Antimicrobials in Food [4 ed.] 2020020320, 9780367178789, 9780429058196

Table of contents :
Cover
Half Title
Title Page
Copyright Page
Table of Contents
Preface
Acknowledgments
Editors
List of Contributors
Chapter 1 Food Antimicrobials – An Introduction
1.1 Role of Additives in Food
1.2 Definition and Function of Chemical Food Preservatives
1.3 Selection of Antimicrobials
1.3.1 Antimicrobial Spectrum
1.3.2 Physico-Chemical Properties of the Antimicrobial
1.3.3 Food-Related Factors
1.3.4 Process Factors
1.3.5 Resistance Development
1.4 Considerations in the Applications of Food Antimicrobials
1.4.1 Toxicological Safety
1.4.2 Labeling with Respect to Antimicrobials
1.5 Future of Antimicrobials
References
Chapter 2 Methods for Activity Assay and Evaluation of Results
2.1 Introduction
2.2 Factors That Affect Antimicrobial Activity
2.3 Current Methodology
2.4 In Vitro Methods
2.4.1 Agar Diffusion
2.4.1.1 Dilution Methods
2.4.1.2 Gradient Plates
2.4.1.3 Automated Methods
2.4.1.4 Vapor-Phase or Volatile Compounds Methods
2.5 Descriptive Methods
2.6 Combined Antimicrobial Systems
2.7 Application Studies
2.7.1 Interpretation of Results
2.8 Acknowledgments
References
Chapter 3 Sodium Benzoate and Benzoic Acid
3.1 Introduction and Historical Background
3.2 Physical and Chemical Properties and Natural Occurrence
3.3 Antimicrobial Activity
3.3.1 Spectrum of Action
3.3.2 Influence of Other Chemicals and Physical Environment
3.3.3 Mechanism of Action
3.3.4 Acquired Resistance to Benzoic Acid
3.4 Microbial Metabolism of Benzoic Acid
3.5 Reaction with Food Constituents
3.6 Regulatory Status
3.7 Applications
3.7.1 Use in Various Food Systems
3.7.2 Use as a Postharvest Fungicide
3.7.3 Other Applications
3.7.4 Storage and Handling
3.8 Toxicology
3.9 Assay
3.10 Acknowledgments
References
Chapter 4 Sorbic Acid and Sorbates
4.1 Introduction
4.2 Chemistry
4.3 Antimicrobial Activity
4.3.1 Inhibition
4.3.2 Selective Action
4.3.3 Degradation
4.3.4 Interactions
4.3.5 Mechanisms of Action
4.4 Applications
4.4.1 Dairy Products
4.4.2 Vegetable Products
4.4.3 Fruit Products
4.4.4 Bakery Products
4.4.5 Meat Products
4.4.6 Miscellaneous Food Products
4.4.7 Other Applications
4.5 Toxicology and Safety
4.6 Detection and Analysis
4.7 Regulatory Status
References
Chapter 5 Organic Acids
5.1 Introduction
5.2 Effective Use
5.3 The Acids
5.3.1 Acetic Acid
5.3.1.1 Antimicrobial Activity in Foods
5.3.1.2 Application and Regulatory Status
5.3.2 Acetates
5.3.2.1 Antimicrobial Properties
5.3.2.2 Application and Regulatory Status
5.3.3 Sodium Diacetate
5.3.3.1 Antimicrobial Activity in Foods
5.3.3.2 Regulatory Approval
5.3.4 Dehydroacetic Acid
5.3.4.1 Antimicrobial Activity against Foodborne Microorganisms
5.3.4.2 Application and Regulatory Status
5.3.5 Adipic Acid
5.3.5.1 Antimicrobial Properties
5.3.5.2 Application and Regulatory Status
5.3.6 Citric Acid
5.3.6.1 Antimicrobial Properties and Activity in Foods
5.3.6.2 Application and Regulatory Status
