christensen and others (2003) provided evidence that quorum sensing plays a role in spoilage of milk samples by Serratia proteamaculans B5a.

Control of Biofilms Developing targeted strategies to combat mixed culture biofilm contamination problems largely depends on understanding the mechanism of its formation, from initiation to detachment. (Nancharaiaha and others 2005). Several studies have shown that adhesion strategies and mechanisms are highly diverse among different microorganisms (Fletcher 1999). SuchettKaye and others (1996) suggested targeting and eliminating specific pathogens by preventing their adhesion to surfaces as a way of controlling biofilms of medical importance. Besides infection due to pathogens forming biofilms on abiotic surfaces such as the environment and implant devices, these organisms also need to attach to mucosal surfaces to initiate successful invasion of the epithelial cells of the host. Therefore, inhibiting initial adhesion could possibly counteract pathogenesis as well. Suchett-Kaye and others (1996) suggested the use of antibodies to disrupt microbial coaggregation as a possible approach for interfering with the adhesion process of oral streptococci. However, the development of these adhesion-blocking strategies would require extensive knowledge on the various antigenic structures involved in microbial coaggregation. Several studies have demonstrated that the use of antibiotics and other biocidal agents in controlling biofilms is more effective when used in combination with low levels of electric current, a phenomenon known

Mixed Culture Biofilms

121

as the bioelectric effect (Costerton and others 1994; Jass and others 1995; Suchett-Kaye and others 1996). Jass and others (1995) reported that the action of tobramycin against P. aeruginosa was markedly higher in the presence of electric current. Tobramycin alone at concentrations of 10 μg/mL had no effect on the biofilm, but its action was significantly enhanced upon application of 9 mA/cm2 electric current. The exact mechanisms of the bioelectric effect are poorly understood, but it still serves as a promising approach for controlling biofilms (SuchettKaye and others 1996). Jass and others (1995) suggested that this bioelectric effect could be due to electrophoresis, iontophoresis, and electrophoresis, thus overcoming the biofilm biomass and cell wall barriers. Other factors such as metabolic activity and growth rate of the bacteria may be important factors as well, and could be critical in optimizing antibiotic efficacy. Since evidences showed that quorum sensing are involved in the development and maturation of many microbial biofilms, the quorum sensing system could be another target for biofilm control (Suntharalingam and Cvitkovitch 2005). Any compound that could disrupt the QS system could possibly inhibit biofilm formation. Recent studies on biofilm control have focused on quorum sensing inhibitors, including furanones, acylases, and lactonases (for a review, see Smith and others 2004). Again, there could be a large spectrum of molecules in a mixed culture biofilms serving as various signals to coordinate metabolic activities. More advanced knowledge on the different types of signal molecules involved would therefore be instrumental in the development of successful biofilm control strategies.

References Amann RI, Ludwig W, Schleifer K-H. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev 59:143–169. Barlaan EA, Sugimori M, Furukawa S, Takeuchi K. 2005. Profiling and monitoring of microbial populations by denaturing high-performance liquid chromatography. J Microbiol Methods 61:399–412. Beloin C, Ghigo J-M. 2005. Finding gene-expression patterns in bacterial biofilms. Trends Microbiol 13:16–19. Bjarnsholt T, Jensen PO, Burmølle M, Hentzer M, Haagensen JA, Hougen HP, Calum H, Madsen KG, Moser C, Molin S, Hoiby N, Givskov M. 2005. Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum-sensing dependent. Microbiology 151:373–383. Brimacombe R. 1978. The sequence of E. coli 16S ribosomal RNA. Nature 276(5687):445. Brogden KA. 2002. Polymicrobial diseases of animals and humans. In Polymicrobial Diseases (KA Brogden, JM Guthmiller, Editors). ASM Press, Washington, DC, pp. 3–20.

122

Biofilms in the Food Environment

Brooun A, Liu S, Lewis K. 2000. A dose–response study of antibiotic resistance in Pseudomonas aeruginosa biofilms. Antimicrob Agents Chemother 44:640–646. Bruhn JB, Christensen AB, Flodgaard LR, Nielsen KF, Larsen TO, Givskov M, Gram L. 2004. Presence of acylated homoserine lactones (AHLs) and AHL-producing bacteria in meat and potential role of AHL in spoilage of meat. Appl Environ Microbiol 70:4293–4302. Christensen AB, Riedel K, Eberl L, Flodgaard LR, Molin S, Gram L, Givskov M. 2003. Quorumsensing-directed protein expression in Serratia proteamaculans B5a. Microbiology 149:471–483. Christensen BB, Sternberg C, Andersen JB, Eberl L, Moller S, Givskov M, Molin S. 1998. Establishment of new genetic traits in a microbial biofilm community. Appl Environ Microbiol 64:2247–2255. Costerton JW, Cheng K-J, Geesey GG, Ladd TI, Nickel JC, Dasgupta M, Marrie TJ. 1987. Bacterial biofilms in nature and disease. Annu Rev Microbiol 41:435–464. Costerton JW, Ellis B, Lam K, Johson F, Khoury AE. 1994. Mechanism of electrical enhancement of efficacy of antibiotics in killing biofilm bacteria. Antimicrob Agents Chemother 38:2803–2809. Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM. 1995. Microbial biofilms. Annu Rev Microbiol 49:711–745. Cvitkovitch DG, Li Y-H, Ellen RP. 2003. Quorum sensing and biofilm formation in streptococcal infections. J Clin Invest 112:1626–1632. Davey ME and O’Toole GA. 2000. Microbial biofilms: From ecology to molecular genetics. Microbiol Mol Biol Rev 64:847–867. 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(5361):295–298. Donlan RM. 2002. Biofilms: Microbial life on surfaces. Emerg Infect Dis 8:881–890. Egland PG, Du LD, Kolenblander PE. 2001. Identification of independent Streptococcus gordonii SspA and SspB functions in coaggregation with Actinomyces naeslundii. Infect Immun 69:512–7516. Egland PG, Palmer Jr. RJ, Kolenblander PE. 2004. Interspecies communication in Streptococcus gordonii–Veillonella atypica biofilms: Signaling in flow conditions requires juxtaposition. Proc Natl Acad Sci USA 101:16917–16922. Federle MJ, Bassler BL. 2003. Interspecies communication in bacteria. J Clin Invest 112:1291–1299. Fletcher M. 1999. Biofilms and biocorrosion. In Manual of Industrial Microbiology and Biotechnology (AL Demain, JE Davies, Editors). American Society for Microbiology, Washington, DC, pp. 704–714. Filoche SK, Anderson SA, Sissons CH. 2004. Biofilm growth of Lactobacillus species is promoted by Actinomyces species and Streptococcus mutans. Oral Microbiol Immunol 19:322–326. Flodgaard LR, Christensen AB, Molin S, Givskov M, Gram L. 2003. Influence of food preservation parameters and associated microbiota on production rate, profile and stability of acylated homoserine lactones from food-derived Enterobacteriaceae. Int J Food Microbiol 84:145–156. Foster TJ, Hook M. 1998. Surface protein adhesins of Staphylococcus aureus. Trends Microbiol 43:484–488. Fuqua C, Parsek MR, Greenberg EP. 2001. Regulation of gene expression by cell-to-cell communication: Acyl-homoserine lactone quorum sensing. Annu Rev Genet 35:439– 468. Fuqua WC, Winans SC, Greenberg EP. 1994. Quorum sensing in bacteria: The LuxR–LuxI family of cell density-responsive transcriptional regulators. J Bacteriol 176:269–275.

Mixed Culture Biofilms

123

Ghigo J-M. 2001. Natural conjugative plasmids induce bacterial biofilm development. Nature 412: 442–445. Hussain MM, Herrmann C, von Eiff Perdreau-Remington F, Peters G. 1997. A 140-kilodalton extracellular protein is essential for the accumulation of Staphylococcus epidermidis strains on surfaces. Infect Immun 65:519–524. Jass JJ, Costerton W, Lappin-Scott HM. 1995. The effect of electrical currents and tobramycin on Pseudomonas aeruginosa biofilms. J Ind Microbiol 15:234–242. Jay JM, Vilai JP, Hughes ME. 2003. Profile and activity of the bacterial biota of ground beef held from freshness to spoilage at 5–7◦ C. Int J Food Microbiol 81:105–111. Jenkinson HF, Lappin-Scott HM. 2001. Biofilms adhere to stay. Trends Microbiol 9:9–10. Kappe R, Fauser C, Okeke CN, Maiwald M. 1996. Universal fungus-specific primer systems and group-specific hybridization oligonucleotides for 18S rDNA. Mycoses 39: 25–30. Kolenbrander PE. 1988. Intergeneric coaggregation among human oral bacteria and ecology of dental plaque. Annu Rev Microbiol 42:627–656. Kolenbrander PE. 1989. Surface recognition among oral bacteria: Multigeneric coaggregations and their mediators. Crit Rev Microbiol 17:137–159. Kolenblander PE. 1997. Biofilm developmental biology. Trends Microbiol 5:475. Kolenblander PE. 2000. Oral microbial communities: Biofilms, Interactions, and Genetic Systems. Annu Rev Microbiol 54:413–437. Kolenblander PE, Egland PG, Diaz PI, Palmer Jr RJ. 2005. Genome–genome interactions: Bacterial communities in initial dental plaque. Trends Microbiol 13:11–15. Kolenblander PE, Parrish KD, Andersen RN, Greenberg EP. 1995. Intergeneric coaggregation of oral Treponema spp. with Fusobacterium spp. and intrageneric coaggregation among Fusobacterium spp. Infect Immun 63:4584–4588. Kolter R. 2005. Surfacing views of biofilm biology. Trends Microbiol 13:1–2. Kroes I, Lepp PW, Relman DA. 1999. Bacterial diversity within the human subgingival crevice. Proc Natl Acad Sci USA 96:14547–14552. Kuramitsu HK, Trapa V. 1984. Genetic exchange between oral streptococci during mixed growth. J Gen Microbiol 130:2497–2500. Kurtzman CP, Blanz PA. 1998. Ribosomal RNA/DNA sequence comparisons for assessing phylogenetic relationships. In The Yeasts: A Taxonomic Study (CP Kurtzman, JW Fell, Editors) Elsevier Science BV, Amsterdam, pp. 69–74. Laramee L, Lawrence JR, Greer CW. 2000. Molecular analysis and development of 16S rRNA oligonucleotide probes to characterize a diclofop-methyl-degrading biofilm consortium. Can J Microbiol 46:133–142. Larue R, Yu Z, Parisi VA, Egan AR, Morrison M. 2005. Novel microbial diversity adherent to plant biomass in the herbivore gastrointestinal tract, as revealed by ribosomal intergenic spacer analysis and rrs gene sequencing. Environ Microbiol 7:530–543. Li YH, Lau PC, Lee JH, Ellen RP, Cvitkovitch DG. 2001. Natural genetic transformation of Streptococcus mutans growing in biofilms. J Bacteriol 183:897–908. Li YH, Tang N, Aspiras MB, Lau PC, Lee JH, Ellen RP, Cvitkovitch DG. 2002. A quorumsensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J Bacteriol 184:2699–2708. Luo H, Wan K, Wang HH. 2005. A high frequency conjugation system facilitated biofilm formation and pAMβ1 transmission in Lactococcus lactis. Appl Environ Microbiol 71:2970– 2978. Lyautey E, Lacoste B, Ten-Hage L, Rols J-L, Garabetian F. 2005. Analysis of bacterial diversity in river biofilms using 16s rDNA PCR-DGGE: Methodological settings and fingerprints interpretation. Water Res 39:380–388.

124

Biofilms in the Food Environment

Mah T-FC, O’Toole GA. 2001. Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol 9:34–39. McNab R, Lamont RJ. 2003. Microbial dinner-party conversations: The role of LuxS in interspecies communication. J Med Microbiol 52:541–545. Molin S, Tolker-Nielsen T. 2003. Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilization of the biofilm structure. Curr Opin Biotechnol 14:255–261. Moretro T, Langsrud S. 2004. Listeria monocytogenes: Biofilm formation and persistence in food-processing environments. Biofilms 1:107–121. Murray AE, Lies D, Li G, Nealson K, Zhou J, Tiedje JM. 2001. DNA–DNA hybridization to microarrays reveals gene-specific differences between closely related microbial genomes. Proc Natl Acad Sci USA 98:9853–9858. Muyzer G, De Waal EC, Uitierlinden AG. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 59:695–700. Nancharaiaha YV, Venugopalan VP, Wuertz S, Wilderer PA, Hausner M. 2005. Compatibility of the green fluorescent protein and a general nucleic acid stain for quantitative description of a Pseudomonas putida biofilm. J Microbiol Methods 60:179–187. Olsen GJ, Lane DJ, Giovannoni SJ, Pace NR, Stahl DA. 1986. Microbial ecology and evolution: A ribosomal RNA approach. Annu Rev Microbiol 40:337–365. Parsek MR, Greenberg EP. 2005. Sociomicrobiology: The connections between quorum sensing and biofilms. Trends Microbiol 13:27–33. Peplies J, Glockner FO, Amann R. 2003. Optimization strategies for DNA microarray-based detection of bacteria with 16S rRNA-targeting oligonucleotide probes. Appl Environ Microbiol 69:1397–1407. Percival SL, Walker JT, Hunter PR. 2000. Microbiological Aspects on Biofilms and Drinking Water. CRC Press LLC, Florida. Petersen FP, Pecharki D, Scheie AA. 2004. Biofilm mode of growth of Streptococcus intermedius is favored by a competence-stimulating signaling peptide. J Bacteriol 186:6327– 6331. Reisner A, Haagensen JA, Schembri MA, Zechner EL, Molin S. 2003. Development and maturation of Escherichia coli K-12 biofilms. Mol Microbiol 48:933–946. Rickard AH, Gilbert P, High NJ, Kolenblander PE, Handley PS. 2003. Bacterial coaggregation: An integral process in the development of multi-species biofilms. Trends Microbiol 11:94–100. Riedel K, Hentzer M, Geisenberger O, Huber B, Steidle A, Wu H, Holby N, Givskov M, Molin S, Eberl L. 2001. N-Acylhomoserine-lactone-mediated communication between Pseudomonas aeruginosa and Burkholderia cepacia in mixed biofilms. Microbiology 147:3249–3262. Rudi K, Flateland SL, Hanssen JF, Bengtsson G, Nissen H. 2002. Development and evaluation of a 16S ribosomal DNA array-based approach for describing complex microbial communities in ready-to-eat vegetable salads packed in a modified atmosphere. Appl Environ Microbiol 68:1146–1156. Sharma A, Inagaki S, Sigurdson W, Kuramitsu HK. 2005. Synergy between Tannerella forsythia and Fusobacterium nucleatum in biofilm formation. Oral Microbiol Immunol 20:39–42. Smith JL, Fratamico PM, Novak JS. 2004. Quorum sensing: A primer for food microbiologists. J Food Prot 67:1053–1070.

Mixed Culture Biofilms

125

Somers EB, Johnson ME, Wong ACL. 2001. Biofilm formation and contamination of cheese by nonstarter lactic acid bacteria in the dairy environment. J Dairy Sci 84:1926–1936. Staley JT, Konopka A. 1985. Measurement of in situ activities of nonphotosynthetic microorganisms in aquatic and terrestrial habitats. Annu Rev Microbiol 39:321–346. Steinmoen H, Knutsen E, Havarstein LS. 2002. Induction of natural competence in Streptococcus pneumonia triggers lysis and DNA release from a subfraction of the population. Proc Natl Acad Sci 99:7681–7686. Stewart PS. 2001. Multicellular resistance: Biofilms. Trends Microbiol 9:204. Sturme MHJ, Kleerebezem M, Nakayama J, Akkermans ADL, Vaughan EE, de Vos WM. 2002. Cell to cell communication by autoinducing peptides in gram-positive bacteria. Antonie Van Leeuwenhoek 81:233–243. Suchett-Kaye G, Morrier J-J, Barsotti O. 1996. Unsticking bacteria: Strategies for biofilm control. Trends Microbiol 4:257–258. Suntharalingam P, Cvitkovitch DG. 2005. Quorum sensing in streptococcal biofilm formation. Trends Microbiol 13:3–6. Sutherland IW. 2001. Biofilm exopolysaccharides: A strong and sticky framework. Microbiology 147:3–9. Thurnheer T, Gmur R, Guggenheim B. 2004. Multiplex FISH analysis of a six-species bacterial biofilm. J Microbiol Methods 56:37–47. Vaudaux PE, Francois P, Proctor RA, McDevitt D, Foster TJ, Albrecht RM, Lew DP, Wabers H, Cooper SL. 1995. Use of adhesion-defective mutants of Staphylococcus aureus to define the role of specific plasma proteins in promoting bacterial adhesion to canine arteriovenous shunts. Infect Immun 63:585–590. von Wintzingerode F, Bocker S, Schlotelburg C, Chiu NHL, Storm N, Jurinke C, Cantor CR, Gobel UB, van den Boom D. 2002. Base-specific fragmentation of amplified 16S rRNA genes analyzed by mass spectrometry: A tool for rapid bacterial identification. Proc Natl Acad Sci USA 99:7039–7044. Wagner M, Loy A, Nogueira R, Purkhold U, Lee N, Daims H. 2002. Microbial community composition and function in wastewater treatment plants. Antonie Van Leeuwenhoek 81:665–680. Wan K, Yousef AE, Schwartz S, Wang HH. 2006. Rapid, specific and sensitive detection of spoilage molds in orange juice using a real-time Taqman® PCR assay. J Food Protect. 69:385–390. Wang BY, Chi B, Kuramitsu HK. 2002. Genetic exchange between Treponema denticola and Streptococcus gordonii in biofilms. Oral Microbiol Immunol 17:108–112. Ward BB. 2002. How many species of prokaryotes are there? Proc Natl Acad Sci USA 99:10234–10236. Watnick P, Kolter R. 2000. Biofilm, city of microbes. J Bacteriol 182:2675–2679. Weisburg WG, Barns SM, Pelletier DA, Lane DJ. 1991. 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173:697–703. Wimpenny J, Manz W, Szewzyk U. 2000. Heterogeneity in biofilms. FEMS Microbiol Rev 24:661–671. Woese CR. 1987. Bacterial evolution. Microbiol Rev 51:221–71.

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 6 PROKARYOTE DIVERSITY OF EPITHELIAL MUCOSAL BIOFILMS IN THE HUMAN DIGESTIVE TRACT Denis O. Krause, H. Rex Gaskins, and Roderick I. Mackie

Summary The gastrointestinal epithelium is covered by a “natural biofilm” or protective mucus gel composed predominantly of mucin glycoproteins that are synthesized and secreted by host goblet cells. Thus, the gut mucosal biofilm is dynamic and interactive with contributions from both the host and resident bacterial community. This arrangement ensures that the indigenous intestinal microbiota are in continuous and intimate contact with host tissues where they outnumber the surrounding host cells by at least an order of magnitude. The protective functions of mucus can be considered as a dynamic defensive barrier rather than a static and constitutive structure. In this chapter we define the chemical composition, distribution, and functional properties of the gut mucosal biofilm. Enzymatic digestion of the mucus coat (mucolysis) by resident microbes provides access to readily available sources of carbon and energy. At present, mucus resident bacterial populations are poorly described for all animal species but our understanding of mucosal and luminal populations has been greatly enhanced by the application of molecular microbial ecology techniques which are cultivation independent. Research using these techniques to study bacterial populations in the intestinal tract according to age, location, and compartment is also compiled and compared in this contribution.

127

128

Biofilms in the Food Environment

Introduction: Toward a Definition of Gut Mucosal Biofilms Biofilms are generally considered as a complex association of microorganisms and microbial products attached to a surface. The bacteria are arranged in a spatial groupings or consortia within the biofilm so that different species are physiologically coordinated with each other. This general view leads to the classical definition of a biofilm as “A film of microorganisms, usually embedded in extracellular polymers, which adheres to surfaces submerged in, or subjected to, aquatic environments” (Singleton and Sainsbury 1987). Adhesion to surfaces provides considerable advantages for the bacteria that live within the biofilm, including protection from antimicrobial agents and the many benefits gained by close association with other organisms such as exchange of nutrients, metabolites, and genetic material. Typically the combination of growth processes, production of metabolites, or the physical presence of the biofilm damages the surface or causes an obstruction and reduces surface efficiency and is termed biofouling and has been observed on a wide range of surfaces such as metal pipelines and corrosion, dental decay, and colonization of medical implants (Lappin-Scott and others 1992). We propose to define these biofilms as “induced” in contrast to the “natural biofilm” lining the gastrointestinal tract of humans and animals of all taxonomic orders. The gastrointestinal epithelium is covered by a protective mucus gel, composed predominantly of mucin glycoproteins, that is synthesized and secreted by host goblet cells. Thus, the gut mucosal biofilm is dynamic and interactive with contributions from both the host and resident bacterial community. This arrangement ensures that the indigenous intestinal microbiota are in continuous and intimate contact with host tissues where they outnumber the surrounding host cells by at least an order of magnitude. Indeed, much of the structure and many of the functions of the mammalian intestine seem to have evolved to enable the host to tolerate the antigenic and chemical challenges associated with the permanent carriage of a complex microbiota (Deplancke and Gaskins 2001).