5.3.7 Fumaric Acid
5.3.7.1 Antimicrobial Properties
5.3.7.2 Application and Regulatory Status
5.3.8 Lactic Acid
5.3.8.1 Antimicrobial Properties
5.3.8.2 Application and Regulatory Status
5.3.9 Lactates
5.3.9.1 Antimicrobial Properties
5.3.9.2 Application and Regulatory Status
5.3.10 Malic Acid
5.3.10.1 Antimicrobial Properties
5.3.10.2 Application and Regulatory Status
5.3.11 Propionic Acid
5.3.11.1 Antimicrobial Properties
5.3.11.2 Application and Regulatory Status
5.3.12 Succinic Acid
5.3.12.1 Antimicrobial Properties
5.3.12.2 Application and Regulatory Status
5.3.13 Tartaric Acid
5.3.13.1 Antimicrobial Properties
5.3.13.2 Application and Regulatory Status
5.4 Mode of Action
5.5 Acid Adaptation and Tolerance
5.6 Toxicology
5.6.1 Toxicity of Organic Acids Post-Ingestion
5.6.2 Irritation and Inhalational Exposure Toxicological Effects
5.6.3 Intravenous Exposure and Toxicological Effects
5.6.4 Mutagenic and Teratogenic Activities of Organic Acids
5.7 Assay Methods
References
Chapter 6 Sulfur Dioxide and Sulfites
6.1 Overview of Sulfite Use in Food
6.2 Chemical Properties of Sulfites Used in Food
6.3 Factors Affecting Antimicrobial Activity
6.4 Antimicrobial Activity of Sulfites
6.4.1 Antibacterial Activity
6.4.2 Activity against Yeasts
6.4.3 Activity against Molds
6.5 Applications
6.5.1 Use in Wine
6.5.2 Use in Beer
6.5.3 Use in Juices
6.5.4 Use in Fruit
6.6 Regulatory Status of Sulfites in Food
6.6.1 Uses and Regulatory Limits of Sulfites in the USA
6.6.2 Uses and Regulatory Limits of Sulfites outside the USA
6.7 Alternatives to Sulfites in Food and Beverages
6.8 Toxicology and Safety
6.9 Detection and Analysis in Food
6.10 Conclusions
References
Chapter 7 Nitrite
7.1 Introduction
7.2 Research History
7.2.1 Before 1950
7.2.1.1 Summary for before 1950
7.2.2 1950–1960
7.2.2.1 Summary for 1950–1960
7.2.3 1960–1970
7.2.3.1 Shelf-Stable Cured Meats
7.2.3.2 Perishable Cured Meats
7.2.3.3 Summary for 1960–1970
7.2.4 1970–1980
7.2.4.1 Shelf-Stable Cured Meat and Perigo Factor
7.2.4.2 Perishable Canned Cured Meat and Slurries of Meat
7.2.4.3 Vacuum-Packaged and Fermented Meats
7.2.4.4 Bacon
7.2.4.5 Test with S. aureus and Other Bacteria
7.2.4.6 Summary for 1970–1980
7.2.5 1980–1990
7.2.5.1 Assessment of the Botulinal Risk in Cured Meats
7.2.5.2 Botulinal Challenge Tests
7.2.5.3 Perigo Factor
7.2.5.4 Alternatives to Nitrite for Botulinal Protection
7.2.5.5 Mechanism of Nitrite Inhibition
7.2.5.6 Miscellaneous Studies in Laboratory Media
7.2.5.7 Spoilage of Cured Meats
7.2.5.8 Summary for 1980–1990
7.2.6 1990–2002
7.2.6.1 Fermented Meats and Dry-Cured Hams
7.2.6.2 Alternatives to the Use of Nitrate or Nitrite
7.2.6.3 Cheese
7.2.6.4 Seafood
7.2.6.5 Effect of Nitrite on Enteric Pathogens
7.2.6.6 Effect of Nitrite on Yeasts and Molds
7.2.6.7 Green Discoloration in Cured Meats
7.2.6.8 Summary for 1990–2002
7.2.7 2002–2019
7.2.7.1 Mechanisms of Nitrite Antimicrobial Activity
7.2.7.2 Antibotulinum Activity of Nitrite
7.2.7.3 Nitrite Effects on Clostridium perfringens
7.2.7.4 Antilisterial Effects of Nitrite
7.2.7.5 Nitrite Controversy and Conflicting Messages
7.2.7.6 Changing Consumer Demands
7.2.7.7 Predictive Models of Microbial Growth or Inactivation That Incorporate Nitrite
7.2.7.8 Summary for 2002–2019
7.3 Regulations Governing the Use of Nitrite and Nitrate
7.3.1 Historical Perspective on Regulations
7.3.2 Nitrite and Nitrate Regulations in the United States
7.3.2.1 US Regulations for Formulation of Cured Meat and Poultry Products
7.3.2.2 US Guidelines for Stabilization (Cooling) of Cooked Cured Meat and Poultry Products
7.3.2.3 US Requirements for Natural Sources of Nitrate or Pre-Converted Nitrite in Meat and Poultry Products
7.3.2.4 US Requirements Regarding Other Uses of Nitrate and Nitrite in Food Products
7.3.2.5 US Regulations on Nitrite and Nitrate in Water
7.3.3 Canadian Regulations Regarding Nitrite and Nitrate in Foods
7.3.4 European Union Regulations Regarding Nitrite and Nitrate in Foods
7.3.5 Regulations in Other Regions Regarding Nitrite and Nitrate Use in Foods
7.4 Assay Method
7.5 Toxicology
References
Chapter 8 Nisin
8.1 Introduction
8.2 Structure
8.3 Physicochemical Properties
8.4 Antimicrobial Activity
8.4.1 Factors Affecting the Antimicrobial Activity in Foods
8.4.1.1 Food Composition
8.4.1.2 Temperature
8.4.1.3 pH
8.4.1.4 Proteolytic Enzymes
8.4.1.5 Process
8.4.1.6 Synergy with Other Preservatives
8.4.1.7 Bacterial Load
8.5 Applications as Food Preservative
8.5.1 Dairy Products
8.5.1.1 General Considerations
8.5.1.2 Main Uses for the Preservation of Dairy Products
8.5.1.3 Uses in Combination with Other Treatments
8.5.2 Meat Products
8.5.3 Seafood Products
8.5.4 Fruits and Vegetables
8.5.5 Beverages
8.5.6 Other Food Products
8.5.7 Food Packaging Containing Nisin
8.5.7.1 General Considerations
8.5.7.2 Non-Biodegradable Films
8.5.7.3 Cellulose-Based Films
8.5.7.4 Chitosan-Based Films
8.5.7.5 Alginate-Based Films
8.5.7.6 Protein-Based Films
8.6 Regulatory Status
References
Chapter 9 Natamycin
9.1 Introduction
9.2 Physical and Chemical Properties
9.3 Antimicrobial Activity
9.4 Mode of Action
9.5 Food Applications
9.5.1 Cheese
9.5.2 Meat
9.5.3 Fruits and Fruit Juices
9.5.4 Miscellaneous Applications
9.5.5 Incorporation into Packaging Films and Coatings for Foods
9.6 Regulatory Status
9.7 Toxicology
9.8 Assay
References
Chapter 10 Lauric Arginate Ethyl Ester
10.1 Chemistry Structure and Characteristics
10.2 Antimicrobial Activity in Microbiological Media
10.3 Antimicrobial Mechanism
10.4 Application in Food Systems
10.4.1 Application Alone or with Other Chemicals on Raw Meat and Poultry Products
10.4.2 Application in Ready-to-Eat Meat Products
10.4.3 Application in Other Food Products
10.4.4 Application within Packaging Material
10.5 Regulatory Status
10.6 Evaluation of Acute Toxicity
10.7 Chemical Assays
10.8 Conclusion
References
Chapter 11 Medium-Chain Fatty Acids (>C8) and Monoesters