The Mucus Layer and Goblet Cell Biology The mucus gel layer overlying the intestinal epithelium is the anatomical site at which the host first encounters gut bacteria. The protective functions of mucus can be considered as a dynamic defensive barrier rather than a static and constitutive structure. The mucus gel layer is an integral structural component of the intestine acting as a medium for protection,

Prokaryote Diversity of Epithelial Mucosal Biofilms

129

lubrication, and transport between the luminal contents and epithelial lining (Forstner and others 1995). The viscoelastic, polymer-like properties of mucus are derived from the major gel-forming glycoprotein components called mucins. Mucins consist of a peptide backbone containing alternating glycosylated and nonglycosylated regions comprising 70–80% of the polymer. N-Acetylglucosamine, N-acetylgalactosamine, fucose, and galactose are the four primary mucin oligosaccharides. Mucin oligosaccharide chains are often terminated with sialic acid or sulfate groups which account for the polyanionic nature of mucins at neutral pH. Oligosaccharide chains are added to mucins individually by specific, membrane-bound glycosyltransferases in the Golgi apparatus whereas sulfate is transferred to peripheral or backbone oligosaccharide chains by Golgi sulfotransferases (Paulsen and Colley 1989; Brockhausen and others 1998). The cytoarchitectural organization of goblet cells, specialized columnar epithelial cells responsible for secretion of mucins, is relatively well described but less is known of factors contributing to glycoprotein heterogeneity. A continuous mucus gel that varies in thickness covers the epithelial lining of the stomach and large intestine (Forstner and others 1995). The mucus layer can reach up to 450 μm in the stomach. In the colon, mucus thickness increases gradually from the ascending colon reaching 285 μm in the rectum. The small intestine is covered with a thinner or discontinuous mucus layer such that Peyer’s patches are apparently not covered with mucus allowing continual sampling of antigenic substances by these immunologic structures. Mucins are classified into neutral or acidic subtypes with acidomucins further differentiated into sulfated (sulfomucin) or nonsulfated (sialomucin) types. Neutral mucins appear to be the predominant subtype expressed in gastric mucosa whereas acidic mucins are expressed throughout the intestinal epithelium and dominate in the large intestine (Sheahan and Jervis 1976). Mucin subtype and goblet cell distribution vary spatially throughout the gastrointestinal tract as well as temporally during postnatal development in all mammalian species studied (Deplancke and Gaskins 2001). The physiologic relevance of distinct mucin subtypes is not well understood but it is suggested that acidic mucins protect against bacterial translocation because sulfated mucins, in particular, appear less degradable by bacterial glycosidases and proteases (Roberton and Wright 1997; Fontaine and others 1996). This concept is consistent with the observation that goblet cells in intestinal compartments densely populated with bacteria express acidic mucins predominantly (Roberton and Wright 1997; Nieuw Amerongen and others 1998; Deplancke and others 2000). Our knowledge of the mucus gel and particularly of bacterial populations that normally or abnormally colonize intestinal mucus is limited

130

Biofilms in the Food Environment

because fixation of intestinal tissues with conventional, dehydrating, cross-linking aldehyde fixatives results in gross morphological alterations as well as detachment and even loss of mucus. Matsuo and others (1997) clearly showed that the use of Carnoy’s fixative, an ethanol and acetic acid based solution, enabled preservation of surface mucus in paraffin sections of human colonic samples. Two distinct layers were identified using conventional mucin histochemical stains with the inner layer attached to the apical epithelial surface and continuous with intra-crypt mucus. Colonic mucins did not mix homogeneously but rather formed a stacked or laminated structure of alternating sialo- and sulfomucins. Bacteria were consistently observed within the laminated arrays of the outer layer, indicating the importance of the mucus gel in preventing direct adherence of even commensal gut bacteria to colonic epithelial cells. A recent paper studying bacterial populations in ileal and colonic biopsy samples with FISH also demonstrates that bacteria are not attached to intestinal epithelial cells but found predominantly attached to the luminal side of the intestinal mucus layer (van der Waaij and others 2005). The photomicrographs presented in this paper clearly show bacteria closely associated with the mucus layer, which seems in conflict with their conclusion that “intestinal commensal bacteria live in suspension in the lumen and that there is no specific mucus-adherent microflora” (van der Waaij and others 2005). By snap-freezing the biopsy sections, the mucus layer did appear to be preserved intact; however, the tissue sections used for FISH analysis were fixed with an aldehyde-based fixative bringing into question the extent to which the fluorescently labeled, 16S rRNA oligonucleotide probes penetrated the mucus layer. Although these technical challenges must be overcome before final conclusions can be reached regarding where precisely bacteria reside in the intestine, there is considerable published data and compelling growth considerations that support bacterial exploitation of the mucus niche, an issue with important implications for many human and animal intestinal disorders.

The Mucus Biofilm Niche Mucus offers numerous ecologic advantages to intestinal bacteria. For example, mucin oligosaccharides represent a direct source of carbohydrates and peptides. Also, exogenous nutrients including vitamins and minerals are likely concentrated within the mucus matrix. Bacteria capable of colonizing mucus can avoid rapid expulsion via the hydrokinetic properties of the intestine and bacterial colonization of mucus would thus impart

Prokaryote Diversity of Epithelial Mucosal Biofilms

131

a growth advantage relative to those in the lumen. In classical biofilm terminology, these two populations are referred to as sessile (bacteria living within the biofilm) and planktonic (free-floating bacteria living in the aqueous phase and not associated with a biofilm). Advantages of the sessile or adherent growth mode are listed below: 1. increased availability of nutrients for growth 2. protection from antimicrobial agents produced by the host or administered exogenously 3. proximity to other bacteria, both closely and distantly related, facilitating transfer of DNA either as plasmids or transposons (conjugation), phage (transduction), or naked DNA (transformation) 4. greater phenotypic plasticity by means of horizontal gene transfer 5. gradation of metabolic activity 6. establishment of complex microbial consortia which enhance advantages listed above In fact, it is difficult to envision a more suitable bacterial niche than host mucus. In this regard, it is not surprising that mucus secretion is typically enhanced in response to intestinal microbes (Deplancke and Gaskins 2001). Both commensal and pathogenic bacteria would derive significant benefit from an ability to regulate mucus synthesis or secretion from host goblet cells. However, our current knowledge of the biochemical basis of goblet cell responsiveness to microbial products is limited and largely restricted to pathogens and their toxins. The most convincing evidence that the intestinal microbiota alters mucin composition comes from histochemical studies in which germ-free animals were compared with conventionally reared controls or were inoculated with mixed bacterial populations. Goblet cells are fewer in number and smaller in size than those of conventionally raised mice (Kandori and others 1996; Ishikawa and others 1989). As a result, the mucus layer may be up to twice as thick in conventionally raised as in germ-free rodents indicating greater mucus production (Szentkuti and others 1990; Enss and others 1992; Kandori and others 1996; Meslin and others 1999). A striking feature of germ-free rodents is pronounced cecal enlargement due to the water retaining capacity of mucins. This morphologic difference is thought to reflect the absence of mucus-degrading intestinal bacteria (Deplancke and Gaskins 2001). Indeed, cecal mucins are rapidly degraded and cecal morphology rapidly normalizes with the introduction of gut bacteria (Gustaffson and CarlstedtDuke 1984). Mucin composition also differs significantly between germfree and conventionally raised animals (Deplancke and Gaskins 2001).

132

Biofilms in the Food Environment

Microbial Mucolysis An ability to enzymatically degrade mucus was documented in pathogens and commensal alike and appears to be a common trait among bacteria (Hoskins and Boulding 1981; Corfield and others 1992; Deplancke and others 2002). Enzymatic digestion of the mucus coat provides readily available sources of carbon and energy and enables access to the epithelial surface. Mucin degradation is a complex process that it not well understood, but is thought to begin with proteolysis of nonglycosylated “naked” regions of mucin glycoproteins by host and microbial proteases (Quigley and Kelly 1995). This initial step markedly reduces mucin gelation and viscosity and results in accumulation of highly glycosylated subunits that are resistant to further proteolytic attack. Mucin glycopeptides are then degraded by various bacterial enzymes specified by the complexity of the oligosaccharide chains which differ in size, degree of branching, type of linkage, and the presence of terminal sulfate or sialic groups. Thus, oligosaccharide side chains are degraded by linkage-specific glycosidases whereas terminal sialic acid or sulfate groups are cleaved by bacterial sialidases and glycosulfatases (Corfield and others 1995). Also of interest is evidence from humans that resident mucolytic bacteria may differ among individuals according to the specific carbohydrate composition of intestinal mucins, such as terminal sugars and branching patterns, which appear to vary by genetic background (Hoskins and others 1985; Hoskins 1992). Evidence of host genetic background influencing bacterial community profiles has also been reported for other mammalian species (Deplancke and others 2000; Simpson and others 1999, 2000, 2002). This is consistent with increasing evidence of stable and host-specific bacterial community profiles (McCartney and others 1996; Zoetendal and others 1998; Simpson and others 1999, 2000, 2002; Toivanen and others 2001; Vaahtovuo and others 2003) and with evidence that endogenous substrates may have a significant influence on the spatial pattern of bacterial population profiles along the gastrointestinal tract (Deplancke and others 2000). Thus the taxonomy, as well as the temporal and spatial distributions, of bacterial groups that preferentially reside within the intestinal mucus must be defined to determine the role of indigenous gut bacteria in mucogenesis and mucolysis. At present, mucus resident bacterial populations are poorly described for all animal species. This limitation derives from the inherent bias of cultivation-dependent microbiological techniques (Mackie and others 1999; Zoetendal and others 2004) as well as the difficulties associated with preserving the mucus layer during tissue fixation (Matsuo and others 1997). Our current understanding of mucosal and luminal

Prokaryote Diversity of Epithelial Mucosal Biofilms

133

populations has been greatly enhanced by the application of molecular microbial ecology techniques which are cultivation independent and are described in the following section.

Bacterial Populations in the Intestinal Tract Traditionally, our interest in the gut microbiota has been restricted either to microbial populations associated with the nutritional physiology of the animal (e.g., ruminants) or to pathogens that cause various forms of gastritis. Even though more than 100 years ago Eli Metchnikoff suggested that the lactic acid bacteria of the digestive tract may have a role in preventing infection (Tauber 2003), little had been done to develop this area of gut microbiology until relatively recently. Early pioneers in this area include R´ene Dubos, Dwane Savage, W. E. C. Moore, L. V. Holdeman, and R. Freter, to name a few, who collectively recognized the importance of the “nonpathogenic” members of the digestive tract in relation to homeostasis of the gut compartment (Savage 1972, 2001). The early studies carried out in this area were generally confounded by the inability to culture the majority of the bacteria in the digestive tract. To try and resolve these problems, studies were carried out with gnotobiotic animals (Gunzer and others 2003). Maintenance of gnotobiotic animals is expensive and rodents have been used most extensively for germ-free and gnotobiotic studies, although a number of useful studies were conducted with the chick (Coates 1986) and some gnotobiotic ruminant studies have been described (Chaucheyras-Durand and Fonty 2001). Gnotobiotic studies have demonstrated that the role of microorganisms on gut function is based on a complex interplay between different species. In an ecological context, these can be seen as either mutualistic (an association between organisms of two different species in which each member benefits) or synergistic (the interaction of two or more organisms so that their combined effect is greater than the sum of their individual effects) (Allen and Banfield 2005). These earlier studies with gnotobiotic rodents clearly indicated the existence of unique microbial associations with the gut wall. Savage (1969a) demonstrated that in some gnotobiotic mice, yeast and lactobacilli populated the secreting stomach of mice in thick layers. However, if penicillin were administered to the mice, the yeasts would dominate and the lactobacilli declined. This situation was reversed when the penicillin was removed from the drinking water. Subsequent studies (Savage 1969b) demonstrated that lactobacilli specifically adhered to the villus of the small intestine.

134

Biofilms in the Food Environment

One of the problems with gnotobiotic rodent models is that typically only monoassociation of the digestive tract with a single bacterium is carried out. This of course removes the complex mutualistic and synergistic associations between microorganisms of the wall of the digestive tract. In 1965, Schaedler and others (1965) colonized germ-free mice with selected bacteria isolated from normal mice. These defined bacteria included aerobic bacteria that were easy to grow and a number of facultative anaerobes. Obligate anaerobes or, as Schaedler called them, extremely oxygen-sensitive (EOS), fusiform bacteria were not included in this cocktail of bacteria (Lee and others 1968, 1971) until appropriate anaerobic techniques had been developed by Hungate (1950) and Bryant (1972). Of the defined bacteria later used in gnotobiotic studies, known as the “Schaedler flora,” eight bacteria were included: Escherichia coli var. mutabilis; Streptococcus faecalis; Lactobacillus acidophilus; Lactobacillus salivarius; group N Streptococcus; Bacteroides distasonis; a Clostridium sp.; and an anaerobic fusiform bacterium (Sarma-Rupavtarm and others 2004). Biofilms of the digestive tract are made up of more than a dozen bacteria as in the Schaedler flora. Similarly, the complexity of various mutualistic and synergistic interactions between bacteria and the host cannot be reduced to studies in gnotobiotic animals. It is now possible to define in far greater detail the composition of the adherent microbiota through the generation of 16S rDNA clone libraries (Eckburg and others 2005; Swidinski and others 2005; Rapp´e and Giovannoni 2004). The studies using ribosomal phylogenies do not impart any function to the clones, but this is now being made possible with the application of metagenomic techniques that allow the reconstruction of genomes from uncultured species (Allen and Banfield 2005). The inventories of the genes present in the metagenomes will give insight into the functional attributes of the adherent gut microbiota. Cultured and Uncultured Representatives The extent of diversity of the gut microbial ecosystem, including those microbes adherent to the wall of the digestive tract, is a fundamental question in gut microbial ecology. Significant progress has been made in this area particularly because of the use of ribosomal genes to assess the uncultured microbiota of the luminal and biofilms of the digestive tract. To help update progress so far, we have interrogated publicly accessible databases of ribosomal sequences. Any analysis of databases is flawed because of the large accumulation of sequences, peer-reviewed and not, and it is limited

Prokaryote Diversity of Epithelial Mucosal Biofilms

135

by the annotations of the sequences. Nevertheless, given the amount of sequence information available, it remains a valuable approach to assess the current status of the evolving picture of gut microbial ecology. The majority of 16S rDNA sequences in the Ribosomal Database Project (RDP; Cole and others 2005) are those found in Genbank. An added benefit is that the sequences in the RDP have been curated and conform to the official taxonomic guidelines outlined in Bergey’s Manual (Garrity and others 2004). We have thus limited our analysis to only those sequences in the RDP. Currently (7 October 2005) there are 169,170 sequence entries in the RDP (Figure 6.1). Of these 33,790, or 20.0%, are from cultured isolates. Some of the largest sequence-based studies have in fact concentrated on the biofilms of the digestive tract (Eckburg and others 2005), and we thus have sufficient information to draw some broad conclusions. The analysis of Rapp´e and Giovannoni (2004) indicates that 52 phyla can be discerned, of which 26 are candidate phyla. Currently, Bergey’s Manual (Garrity and others 2004) does not recognize all these phyla. Use of the search term “gut” in the RDP hierarchical search engine generated 12 recognized phyla that had either cultured or uncultured bacterial members. The dominant bacterial phyla in the gut (Figure 6.2) are the Bacteriodetes, Firmicutes, and the Proteobacteria. In each of these dominant groups, the uncultured to cultured ratios are 1:75 (Bacteriodetes), 1:27 (Firmicutes), and 1:12 (Proteobacteria). There are also some underrepresented phyla: Cyanobacteria, Deferribacteres, Fusobacteria, Planctomyces, and Verrucomicrobia, which have few or no cultured representatives. Microbiota of the Digestive Tract Bacterial diversity of gut biofilms is best viewed in the context of 16S rDNA phylogeny. Woese (1987) originally proposed 12 taxa based on oligonucleotide cataloguing. Indeed, a large proportion of these original taxa are represented in the digestive tract (Backhed and others 2005). This was largely due to the fact that many of the cultured bacteria in collections were pathogens and fell within the Bacteroides, Proteobacteria, or Firmicute groupings (Backhed and others 2005; Eckburg and others 2005; Swidinski and others 2005; Savage 1972). The Proteobacteria, in particular, are highly represented in culture collections, are Gram-negative, and generally recognized as one of the most successful microbial groups on the planet Earth (White and others 2004; Hornef and others 2005). This phylum includes well-studied genera like Escherichia, which form biofilms in the digestive tract (Galvez and others 1998).

0

753

20,000

40,000

33,790

60,000 Numbers

80,000

100,000

120,000

140,000

135,380

160,000

Figure 6.1. Graph depicting the number of 16S rRNA gene sequences currently deposited in the RDP database. The analysis compares the total number of sequences with those derived from gut ecosystems and the number of sequences obtained from cultured isolates. Currently (7 October 2005) there are 169,170 sequence entries and 33,790 of these (20.0%) are from cultured isolates. Some of the largest sequence based studies have in fact concentrated on the biofilm of the digestive tract and 10.1% of total sequences are gut derived.

All

Gut

16,333

Cultured

Uncultured

2 0

Deferribacteres

0

8 0

Cyanobacteria

19 0

Planctomycetes

25

Unclassified

340

207

261

214

1000

684

2000

2363

2226

Number

3000

4000

5000

6000

5853

Gut uncultured Gut isolates

Figure 6.2. Graph representing phylogenetic affiliation of 16S rRNA gene sequences from gut ecosystems deposited in the RDP database. The analysis compares the number of sequences obtained from cultured to uncultured bacteria within each phylum. Bacteria in the gut are not found in all 26 recognized phyla but are restricted to 12 named divisions, some with very few deposited sequences. The dominant phyla are the Bacteriodetes, Firmicutes, and the Proteobacteria.

4 0

24 0

TM7

Verrucomicrobia

29

Spirochaetes

Proteobacteria

20 0

Fusobacteria

Firmicutes

24 0

33

Fibrobacters

70

Bacteroidetes

2 3

Actinobacteria

Acid obacteria

s

138

Biofilms in the Food Environment

The Gram-positive bacteria were originally defined as one phylum but are now divided into two phyla (Rapp´e and Giovannoni 2004), the Firmicutes (low G + C) and the Actinobacteria (high G + C), both of which are represented in gut biofilms, with the Firmicutes being one of the most dominant phyla in the digestive tract (Eckburg and others 2005; Ley and others 2004). The Actinobacteria contain some well-known human pathogens, and some, like the Mycobacterium avium complex, are found in the gut (Martins and others 2005; Hermon-Taylor and others 2000). Interestingly, bacteria such as Mycobacterium are underrepresented in clone libraries of the gut, but well represented in culture collections (Eckburg and others 2005; Ley and others 2005). Biofilm formers in this genus are M. paratuberculosis that invade the gut wall and are the causative agent of Johne’s disease in ruminants (Li and others 2005), and have also been isolated from gut biopsy tissue of Crohn’s patients (Hermon-Taylor and others 2000). Of the original 12 phyla, the genera Cytophaga, Bacteroides, and Flavobacterium formed a major lineage (Paster and others 1985), known now as the Bacteroidetes phylum (Boone and others 2001), which are foremost representatives of the intestinal tract. No members of the phyla Thermotogae, Chloroflexi, Chlorobi, or Deinococcus-Thermus have been found in gut ecosystems. According to Rapp´e and Giovannoni (2004), an additional 14 phyla with cultivated representatives have been identified since 1987, but not all of these are recognized by Bergey’s Manual. This group includes Aquificae (in Bergey’s Manual), Thermodesulfobacteria (in Bergey’s), Dictyoglomi (in Bergey’s), Nitrospira (in Bergey’s), Coprothermobacteria (not in Bergey’s), Caldithrix (not in Bergey’s), and Desulfurobacteria (not in Bergey’s). None of these phyla have yet been identified in the digestive tract. However, phyla with cultured representatives that have been found in the digestive tract since 1987 include the Acidobacteria and Fibrobacters. Acidobacteria (Rapp´e and Giovannoni 2004) is apparently ubiquitous and abundant in nature, but has thus far only been identified in the termite gut (Stevenson and others 2004), and not that of higher mammals or humans. Fibrobacter was also identified as a separate pylum since 1987 but is noteworthy because it is highly abundant in the digestive tract of herbivores (reviewed by Krause and others 2003), but is present in clone libraries in the human digestive tract (Figure 6.2). Fibrobacter was previously classified as Bacteriodes, but based on 16S rRNA sequence information could be divided into two genera, F. succinogenes and F. intestinalis. F. succinogenes is a classic example of a biofilm-forming organism that densely covers the surface of plant material in the rumen and, together with Ruminococcus albus and R. flavefaciens (members of the phylum

Prokaryote Diversity of Epithelial Mucosal Biofilms

139

Firmicutes), actively degrades cellulose in a mutualistic association with the host animal (reviewed by Krause and others 2003). Members of Underrepresented Phyla Associated with Intestinal Tract Biofilms With the ability to extensively sequence rRNA genes, and with the more recent occurrence of high throughput sequencing studies of gut microbiota, a great deal can now be concluded about the diversity of biofilm composition in the human gut, and also about the distribution of species across gut ecosystems. Clearly, there are some fascinating observations that can be made about the allotment of species and phyla represented in the gut in comparison to other ecosystems. Unfortunately, it is beyond the scope of this chapter to address all these observations and we will restrict ourselves to some specific cases pertinent to underrepresented phyla. Clone library studies identify a number of phyla that are present in the human gut but which are not represented by isolates, and these include Cyanobacteria, Deferribacteres, Fibrobacters, Fusobacteria, Planctomycetes, and Verrucomicrobia. Members of the Cyanobacteria, Acidobacteria, and Planctomyctes are gut associated but apparently do not reside in mammalian digestive tracts. However, they were identified with our search criteria because they are gut bacteria. Cyanobacteria were identified in a marine bivalve (Blankenship and others 2004), Acidobacteria in termites (see RDP), and Planctomyctes seem to be associated with both marine bivalves and termites (see RDP). Fibrobacters are found in abundance in ruminants but infrequently in the human digestive tract, and Verrucomicrobia (in pigs) and Deferribacteres (in humans) are both associated with the nonruminant digestive tract. In addition to poorly represented phyla, there are “Candidate phyla” that can be defined as phyla that have deeply diverging clusters of sequences equivalent to those of recognized phyla. Additionally, there are no cultured representatives of these phyla and they are typically not recognized by Bergey’s Manual. From our analysis, it can be seen that at least one of these phyla, TM7, is represented in the digestive tract. Presently, clone library studies are not extensive enough to determine if they are ubiquitous in a variety of gut ecosystems. One of the issues that must be addressed with phyla that are in low abundance in biofilms of the gut, or the gut in general, is whether they are indeed true members of the gut ecosystem. This issue arises from the fact that the digestive tract is open, and animals and humans are continually inoculating themselves, usually from food sources with microorganisms