11.1 Introduction and History
11.2 Antimicrobial Activity of Fatty Acids
11.2.1 Antibacterial Activity
11.2.2 Antifungal Activity
11.2.3 Effect of Chemical Modifications on Antimicrobial Activity
11.2.4 Structure–Function Relationship
11.3 Fatty Acid Monoesters
11.3.1 Sucrose Monoesters
11.3.2 Polyglycerol Esters
11.3.3 Glycerol Monoesters
11.3.3.1 Antibacterial Activity
11.3.3.2 Antifungal Activity
11.3.3.3 Antiviral Activity
11.3.3.4 Combinations with Other Antimicrobials
11.3.3.5 Combination with Heat and Cold
11.4 Food Applications
11.4.1 Dairy Products
11.4.2 Meat Products
11.4.3 Seafood
11.4.4 Other Foods
11.5 Mechanism of Action
References
Chapter 12 Parabens
12.1 Chemical and Physical Properties
12.2 Antimicrobial Activity
12.2.1 Bacteria
12.2.2 Fungi
12.3 Mechanism of Action
12.4 Applications
12.5 Regulatory Status
12.6 Toxicology
12.7 Assay
References
Chapter 13 Dimethyl Dicarbonate and Diethyl Dicarbonate
13.1 Introduction
13.2 Chemistry
13.2.1 Description
13.2.2 Ethyl Carbamate Formation
13.2.3 Synthesis
13.2.4 Reactions
13.3 Antimicrobial Activity
13.3.1 Yeast
13.3.2 Bacteria
13.3.3 Molds
13.3.4 Physical and Chemical Effects on Activity
13.3.4.1 Temperature
13.3.4.2 pH
13.3.4.3 Ethanol and Other Constituents
13.4 Mechanism of Action
13.5 Applications
13.5.1 Fruit Juices
13.5.2 Grape Juice and Wines
13.5.3 Soft Drinks
13.5.4 Foods
13.5.5 Beer
13.6 Regulatory Status
13.7 Toxicology
13.7.1 Foods
13.7.2 Direct Exposure
13.7.3 Indirect Exposure
13.8 Analysis
13.8.1 Direct
13.8.2 Indirect
References
Chapter 14 Lysozyme
14.1 Background and General Properties of Lysozyme
14.2 Properties
14.2.1 Chromatographic Elution Characteristics
14.2.2 Basic Enzymatic and Hydrolytic Properties
14.2.3 Common Properties amongst Differing Lysozyme Types
14.3 Uses and Stability in Foods
14.3.1 Thermal Stability in Solution and in Food Products during Heat-Processing
14.3.2 Lysozyme Stability to Non-Thermal Processing and Food Preservatives
14.4 Antimicrobial Spectrum of Activity
14.4.1 Gram-Positive and Gram-Negative Bacteria Sensitivity to Lysozyme
14.4.2 Lysozyme Sensitivity as a Function of Microbial Physiological Status
14.5 Activity against Food-Related Microorganisms
14.5.1 Antimicrobial Activity of Lysozyme against Gram-Positive Rods in Foods
14.5.2 Antifungal Activity against Foodborne Yeasts and Spoilage Bacteria in Foods
14.6 Effect on Heat Resistance of Bacterial Spores
14.7 Enhancement of Activity by Other Chemical Agents
14.7.1 Enhancement of Lysozyme by Chelators against Foodborne Bacteria
14.7.2 Lysozyme Use in Combination with Antimicrobial Polypeptides and Bacteriocins
14.8 Enhancement by Physical Processes
14.9 Food Applications
14.9.1 Functionality in Ready-to-Eat Further Processed Foods
14.9.2 Lysozyme Utility on Animal Carcass and Derived Product Surfaces
14.9.3 Lysozyme Applications for Food Contact Surfaces Disinfection
14.10 Non-Enzymatic Antimicrobial Activity of Lysozyme and Lysozyme-Peptides
14.10.1 Bactericidal Characteristics of Lysozyme
14.10.2 Lysozyme-Derived Antimicrobial Peptides
14.11 Recombinant Lysozymes for Use as Food Antimicrobials
14.12 Regulatory Status and Toxicology
14.13 Summary and Perspectives
References
Chapter 15 Bacteriocins and Their Applications in Foods
15.1 Introduction
15.2 Classification of Bacteriocins
15.3 Genetics, Biosynthesis, and Mode of Action
15.3.1 Organization of Gene Clusters
15.3.2 Biosynthetic Pathway
15.3.3 Post-Translational Modification, Activation, and Transport
15.3.4 Regulation of Biosynthesis
15.3.5 Producer Immunity
15.3.6 Mode of Action
15.4 Production and Modeling
15.5 Resistance
15.6 Examples of Activity Spectra and Biochemical Properties
15.6.1 Lactococcus
15.6.2 Pediococcus
15.6.3 Lactobacillus
15.6.4 Carnobacterium
15.6.5 Leuconostoc and Other Lactic Acid-Associated Bacteria
15.7 Applications in the Food Industry
15.7.1 Dairy Products
15.7.2 Meat Products
15.7.3 Fruits and Vegetables
15.8 Summation
References
Chapter 16 Bacteriophages
16.1 History of Bacteriophages
16.2 The Diversity of Bacteriophages: Phage Morphology Forms the Basis for Classification
16.3 Phage Lifecycle
16.3.1 Lytic Pathway
16.3.2 Lysogenic Pathway
16.4 Phage Application within the Food Industry
16.4.1 Fresh Produce
16.4.2 Meat and Dairy Food Products
16.4.3 Food-Contact Surfaces
16.4.4 Pre-Harvest Control
16.4.5 Bacteriophage Insensitive Mutants
16.5 Where Does Phage Technology Stand Today?
16.6 Conclusion
References
Chapter 17 Naturally Occurring Compounds – Plant Sources
17.1 Introduction
17.2 Sources of Natural Antimicrobials from Plants
17.2.1 Natural Phenolic Compounds
17.2.2 Essential Oils
17.2.3 Phytoalexins
17.2.4 Phytoanticipins
17.2.5 Antimicrobial Peptides (AMPs)
17.2.6 Other Sources of Plant Antimicrobial Agents
17.3 Testing the Efficacy of Antimicrobials
17.3.1 In Vitro Testing
17.3.2 Vapor Phase
17.4 Mechanisms of Action
17.5 Factors Affecting Activity
17.5.1 Plant Source Variation
17.5.2 Extraction Methods
17.5.3 Interaction with Food Matrix
17.6 Increasing the Efficacy of Natural Antimicrobials from Plants
17.6.1 Application of Essential Oils in Gaseous Phase
17.6.2 Combination with Other Antimicrobials and/or Preservation Factors
17.6.3 Delivery Systems
17.6.3.1 Emulsions
17.6.3.2 Nanosized Carriers/Coatings
17.6.3.3 Packaging Films/Coatings
17.7 Toxicity of Natural Antimicrobials and Its Evaluation and Regulatory Aspects
17.8 Application in Food and Sensory Analysis
17.9 Final Remarks
17.10 Acknowledgments
References
Chapter 18 Naturally Occurring Compounds – Animal Sources
18.1 Introduction
18.2 Lactoperoxidase
18.2.1 Molecular Properties
18.2.1.1 Occurrence and Biosynthesis
18.2.1.2 Chemistry and Structure
18.2.1.3 Stability
18.2.2 Antimicrobial Activity
18.2.2.1 Mode of Action
18.2.2.2 Specificity
18.2.3 Applications in Food
18.3 Transferrins
18.3.1 Lactoferrin, Lactoferricin B, and Activated Lactoferrin
18.3.1.1 Molecular Properties
18.3.1.2 Antimicrobial Activity