140

Biofilms in the Food Environment

that pass through the gut. Some of these microorganisms may in fact be normal members of the gut population, while others may be contaminants from substrates such as grass or soil. These questions will only be resolved as more sequence information becomes available. Successional Colonization of the Digestive Tract The formation of biofilms in the digestive tract begins at birth. During the birth process, and soon thereafter, bacterial colonization of a previously germ-free infant intestinal tract begins. In general, the gut is initially colonized by facultative aerobic bacteria, probably by inoculation from the environment, and is composed of bacteria such as streptococci, coliforms, and lactobacilli. As the infant animal becomes older, and approaches adulthood, a more anaerobic microbiota is established, composed predominantly of members of the phyla Bacteroidetes, Firmicutes, and Proteobacteria. Initial studies on infant microbiota succession were carried out by Tissier (1900), and focused largely on aerobic bacteria of stool samples. The succession and composition of the microbiota are significantly influenced by the diet of the infant (Cooperstock and Zedd 1983; Dai and Walker 1999; Mackie and others 1999). For example, in full-term naturally delivered babies, colonization is initiated at birth and enterobacteria and streptococci appear in the feces within a few days. In breast-fed infants the lactic acid bacterium, Bifidobacterium, increases rapidly in abundance, but its numbers decline, and bacterial diversity increases, only when the infants start consuming a diet with more varied substrates. Formula-fed infants have a more diverse microbiota consisting of Streptococcus, Bacteroides, and Clostridium, in addition to members of the genus Bifidobacterium. Molecular-based studies have to a large extent confirmed these earlier culture-based investigations, but with a higher degree of taxonomic resolution. Recently, denaturing gradient gel electrophoresis (DGGE) of infant fecal microbiota demonstrated that initially the microbiota is relatively simple and increases in complexity as the infant becomes older (Favier and others 2002). Clone libraries of amplified 16S rDNA fragments from infant feces indicate that the predominant genera were Bifidobacterium, Ruminococcus, Enterococcus, Clostridium, and Enterobacter. Species most closely related to the genera Bifidobacterium, and Ruminococcus in particular, dominate the intestinal microbiota and form a stable community. Studies using DGGE in preterm infants (Schwiertz and others 2003) largely confirmed the results by Favier and others (2002), but within individual variation was much lower in the preterm than in full-term breast-fed

Prokaryote Diversity of Epithelial Mucosal Biofilms

141

infants, suggesting that environment is an important arbiter of initial composition. Once a climax community has been reached it appears to be relatively stable in the normal healthy individual. Zoetendal and others (1998) monitored the composition of the fecal microbiota of two normal healthy adult volunteers over a period of 6 months using temperature gradient gel electrophoresis. Each individual appeared to have a unique microbiota that was stable over time. Given that each individual differed by diet and age, and they were genetically unrelated, it can be hypothesized that host genetic factors have a significant effect on composition of the gut microbiota. To explore the effect of host genetic relationships on fecal microbiota composition, Zoetendal and others (2001) assessed the DGGE profiles of homozygotic twins and found them to have a higher degree of similarity than unrelated individuals. In addition, they demonstrated that the fecal microbiota profiles of spouses, sharing similar diets, and environment, were not more similar than unrelated individuals living separately. The concept of genetic predisposition can be carried further to include specific genetic predispositions. Diseases that can be traced to specific gene defects may reduce the complexity of the gut microbiota. A recent study (Ley and others 2005) analyzed 5,088 bacterial 16S rRNA gene sequences from the distal intestinal contents of genetically obese ob/ob mice, lean ob/+ and wild-type siblings, and their ob/+ mothers. Microbial-community composition could be shown to be an inherited trait, and obese mice, regardless of kinship, exhibited a 50% reduction in the abundance of Bacteroidetes and a proportional increase in Firmicutes. In contrast, Crohn’s disease, a chronic intestinal inflammation of the human digestive tract, can be explained in some individuals by the NOD2 mutation, which is expressed intracellularly in enterocytes (Abreu 2005). However, the occurrence of the single mutation did not correlate with a decrease in intestinal tract biofilm diversity when compared to normals (Ott and others, 2004). On the other hand, in swine, selection against a SNP representing an adherence factor for E. coli K88+ (Jorgensen and others 2003) can dramatically reduce the adherence of K88+ E. coli (Bertels and others 1997). Stool Versus Mucosal Populations A key question is whether the microbial composition of the gut biofilm is fundamentally different from that of the lumen. A recent study by Eckburg and others (2005) conducted a 16S rDNA census of three normal human subjects with biopsy tissue from the sigmoid colon, cecum, rectum, and stool. In total, 13,355 sequences, representing 395 bacterial phylotypes

142

Biofilms in the Food Environment

(distinguished at the 99% similarity level), were distinguished. A single archaeal species was identified. There were differences in the phylotype composition by site, and between subjects, but it appeared that the stool phylotypes were a composite of the mucosal populations. Zoetendal and others (2002) assessed the bacterial communities in feces and biopsy samples from the ascending, transverse, and descending colons of 10 subjects using DGGE. Surprisingly no significant differences were observed between healthy and diseased individuals, given that some of these diseased subjects had ulcerative colitis in which diversity typically declines (Ott and others 2004). Host-specific DGGE profiles of the mucosa-associated bacterial community in the colon support the hypothesis that genetic predisposition is important in determining the microbial composition. Zoetendal and others (2002) found very little difference in the microbial diversity between sites, which is in contrast to that the findings of Ley and others (2005) but not of Nielsen and others (2003). Using a slightly different approach based on 16S rDNA RFLP analysis, Pryde and others (1999) demonstrated that the lumen of the colon and cecum, and cecum wall were largely similar although there appeared to be differences in the Streptococcus and Ruminococcus species abundance. Terminal restriction length polymorphism (T-RFLP) is becoming a robust method for analyzing community structure and has a bioinformatic advantage over techniques like DGGE (Liu and others 1997). T-RFLP is based on restriction digestion of 16S rDNA amplicons with one fluorescently labeled primer. In theory, every species has a different terminal fragment length, which can be rapidly resolved on a high-throughput sequencing instrument. The fragments can then be compared to databases like the RDP, which currently contains over 200,000 entries, and species annotation can be attained without sequencing. Recently, Hayashi and others (2005) demonstrated species heterogeneity between the jejunal, ileal, cecal, and recto-sigmoidal sections of the mucosa of elderly human cadavers. The largest differences occurred within the low G + C Gram-positive bacteria. We have recently demonstrated that there are significant differences in mucosal microbiota between sites within the same patient and within patients with Crohn’s disease (Figure 6.3), but the overall similarity among the healthy and Crohn’s groups was greater than between-group variation (Kotlowski and others 2006). Taken together, the studies cited indicate that there are some differences in composition of gut microbiota when the different regions of the digestive tract are compared, but these differences are not large. Furthermore, the composition of the lumen or stool is largely a reflection of the mucosal population and is a product of sloughed off epithelial cells. A far more important factor is the genotype of the host. One critical aspect that

0

0

100

150

200

250

300

Size (nt)

350

400

450

500

550

600

Figure 6.3. DNA fingerprinting techniques provide an index of total bacterial diversity from different gut locations. This figure presents a T-RFLP chromatogram of biopsy rDNA extracted from the terminal ileum (a) and the rectum (b) of a Crohn’s disease patient. PCR amplicons were obtained by amplifying rDNA with conserved primers, 27f and 342r, and digesting the product with the restriction enzyme HahI. Terminal fragments which differ in length and intensity (peak height) demonstrate qualitative and quantitative differences in bacterial populations between the ileum and rectum.

5

b

(b)

a

(a)

144

Biofilms in the Food Environment

has not been addressed is whether the composition of mucosal associated populations changes over time. In humans, and to some extent in all animal studies, this is a difficult experiment to conduct because the same site, and the same individual, must be subjected to repeated biopsies. Having said this, it is of course also true that all the cited studies have been conducted using 16S rDNA as a phylogenetic marker, and it is well known that 16S rDNA is not necessarily an indicator of function. Thus, even though the overall composition of the lumen and gut microbial biofilm may be similar, this does not mean that they are functionally identical. Bacterial Growth Rate in Situ The classical view of growth rate in the intestine is that resident bacteria grow at a rate sufficient to resist being washed out by intestinal peristaltic movements and fluid flow, but without bacterial overgrowth. Accurate calculations of the rates of bacterial proliferation in the intestine have so far been represented only by average estimates at the level of populations. For example, the growth rate of E. coli has been estimated in vivo with generation times of 30 min to 40 h (Gibbons and Kapsimalis 1967; Freter and others 1983a; Miller and Wolin 1981). Also, continuous-flow cultures have been developed to mimic bacterial interactions in the gut and estimate overall growth rates (Miller and Wolin 1981; Freter and others 1983b). These systems, however, do not reflect the physiological conditions in the gut, where entrapping of the bacteria in the mucus gel plays an important role. Recently developed methods based on hybridization to whole cells with fluorophore-labeled oligonucleotide probes targeting the rRNA, and epifluorescence microscopy coupled to digital image analysis, allow estimations of the concentration of rRNA in single cells. The ribosomal contents of the bacteria isolated from the environment may then be correlated with the ribosomal contents of bacteria growing at defined rates (DeLong and others 1989; Poulsen and others 1993). Using the streptomycin-treated mouse as the animal model, growth rates of E. coli BJ4 colonizing the large intestine of mice were estimated by quantitative hybridization with rRNA target probes and by epifluorescence microscopy (Poulsen and others 1995). The ribosomal contents in bacteria isolated from the cecal mucus, cecal contents, and feces were measured and correlated with the ribosomal contents of bacteria growing in vitro at defined rates. The data suggest that E. coli BJ4 grows at an overall high rate in the mouse intestine. However, when taking into account the total intestinal volume and numbers of bacteria present in cecal mucus, cecal contents, and feces, and factors such as washout, mucosal turnover, and release of bacteria from the mucus layer, Poulsen and others (1995) suggest

Prokaryote Diversity of Epithelial Mucosal Biofilms

145

that E. coli BJ4 in the intestine consists of two populations, one situated in the mucus layer, which has an apparent generation time of 40 to 80 min, and one sloughed into the luminal contents and feces, which contributes little to growth. These conclusions may be true only for streptomycintreated mice, and future experiments with germ-free and conventional mice need to be carried out. The reason why bacteria resident in the gut mucus gel grow faster than those sloughed into the intestine has not been studied, but most likely involves higher nutrient concentrations and other advantages of growing in a consortium listed previously. Many intestinal microorganisms use mucins as carbon, nitrogen, and energy sources (Macfarlane and Gibson 1991; Macfarlane and others 1992, 2005). These molecules are likely to be important sources of carbohydrate for saccharolytic bacteria growing in the large intestine, particularly in the distal bowel, where the supply of fermentable carbohydrate is usually limiting (Macfarlane and others 1992). The implications of these combined findings are far-reaching since it means that the “true” resident gut population must be studied at the level of the mucosal biofilm and that luminal and fecal populations are inadequate for studying microbial ecology of the gut.

Conclusion The gut mucosal biofilm can be considered as a dynamic defensive barrier with contributions from both the host and resident bacterial community. This arrangement ensures that the indigenous intestinal microbiota are in continuous and intimate contact with host tissues where they outnumber the surrounding host cells by at least an order of magnitude. The mucus gel layer overlying the intestinal epithelium is the anatomical site at which the host first encounters gut bacteria. Indeed, much of the structure and many of the functions of the mammalian intestine seem to have evolved to enable the host to tolerate the antigenic and chemical challenges associated with the permanent carriage of a complex microbiota. Despite its importance, our knowledge of the mucus gel and particularly of bacterial populations that normally or abnormally colonize intestinal mucus is limited. This limitation derives from the inherent bias of cultivation-dependent microbiological techniques, as well as the difficulties associated with preserving the mucus layer during tissue fixation. Current understanding of mucosal and luminal populations has been greatly enhanced by the application of molecular microbial ecology techniques which are cultivation independent and are described in this contribution with emphasis on bacteria which are currently unculturable and greatly

146

Biofilms in the Food Environment

underestimated. We predict a renaissance of interest and research activity focusing on interactions between the host and its resident microbiota. This resurgence of activity will be largely based on new “omics” technologies that enable complex interacting and signaling pathways to be elucidated and contribute to our understanding of complex mutual interactions as well as mechanisms controlling immunologic, inflammatory and allergic reactions.

References Abreu MT. 2005. Nod2 in normal and abnormal intestinal immune function. Gastroenterology 129:1302–1314. Allen EE, Banfield JF. 2005. Community genomics in microbial ecology and evolution. Nat Rev Microbiol 3:489–498. Backhed F, Ley RE, Sonnenburg JL, Peterson DA, Gordon JI. 2005. Host-bacterial mutualism in the human intestine. Science 307:1915–1920. Bertels A, Jourquin J, Buys N, Van Zeveren A, De Greve H, Hernalsteens JP, Bouquet Y. 1997. Selection of swine resistant to F4-positive Escherichia coli. Adv Exp Med Biol 412:427–439. Blankenship LE, Duplessis MR, Dufour SC, Yayanos AA, Felbeck H. 2004. Anatomical and experimental evidence for particulate feeding in Lucinoma aequizonata and Parvilucina tenuisculpta (Bivalvia: Lucinidae) from the Santa Barbara Basin. Genbank Accession Number AY536229. Boone DR, Castenholz RW, Garrity GM, Editors. 2001. Bergey’s Manual of Systematic Bacteriology, Vol. 1. Springer, New York. Brockhausen I, Schutzbach J, Kuhns W. 1998. Glycoproteins and their relationship to human disease. Acta Anat (Basel) 161:36–78. Bryant MP. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria. Am J Clin Nutr 25:1324–1328. Chaucheyras-Dur F, Fonty G. 2001. Establishment of cellulolytic bacteria and development of fermentative activities in the rumen of gnotobiotically-reared lambs receiving the microbial additive Saccharomyces cerevisiae CNCM I-1077. Reprod Nutr Dev 41:57– 68. Coates ME. 1986. Gordon memorial lecture. The biologists’ debt to the domestic fowl. Br Poult Sci 27:3–10. Cole JR, Chai B, Farris RJ, Wang Q, Kulam SA, McGarrell DM, Garrity GM, Tiedje JM. 2005. The Ribosomal Database Project (RDP-II): sequences and tools for high-throughput rRNA analysis. Nucl Acids Res 33:D294–D296. Cooperstock MS, Zedd AJ. 1983. Intestinal flora of infants. In Human Intestinal Microflora in Health and Disease (DJ Hentges, Editor). Academic Press, New York, pp. 79– 99. Corfield AP, Wagner SA, Clamp JR, Kriaris MS, Hoskins LC. 1992. Mucin degradation in the human colon: production of sialidase, sialate O-acetylesterase, N-acetylneuraminate lyase, arylesterase, and glycosulfatase activities by strains of fecal bacteria. Infect Immun 60:3971–3978. Corfield AP, Myerscough N, Gough M, Brockhausen I, Schauer R, Paraskeva C. 1995. Glycosylation patterns of mucins in colonic disease. Biochem Soc Trans 23:840–845.

Prokaryote Diversity of Epithelial Mucosal Biofilms

147

Dai D, Walker WA. 1999. Protective nutrients and bacterial colonization in the immature human gut. Adv Pediatr 46:353–382. DeLong EF, Wickham GS, Pace NR. 1989. Phylogenetic stains: ribosomal RNA-based probes for the identification of single cells. Science 243(4896):1360–1363. Deplancke B, Gaskins HR. 2001. Microbial modulation of innate defense: goblet cells and the intestinal mucus gel. Am J Clin Nutr 73(Suppl):1131S–1141S. Deplancke B, Hristova KR, Oakley HA, McCracken VJ, Aminov RI, Mackie RI Gaskins HR. 2000. Molecular ecological analysis of dissimilatory sulfidogen succession and diversity in the mouse gastrointestinal tract. Appl Environ Microbiol 66:2166–2174. Deplancke B, Vidal O, Ganessunker D, Donovan SM, Mackie RI, Gaskins HR. 2002. Selective growth of mucolytic bacteria including Clostridium perfringens in a neonatal piglet model of total parenteral nutrition. Am J Clin Nutr 76:1117–1125. Eckburg PB, Bik EM, Bernstein CN, Purdom E, Dethlefsen L, Sargent M, Gill SR, Nelson KE, Relman DA. 2005. Diversity of the human intestinal microbial flora. Science 308:1635– 1638. Enss ML, Grosse-Siestrup H, Schmidt-Wittig U, Gartner K. 1992. Changes in colonic mucins of germfree rats in response to the introduction of a “normal” rat microbial flora. Rat colonic mucin. J Exp Anim Sci 35:110–119. Favier CF, Vaughan EE, De Vos WM, Akkermans AD. 2002. Molecular monitoring of succession of bacterial communities in human neonates. Appl Environ Microbiol 68:219– 226. Fontaine N, Meslin JC, Lory S, Andrieux C.1996. Intestinal mucin distribution in the germfree rat and in the heteroxenic rat harbouring a human bacterial flora: effect of inulin in the diet. Br J Nutr 75:881–892. Forstner JF, Oliver MG, Sylvester FA. 1995. Production, structure, and biologic relevance of gastrointestinal mucins. In Infections of the Gastrointestinal Tract (MJ Blaser, PD Smith, JI Ravdin, HB Greenberg, RL Guerrant, Editors). Raven Press, New York, pp. 71–88. Freter R, Brickner H, Botney M, Cleven D, Aranki A. 1983a. Mechanisms that control bacterial populations in continuous-flow culture models of mouse large intestinal flora. Infect Immun 39(2):676–685. Freter R, Stauffer E, Cleven D, Holdeman LV, Moore WE. 1983b. Continuous-flow cultures as in vitro models of the ecology of large intestinal flora. Infect Immun 39(2):666–675. Galvez A, Maqueda M, Martinez-Bueno M, Valdivia E. 1998. Publication rates reveal trends in microbiological research. ASM News 64:269–275. R Garrity GM, Bell JA, Lilburn TG. 2004. Taxonomic outline of the prokaryotes. Bergey’s Manual of Systematic Bacteriology. Release 5.0. Gibbons RJ, Kapsimalis B. 1967. Estimates of the overall rate of growth of the intestinal microflora of hamsters, guinea pigs, and mice. J Bacteriol 93:510–512. Gunzer F, Hennig-Pauka I, Waldmann KH, Mengel M. 2003. Gnotobiotic piglets as an animal model for oral infection with O157 and non-O157 serotypes of STEC. Methods Mol Med 73:307–327. Gustaffson BE, Carlstedt-Duke B. 1984. Intestinal water soluble mucins in germfree, exgermfree and conventional animals. Acta Pathol Microbiol Immunol Scand 92:247– 252. Hayashi H, Takahashi R, Nishi T, Sakamoto M, Benno Y. 2005. Molecular analysis of jejunal, ileal, caecal and recto-sigmoidal human colonic microbiota using 16S rRNA gene libraries and terminal restriction fragment length polymorphism. J Med Microbiol 54:1093–1101. Hermon-Taylor J, Bull TJ, Sheridan JM, Cheng J, Stellakis M, Sumar N. 2000. Causation of Crohn’s disease by Mycobacterium avium subspecies paratuberculosis. Can J Gastroenterol 14:521–539.

148

Biofilms in the Food Environment

Hornef MW, Normark S, Henriques-Normark B, Rhen M. 2005. Bacterial evasion of innate defense at epithelial linings. Chem Immunol Allergy 86:72–98. Hoskins LC. 1992. Mucin degradation in the human gastrointestinal tract and its significance to enteric microbial ecology. Eur J Gastroenterol Hepatol 5:205–212. Hoskins LC, Boulding ET. 1981. Mucin degradation in human colon ecosystems. Evidence for the existence and role of bacterial sub-populations producing glycosidases as extracellular enzymes. J Clin Invest 67:163–172. Hoskins LC, Augustines M, McKee WB, Boulding ET, Kriaris M, Niedermeyer G. 1985. Mucin degradation in human colon ecosystems. Isolation and properties of fecal strains that degrade ABH blood group antigens and oligosaccharides from mucin glycoproteins. J Clin Invest 75:944–953. Hungate RE. 1950. The anaerobic mesophilic cellulolytic bacteria. Bacteriol Rev 14:1–49. Ishikawa K, Satoh Y, Oomori Y, Yamano M, Matsuda M, Ono K. 1989. Influence of conventionalization on cecal wall structure of germ-free Wistar rats: quantitative light and qualitative electron microscopic observations. Anat Embryol (Berl) 180:191–198. Jorgensen CB, Cirera S, Anderson SI, Archibald AL, Raudsepp T, Chowdhary B, EdforsLilja I, Andersson L, Fredholm M. 2003. Linkage and comparative mapping of the locus controlling susceptibility towards E. coli F4ab/ac diarrhoea in pigs. Cytogenet Genome Res 102:157–162. Kandori H, Hirayama K, Takeda M, Doi K. 1996. Histochemical, lectin-histochemical and morphometrical characteristics of intestinal goblet cells of germfree and conventional mice. Exp Anim 45:155–160. Krause DO, Denman SE, Mackie RI, Morrison M, Rae AL, Attwood GT, McSweeney CS. 2003. Opportunities to improve fiber degradation in the rumen: microbiology, ecology, and genomics. FEMS Microbiol Rev 27:663–693. Kotlowski R, Bernstein CN, Sepehri S, Krause DO. 2006. Serine protease autotransporters (SPATE) and autoaggregative adhesins from Escherichia coli are potential virulence factors in inflammatory bowel disease. In Gut Microbiology: Research to Improve Health, Immune Response and Nutrition. Aberdeen 21–23 June 2006. Lappin-Scott HM, Costerton JM, Marrie TJ. 1992. Biofilms and biofouling. In Encyclopedia of Microbiology, Vol. 1 (JL Lederberg, Editor-in-Chief). Academic Press, San Diego, CA, pp. 277–284. Lee A, Gordon J, Dubos R. 1968. Enumeration of the oxygen sensitive bacteria usually present in the intestine of healthy mice. Nature 220:1137–1139. Lee A, Gordon J, Dubos R. 1971. The mouse intestinal flora with emphasis on the strict anaerobes. J Exp Med 133:339–352. Ley RE, Backhed F, Turnbaugh P, Lozupone CA, Knight RD, Gordon JI. 2005. Obesity alters gut microbial ecology. Proc Natl Acad Sci USA 102:11070–11075. Li L, Bannantine JP, Zhang Q, Amonsin A, May BJ, Alt D, Banerji N, Kanjilal S, Kapur V. 2005. The complete genome sequence of Mycobacterium avium subspecies paratuberculosis. Proc Natl Acad Sci U S A 102:12344–12359. Liu WT, Marsh TL, Cheng H, Forney LJ. 1997. Characterization of microbial diversity by determining terminal restriction fragment length polymorphisms of genes encoding 16S rRNA. Appl Environ Microbiol 63:4516–4522. McCartney AL, Wenzhi W, Tannock G. 1996. Molecular analysis of the composition of the bifidobacterial and Lactobacillus microflora of humans. Appl Environ Microbiol 62:4608–4613. Macfarlane GT, Gibson GR. 1991. Formation of glycoprotein degrading enzymes by Bacteroides fragilis. FEMS Microbiol Lett 77:289–294.

Prokaryote Diversity of Epithelial Mucosal Biofilms

149

Macfarlane GT, Gibson GR, Cummings JH. 1992. Comparison of fermentation reactions in different regions of the human colon. J Appl Bacteriol 72:57–64. Macfarlane S, Woodmansey S, Macfarlane GT. 2005. Colonization of mucin by human intestinal bacteria and establishment of biofilm communities in a two-stage continuous culture system. Appl Environ Microbiol 71:7483–7492. Mackie RI, Sghir A, Gaskins HR. 1999. Developmental microbial ecology of the neonatal gastrointestinal tract. Am J Clin Nutr 69:1035S–1045S. Martins AB, Matos ED, Lemos AC. 2005. Infection with the Mycobacterium avium complex in patients without predisposing conditions: a case report and literature review. Braz J Infect Dis 9:173–179. Matsuo K, Ota H, Akamatsu T, Sugiyama A, Katsuyama T. 1997. Histochemistry of the surface mucous gel layer of the human colon. Gut 40:782–789. Meslin JC, Fontaine N, Andrieux C. 1999. Variation of mucin distribution in the rat intestine, caecum and colon: effect of the bacterial flora. Comp Biochem Physiol 123:235–239. Miller TL, Wolin MJ. 1981. Fermentation by the human large intestine microbial community in an in vitro semicontinuous culture system. Appl Environ Microbiol 42:400–407. Nielsen DS, Moller PL, Rosenfeldt V, Paerregaard A, Michaelsen KF, Jakobsen M. 2003. Case study of the distribution of mucosa-associated Bifidobacterium species, Lactobacillus species, and other lactic acid bacteria in the human colon. Appl Environ Microbiol 69:7545–7548. Nieuw Amerongen AV, Bolscher JGM, Bloemena E, Veerman ECI. 1998. Sulfomucins in the human body. Biol Chem 379:1–18. Ott SJ, Musfeldt M, Wenderoth DF, Hampe J, Brant O, Folsch UR, Timmis KN, Schreiber S. 2004. Reduction in diversity of the colonic mucosa associated bacterial microflora in patients with active inflammatory bowel disease. Gut 53:685–693. Paster BJ, Ludwig W, Weisburg WG, Stackebrandt E, Hespell RB, others 1985. A phylogenetic grouping of the Bacteroides, Cytophagas, and certain Flavobacteria. Syst Appl Microbiol 6:34–42. Paulson JC, Colley KJ. 1989. Glycosyltransferases. Structure, localization, and control of cell type-specific glycosylation. J Biol Chem 264:17615–17618. Poulsen LK, Ballard G, Stahl DA. 1993. Use of rRNA fluorescence in situ hybridization for measuring the activity of single cells in young and established biofilms. Appl Environ Microbiol 59:1354–1360. Poulsen LK, Licht TR, Rang C, Krogfelt KA, Molin S. 1995. Physiological state of Escherichia coli BJ4 growing in the large intestines of streptomycin-treated mice. J Bacteriol 177:5840–5845. Pryde SE, Richardson AJ, Stewart CS, Flint HJ. 1999. Molecular analysis of the microbial diversity present in the colonic wall, colonic lumen, and cecal lumen of a pig. Appl Environ Microbiol 65:5372–5377. Quigley ME, Kelly SM. 1995. Structure, function, and metabolism of host mucus glycoproteins. In Human Colonic Bacteria (GR Gibson, GT Macfarlane, Editors). CRC Press, Boca Raton, FL, pp. 175–201. Rapp´e MS, Giovannoni SJ. 2004. The uncultured microbial majority. Annu Rev Microbiol 57:369–394. Roberton AM, Wright DP. 1997. Bacterial glycosulphatases and sulphomucin degradation. Can J Gastroenterol 11:361–366. Sarma-Rupavtarm RB, Ge Z, Schauer DB, Fox JG, Polz MF. 2004. Spatial distribution and stability of the eight microbial species of the altered schaedler flora in the mouse gastrointestinal tract. Appl Environ Microbiol 70:2791–2800.

150

Biofilms in the Food Environment

Savage DC. 1969a. Microbial interference between indigenous yeast and lactobacilli in the rodent stomach. J Bacteriol 98:1278–1283. Savage DC. 1969b. Localization of certain indigenous microorganisms on the ileal villi of rats. J Bacteriol 97:1505–1506. Savage DC. 1972. Associations and physiological interactions of indigenous microorganisms and gastrointestinal epithelia. Am J Clin Nutr 25:1372–1379. Savage DC. 2001. Microbial biota of the human intestine: a tribute to some pioneering scientists. Curr Issues Intest Microbiol 2:1–15. Schaedler RW, Dubos R, Costello R. 1965. Association of germfree mice with bacteria isolated from normal mice. J Exp Med 122:77–82. Schwiertz A, Gruhl B, Lobnitz M, Michel P, Radke M, Blaut M. 2003. Development of the intestinal bacterial composition in hospitalized preterm infants in comparison with breast-fed, full-term infants. Pediatr Res 54:393–399. Sheahan DG, Jervis HR. 1976. Comparative histochemistry of gastrointestinal mucosubstances. Am J Anat 146:103–131. Simpson JM, McCracken VJ, White BA, Gaskins HR, Mackie RI. 1999. Application of denaturant gradient gel electrophoresis for the analysis of the porcine gastrointestinal microbiota. J Microbiol Methods 36:167–179. Simpson JM, McCracken VJ, Gaskins HR, Mackie RI. 2000. Denaturant gradient gel electrophoresis based analysis of 16S ribosomal DNA amplicons to monitor changes in fecal bacterial populations of weaning pigs following introduction of Lactobacillus reuteri strain MM53. Appl Environ Microbiol 66:4705–4714. Simpson JM, Martineau B, Jones WE, Ballam JM, Mackie RI. 2002. Characterization of fecal bacterial populations in canines: effects of age, breed and dietary fiber. Microb Ecol 44:186–197. Singleton P, Sainsbury D. 1987. Dictionary of Microbiology and molecular Biology, 2nd Edition. Wiley-Interscience, New York. Stevenson BS, Eichorst SA, Wertz JT, Schmidt TM, Breznak JA. 2004. New strategies for cultivation and detection of previously uncultured microbes. Appl Environ Microbiol 70:4748–4755. Swidsinski A, Weber J, Loening-Baucke V, Hale LP, Lochs H. 2005. Spatial organization and composition of the mucosal flora in patients with inflammatory bowel disease. J Clin Microbiol 43:3380–3389. Szentkuti L, Riedesel H, Enss ML, Gaertner K, Von Engelhardt W. 1990. Pre-epithelial mucus layer in the colon of conventional and germ-free rats. Histochem J 22:491–497. Tauber AI. 2003. Metchnikoff and the phagocytosis theory. Nat Rev Mol Cell Biol 4:897–901. Tissier H. 1900. Recherches sur la flore intestinale des nourrissons. In E´tat normal et pathologique (G Carre, C Naud, Editors). Facult´e de M´edecine de Paris, Paris, France (th`ese). Toivanen P, Vaahtovuo J, Eerola E. 2001. Influence of major histocompatibility complex on bacterial composition of fecal flora. Infect Immun 69:2372–2377. Vaahtovuo J, Toivanen P, Eerola E. 2003. Bacterial composition of murine fecal microflora is indigenous and genetically guided. FEMS Microbiol Ecol 44:131–6. van der Waaij LA, Harmsen HJ, Madjipour M, Kroese FG, Zwiers M, van Dullemen HM, de Boer NK, Welling GW, Jansen PL. 2005. Bacterial population analysis of human colon and terminal ileum biopsies with 16S rRNA-based fluorescent probes: commensal bacteria live in suspension and have no direct contact with epithelial cells. Inflamm Bowel Dis 11:865–871.

Prokaryote Diversity of Epithelial Mucosal Biofilms

151

White DG, Zhao S, Singh R, McDermott PF. 2004. Antimicrobial resistance among gramnegative foodborne bacterial pathogens associated with foods of animal origin. Foodborne Pathog Dis 1:137–152. Woese CR. 1987. Bacterial evolution. Microbiol Rev 51:221–271. Zoetendal EG, Akkermans ADL, de Vos WM. 1998. Temperature gradient gel electrophoresis analysis from human fecal samples reveals stable and host-specific communities of active bacteria. Appl Environ Microbiol 64:3854–3859. Zoetendal EG, Akkermans ADL, Akkermans van-Vliet WM, de Visser JAGM, de Vos WM. 2001. The host genotype affects the bacterial community in the human gastrointestinal tract. Microbiol Ecol. Health Dis 13:129–134. Zoetendal EG, von Wright A, Vilpponen-Salmela T, Ben-Amor K, Akkermans AD, de Vos WM. 2002. Mucosa-associated bacteria in the human gastrointestinal tract are uniformly distributed along the colon and differ from the community recovered from feces. Appl Environ Microbiol 68:3401–3407. Zoetendal EG, Collier CT, Koike S, Mackie RI, Gaskins HR. 2004. Molecular ecological analysis of the gastrointestinal microbiota: a review. J Nutr 134:465–472.

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 7 BENEFICIAL BACTERIAL BIOFILMS Gregor Reid, Pirkka Kirjavainen, and Bryan Richardson

Introduction Governments create agencies to generate and enforce policies and procedures, and to adjudicate and control the spending of funds. In most countries, activities associated with “food” are placed in an agency covering agriculture, food, and/or fisheries, while a separate agency is set up to cover “health.” Such bureaucratic separations can result in the perception that activities associated with “food” begins at the farm/soil and end at the table. Just so, one might wonder why a chapter on beneficial bacterial biofilms focused on the oral, intestinal, and urogenital tract of humans would legitimately be placed in a book on “Biofilms in the food environment”? The reasons are relatively simple, albeit varying from the norm of some people. Humans would not be alive today if not for their microbiota. As will be discussed below, these organisms enter our system primarily through the food chain, and play a major role in many aspects of our lives. On a daily basis, many such bacteria also die within our system and leave through defecation, urination and in some cases spitting. Ingested nutrients influence, to a large extent, which organisms survive and thrive, and subsequently how healthy the host becomes. Thus, it is clear that more thought needs to be given to which bacteria we should ingest at various life stages, which we should replenish on a daily basis and how our body’s defenses cope with this diverse array of microbes. Given the critical nature of our microbial content, it is mystifying how we spend so much time trying to kill them, and so little time understanding how to nurture and live in harmony with them. In terms of biofilms, it is clear that the majority of microbes prefer to live within biofilms, and many have been well described in the mouth, intestine, and urogenital tracts (Reid and others 2001a; Probert and 153

154

Figure 7.1.

Biofilms in the Food Environment

The gut microbiota is a dynamic multi-species biofilm.

Gibson 2002; Sbordone and Bortolaia 2003; Anderson and others 2004) (Figure 7.1). Several studies have described the formation and metabolism of biofilms, and methods designed to eradicate or control them or use them to benefit food production or the environment. In this chapter, the emphasis will be on the organisms that contribute positively to our health, and the food that can either deliver them or enhance their survival.

The Beginning of Our Lifelong Association Having evolved from microbes through evolution, it is no surprise that within minutes of our entry into the world, microbes establish a foothold. Either through exposure to vaginal or fecal organisms or through handling, suckling, being kissed or feeding, we each become colonized by bacteria. Little is known about the organisms we inherit, the role the host has in selecting them, the outcome to health if certain species are not ingested, and

Beneficial Bacterial Biofilms

155

the extent to which breast milk and different formula milks alter the levels of species. One recent study has shown that clostridia levels are higher in babies born vaginally compared to by caesarian, and this seems to reduce later allergy (Salminen and others 2004). A study of two babies showed that the stool microbiota became more complex with time and was altered by formula feeding and weaning (Favier and others 2002). Intriguingly, another study of the first four weeks of life in 29 premature babies showed that the actual bacterial composition of hospitalized preterm infants was similar to breast-fed, full-term infants and was not necessarily influenced by birth weight, diet, or antibiotic treatment (Schwiertz and others 2003). This latter observation suggests a remarkable similarity in exposure of babies to certain organisms, plus an ability of the gut to retain them. The similarity may even extend across continents, following studies in which we have shown that the vaginal microbiota in women from Nigeria is very similar to that of Canadian women (Anukam and others 2005). The use of extensively hydrolyzed whey formula supplemented with Bifidobacterium lactis BB12 in a group of infants led to fewer Bacteroides associated with atopic sensitization, and greater numbers of lactobacilli and enterococci (Kirjavainen and others 2002). Thus, there are mounting data to indicate that the microbiota of the infant gut can be manipulated by selective bacterial ingestion, and this in turn can provide health benefits. In some countries, lactobacilli or bifidobacteria are administered in formula milk to premature babies, born by caesarian section, devoid of a fully formed gut and at high risk of infection, in the hope of preventing necrotizing enterocolitis (NEC) and other deadly diseases. This purposeful administration of microbes is referred to as probiotics, defined as “live microorganisms which when administered in adequate amounts confer a health benefit on the host” (FAO/WHO 2002). The evidence that probiotic use immediately after child birth is beneficial, is growing, since the clinical trial that showed the use of Lactobacillus acidophilus and Bifidobacterium sp. led to a major reduction in cases of NEC and resultant death in an intensive care unit in Columbia (Hoyos 1999). Another study in which mothers and babies were administered with L. rhamnosus GG showed a significant reduction in atopic dermatitis and therefore a positive benefit to the immune status of the host (Kalliomaki and others 2001). However, the same GG strain failed to prevent NEC in another study (Dani and others 2002) and appeared to increase asthma rates in the dermatitis study group (Kalliomaki and others 2003) raising doubts about probiotics being suitable in formula milk, at least for premature and “at risk” babies. Still, there is a growing literature on the benefits of probiotics in infants to prevent and treat diarrhea (Nopchinda and others 2002; Rosenfeldt and others 2002; Erdeve and others 2004) and stunting associated with

156

Biofilms in the Food Environment

diarrhea (Saran and others 2002). More dramatic effects are being considered feasible, such as the delay in onset of type 1 diabetes (Matsuzaki and others 1997; Calcinaro and others 2005), the use of select strains to form a healthy intestinal tract through angiogenesis (Stappenbeck and others 2002), the manipulation of the gut microbiota to reduce the risk of obesity (Backhed and others 2004), and even implantation of organisms to increase life longevity (Brummel and others 2004)! Food itself, of course, plays an important role in how the microbiota develops (Mai 2004). Oligo-saccharides and glycoconjugates, compounds found in human milk, can interfere with attachment of enteropathogens. Other saccharides not metabolized by humans, and referred to as prebiotics, can influence the gut microbiota. Prebiotics are defined as “nondigestible substances that provide a beneficial physiological effect on the host by selectively stimulating the favorable growth or activity of a limited number of indigenous bacteria” (Reid and others 2003). These compounds are known to influence microbial growth, and a study has shown that when fructooligosaccharides (FOS) and transbeta-galactooligosaccharides (TOS) are added to baby formula, the net result is an increased number of lactobacilli and bifidobacteria (Miniello and others 2003). Another study in 14 adults who supplemented their diet for 2 weeks with a mixture of 7.5 g of oligofructose and 7.5 g inulin showed a resultant increase in mucosal bifidobacteria and lactobacilli, with significantly more eubacteria but no changes in total anaerobes clostridia, bacteroides, or coliforms (Langlands and others 2004). Such trials rely mostly on stool sampling, whereas many bacteria, for example bifidobacteria (Ouwehand and others 2004), bind in large numbers to the mucus of the intestine, and others are retained in the small intestine. Diet can clearly affect the gut microbiota, but extended feeding changes may be required to disrupt a well-established microbiota (McBain and others 2003), perhaps in part due to the recalcitrant nature of biofilms formed within the mucus layer and on the surface of cells. The ability of so-called beneficial organisms to form biofilms and then confer benefits on the host as a result of such biofilm structures has not been studied to any great extent. In the oral cavity, complex biofilms form through sequential adhesion to surfaces, coaggregation between different pairs of species, and use of primary nutrients and secondary bacterial metabolites (Kolenbrander 2000). For example, an aggressive periodontal pathogen Porphyromonas gingivalis first establishes itself in a mixed-species plaque biofilm on tooth surfaces by binding to commensal Streptococcus gordonii (Lamont and others 2002). In the mouth as in other orifices inhabited by bacterial biofilms, it is difficult to dislodge or change the biofilm structure and composition. Part of this is caused by the

Beneficial Bacterial Biofilms

157

firm binding of the biofilms to the surfaces (Busscher and others 1995), the ability of the glycocalyx to “hide” the structure or protect it from host immune attack (Jesaitis and others 2003; Vuong and others 2004), but it may also be due to programming at the mucosal sites such that only certain species are “accepted” by the host (Gibbons and others 1990), as well as an immune memory system that latterly recognizes new bacterial entrants as friend or foe (Jump and Levine 2004).

Formation of, Communication within, and Displacement of Biofilms Two studies provide an illustration of how lactobacilli can influence pathogenic biofilms in the mouth and throat. In the first study, 594 children, 1–6 years old, received regular milk or one that was supplemented with L. rhamnosus GG for 5 days a week for 7 months with a net decrease in caries (Nase and others 2001). The mechanisms were not investigated and could have been due to displacement or inhibition of pathogens or influence on local immunity causing an environment less conducive to S. mutans growth. In the second study, voice prostheses were exposed three times daily to a Lactococcus lactis or Streptococcus thermophilus suspension resulting in significant reduction in yeast and bacterial pathogen colonization (Free and others 2001). As stated in the introduction, food intake affects biofilms in a number of body sites, not the least of which is the urogenital tract in women. The effects start in the bowel where the excretion of excess food and waste is accompanied by significant loss of bacteria. The anal opening is only 4 cm from the urogenital skin, and therefore on a daily basis, the opportunity arises for many microbial species to colonize the perineum, vulva, and vagina. However, relatively few species actually are able to do this, and the vaginal microbiota generally comprises between 1 and 10 species (Heinemann and Reid 2005). The development, breakdown or displacement of biofilms in this region are not well understood, but appear to be different from the more stable biofilms in the mouth and gut. In women deemed “healthy,” the microbiota is dominated by lactobacilli, often a single species. These organisms can be transferred to the area by ingestion in milk and capsules (Gardiner and others 2002; Reid and others 2001a; Morelli and others 2004) and subsequent passage from the anus to the vagina by passive contact. They can bind to the mucosa using a number of mechanisms including electrostatic and hydrophilic binding, lipoteichoic acid, fimbriae or similar cell wall structures, collagen binding proteins, capsules, and an elongation factor (Chan and others

158

Biofilms in the Food Environment Fimbriated E. coli and streptococci Encapsulated Gram negative pathogens and lactobacilli Mucus layer on top of epithelial surface

Bacterial receptors below and protruding through mucus Figure 7.2. The vaginal microbiota is less complex than the gut, but bacteria-bacteria and bacteria-host communication likely plays an important role in how it changes between a low density lactobacilli dominated microbiota associated with health, and a high density pathogen dominated environment associated with urinary and vaginal infections. Pathogens express various capsule, fimbriae and virulence factors, while lactobacilli can also produce capsules, as well as hydrogen peroxide, acids and bacteriocins capable of killing pathogens, plus biosurfactants, cell-signaling molecules and collagen binding proteins that interfere with pathogenesis.