18.3.1.3 Applications in Foods
18.3.1.4 Safety and Tolerance
18.3.2 Ovotransferrin
18.4 Immunoglobulins
18.4.1 Lactoglobulins
18.4.1.1 Molecular Properties
18.4.1.2 Antimicrobial Activity
18.4.1.3 Applications in Food
18.4.2 Ovoglobulins
18.4.2.1 Molecular Properties
18.4.2.2 Antimicrobial Activity
18.4.2.3 Applications in Food
18.5 Avidin
18.5.1 Molecular Properties
18.5.1.1 Occurrence and Biosynthesis
18.5.1.2 Chemistry and Structure
18.5.1.3 Stability
18.5.2 Antimicrobial Activity
18.5.2.1 Mode of Action
18.5.2.2 Specificity
18.5.3 Applications in Food
18.6 Lactolipids
18.6.1 Molecular Properties
18.6.1.1 Occurrence and Biosynthesis
18.6.1.2 Chemistry and Structure
18.6.1.3 Stability
18.6.2 Antimicrobial Activity
18.6.2.1 Mode of Action
18.6.2.2 Specificity
18.6.3 Applications in Food
18.7 Defensins
18.7.1 Molecular Properties
18.7.1.1 Occurrence and Biosynthesis
18.7.1.2 Chemistry and Structure
18.7.2 Antimicrobial Activity
18.7.3 Applications in Food
18.8 Chitosan
18.9 Other Antimicrobials of Animal Origin
18.9.1 Pleurocidin
18.9.2 Casocidin
18.9.3 Lysozyme (Consult Chapter 14, “Lysozyme” for More Information)
18.9.4 Lipids
References
Chapter 19 Use of Antimicrobials as Processing Aids in Food Processing
19.1 Importance
19.2 Regulatory Perspectives
19.3 Types of Processing Aids
19.4 Processing Aids in Meat and Poultry Products
19.4.1 Foodborne Illness and Microorganisms of Concern in Meat and Poultry
19.4.2 Animal Post-Evisceration Carcass Decontamination
19.4.2.1 Carcass Washes
19.4.2.2 Processing Aids in Scald and Chilling Water
19.4.2.3 Post-Chill Processing Aids Added for Antimicrobial Activity
19.4.3 Addition of Processing Aids During Further Processing of Meat and Poultry Products
19.4.3.1 Non-RTE Meat and Poultry Products
19.4.3.2 RTE Meat and Poultry Products
19.4.4 The Role of Acidifiers in Microbial Reduction of Meat and Poultry Products
19.4.5 Summary
19.5 Decontamination of Fresh and Fresh-Cut Produce
19.5.1 Fresh Produce – Benefits and Risks
19.5.2 Fresh Produce – Foodborne Illness and Microorganisms of Concern
19.5.3 Washing and Sanitizing Treatments for Fruits and Vegetables
19.5.3.1 Chlorine
19.5.3.2 Peroxyacetic Acid (PAA)
19.5.4 Summary
References
Chapter 20 Antimicrobial Delivery Systems
20.1 Introduction
20.2 The Need for Delivery Systems
20.3 Emulsion-Based Delivery Systems
20.3.1 Preparation Methods
20.3.2 Applications
20.4 Lipid-Based Delivery Systems
20.4.1 Preparation Methods
20.4.2 Applications
20.5 Polymer-Based Delivery Systems
20.5.1 Preparation Methods
20.5.2 Applications
20.6 Inorganic Particles-Based Delivery Systems
20.6.1 Preparation Methods
20.6.2 Applications
20.7 Important Properties for Antimicrobial Delivery Systems and Methods of Characterization
20.7.1 Antimicrobial Loading and Encapsulation Efficiency
20.7.2 Size, Zeta Potential, and Morphology
20.7.3 Controlled-/Triggered-Release Properties
20.7.4 Antimicrobial Delivery System Efficacy Evaluation