1985; Conway and Kjelleberg, 1989; Eisen and Reid, 1989; McGroarty, 1994; Adlerberth and others 1996; Heinemann and others 2000; Granato and others 2004). Certain strains of Lactobacillus can also coaggregate with a range of Gram-positive and Gram-negative bacteria (Reid and others 1990) potentially aiding their establishment in the urogenital area (Figure 7.2). Biofilms observed by Gram stain of sloughed vaginal cells tend to be sparsely distributed and not dense like the mushroom-shaped biofilms produced by organisms such as Pseudomonas sp. This structural difference may be due to turnover of the cells and relatively slow multiplication of lactobacilli, or an absence of biofilm forming genes. Organisms such as E. coli have been shown to possess biofilm associated genes, such as the morphogene bolA (Vieira and others 2004) as well as genes for stress response (hslS, hslT, hha, and soxS), adhesion (fimG for type I fimbriae), metabolism (metK), and others of unknown function (Ren and others 2004). Of the Lactobacillus genomes sequenced to date, no biofilm genes have yet been described, albeit none may have been specifically looked for. If these organisms lack such genes, it may explain why an in vitro study found that L. rhamnosus formed a pallid, very thin (average depth < 20 m/mm2 ) biofilm (Filoche and others 2004). Interestingly, a 4- to 7-fold increase in biofilm size occurs when L. rhamnosus is found in the presence of a second organism (Filoche and others 2004). A Lactobacillus coryniformis strain DSM 20001, not found in the vagina, has been shown to produce a coaggregation-promoting factor (Cpf) that allows it to bind to E. coli (Schachtsiek and others 2004);

Beneficial Bacterial Biofilms

159

Lactobacillis amylovorus also coaggregates with Candida (Hsu and others 1990). In both cases, the heterogeneous coaggregates would contribute to the formation of larger biofilms than single strain cultures. Thus, candida and/or E. coli and lactobacilli are commonly found together in the vagina. Studies are needed to determine if there is any symbiotic relationship. On the other hand, an Enterococcus sp. can also coaggregate with lactobacilli (McGroarty and others 1992) and produce bacteriocins that can kill them (Kelly and others 2003), thereby potentially breaking down existing biofilms. This latter example could help us explain why the microbiota changes from being dominated by lactobacilli to the one that is dominated by dense bacterial vaginosis associated biofilms lacking in lactobacilli. This displacement of lactobacilli can actually be reversed by exogenous application of probiotic lactobacilli as shown in a human study, where Gardnerella vaginalis were found, as monitored by molecular typing, to be displaced for up to 21 days (Burton and others 2003). The mechanisms of the displacement have not been elucidated, although in vitro experiments have shown that L. rhamnosus GR-1 is able to penetrate and overcome E. coli biofilms (Reid and others 2001b; Reid 2005). No studies, to date, have been reported on cell–cell communication within commensal biofilms in the urogenital tract, with one exception. Our group has investigated signaling from L. rhamnosus GR-1 and L. reuteri RC-14 to E. coli and S. aureus, respectively. The GR-1 strain is able to displace the pathogens through its lactic acid production, while RC-14 causes a significant down regulation of exotoxin production (unpublished findings). Both these processes illustrate the complexity of bacterial colonization and suggest that we are only at the cusp of understanding, or trying to alter, events in favor of health over disease. Harnessing such signaling effects in food grade lactobacilli could have implications for reducing the risk of food poisoning or improving the health benefits provided by foods. Other cross-talk studies have demonstrated that lactobacilli can signal mucus production by host intestinal cells by a mechanism not yet determined (Mack and others 2003). The net effect is hypothesized to be a benefit to the host through production or enhancement of a physical mucin barrier to pathogenic colonization. Given that mucin hosts a large proportion of the total fecal microbiota, increasing its production must influence gut activity, immunity, and health (Ouwehand and others 2004). Other evidence of communication with the host comes from studies showing that strains can activate transcription factors involved in cytokine signaling directly leading to NF-kappa B activation, and indirectly via cytokines, leading to STAT activation (Miettinen and others 2000). Living Lactobacillus reuteri has a potent antiinflammatory activity on human

160

Biofilms in the Food Environment

epithelial cells, up-regulating NGF, and inhibiting NF-kappaB translocation to the nucleus (Ma and others 2004). Such experiments are interesting, but they do not elucidate if and how lactobacilli can signal the host from within a biofilm, or in the presence of existing indigenous host biofilms. Pursuit of studies in this area will contribute greatly to our understanding of how beneficial bacteria can promote health. This is particularly important given the massive rise in obesity and diabetes rates. A recent paper considered obesity as a “state of chronic inflammation” induced by glucose and macronutrient intake leading to oxidative stress (Dandona and others 2004). The increased levels of IL-6 and TNF-alpha found in obesity and type 2 diabetes were hypothesized to interfere with insulin action by suppressing insulin signal transduction and causing insulin resistance. This relates back to bacterial biofilms through a study, which showed that use of bacteria to modulate the intestinal microbiota could delay onset of type 1 diabetes (Rozanova and others 2002). Henke and Bassler (2004) think of biofilms as “bacterial social engagements.” The signaling effects mentioned above for bacteria–bacteria communication are different from those that operate between bacteria and host cells. The end goal of quorum sensing is to send out sensors into the environment, receive them back, and alter metabolism and activity accordingly, or for the host to detect and respond to the signals. As alluded to earlier, the creation of plaque is perhaps the best documented example of the end result of bacterial communication with both pathogenic and mutualistic bacteria coexisting in homeostasis (Kolenbrander and others 2002). For S. gordonii the attachment process begins when it perceives the presence of salivary agglutinins, to which it expresses surface proteins that allow it to bind to the pellicle on tooth surfaces. This is but one example of a series of signaling reactions leading to dense oral biofilms that comprise organisms essentially feeding off each other. Food intake obviously influences the oral microbiota and plays a role in swaying the extent to which the biofilms comprise organisms associated with health to those associated with caries (Sbordone and Bortolaia 2003). While Henke and Bassler (2004) describe just three signaling processes, it is likely that more will be found, for example species or strain specific signals. Why one strain of a species (L. reuteri RC-14) sends such influential signals to pathogenic S. aureus, while another strain of the same family also found in the human gut and vagina where they are exposed to staphylococci and similar organisms (L. rhamnosus GR-1) does not, is not only an evolutionary puzzle but also a physiological one. In the case of GR-1, of the signals it does release in quorum sensing activity within its environment, clearly the one influencing S. aureus is absent. This was one reason why a two-strain probiotic combination (GR-1 and RC-14) was

Beneficial Bacterial Biofilms

161

chosen for human probiotic use (Reid and others 2001a), namely as the strains can affect the host and microbes in different ways. Clearly, some researchers are looking to identify and synthesize quorumsensing molecules with a view to using them clinically to trick pathogens into performing in such a way as not to infect the host, or to make the bacteria more susceptible to attack by antimicrobial agents. Part of this approach will involve incorporating probiotic organisms into foods. The next 5 to 10 years will prove to be extremely exciting for scientists and students investigating probiotics and the indigenous microbiota. The application of prebiotics and probiotics in foods will, without question, have a major impact on human health. Acknowledgement We appreciate the support provided by NSERC of Canada and Finnish Academy of Finland. References Adlerberth I, Ahrne S, Johansson ML, Molin G, Hanson LA, Wold AE. 1996. A mannosespecific adherence mechanism in Lactobacillus plantarum conferring binding to the human colonic cell line HT-29. Appl Environ Microbiol. 62(7):224451. Anderson GG, Martin SM, Hultgren SJ. 2004. Host subversion by formation of intracellular bacterial communities in the urinary tract. Microbes Infect. 6(12):1094–1101. Anukam KC, Osazuwa EO, Ahonkhai I, Reid G. 2006. Lactobacillus vaginal microbiota of women attending a reproductive health care service in Benin City, Nigeria. Sex. Transm. Dis. 33(1):59–62. Backhed F, Ding H, Wang T, Hooper LV, Koh GY, Nagy A, Semenkovich CF, Gordon JI. 2004. The gut microbiota as an environmental factor that regulates fat storage. Proc Natl Acad Sci U S A 101(44):15718–15723. Brummel T, Ching A, Seroude L, Simon AF, Benzer S. 2004. Drosophila lifespan enhancement by exogenous bacteria. Proc Natl Acad Sci U S A 101(35):12974–12979. Burton JP, Cadieux P, Reid G. 2003. Improved understanding of the bacterial vaginal microbiota of women before and after probiotic instillation. Appl Environ Microbiol 69:97– 101. Busscher HJ, Bos R, van der Mei HC. 1995. Initial microbial adhesion is a determinant for the strength of biofilm adhesion. FEMS Microbiol Lett 128(3):229–234. Calcinaro F, Dionisi S, Marinaro M, Candeloro P, Bonato V, Marzotti S, Corneli RB, Ferretti E, Gulino A, Grasso F, De Simone C, Di Mario U, Falorni A, Boirivant M, Dotta F. 2005. Oral probiotic administration induces interleukin-10 production and prevents spontaneous autoimmune diabetes in the non-obese diabetic mouse. Diabetologia 48(8):1565– 1575. Chan RCY, Reid G, Irvin RT, Bruce AW, Costerton JW. 1985. Competitive exclusion of uropathogens from uroepithelial cells by Lactobacillus whole cells and cell wall fragments. Infect Immun 47:84–89.

162

Biofilms in the Food Environment

Conway PL, Kjelleberg S. 1989. Protein-mediated adhesion of Lactobacillus fermentum strain 737 to mouse stomach squamous epithelium. J Gen Microbiol 135(Pt 5):1175– 1186. Dandona P, Aljada A, Bandyopadhyay A. 2004. Inflammation: the link between insulin resistance, obesity and diabetes. Trends Immunol 25:4–7. Dani C, Biadaioli R, Bertini G, Martelli E, Rubaltelli FF. 2002. Probiotics feeding in prevention of urinary tract infection, bacterial sepsis and necrotizing enterocolitis in preterm infants. A prospective double-blind study. Biol Neonate 82(2):103–108. Eisen A, Reid G. 1989. Effect of culture media on Lactobacillus hydrophobicity and electrophoretic mobility. Microbial Ecol 17(1):17–25. Erdeve O, Tiras U, Dallar Y. 2004. The probiotic effect of Saccharomyces boulardii in a pediatric age group. J Trop Pediatr 50(4):234–236. FAO/WHO. 2002. Guidelines for the evaluation of probiotics in food. Food and Agriculture Organization of the United Nations and World Health Organization Working Group Report. http://www.fao.org/es/ESN/Probio/probio.htm. Favier CF, Vaughan EE, De Vos WM, Akkermans AD. 2002. Molecular monitoring of succession of bacterial communities in human neonates. Appl Environ Microbiol 68(1):219– 226. Filoche SK, Anderson SA, Sissons CH. 2004. Biofilm growth of Lactobacillus species is promoted by Actinomyces species and Streptococcus mutans. Oral Microbiol Immunol 19(5):322–326. Free RH, Busscher HJ, Elving GJ, van der Mei HC, van Weissenbruch R, Albers FW. 2001. Biofilm formation on voice prostheses: in vitro influence of probiotics. Ann Otol Rhinol Laryngol 110(10):946–951. Gardiner G, Heinemann C, Baroja ML, Bruce AW, Beuerman D, Madrenas J, Reid G. 2002. Oral administration of the probiotic combination Lactobacillus rhamnosus GR-1 and L. fermentum RC-14 for human intestinal applications. Int Dairy J 12(2–3):191–196. Gibbons RJ, Hay DI, Childs WC 3rd, Davis G. 1990. Role of cryptic receptors (cryptitopes) in bacterial adhesion to oral surfaces. Arch Oral Biol 35(Suppl):107S–114S. Granato D, Bergonzelli GE, Pridmore RD, Marvin L, Rouvet M, Corthesy-Theulaz IE. 2004. Cell surface-associated elongation factor Tu mediates the attachment of Lactobacillus johnsonii NCC533 (La1) to human intestinal cells and mucins. Infect Immun 72(4):2160–2169. Heinemann C, Van Hylckama Vlieg JET, Janssen DB, Busscher HJ, van der Mei HC, Reid G. 2000. Purification and characterization of a surface-binding protein from Lactobacillus fermentum RC-14 inhibiting Enterococcus faecalis 1131 adhesion. FEMS Microbiol Lett. 190:177–180. Heinemann C, Reid G. 2005. Vaginal microbial diversity among postmenopausal women with and without hormone replacement therapy. Can J Microbiol 51(9):777–781. Henke JM, Bassler BL. 2004. Bacterial social engagements. Trends Cell Biol 14(11):648–656. Hoyos AB. 1999. Reduced incidence of necrotizing enterocolitis associated with enteral administration of Lactobacillus acidophilus and Bifidobacterium infantis to neonates in an intensive care unit. Int J Infect Dis 3(4):197–202. Hsu LY, Minah GE, Peterson DE, Wingard JR, Merz WG, Altomonte V, Tylenda CA. 1990. Coaggregation of oral Candida isolates with bacteria from bone marrow transplant recipients. J Clin Microbiol 28(12):2621–2626. Jesaitis AJ, Franklin MJ, Berglund D, Sasaki M, Lord CI, Bleazard JB, Duffy JE, Beyenal H, Lewandowski Z. 2003. Compromised host defense on Pseudomonas aeruginosa biofilms: characterization of neutrophil and biofilm interactions. J Immunol 171(8): 4329–4339.

Beneficial Bacterial Biofilms

163

Jump RL, Levine AD. 2004. Mechanisms of natural tolerance in the intestine: implications for inflammatory bowel disease. Inflamm Bowel Dis 10(4):462–478. Kalliomaki M, Salminen S, Arvilommi H, Kero P, Koskinen P, Isolauri E. 2001. Probiotics in primary prevention of atopic disease: a randomised placebo-controlled trial. Lancet 357(9262):1076–1079. Kalliomaki M, Salminen S, Poussa T, Arvilommi H, Isolauri E. 2003. Probiotics and prevention of atopic disease: 4-year follow-up of a randomised placebo-controlled trial. Lancet 361(9372):1869–1871. Kelly MC, Mequio MJ, Pybus V. 2003. Inhibition of vaginal lactobacilli by a bacteriocin-like inhibitor produced by Enterococcus faecium 62-6: potential significance for bacterial vaginosis. Infect Dis Obstet Gynecol 11(3):147–156. Kirjavainen PV, Arvola T, Salminen SJ, Isolauri E. 2002. Aberrant composition of gut microbiota of allergic infants: a target of bifidobacterial therapy at weaning? Gut 51(1): 51–55. Kolenbrander PE. 2000. Oral microbial communities: biofilms, interactions, and genetic systems. Annu Rev Microbiol 54:413–437. Kolenbrander PE, Andersen RN, Blehert DS, Egland PG, Foster JS, Palmer RJ Jr. 2002. Communication among oral bacteria. Microbiol Mol Biol Rev 66(3):486–505. Lamont RJ, El-Sabaeny A, Park Y, Cook GS, Costerton JW, Demuth DR. 2002. Role of the Streptococcus gordonii SspB protein in the development of Porphyromonas gingivalis biofilms on streptococcal substrates. Microbiology 148(Pt 6):1627–1636. Langlands SJ, Hopkins MJ, Coleman N, Cummings JH. 2004. Prebiotic carbohydrates modify the mucosa associated microflora of the human large bowel. Gut 53(11):1610– 1616. Mack DR, Ahrne S, Hyde L, Wei S, Hollingsworth MA. 2003. Extracellular MUC3 mucin secretion follows adherence of Lactobacillus strains to intestinal epithelial cells in vitro. Gut 52(6):827–833. Mai V. 2004. Dietary modification of the intestinal microbiota. Nutr Rev 62(6 Pt 1):235–242. Matsuzaki T, Nagata Y, Kado S, Uchida K, Kato I, Hashimoto S, Yokokura T. 1997. Prevention of onset in an insulin-dependent diabetes mellitus model, NOD mice, by oral feeding of Lactobacillus casei. APMIS 105(8):643–649. McBain AJ, Bartolo RG, Catrenich CE, Charbonneau D, Ledder RG, Gilbert P. 2003. Growth and molecular characterization of dental plaque microcosms. J Appl Microbiol 94(4):655–664. McGroarty JA. 1994. Cell surface appendages of lactobacilli. FEMS Microbiol Lett 124(3):405–409. McGroarty JA, Lee V, Reid G, Bruce AW. 1992. Modulation of adhesion of uropathogenic Enterococcus faecalis to human epithelial cellsin vitro by Lactobacillus species. Microbial Ecol Health Dis 5:309–314. Ma D, Forsythe P, Bienenstock J. 2004. Live Lactobacillus reuteri is essential for the inhibitory effect on tumour necrosis factor alpha-induced interleukin-8 expression. Infect Immun 72:5308–5314. Miettinen M, Lehtonen A, Julkunen I, Matikainen S. 2000. Lactobacilli and streptococci activate NF-kappa B and STAT signaling pathways in human macrophages. J Immunol 164(7):3733–3740. Miniello VL, Moro GE, Armenio L. 2003. Prebiotics in infant milk formulas: new perspectives. Acta Paediatr Suppl 91(441):68–76. Morelli L, Zonenenschain D, Del Piano M, Cognein P. 2004. Utilization of the intestinal tract as a delivery system for urogenital probiotics. J Clin Gastroenterol 38(6 Suppl):S107– S110.

164

Biofilms in the Food Environment

Nase L, Hatakka K, Savilahti E, Saxelin M, Ponka A, Poussa T, Korpela R, Meurman JH. 2001. Effect of long-term consumption of a probiotic bacterium, Lactobacillus rhamnosus GG, in milk on dental caries and caries risk in children. Caries Res 35(6):412– 420. Nopchinda S, Varavithya W, Phuapradit P, Sangchai R, Suthutvoravut U, Chantraruksa V, Haschke F. 2002. Effect of Bifidobacterium Bb12 with or without Streptococcus thermophilus supplemented formula on nutritional status. J Med Assoc Thai 85(Suppl 4):S1225–S1231. Ouwehand AC, Salminen S, Arvola T, Ruuska T, Isolauri E. 2004. Microbiota composition of the intestinal mucosa: association with fecal microbiota? Microbiol Immunol 48(7):497– 500. Probert HM, Gibson GR. 2002. Bacterial biofilms in the human gastrointestinal tract. Curr Issues Intest Microbiol 3(2):23–27. Reid G. 2005. Colonization of the vagina and urethral mucosa. In Colonization of Mucosal Surfaces ( JP Nataro, PS Cohen, HLT Mobley, JN Weiser, Editors). ASM Press, Washington, DC, pp. 431–448. Reid G, Bruce AW, Fraser N, Heinemann C, Owen J, Henning B. 2001a. Oral probiotics can resolve urogenital infections. FEMS Microbiol. Immunol 30:49–52. Reid G, Heinemann C, Howard J, Gardiner G, Gan BS. 2001b. Understanding urogenital biofilms and the potential impact of probiotics. Methods Enzymol 336:403– 410. Reid G, McGroarty JA, Domingue PAG, Chow AW, Bruce AW, Eisen A, Costerton JW. 1990. Coaggregation of urogenital bacteria in vitro and in vivo. Current Microbiol 20: 47–52. Reid G, Sanders ME, Gaskins HR, Gibson GR, Mercenier A, Rastall R, Roberfroid M, Rowland I, Cherbut C, Klaenhammer TR. 2003. New scientific paradigms for probiotics and prebiotics. J Clin Gastroenterol 37(2):105–118. Ren D, Bedzyk LA, Thomas SM, Ye RW, Wood TK. 2004. Gene expression in Escherichia coli biofilms. Appl Microbiol Biotechnol 64(4):515–524. Rosenfeldt V, Michaelsen KF, Jakobsen M, Larsen CN, Moller PL, Tvede M, Weyrehter H, Valerius NH, Paerregaard A. 2002. Effect of probiotic Lactobacillus strains on acute diarrhea in a cohort of nonhospitalized children attending day-care centers. Pediatr Infect Dis J 21(5):417–419. Rozanova GN, Voevodin DA, Stenina MA, Kushnareva MV. 2002. Pathogenetic role of dysbacteriosis in the development of complications of type 1 diabetes mellitus in children. Bull Exp Biol Med 133(2):164–166. Salminen S, Gibson GR, McCartney AL, Isolauri E. 2004. Influence of mode of delivery on gut microbiota composition in seven year old children. Gut 53(9):1388–1389. Saran S, Gopalan S, Krishna TP. 2002. Use of fermented foods to combat stunting and failure to thrive. Nutrition 18(5):393–396. Sbordone L, Bortolaia C. 2003. Oral microbial biofilms and plaque-related diseases: microbial communities and their role in the shift from oral health to disease. Clin Oral Investig 7(4):181–188. Schachtsiek M, Hammes WP, Hertel C. 2004. Characterization of Lactobacillus coryniformis DSM 20001T surface protein Cpf mediating coaggregation with and aggregation among pathogens. Appl Environ Microbiol 70(12):7078–7085. Schwiertz A, Gruhl B, Lobnitz M, Michel P, Radke M, Blaut M. 2003. Development of the intestinal bacterial composition in hospitalized preterm infants in comparison with breast-fed, full-term infants. Pediatr Res 54(3):393–399.

Beneficial Bacterial Biofilms

165

Stappenbeck TS, Hooper LV, Gordon JI. 2002. Developmental regulation of intestinal angiogenesis by indigenous microbes via Paneth cells. Proc Natl Acad Sci U S A 99(24):15451– 15455. Vieira HL, Freire P, Arraiano CM. 2004. Effect of Escherichia coli morphogene bolA on biofilms. Appl Environ Microbiol 70(9):5682–5684. Vuong C, Kocianova S, Voyich JM, Yao Y, Fischer ER, Deleo FR, Otto M. 2004. A crucial role for exopolysaccharide modification in bacterial biofilm formation, immune evasion, and virulence. J Biol Chem 279(52):54881–54886.

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 8 APPLICATIONS OF BIOFILM REACTORS FOR PRODUCTION OF VALUE-ADDED PRODUCTS BY MICROBIAL FERMENTATION Ali Demirci, Thunyarat Pongtharangkul, and Anthony L. Pometto III

Introduction Microbial film can cause many detrimental effects on human health, such as infections on prosthetic implants (Bayston 2000) and in patients with cystic fibrosis (Jackson and others 2003). It also causes biofouling in engineered materials and systems (Gassey and Bryers 2000; Wirtanen and others 2000; Chmielewski and Frank 2003; Patching and Fleming 2003). However, biofilm can be very useful in many applications, for example, wastewater treatment (Lazarova and Manem 2000; O’Flaherty 2003), bioremediation (Von Canstein and others 2002), and degradation of toxic pollutants (Ebihara and Bishop 2002; Jin and others 2003). Unlike the intensively studied, commercial-scale applications in wastewater treatment, biofilm reactors in production of value-added products have been studied only in bench-scale and pilot-scale productions during the past few decades. In spite of this, they have been proved to enhance productions of various value-added products such as ethanol, organic acids, antibiotics, and enzymes as they can generate increased volumetric productivity rates by maintaining high biomass concentration in the bioreactors. Given their potential for development into a continuous culture, their exceptional stability, and their lower nutrient requirements, biofilm reactors have great potential for applications in the large-scale production of many value-added products. In this chapter, the brief background and 167

168

Biofilms in the Food Environment

some recent successful applications of biofilm reactors for productions of various value-added products, mainly ethanol, organic acids, antibiotics, and enzymes, are presented. Biofilm Formation and Structure During the past decade, our understanding on the properties and formation of biofilm has been tremendously expanded not only to the molecular and genetic aspects, but also to the mathematical modeling of the biofilm structure. The following section will briefly explore only the molecular and modeling aspects of the biofilm formation and structure. Biofilm Formation Biofilm is an accumulation of cells embedded in an organic polymer matrix of microbial origin (Characklis and Marshall 1990). More than 90% of wet biofilm mass is water, while the extracellular polymeric substances (EPS), containing polysaccharides and glycoproteins, correspond to ≥70% of the dry biofilm mass (Melo and Oliveira 2001). Biofilm thickness can vary from a few microns to even a few centimeters, depending on factors such as microbial species, biofilm age, available nutrients, and liquid shear stresses. A formation of biofilm takes several steps (Bryers 2000; Busscher and van der Mei 2000) as shown in Figure 8.1. First, the conditioning of the

Products

Bulk Fluid

Nutrients Oxygen

7

2 4 6 1 3

Figure 8.1.

5

Formation of biofilm (see text for description). (Adapted from Bryers 2000.)

Applications of Biofilm Reactors

169

substratum is formed either by macromolecules in bulk liquid or by intentionally coated material (step 1). The microorganisms suspended in the liquid are then transported to the surface by diffusion, convection, or self-motility (step 2). These microorganisms then form weak reversible adhesions with the solid surface (steps 3 and 4). Later, irreversible adhesions take place as a result of formations of polymer bridges between the conditioning layer (an adsorbed layer of macromolecules on the solid surface) and the EPS excreted by the microbes (step 5). Detailed studies on the adhesion mechanism of microbial cells onto the support surface can be found in the literature (Busscher and others 1995; Azeredo and others 1999; Teixeira and Oliveira 1999; Bryers 2000). At this step, a coadhesion, which is a process in which the secondary colonizers coadhere with organisms already adhering to the surface, occurs simultaneously with the initial adhesion. Coaggregates of organisms may also form in bulk liquid and then adhere to the biofilm surface in a process called coaggregation. After these initial steps, the growth of microorganisms (step 6), which depends on substrate availability, plays a more important role in biofilm build-up than does the transport and adhesion of microorganisms to the biofilm (Bott 1995). Detachment processes (step 7), erosion or disruption or sloughing off, occur simultaneously and in response to fluid shear forces, weak internal cohesion, and depletion of nutrients or oxygen in the biofilm. Erosion, a continuous process resulted by liquid shear forces, is the removal of small portions of biofilm. Sloughing, by contrast, is the random detachment of large portions of the biofilm as a result of rapid change or depletion of nutrients (Howell and Atkinson 1976). Last, when growth balances with detachment, the maximum average thickness of biofilm is reached and the system is considered as pseudo-steady state. Although the condition inside the biofilm may be different from the liquid outside, biofilm formation appears to be favored when the temperature and pH of the outside liquid approach the optimum values for microbial growth (Melo and Oliveira 2001). Biofilm Structure It has long been known that the structure of biofilm is not uniform in time or space (Characklis and Marshall 1990). Many detailed analyses show that even in a well-mixed biofilm reactor, different types of biofilm structure can be found (Gjaltema and others 1994). The structure of biofilm is not entirely understood, but at least three different models have been proposed as the structures of biofilm (Wimpenny and others 2000). The first model is the “heterogeneous model”

170

(a)

Biofilms in the Food Environment

(b)

(c)

Liquid Flow

Figure 8.2. Models of biofilm structure: (a) heterogeneous model, (b) heterogeneous mosaic model, and (c) mushroom model. (After Atkinson and Swilley, 1963; van Loosdrecht and others, 2002.)

(Figure 8.2a) in which the microorganisms form a dense, planar, homogeneous biofilm exposed to the flowing liquid (Atkinson and Swilley 1963; Nyvad and Fejerskov 1997). The second model called “heterogeneous mosaic model” or “pseudo-homogeneous model” (Figure 8.2b) describes stacks consisting of cells hold together by EPS and appeared as columns separated by water channels over a layer of cells about 5 μm thick on the substratum (Atkinson and Swilley 1963; Keevil and Walker 1992; Stewart and others 1995). The third model called “mushroom” or “tulip model” (Figure 8.2c) is the most recent model revealed by the use of a confocal laser scanning microscopy and the molecular probe like fluorescent markers. In this model, the biofilm was formed in a mushroom-shaped column surrounded by water channels through which oxygen and nutrients were carried with the liquid flow. Hermanowicz (1998) and Wimpenny and others (2000) have pointed out that the unifying factors of the three types of structure were the substrate concentration in bulk liquid and the external mass transfer limitation (presented in a form of the dimensionless boundary layer thickness by Hermanowicz 1998). Overall, biofilm structure is affected by several factors as summarized in Figure 8.3. Apart from the effects of nutrient depletion and shear forces exerted by the bulk liquid as previously described, nutrient compositions and concentrations in bulk liquid also affect the structure and development of biofilm (Veiga and others 1992; Acuna and others 2002; Allan and others 2002). Van Loosdrecht and others (2002) proposed that biofilm structure depends primarily on an interaction between mass transfer, conversion rates, and detachment forces. When the substrate consumption rate exceeds its maximum mass transfer rate, limited diffusion results in a strongly porous or even filamentous biofilm. Conversely, when the conversion rate or growth is a rate-limiting step, the biofilm becomes more

171

Applications of Biofilm Reactors

ADHESION van der Waals forces Hydrophobicity EPS Production Nutrients availability Nutrients diffusion Temp & pH

BIOFILM FORMATION & STRUCTURE

GROWTH

DETACHMENT

Shear forces Internal cohesion Lacking of nutrient

Figure 8.3. Factors affecting biofilm formation and structure.

homogeneous and compact. Another factor that has a significant effect on biofilm structure is detachment force. The system with low detachment forces results in a detachment in a form of sloughing off and a more porous biofilm, whereas the one with higher detachment forces results in detachment as an erosion and finally yields a more compact biofilm.

Biofilm Reactors Although the use of biofilm reactors is not confined only to microorganisms but also comprises plant and animal cell applications, this chapter focuses only on applications in microbial (bacterial and fungal) fermentations. Such biofilm reactors are usually operated either as packed- or fluidized-bed reactors, with either active or passive immobilized cells. Advantages of Biofilm Reactors One way to increase the productivity of fermentation is to increase biomass in the reactor by using methods such as cell-recycle reactors,

172

Biofilms in the Food Environment

hollow-fiber reactors, and cell immobilization. Hollow-fiber reactors and cell-recycle reactors have their limitations in their high capital and operation cost as well as in the potential for membrane fouling during fermentation. On the other hand, immobilized-cell reactors like biofilm reactors are excellent examples of high-biomass density systems with lesser tendencies to develop membrane fouling and lower required capital costs. In general, cell immobilization methods can be divided into two major categories (Fukuda 1995): (1) active or artificial immobilization, and (2) passive or natural immobilization. Active immobilizations, including covalent bonding to surfaces using various coupling agents and entrapments in polymer matrix, can be achieved by chemical agents. Several disadvantages of active immobilization include (1) toxicity of coupling agents and cross-linking agents on cell viability and activity; (2) instability of the polymer matrix (e.g., calcium alginate gel) with various anions including phosphate, citrate, EDTA, and lactate; (3) cell leakage from the gel matrix; (4) limited mass transfer across the beads; (5) poor operational stability; and (6) high cost of the carrier. On the other hand, passive immobilization occurs by natural adsorptions and colonizations of films or flocculants of microorganisms around or within the solid support materials (Fukuda 1995). The adsorption phenomenon is mainly based on electrostatic interactions between cell surface and the support materials, including both natural materials (such as cellulose and dextran) and synthetic resins. Colonization, by contrast, is based on a technique using porous biomass support particles (BSPs) which was developed by Atkinson and others (1979). The immobilized cells or biofilm is developed on BSPs during the initial batch period due to microbial formation of EPS. Biofilm reactors show many advantages over suspended cell reactors, especially in their higher biomass density and operation stability. Biofilm reactors are able to retain 5 or 10 times more biomass per unit volume of reactor, increasing production rates, reducing the risk of washing out when operating at high dilution rates during continuous fermentation, and eliminating need for reinoculation during repeated-batch fermentation (Fukuda 1995). In the case of filamentous microorganisms, such as Aspergillus niger, the immobilization in biofilm reactors leads to decreased medium viscosity and enhanced nutrient and oxygen transfer (Wang 2000). Moreover, the structure of the biofilm matrix contributes to high resistance of microorganisms to extreme conditions of pH and temperature, contaminations, hydraulic shocks, antibiotics, and toxic substances (White 1984; Norwood and Gilmour 2000). Products from biofilm reactor can be easily recovered (in comparison to product from suspended cell systems, for instance), resulting in more efficient downstream processes.

Applications of Biofilm Reactors

173

Types of Biofilm Reactors In general, biofilm reactors can be categorized into two categories: fixedbed and expanded-bed reactors. Fixed-bed reactors, which include all processes in which the biofilms develop on static media, can be divided into (1) submerged beds, in which the biofilm particles are completely immersed in the liquid (Figures 8.4a and 8.4b); (2) trickling filters, in which the liquid flows downward through the biofilm bed, while the gas flows upward; (3) rotating disk reactors, in which the biofilm develops on the surface of a vertical disk that is partially submerged and rotates within the liquid (Figure 8.4c); and (4) membrane biofilm reactors, in which the microbial layer is attached to a porous gas-permeable membrane (Figure 8.4f). Expanded-bed reactors, by contrast, which include all biofilm processes with continuously moving media maintained by high air or liquid velocity or by mechanical stirring, can be divided into

Figure 8.4. Most common types of biofilm reactors: (a) stirred-tank reactor, (b) fixed-bed reactor, (c) rotating-disk reactor, (d) fluidized-bed reactor, (e) airlift reactor, (f) membranebiofilm reactor. (Adapted from Fukuda 1995; Melo and Oliveira 2001.)

174

Biofilms in the Food Environment

(1) fluidized beds, in which particles move up and down within the expanded bed in the well-defined zone of the reactor (Figure 8.4d); and (2) moving beds, in which the whole expanded bed circulates throughout the reactors such as airlift reactor (Figure 8.4e) and circulating-bed reactors. In expanded beds, the bed is usually expanded by the upward flow of liquid and gas bubbles. However, the inverse fluidized bed, which was developed later, is based on circulation of low-density support material by downward flow of liquid (Karamanev and Nikolov 1992, 1996). Detailed descriptions and comparative analysis of the advantages and disadvantages of various types of biofilm reactors can be found in many publications (Cabral and Tramper 1994; Fukuda 1995; Willaert and others 1996; Lazarova and Manem 2000; Melo and Oliveira 2001). Biomass Support Particles In the selection of solid supports for biofilm reactors, following factors need to be considered. The support must (1) favor microorganism adhesion; (2) have a high mechanical resistance to liquid shear forces and particle collision; (3) be inexpensive; and (4) be widely available. Some of the properties of solid support, such as surface charge, hydrophobicity, porosity, roughness, particle diameter, and density, dramatically affect the adhesion of microorganisms. A great variety in solid support shapes, dimensions, and materials have been developed and used in biofilm reactors. Over the last few decades, the solid support has been developed and designed in order to increase the specific surface area per volume of reactor. Today, biofilm reactors achieve higher efficiency and compactness than in the past. The interaction between microorganism cells and solid support is a result of a balance between the van der Waals forces of attraction and repulsive forces (Oliveira 1992). Generally, the bacterial cell surfaces and most of the existing solid materials display a net negative charge when immersed in aqueous solution with pH near neutrality (Melo and Oliveira 2001). Recent studies on selections of support materials for different types of biofilm reactors and microorganisms suggested that the higher degree of hydrophobicity of solid surfaces strongly enhances adhesions of microorganisms (Sousa and others 1997; Teixeira and Oliveira 1999; Pereira and others 2000). In contrast, Ho and others (1997c) reported that Lactobacillus casei, which was considered to be hydrophilic, was more readily attached to the less hydrophobic support materials. This finding was in agreement with the conclusion of van Loosdrecht and others (1987) that hydrophobic bacteria adhered to hydrophobic surfaces more readily than hydrophilic bacteria. However, the leaching of nutrients can compensate for the hydrophobic nature of solid supports in some cases,

Applications of Biofilm Reactors

175

resulting in a higher biofilm population in more hydrophobic support materials (Ho and others 1997b). To achieve a larger surface area on solid support, a smaller diameter of carrier particles and rough and/or porous surface materials are used. Also, a porous matrix of materials provides niches sheltered from hydraulic shear forces (Bryers 1987; Massol-Deya and others 1995; Ho and others 1997c). If the particles are porous, the film is formed not only on the surface, but also within the pores. Many studies confirmed that surface colonization is favored when crevices are microbially sized (Messing and Opperman 1979; Shimp and Pfaender 1982). Moreover, the problems of deficient nutrient diffusion to the inner area and accumulation of gaseous metabolites inside porous carriers, resulting in carrier washout, can be overcome by using materials with adequately large pores and internal porous volume (Melo and Oliveira 2001). Plastic composite support (PCS), used in many studies provides not only ideal physical structure, but also slow-released nutrients during fermentation (Figure 8.5). PCS, developed at Iowa State University (U.S. Patent Number: 5,595,893), is an extrusion product of polypropylene and several agricultural products which can be custom-made for specific microorganism. For example, PCS ingredients for L. casei consisted of 50% (wt/wt) polypropylene (Quantum USI Division, Cincinnati, OH), 35% soybean hulls (Cargill Soy Processing Plant, Iowa Falls, IA), 5% yeast extract (Ardamine Z, Champlain Industries, Clifton, NJ), 5% defatted soybean flour (Archer Daniels Midland, Decatur, IL), 5% bovine albumin (American Protein Corp., Ames, IA), and mineral salts, which were mixed and extruded with twin screw corotating Brabender PL2000 extruder (model CTSE-V; Brabender Instruments, South Hackensack, NJ) at a rate of 11 rpm with a barrel temperature of 200◦ C and a die temperature of 167◦ C to form a continuous tube. The tubes with a wall thickness of 3.5 mm and an outer diameter of 10.5 mm can be cut into desired dimension, e.g., 10-cm length 0.3-10 cm

0.35 cm

Figure 8.5. Dimensions of the PCS.

176

Biofilms in the Food Environment

Agitator shaft

Packed-bed reactor with PCS rings Figure 8.6.

Packed-bed reactor with PCS tubes

PCS biofilm reactors.

tubes (PCS tubes) or 3-mm slices (PCS rings) (Ho and others 1997c). The PCS rings and tubes were used as solid supports for biofilm formation in packed-bed reactors (Ho and others 1997a) or as PCS tubes attached to the bioreactor agitator shaft (Cotton and others 2001) as shown in Figure 8.6.

Applications of Biofilm Reactors Although biofilm reactors have applications in many areas, including waste treatment and bioremediation, this section will focus on recent applications in the production of value-added products (also summarized in Table 8.1).

Airlift reactor Packed-bed reactor with PCS

A. succinogenes C. acremonium

C. acremonium L. lactis

Nisin

L. casei

Succinic acid Cephalosporin C

Packed-bed reactor with PCS

Demirci and others (1995, 1997); Kunduru and Pometto (1996a, b) Park and Toda (1992) Cao and others (1996, 1997) Lee and others (1989)

Packed-bed reactor with PCSa ring and disk Multistage shallow flow biofilm reactor Rotating-disk reactor Polyurethane foam, bubble column reactor Polyurethane foam, fluidized-bed reactor Polyurethane foam particle Rotating-disk reactor Activated carbon, fluidized-bed reactor Packed-bed reactor with PCS

(continued )

Urbance and others (2003, 2004) Park and Seo (1988); Park and others (1989) Srivastava and Kundu (1999) Bober (2002)

Ho and others (1997a–c); Velazquez and others (2001) Cotton and others (2001)

Ricciardi and others (1997) Wang (2000) Andrews and Fonta (1989) Demirci and others (1993a, b, 1995)

Sanroman and others (1994, 1996)

References

Support Materials/Reactor Types

PCS biofilm reactor with grid-like orientation of PCS Packed-bed reactor with PCS Celite, fluidized-bed reactor

Lactic acid

A. niger A. niger Strep. thermophilus L. amylophilus, L. casei, L. delbrueckii L. casei

A. niger

Z. mobilis and Sacch. cerevisiae A. aceti M7 R. oryzae A. niger

Ethanol

Acetic acid Fumaric acid Citric acid

Organisms

Recent Applications of Biofilms in the Production of Value-Added Products

Product

Table 8.1.

P. chrysosporium P. chrysosporium

P. chrysosporium

Lignin peroxidase

Lignin peroxidase and manganese peroxidase Fermentable levoglucosan from corn stover pyrolysis liquor

U.S. Patent Number: 5,595,893.

Recombinant E. coli R. chinensis

Amylase Lipase

a

T. viride

Cellulase

Pseudomonas putida and Streptomyces setonii 75Vi2 mixed culture; P. chrysosporium

Organisms

PCS tubes attached to the agitator shaft

Nilon web Ultrafiltration capillary membrane, membrane gradostat reactor PCS tubes attached to the agitator shaft

Stainless steel particle, spouted-bed reactor Silicone foam Polyurethane foam

Support Materials/Reactor Types

Khiyami and others (2005a) Khiyami and others (2005b)

Khiyami and others (2006)

Fukuda and others (1986); Webb and others (1986) Oriel (1988) Kyotani and others (1991); Hara and Nakashima (1996, 1998) Linko (1992) Govender and others (2003)

References

Recent Applications of Biofilms in the Production of Value-Added Products (Continued )

Product

Table 8.1.

Applications of Biofilm Reactors

179

Alcohol Production Biofilm reactors are used in alcohol production, mainly ethanol production, in order to overcome cell washout and low productivity in continuous fermentation. Chung and Park (1983) developed a fixed-bed reactor packed vertically with ceramic rods for continuous ethanol production of Saccharomyces cerevisiae and achieved a maximum productivity of 27.5 g/(L h) at glucose concentration of 150 g/L and dilution rate of 1.5/h. They also reported a better ethanol tolerance of the yeast in the fixed-bed biofilm reactor compared to a continuous stirred-tank reactor. Dempsey (1990) developed a fluidized-bed reactor for continuous ethanol production using Zymomonas mobilis naturally immobilized on 0.7–1.0-mm coke particles. The maximum productivity of 100 g/(L h) was obtained at glucose concentration of 100 g/L and dilution rate of 2.25/h. Kunduru and Pometto (1996a, 1996b) applied PCS chips, consisting of polypropylene with up to 25% wt/wt of various agricultural materials (corn hulls, cellulose, oat hulls, soybean hulls, or starch) and nutrients (soybean flour and mineral salts), as packed-bed reactors in batch and continuous ethanol production. The PCS biofilm packed-bed reactors were operated using pure and mixed cultures of Z. mobilis or Sacch. cerevisiae and mixed cultures of either of these ethanol-producing microorganisms and the biofilm-forming Streptomyces viridosporus T7A. In batch fermentation of Z. mobilis, pure culture yielded a higher maximum productivity of 374 g/(L h) (44% yield) as compared to a maximum productivity of 148 g/(L h) from mixed culture with Strep. viridosporus T7A. On the other hand, mixed cultures of Sacch. cerevisiae resulted in a higher maximum productivity of 190 g/(L h) (35% yield) as compared to an obtained value of 40 g/(L h) (47% yield) from pure culture. Overall, ethanol productivities in batch fermentation of PCS biofilm reactors with Z. mobilis and Sacch. cerevisiae were three- and eightfolds higher than suspension cultures, which yielded maximum productivities of 124 and 5 g/(L h) for Z. mobilis and Sacch. cerevisiae, respectively (Kunduru and Pometto 1996a). Continuous ethanol fermentations using PCS chip biofilm reactors resulted in much higher productivities of 15–100-fold compared to suspension cell reactors, 96 g/(L h) with 50% yield for Z. mobilis, and 76 g/(L h) with 45% yield for Sacch. cerevisiae (Kunduru and Pometto 1996b). The PCS reactors were further developed using higher compositions of nutrients (40% ground soybean hulls, 5% soybean flour, 5% yeast extract, mineral salts, and 50% polypropylene) and applied in continuous ethanol production of Sacch. cerevisiae (Demirci and others 1995, 1997). Two to 10 times higher continuous ethanol production was achieved in

180

Biofilms in the Food Environment

PCS ring biofilm reactors as compared to polypropylene-alone support (PPS) control. Moreover, no significant reduction in ethanol production was found in repeated-batch fermentation even in 0% nitrogen medium with PCS, whereas the ethanol production when using PPS was reduced significantly. Overall, PCS biofilm reactors exhibited high potential in both repeated-batch and continuous ethanol fermentation. This potential stems not only from enhanced productivity but also from the reduced nutrient requirements of the microorganisms. Organic Acids Production One of the earliest applications of biofilm reactors was in the production of vinegar or acetic acid. The vinegar process used a trickling-bed reactor in which an alcoholic solution trickled down through a packed bed of wood chips where a biofilm of Acetobacter spp. developed (Bailey and Ollis 1986). In a later study, Park and Toda (1992) developed a multistage biofilm reactor composed of 10 shallow-flow horizontal reactors for continuous acetic acid production. They reported a very high production rate of 4.3 g/(L h) when a step feeding of ethanol-rich medium was applied. Apart from acetic acid, various organic acids, such as lactic acid, fumaric acid, and citric acid, have been produced using biofilm reactors. Rotating-disk reactors (with self-immobilized Rhizopus mycelia on the plastic disks) have been used for fumaric acid production in Rhizopus oryzae (Cao and others 1996, 1997). Compared to a stirred-tank reactor, the volumetric productivity from biofilm reactor was about threefolds higher (3.78 g/(L h)), while the fermentation time was only one-third of the stirred-tank system. Furthermore, the immobilized biofilm was active for more than 2 weeks with repetitive use and without loss of activity. Citric acid production from biofilm reactors was firstly developed by Lee and others (1989) using immobilized A. niger on polyurethane foam (PUF) in a bubble column reactor. However, this reactor’s productivity was nearly the same as that of suspended cell. Sanroman and others (1994, 1996) studied citric acid production from immobilized A. niger in PUF using fluidized-bed reactor and reported that the adsorption technique was superior to entrapment technique. Ricciardi and others (1997) obtained equivalent levels of citric acid production from biofilm reactors with A. niger immobilized on PUF particles and suspension cell reactor (0.107 g/(L h), 63% yield), similar to Lee and others (1989). However, more recent studies using a rotating-disk contactor (RDC) exhibited a higher potential for a biofilm reactor in citric acid production. Sakurai and others (1999) reported that RDC had a 1.7-times higher productivity than the submerged reactor. A similar study by Wang (2000) used a

Applications of Biofilm Reactors

181

rotating-disk reactor consisting of plastic disks mounted on a horizontal shaft with PUF as a solid support. Wang reported a 3-times higher productivity (0.896 g/(L h)) for this system as compared to their stirred-tank system. As for lactic acid production, Andrews and Fonta (1989) employed a fluidized bed of monosized activated carbon coated with a biofilm of the homolactic fermentative organism Streptococcus thermophilus. They reported that a productivity of 12 g/(L h) was achieved without any pH control and that the outlet product concentration was higher than the completely inhibiting concentration. They also suggested that the form of lactic acid that inhibited the metabolism was adsorbed on the activated carbon. Demirci and others (1993a, 1993b) developed PCS chips, consisting of polypropylene and 25% agricultural products (e.g., corn starch, oat hulls), and then compared the PCS with other solid supports (pea gravels and 3M-macrolite ceramic spheres) for biofilm formation using various biofilm-forming bacterial strains. PCS, which consistently yielded the best biofilm, was used in continuous lactic acid production using three different strains of lactic acid bacteria (Lactobacillus amylophilus, L. casei, and L. delbrueckii mutant DP3, grown optimally at 25, 37, and 45◦ C, respectively) in forms of pure and mixed cultures with Strep. viridosporus T7A, which was used to form a biofilm. Mixed-culture biofilm reactors produced higher levels of lactic acid than lactic acid bacteria alone. The best combination was L. casei subsp. rhamnosus with Strep. viridosporus T7A. Overall, lactic acid production rates were two- to fivefolds higher in pure- and mixed-culture biofilm reactors than in those with suspension cultures. The PCS biofilm reactor was also evaluated for lactic acid production in pure and mixed culture of repeated-batch fermentation (Demirci and others 1995). Compared with suspension culture and PPS (48 g/L), higher concentrations of lactic acid were produced by pureand mixed-culture PCS biofilm reactors (60 and 55 g/L, respectively). PCS was recommended for pure-culture lactic acid production in long-term repeated-batch biofilm fermentation. PCS rings were developed further and evaluated for the effects of different agricultural components on biofilm formation and lactic acid production (Ho and others 1997c). The addition of soybean hulls, yeast extract, or mineral salts in PCS resulted in less hydrophobic supports and enhanced cell attachment. Moreover, the presence of yeast extract in PCS significantly enhanced growth of free and attached cells in the system. The leaching rate of nutrients and lactic acid accumulation of PCS were also studied for application in large-scale and long-term lactic acid production (Ho and others 1997b). PCS blended with dried bovine albumin, dried

182

Biofilms in the Food Environment

bovine erythrocytes, and/or soybean flour showed slower leaching rates than PCS with only yeast extract, and could even still maintain production rate of 1 g/L at the twentieth repeated-batch fermentation. Moreover, lactic acid accumulation in PCS was mainly due to absorption and had no correlation with lactic acid production or biofilm formation. Furthermore, the performance of PCS biofilm reactor in long-term repeated-batch fermentation was compared with that of suspended cell reactors (Ho and others 1997a). It was reported that PCS could stimulate biofilm formation, supply nutrients to attached and free suspended cells, and reduce medium channeling for lactic acid production. Furthermore, compared with conventional repeated-batch fermentation, PCS biofilm reactors shortened lag time by three- to sixfold, increased lactic acid production by 40–70%, and shortened total fermentation time by 28–61% (at yeast extract concentrations of 0.4 and 0.8%, respectively). Overall, PCS biofilm reactors significantly improved the lactic acid production with reduced complexnutrient addition. Velazquez and others (2001) employed periodic spiking of concentrate glucose solution for lactic acid production using repeated fed-batch PCS rings biofilm reactors and suspended cell reactors. The obtained results were similar to previous studies in that the PCS biofilm reactors consistently outperformed the suspended cell reactors, requiring less yeast extract and yielding higher lactic acid production rates. Also, Cotton and others (2001) developed a different design of PCS orientation in which PCS tubes were fixed on agitator shaft instead of a packed bed of small pieces of PCS rings (Figure 8.6) for continuous lactic acid production in PCS biofilm reactors. The new design attempted to solve problems that frequently occurred in packed- and fluidized-bed and cell recycling bioreactors, such as reduced flows and membrane fouling. This PCS design demonstrated positive effects on cell density, production rates, and yields in the continuous lactic acid production (9.0 g/(L h), 70% yield). For succinic acid production using Actinobacillus succinogenes, Urbance and others (2003) screened 20 different PCS blends for biofilm formation and succinic acid production in repeated-batch flask studies with no observed correlation between biomass formation and succinic acid production. However, selected PCS blends demonstrated higher succinic production over suspended cell fermentation. In continuous and repeated-batch biofilm fermentations using Actin. succinogenes, PCS containing 50% (wt/wt) polypropylene, 35% (wt/wt) ground soybean hulls, 5% (wt/wt) dried bovine albumin, 2.5% (wt/wt) soybean flour, 2.5% (wt/wt) yeast extract, 2.5% (wt/wt) dried red blood cells, and 2.5% (wt/wt) peptone, or polypropylene tubes (control), 8.5 cm in length, was arranged around the agitator shaft in a grid formation (Urbance and others 2004).

Applications of Biofilm Reactors

183

For PCS continuous biofilm reactors, the highest final succinic acid concentrations (10.1 and 10.4 g/L) and percentage yields (62.6 and 71.6%) occurred at the dilution rate of 0.2/h, whereas the suspended cell bioreactor continuous fermentation achieved 7.0 g/L succinic acid at a dilution rate of 1.0/h with 76% yield. This still needs to be verified in long-term (3–5 months) continuous fermentations. On the other hand, in repeatedbatch PCS biofilm reactors, succinic acid production rate is halted when concentrations of 40 g/L is reached. Thus, glucose concentration of 40 g/L results in high percentage yield of succinic acid (86.7 %) and 38 g/L succinic acid in PCS biofilm reactors, while the suspended cell reactors illustrated significantly lower percentage yields (45%) at 40 g/L glucose. Therefore, to achieve high succinic acid percentage yields in continuous and repeated-batch fermentations of Actin. succinogenes, suspended cell continuous fermentations at a dilution rate of 1.0/h and PCS repeatedbatch fermentations with initial glucose concentrations of 40 g/L were recommended, respectively. Antibiotic Production Comparing to organic acids productions, biofilm reactors have been sparingly employed in the production of antibiotics. Three-phase fluidized-bed biofilm reactors have been used for production of penicillin and reported that a complete-mixed contacting pattern yielded better results than a plug flow pattern in both product concentrations and specific productivities (Park and Wallis 1984). Moreover, the inhibitory effect of the carbon source was less pronounced in complete-mixed contacting pattern. Another antibiotic investigated using biofilm reactor was cephalosporin. C. Park and Seo (1988) used a fluidized-bed reactor with Cephalosporium acremonium entrapped in celite particles. Despite an unstable biofilm due to morphological differentiation, the overall performance of the fluidized-bed reactor for cephalosporin C production was improved by 1.9-fold compared to their suspended cell reactor. Park and others (1989) found no improvement in biofilm stability when methionine was supplemented into the system. A similar result was also obtained from a more recent study (Srivastava and Kundu 1999) using airlift reactor with pellets and Siran carriers. Although the productivity was increased by 18–28% probably due to enhanced mass transfer in fermentation broth, instability of biofilm was observed. As for bacteriocin production, a recent study from Bober (2002) using packed-bed PCS rings biofilm reactor with Lactococcus lactis in nisin production showed that the nisin production rate was not significantly different from the value obtained from the suspended cell reactor. However,

184

Biofilms in the Food Environment

inadequate mixing in the reactor was observed. At present, our research group is working on an optimization of nisin production in PCS biofilm reactor by adjusting culture conditions, such as nutrient sources, agitation, and pH control. Enzyme Production An enhancement for enzyme production using biofilm reactor has been reported in many studies. The continuous production of cellulase using biofilm reactor using Trichoderma viride QM9123 (using stainless steel biomass support particles in spouted bed fermenter) was reported to exhibit higher yield and productivity compared to suspended cell reactor (Fukuda and others 1986; Webb and others 1986). Oriel (1988) reported a fivefold increase in thermostable amylase production using immobilized recombinant Escherichia coli (EC147) on silicone foam support. Fukuda and others (1986) reported an application of acetone-dried cells of Rhizopus chinensis immobilized within PUF particles as a lipase catalyst in industrial interesterification of fats and oils (Kyotani and others 1991; Hara and Nakashima 1996, 1998). Overall, the specific activity of dried cells within BSPs increased sevenfold over that of freely suspended cells, and the biofilm reactor using biomass support particles demonstrated higher lipase activity over suspended cell reactor in batch, fed-batch, and continuous processes. The potential of the BSP-immobilized fungus system for practical applications in transesterification of phospholipids has been discussed in detail (Hara and Nakashima 1998). Production of lignin peroxidase was found to be increased when nilon web was used in immobilized system (Linko 1992). Recently, the first demonstration of process scale-up of a membrane gradostat reactor for continuous ligninolytic enzymes production, using Phanerochaete chrysosporium, had been reported. The fungus, P. chrysosporium ME446, was immobilized by reverse filtration on an external unskinned ultrafiltration capillary membranes and sevenfold increase in enzyme productivity was achieved (Govender and others 2003). On the other hand, Khiyami and others (2006) reported that P. chrysosporium (ATCC 24725) produced lignin peroxidase (LiP) and manganese peroxidase (MnP) in defined medium (pH 4.5) with PCS tubes attached to the agitator shaft of stirred-tank biofilm reactors. The formation of P. chrysosporium biofilm on PCS was essential for the production of MnP and LiP. The production of both enzymes’ MnP activity of 0.0473 and 0.0493 U/mL and Lip activity of 0.03 and 0.0277 U/mL, respectively, was observed. The bioreactor was operated as a repeat batch, and no reinoculation was required between 14 batches. Peroxidases production was influenced by

Applications of Biofilm Reactors

185

purging of the bioreactor at 0.005 vvm with pure oxygen for 5 min or continuous aerating with mixture of air and oxygen. Addition of veratryl alcohol and MnSO4 on day 0 with 300 rpm agitation and continuous aeration at 0.005 vvm produced MnP on day 3.

Summary Many laboratory and pilot-scale studies have demonstrated several advantages of biofilm reactors in the productions of various value-added products, especially in alcohol and organic acid productions. Biofilm reactors, like other immobilized reactors, not only yield higher production rates, but also possess greater potential compared with suspended cell reactors for development into continuous culture (which is significantly more productive in terms of fermenter downtime per unit of product manufactured). Combining exceptional stability and lower nutrient requirements, biofilm reactors exhibit high potential for applications in the large-scale production of many products such as alcohols, organic acids, antibiotics, and enzymes. Nevertheless, before biofilm reactors can be applied in industrial-scale production, many additional scale-up studies on several parameters such as culture conditions, mass and heat transfer constraints, and kinetics are necessary.

References Acuna ME, Villanueva C, Cardenas B, Christen P, Revah S. 2002. The effect of nutrient concentration on biofilm formation on peat and gas phase toluene biodegradation under biofiltration conditions. Proc Biochem 38:7–13. Allan VJM, Callow ME, Macaskie LE, Paterson-Beedle M. 2002. Effect of nutrient limitation on biofilm formation and phosphatase activity of a Citrobacter sp. Microbiology 148:277– 288. Andrews GF, Fonta JP. 1989. A fluidized-bed continuous bioreactor for lactic acid production. Appl Biochem Biotechnol 20–21:375–390. Atkinson B, Black GM, Lewis PJS, Pinches A. 1979. Biological particles of given size, shape, and density for use in biological reactors. Biotechnol Bioeng 21:193–200. Atkinson B, Swilley EL. 1963. A mathematical model for the trickling filter. In Proc. 18th Ind Waste Conf., Purdue University, Lafayatte, IN, pp. 706–737. Azeredo J, Visser J, Oliveira R. 1999. Exopolymers in bacterial adhesion: Interpretation in terms of DLVO and XDLVO theories. Colloids Surf B Biointerfaces 14:141–148. Bailey JE, Ollis D. 1986. Biochemical Engineering Fundamentals, 2nd Edition, McGrawHill Book Company, New York. Bayston R. 2000. Biofilm infections on implant surfaces. In: Evans, L.V. (ed) Biofilms: Recent Advances in Their Study and Control. Harwood Academic, Amsterdam, pp. 117–131.

186

Biofilms in the Food Environment

Bober JA. 2002. Nisin Fermentation by Lactococcus lactis supsp. Lactis in Biofilm Reactors Using Plastic Composite Supports (PCS). Master Thesis, Pennsylvania State University. Bott TR. 1995. Biological growth on heat exchanger surfaces. In Fouling of Heat Exchangers (TR Bott, Editor). Elsevier, Amsterdam, pp. 223–267. Bryers JD. 1987. Biologically active surfaces: Processes governing the formation and persistence of biofilms. Biotechnol Prog 3:57–68. Bryers JD. 2000. Biofilm formation and persistence. In Biofilms II: Process Analysis and Applications (JD Bryers, Editor). Wiley-Liss, Inc., New York, pp. 45–88. Busscher HJ, Bos R, van der Mei HC. 1995. Initial microbial adhesion is determinant for the strength of biofilm adhesion: Hypothesis. FEMS Microbiol Lett 128:229–234. Busscher HJ, van der Mei HC. 2000. Initial microbial adhesion events: Mechanisms and implications. In Community Structure and Co-operation in Biofilms (D Allison, P Gilbert, H Lappin-Scott, M Wilson, Editors). Cambridge University Press, Cambridge, pp. 25–36. Cabral JMS, Tramper J. 1994. Bioreactor design. In Applied Biocatalysis (JMS Cabral, D Best, L Boross, J Tramper, Editors). Harwood Academic Publishers, Switzerland, pp. 333–370. Cao N, Du J, Chen C, Gong CS, Tsao GT. 1997. Production of fumaric acid by immobilized Rhizopus using rotary biofilm contactor. Appl Biochem Biotech 63–65:387–394. Cao N, Du J, Gong CS, Tsao GT. 1996. Simultaneous production and recovery of fumaric acid from immobilized Rhizopus oryzae with a rotary biofilm contactor and an adsorption column. Appl Environ Microbiol 62:2926–2931. Characklis WG, Marshall KC. 1990. Biofilms. John Wiley and Sons, New York. Chmielewski RAN, Frank JJ. 2003. Biofilm formation and control in food processing facilities. Compr Rev Food Sci Food Saf 2:22–32. Chung IJ, Park YS. 1983. Ethanol fermentation by S. cerevisiae in a bioreactor packed vertically with ceramic rods. In Proc.-Pac. Chem. Eng. Congr., 3rd, Korean Institute of Chemical Engineering, Seoul, Korea, Volume 4, pp. 174–179. Cotton JC, Pometto AL III, Gvozdenovic-Jeremic J. 2001. Continuous lactic acid fermentation using a plastic composite support biofilm reactor. Appl Microbiol Biotechnol 57:626–630. Demirci A, Pometto AL III, Ho K-LG. 1995. Continuous ethanol production in biofilm reactors containing plastic composite rings and disks. In Proc.-Biomass Conference of the Americas: Energy, Environment, Agricultural and Industry, 2nd, 21–24 August 1995, Portland, OR, National Renewable Energy Laboratory, Golden, CO. Demirci A, Pometto AL III, Ho K-LG. 1997. Ethanol production by Saccharomyces cerevisiae in biofilm reactors. J Ind Microbiol Biotechnol 19:299–304. Demirci A, Pometto AL III, Johnson KE. 1993a. Evaluation of biofilm reactor solid support for mixed-culture lactic acid production. Appl Microbiol Biotechnol 38:728–733. Demirci A, Pometto AL III, Johnson KE. 1993b. Lactic acid production in a mixed-culture biofilm reactor. Appl Microbiol Biotechnol 59:203–207. Dempsey MJ. 1990. Ethanol production by Zymomonas mobilis in a fluidized bed fermenter. In Physiology of Immobilized Cells (JAM de Bong, J Visser, B Matiasson, J Tramper, Editors). Elsevier Science Publishers BV, Amsterdam, pp. 137–148. Ebihara T, Bishop PL. 2002. Effect of acetate on biofilms utilized in PAH bioremediation. Environ Eng Sci 19:305–319. Fukuda H. 1995. Immobilized microorganism bioreactors. In Bioreactor System Design (JA Asenjo, JC Merchuk, Editors). Marcel Dekker, New York, pp. 339–375. Fukuda H, Webb C, Atkinson B. 1986. Continuous cellulase production using immobilized cells. In Process Engineering Aspects Immobilised Cell Systems (C Webb, Editor). Institute of Chemical Engingeering, Rugby, U.K., pp. 309–315.

Applications of Biofilm Reactors

187

Gassey GG, Bryers, JD. 2000. Biofouling of engineered materials and systems. In Biofilms II: Process Analysis and Applications (JD Bryers, Editor). Wiley-Liss, Inc., New York, pp. 237–279. Gjaltema A, Arts PAM, van Loosdrecht MCM, Kuenen JG, Heijnen JJ. 1994. Heterogeneity of biofilm in rotating annular reactors: Occurrence, structure, and consequences. Biotechnol Bioeng 44:194–204. Govender S, Jacobs EP, Leukes WD, Pillay VL. 2003. A scalable membrane gradostat reactor for enzyme production using Phanerochaete chrysosporium. Biotechnol Lett 25:127– 131. Hara F, Nakashima T. 1996. Transesterification of phospholipids by acetone-dried cells of a Rhizopus species immobilized on biomass support particles. J Am Oil Chem Soc 73:657–659. Hara F, Nakashima T. 1998. Enzymic conversion of phospholipids using an acetone-dried fungus immobilized on biomass support particles. Recent Res Dev Oil Chem 2:15– 29. Hermanowicz SW. 1998. A model of two-dimensional biofilm morphology. Water Sci Technol 37(4–5):219–222. Ho K-LG, Pometto AL III, Hinz PN. 1997a. Optimization of L-(+)-lactic acid production by ring and disk plastic composite supports through repeated-batch biofilm fermentation. Appl Environ Microbiol 63:2533–2542. Ho K-LG, Pometto AL III, Hinz PN, Demirci A. 1997b. Nutrient leaching and end product accumulation in plastic composite supports for L-(+)-lactic acid biofilm fermentation. Appl Environ Microbiol 63:2524–2532. Ho K-LG, Pometto AL III, Hinz PN, Dickson JS, Demirci A. 1997c. Ingredient selection for plastic composite supports for L-(+)-lactic acid biofilm fermentation by Lactobacillus casei subsp. Rhamnosus. Appl Environ Microbiol 63:2516–2523. Howell JA, Atkinson B. 1976. Sloughing of microbial film in trickling filters. Water Res 10:307–316. Jackson K, Keyser R, Wozniak DJ. 2003. The role of biofilms in airway disease. Semin Respir Crit Care Med 24(6): 663–670. Jin G, Englande AJ Jr, Qiu YL. 2003. An integrated treatability protocol for biotreatment/bioremediation of toxic pollutants generated by chemical industries. J Environ Sci Health A Toxic/Hazard Subst Environ Eng 38:597–607. Karamanev DG, Nikolov LN. 1992. Bed expansion of liquid–solid inverse fluidization. AlChE J 38:1916–1922. Karamanev DG, Nikolov LN. 1996. Application of inverse fluidization in wastewater treatment: From laboratory to full-scale reactors. Environ Prog 15:194–196. Keevil CW, Walker JT. 1992. Normarski DIC microscopy and image analysis of biofilms. BINARY 4:93–95. Khiyami MA, Pometto AL III, Brown RC. 2005a. Detoxification of corn stover and corn starch pyolysis liquors by Pseudomonas putida and Streptomyces setonii suspended cells and PCS biofilms. J Agric Food Chem 53:2978–2987. Khiyami MA, Pometto AL III, Brown RC. 2005b. Detoxification of Corn stover and corn starch pyrolysis liquors by ligninolytic enzymes of Phanerochaete chrysporium. J Agric Food Chem 53:2969–2977. Khiyami MA, Pometto AL III, Kennedy WJ. 2006. Lignolytic enzyme production by Phanerochaete chrysosporium in PCS biofilm sitrred tank bioreactor. J Agric Food Chem 54:1693–1698. Kunduru RM, Pometto AL III. 1996a. Evaluation of plastic composite supports for enhanced ethanol production in biofilm reactors. J Ind Microbiol 16:241–248.

188

Biofilms in the Food Environment

Kunduru RM, Pometto AL III. 1996b. Continuous ethanol production by Zymomonas mobilis and Saccharomyces cerevisiae in biofilm reactors. J Ind Microbiol 16:249–256. Kyotani S, Nakashima T, Izumoto E, Fukuda H. 1991. Continuous interesterification of oils and fats using dried fungus immobilized in biomass support particles. J Ferm Bioeng 71:286–288. Lazarova V, Manem J. 2000. Innovative biofilm treatment technologies for water and wastewater treatment. In Biofilms II: Process Analysis and Applications (JD Bryers, Editor). Wiley-Liss, Inc., New York, pp. 159–206. Lee YH, Lee CW, Chang HN. 1989. Citric acid production by Aspergillus niger immobilized on polyurethane foam. Appl Microbiol Biotechnol 30:141–143. Linko S. 1992. Production of Phanerochaete chrysosporium lignin peroxidase. Finland Biotechnol Adv 10:191–236. Massol-Deya AA, Whallon J, Hickey RF, Tiedje JM. 1995. Channel structures in aerobic biofilms of fixed-film reactors treating contaminated groundwater. Appl Environ Microbiol 61:79–777. Melo LF, Oliveira R. 2001. Biofilm reactors. In Multiphase Bioreactor Design (JMS Cabral, M Mota, J Tramper, Editors). Taylor & Francis Inc, New York, pp. 271–308. Messing RA, Opperman RA. 1979. Pore dimensions for accumulating biomass. I: Microbes that reproduce by fissing or by budding. Biotechnol Bioeng 21:49–58. Norwood DE, Gilmour A. 2000. The growth and resistance to sodium hypochlorite of Listeria monocytogenes in a steady-state multispecies biofilm. J Appl Microbiol 88:512– 520. Nyvad B, Fejerskov O. 1997. Assessing the stage of caries lesion activity on the basis of clinical and microbiological examination. Commun Dent Oral Epiderm 25:69–75. O’Flaherty V. 2003. Biofilms in wastewater treatment systems. In Biofilms in Medicine, Industry and Environmental Biotechnology: Characteristics, Analysis and Control (P Lens, Editor). IWA Publishing, London, pp. 132–159. Oliveira R. 1992. Physico-chemical aspects of adhesion. In Biofilms—Science and Technology (LF Melo, TR Bott, M Fletcher, B Capdeville, Editors). Kluwer Academic Publishers, Dordrecht, pp. 45–58. Oriel P. 1988. Immobilization of recombinant Eschericia coli in silicone polymer beads. Enzyme Microb Technol 10:518–523. Park YH, Kim EY, Seo WT, Jung KH, Yoo YJ. 1989. Production of cephalosporin C in a fluidized-bed bioreactor. J Ferm Bioeng 67:409–414. Park YH, Seo WT. 1988. Production of cephalosporin C in a fluidized-bed bioreactor. Sanop Misaengmul Hakhoechi 16:25–32. Park YS, Toda K. 1992. Multi-stage biofilm reactor for acetic acid production at high concentration. Biotechnol Lett 14:609–612. Park YH, Wallis DA. 1984. Steady-state performance of a continuous biofilm fermentor system for penicillin production. Korean J Chem Eng 1:119–128. Patching JW, Fleming GTA. 2003. Industrial biofilms: Formation, problems and control. In Biofilms in Medicine, Industry and Environmental Biotechnology: Characteristics, Analysis and Control (P Lens, Editor). IWA Publishing, London, pp. 568–590. Pereira MA, Alves MM, Azeredo J, Mota M, Oliveira R. 2000. Physico-chemical properties of porous microcarriers in relation with the adhesion of an anaerobic consortium. J Ind Microbiol Biotechnol 24:181–186. Ricciardi A, Parente E, Volpe F, Clementi F. 1997. Citric acid production from glucose by Aspergillus niger immobilized in polyurethane foam. Microbiol Enzimol 47:63–76. Sakurai A, Imai H, Sakakibara M. 1999. Citric acid production using biofilm of Aspergillus niger. Recent Res Dev Biotechnol Bioeng 2:1–13.

Applications of Biofilm Reactors

189

Sanroman A, Feijoo G, Lema JM. 1996. Immobilization of Aspergillus niger and Phanerochaete chrysosporium on polyurethane foam. Prog Biotechnol 11:132–135. Sanroman A, Pintado J, Lema JM. 1994. A comparison of two techniques (adsorption and entrapment) for the immobilization of Aspergillus niger in polyurethane foam. Biotechnol Tech 8:389–394. Shimp RJ, Pfaender FK. 1982. Effect of surface area and flow rate on marine bacterial growth on activated carbon columns. Appl Environ Microbiol 44:471–476. Sousa M, Azeredo J, Feijo J, Oliveira R. 1997. Polymeric supports for the adhesion of a consortium of autotrophic nitrifying bacteria. Biotechnol Tech 11:751–754. Srivastava P, Kundu S. 1999. Studies on cephalosporin C production in an air lift reactor using different growth modes of Cephalosporium acremonium. Proc Biochem 34:329–333. Stewart PS, Murga R, Srinivasan R, de Beer D. 1995. Biofilm structural heterogeneity visualized by three microscopic methods. Water Res 29:2006–2009. Teixeira P, Oliveira R. 1999. Influence of surface characteristics on the adhesion of Alcaligenes denitrificans to polymeric substrates. J Adhes Sci Tech 13:1287–1294. Urbance SE, Pometto AL III, DiSpirito AA, Demirci A. 2003. Medium evaluation and plastic composite support ingredient selection for biofilm formation and succinic acid production by Actinobacillus succinogenes. Food Biotechnol 17:53–65. Urbance SE, Pometto AL III, DiSpirito AA, Denli Y. 2004. Evaluation of succinic acid continuous and repeated-batch biofilm fermentation by Actinobacillus succinogenes using plastic composite support bioreactors. Appl Microbiol Biotechnol 65:664–670. van Loosdrecht MCM, Heijnen JJ, Eberl H, Kreft J, Picioreanu C. 2002. Mathematical modeling of biofilm structure. Antonie Van Leeuwenhoek 81:245–256. van Loosdrecht MCM, Lyklema J, Norde W, Schraa G, Zehnder AJB. 1987. The role of bacterial cell wall hydrophobicity in adhesion. Appl Environ Microbiol 53:1893–1897. Veiga MC, Mendez R, Lema JM. 1992. Development and stability of biofilms in bioreactors. In Biofilms—Science and Technology (LF Melo, TR Bott, M Fletcher, B Capdeville, Editors). Kluwer Academic Publishers, Dordrecht, pp. 421–434. Velazquez AC, Pometto AL III, Ho K-LG, Demirci A. 2001. Evaluation of plastic-composite supports in repeated fed-batch biofilm lactic acid fermentation by Lactobacillus casei. Appl Microbiol Biotechnol 55:434–441. Von Canstein H, Li Y, Leonhauser J, Haase E, Felske A, Deckwer W-D, Wagner-Dobler I. 2002. Spatially oscillating activity and microbial succession of mercury-reducing biofilms in a technical-scale bioremediation system. Appl Environ Microbiol 68:1938–1946. Wang J. 2000. Production of citric acid by immobilized Aspergillus niger using a rotating biological contactor (RBC). Bioresour Technol 75:245–247. Webb C, Fukuda H, Atkinson B. 1986. The production of cellulase in a spouted bed fermentor using cells immobilized in biomass support particles. Biotechnol Bioeng 28:41–50. White DC. 1984. Chemical characterization of films. Life Sci Res Rep 31:159–176. Willaert RG, Baron GV, de Backer L. 1996. Immobilised Living Cell Systems: Modelling and Experimental Methods. John Wiley and Sons, Chichester, England, Chap. 3. Wimpenny J, Manz W, Szewzyk U. 2000. Heterogeneity in biofilms. FEMS Microbiol Rev 24:661–671. Wirtanen G, Saarela M, Mattila-Sandholm T. 2000. Biofilms-impact on hygiene in food industries. In Biofilms II: Process Analysis and Applications (JD Bryers, Editor). Wiley-Liss, Inc., New York, pp. 327–372.

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

Index

acid sanitizers, 95 acid tolerance, 37 acid-sensitive, S.flexneri mutants, 38 active immobilization, disadvantages, 172 active or artificial immobilization, 172 acyl-homoserine lactone, autoinducer, 5 adsorption phenomenon, 172 alcohol production, 179 alkaline FitTM produce wash, 42 animal model, streptomycin-treated mouse, 144 anionic sanitizers, 95 antibiotic production, 183 antimicrobial resistance, 5 atomic force microscopy (AFM), 12 autoaggregation, 112 autoinducer. See also N-acyl-homoserine lactones (AHL) AI-2, 10, 11 in Gram-negative microorganism, 10 synthesis of, 11 Bacillus cerus, ecology of microbes, 21 Bacillus flavothermus, 7. See also disinfectants bacteria–bacteria communication, 160 bacterial growth rate in situ, 144 bacterial populations, in intestinal tract, 133 bacterial strain identification, parameters, 107 beneficial bacterial biofilms, 153–161 biofilm communities, intercellular communication, 115–116 biofilm development role of conjugation, 114–115 use of antibiotics, 120 biofilm ecosystem development, 111–112 biofilm formation, 168 biofilm reactors advantages of, 171–172

applications of, 176 biomass support particles, 174 immobilization, 172 types of, 173 biofilm structure model, 170 biofilm-forming abilities, degrees, 11 biofilms antimicrobial resistance, 5, 6 communication within, 157 control, 120 displacement of, 157 formation of, 157, 168 heterogeneous nature, 6 image of, 12 structure of, 168, 170 biofilms, microbial diversity analysis, 106–111 by Denaturing Gradient Gel Electrophoresis, 108 by DNA sequencing, 106 by fluorescent in situ hybridization, 110 by microarray, 109 by molecular probes, 109 biofilms, microscopic examination, 11 atomic force microscopy (AFM), 12 confocal scanning laser microscopy (CSLM), 12–14 differential interference contrast (DIC), 14 environmental SEM, 12–13 fluorescence microscope, 14 modulation contrast microscopy, 13 scanning electron microscopy (SEM), 12–13 transmission electron microscopy (TEM), 12–13 biofouling, 3, 128, 167 Biomass Support Particles (BSPs), 172, 184 BSP-immobilized fungus system, 184

191

192

Index

C. perfringens, ecology of microbes on produce, 21 cabbage, S. flexneri levels, 33 Campylobacter, in parsley sample, 20, 35 carrot salad, S. flexneri level, 33 carrots, psychrotrophs, 25 cell immobilization methods, 172 cell-to-cell signaling systems, 11, 115–116 CFR. See Code of Federal Regulations chemical sanitizers effectiveness and inactivation of biofilms, 82–90 chlorine as enterotoxigenic E. coli, 28 concentration in wash water, 26 disinfection of fresh produce, 27–28 cilantro (Coriandrum sativuum) Salmonella population, 36 Salmonella serovar Thompson, 36–37 cleaning and biofilm control, 74 and sanitizing, 25 compounds in, 74–75 cleaning agents effectiveness, 76–78 and concentration, 79 cleaning methods for biofilms removal, 79 Clostridium botulinum, ecology of microbes on produce, 21 Coaggregation between genetically distinct microorganisms, 112 Code of Federal Regulations, 39 coleslaw, S. flexneri levels, 33 commercial FitTM produce wash, efficacy of, 40, 42 confocal scanning laser microscopy (CSLM), 12–14 conjugation, 114, 119, 131 cooperative behavioral patterns, 116 crab salad, S. flexneri level, 33 cucumbers, psychrotroph population, 25 cultured and uncultured representatives, 134 denaturing Gradient Gel Electrophoresis (DGGE), 108 differential interference contrast (DIC), 14 disinfectants, 7 DNA sequencing, 106 DsRed P. agglomerans. See also cilantro, 37 D-values, 98 E. coli BJ4, in intestine, 145 efficacy of spray, 29

solubility of, 28 environmental persistence, 117 Environmental Protection Agency (EPA), 39 enzyme production, 184 Erwinia Gram-negative rods, 25 exopolymeric substances (EPS), 5, 7, 9, 10, 15 exopolysaccharides, 9 extracellular polymeric substances (EPS), 50 fatty acid sanitizers, 96 fecal flora coliforms, 21 contamination, 21–22 FitTM treatments, 42 Fixed-bed reactors, 173 Flavobacterium, 7, 61. See also disinfectants Fluorescens, 9. See also food processing surfaces fluorescent in situ hybridization, 110 food environment cell-to-cell communications in, 119–120 mixed culture biofilms, 119–120 food industry impact of biofilms in, 3–5 food processing surfaces, 8 foodborne outbreaks, 21 involving Shigella, 30–32 fresh-processed produce colonization of microorganisms on surface of, 22 consumption, 19 efficacy of sanitizer on microorganisms of, 39 as growing industry, 19 pathogen growth on, 22 produce related outbreaks, 20 spoilage of, 23 Generally recognized as safe (GRAS) status. See also CFR, 39 GFP Salmonella serovar Thompson on cilantro, 37 Gram-negative rods Enterobacter, 24 Erwinia, 24 Pseudomonas, 24–25 Growth-phase-dependent sigma factorδ 38 , 38 Gut mucosal biofilms, definition, 128 2-Heptyl-3-hydroxy-4-quinolone (PQS), 10 heterogeneous model. See also biofilms, 170 heterogeneous mosaic model, 170 horizontal gene transfer, 119

Index hot water sanitation, 97, 98 human digestive system inactivation of, 73–101 initial attachment of, 49 iodine efficacy, 94 molecular attributes, 58–59 ozone as sanitizer, 93 prediction model for, 99–100 prokaryotic diversity of epithelial mucosal biofilm, 127–145 regulatory pathways, 58–59 sequential treatment with acetic acid and sodium hydroxide, 95 structure of, 50 initiation, in biofilm formation, 49 intestinal tract biofilms, members of underrepresented Phyla, 139 irrigation water, pathogen detection, 21, 22 Klebsiella pneumoniae, mixed species biofilms, 6, 21 Knockout mutants, biofilm production, 11 Lactic acid bacteria, ecology of microbs on produce, 24 Lactococcal CluA protein, 115 lasI gene product, 11 lasR-lasI. See also cell-to-cell signaling systems, 11 lettuce bacteria count, 25 E. coli population, 25 microbial quality, 24 psychrotroph populations, 25 shredding temperature, 32 Leuconostocs ecology of microbes on produce, 24 lignin peroxidase, production, 184 liquid shear forces, 168, 169, 174. See also biofilm Listeria monocytogenes adsorption of nisin, 64 biofilm development, 47–67 chlorine dioxide (ClO2 ) sanitizer, 91 effects of halogens, 81 biofilm, prevention and control of, 64–67 effectiveness of chlorine, 66 effectiveness of peracetic acid, 66 electrolyzed (EO) water sanitizer, 92 environmental factors impact, 54 acidity, 57 five-strain cocktail of, 100

193

nutrients, 55 surface, 55 temperature, 56 EPS formation, 51 food-borne pathogen, 47 inactivation of biofilm, 73, 74 strain variability, 52–54 structural characteristics, 50–52 Listeria Salmonella typhimurium, 28. See also chlorine Listeria Staphylococcus aureus, produce wash, 40 Listeria, food processing environments, 9 Manganese peroxidase (MnP), 184 MexABOprM pump, 118 microarray, 109 microbes on produce, ecology of, 21 microbial biofilms basic units of, 4 cells in, 5–6 definition of, 3 formation of, 4 microbial heat inactivation predictive models, 98 microbial mucolysis, 132 microbiota, of digestive tract, 135 microorganisms, potential sources, 21 minimal processing of vegetables, 23 minimally processed produce deterioration of, 19 features, 19 microflora of, 24 storage temperature of, 22 mixed culture biofilms, 59 public health risks, 117 safety, 117 modeling heat inactivation of biofilms, 98 modified Welshimer’s broth (MWB), 54 molecular probes, 109 mucus biofilm niche, 130–131 mucus layer and goblet cell biology, 128 multispecies Biofilms, 6 multispecies biofilms, 6, 7 mushroom or tulip model, 170 N-(3-oxododecanoyl)-L-homoserine lactone, autoinducer. See also cell-to-cell signaling systems; quorum sensing, 10, 11, 113 N-acyl-homoserine lactones (AHL), autoinducer in G (-ve). See also cell-to-cell signaling systems; quorum sensing, 10, 113

194

Index

ofloxacin resistance, P. aeruginosa biofilms, 118 organic acids production, 180 outbreaks foodborne, 21 produce-related, 20 Pseudomonas aeruginosa biofilms, 6, 10, 40, 59 P. cepacia, produce wash, 40 P. chloroaphis, on Cilantro, 36 P. fluorescens, food processing surfaces, 9 P. fluorescens, food processing surfaces, 9 P. fragi, mixed cultured biofilms, 60 Parsley herb, 34 Parsley, S. sonnei population, 34 Passive or natural immobilization, 172 PCS (plastic composite support) biofilm reactors, 175, 176 pectinolytic strains, of Pseudomonas, 24 peroctanoic acid-based sanitizer, 95 peroxygen compounds, 94 produce contamination of, 24 pathogenic microorganisms, 22 washing of, 26 produce sanitizers, efficacy of, 42 produce wash, 39 produce-related outbreaks, 20 pseudo-homogeneous model, 170 quorum sensing, 10, 113. See also cell-to-cell signaling systems 16S rRNA gene, 107–108 ribosomal intergenic spacer analysis (RISA), 108 rpoS-dependent, process in Gram-negative bacteria, 118 Saccharomyces cerevisiae, surface sanitizer, 95 salad vegetables, E. coli O157:H7, 25 Salmonella Typhimurium, surface sanitizer, 95 sanitizing agents and biofilm inactivation, 81 Scallions, consumption, 31. See also Shigella flexneri 6A scanning electron microscopy (SEM), 12–13 Serratia mixed culture biofilms, 25 shear conditions impacts on biofilm formation, 3, 4, 10, 168 Shigella flexneri 6A, multistate outbreak, 31

Shigella sonnei, outbreaks, 30–31 shredded cabbage, S. sonnei survival, 33 signaling processes, 160 single culture L. monocytogenes biofilm, 48 specific genetic predispositions, 141 Staphylococcus, 9, 10, 34. See also Food Processing Surfaces S. aureus, 10, 41. See also cell–cell communication S. boydii, stool culture, 30 S. choleraesuis, produce wash, 40 S. enterica Derby on Cilantro, 37 Enteritidis on Cilantro, 37 Newport on Cilantro, 37 Thompson on Cilantro, 37 S. epidermis, 10 S. flexneri, 6A outbreak, 31–32, 38 and survival on salads, 33 S. Montevideo, on tomato surfaces, 29 S. sciuri biofilm, 61 S. sonnei, outbreaks, 30–32, 34 S. Thompson outbreak, 36 S. typhimurium, on Parsley, 35, 38 S. xylosus biofilms, 62 stationary phase E. coli K-12, 37 stool versus mucosal populations, 141 strain variability, 52 successional colonization of digestive tract, 140 surface type, 80 surfactant sanitizers, 96 survivability on fresh produce, 32 temperature and contact time, 79–80 theory of biofilms, 3 theory of, 3 thermophile sporeforming, 7 thermotolerant coliforms, 21 tomatoes survival of Salmonella Montevideo, 29 uptake of Salmonella, 26 transmission electron microscopy (TEM), 12–13 vaginal microbiota, 157–158 Vibrio, 10, 11, 116. See also autoinducer virulence genes, transcription, 11 Water in cleaning, role of, 80 Yersinia enterocolitica, chlorine effect, 28, 34