20.7.5 Comparison among Delivery Systems – Advantages/Disadvantages
20.8 Conclusions and Future Trends
References
Chapter 21 Hurdle Technology – or Is It? Multifactorial Food Preservation for the Twenty-First Century
21.1 Introduction
21.2 Benefits of the Hurdle Concept
21.3 Drawbacks of the Hurdle Concept
21.4 Alternative “Multifactorial” Metaphors
21.4.1 The Swiss Cheese Model
21.4.2 The Pole Vault: Hurdles Upended
21.5 Risks
21.6 Microbial Modeling to Quantify Factors and Control Risk
21.6.1 Pasteurized Processed Cheese Spread and Control of Clostridium botulinum
21.6.2 Ready-to-Eat Meat Products and Listeria Control
21.7 Multifactorial Preservation in Action
21.7.1 Market Examples of Multifactorial Products
21.7.1.1 Co-Extruded Cereal Bar with Fruit Core
21.7.1.2 Optimization of “Non-C. botulinum” Thermophilic Cook with Use of Nisin in Canned Foods
21.7.1.3 Bottled Water
21.7.1.4 Refrigerated Liquid Egg Containing Vegetable Blend
21.7.1.5 High-Pressure Processed Products
21.8 Future Development of Novel Opportunities
21.9 Concluding Remarks
21.10 Acknowledgments
21.11 Disclaimer
References
Chapter 22 Applications of Antimicrobials to Foods – A Food Industry Perspective
22.1 Introduction
22.2 Research and Development Perspectives
22.2.1 Combination Studies
22.2.2 Standardization of Efficacy Determination
22.3 Considerations for Commercial Application of Antimicrobials in Food
22.3.1 Phase 1 – Discovery or Proof of Concept
22.3.2 Phase 2 – Technology Development
22.3.2.1 Efficacy
22.3.2.2 Sensory Impact
22.3.2.3 Cost-in-Use
22.3.2.4 Capital Expenditure
22.3.2.5 Minimum Order Quantity (MOQ)
22.3.2.6 Patent Landscape
22.3.2.7 Regulatory Assessment
22.3.2.8 Challenge Studies
22.3.2.9 Sensory and Shelf Life Study
22.3.2.10 The Production Process
22.3.2.11 Occupational Safety and Health Act (OSHA)
22.3.3 Phase 3 – Technology Transfer (Scale-up and Commercialization)
22.3.3.1 Food Product Matrix
22.3.3.2 Beverages
22.3.3.3 Bakery Products
22.3.3.4 Fresh and Minimally Processed Fruits and Vegetables
22.3.3.5 Dairy Products
22.3.3.6 Meat and Poultry Products
22.3.3.7 Sauces and Condiments
22.4 Conclusion
22.5 Acknowledgments
Disclaimer
References
Chapter 23 Mechanisms of Action, Resistance, and Stress Adaptation
23.1 Introduction
23.2 Mechanism of Antimicrobial Action on Potential Target Sites of Vegetative Bacteria
23.2.1 Antimicrobial Interactions with the Outer Membrane of Gram-Negative Bacteria
23.2.1.1 The Porin Pathway
23.2.1.2 The Hydrophobic Pathway
23.2.1.3 Self-Promoted Uptake Pathway
23.2.2 The Peptidoglycan Layer as a Potential Target Site
23.2.2.1 Peptidoglycan Degrading Compounds
23.2.2.2 Diffusion through the Peptidoglycan
23.2.3 Potential Targets and Mechanisms Associated with Cytoplasmic Membranes
23.2.3.1 Dissipation of Proton Motive Force
23.2.3.2 Membrane Permeabilization
23.2.3.3 Inhibition of Membrane-Embedded Proteins/Enzymes
23.2.4 Antimicrobial Mechanisms on Cytoplasmic Targets
23.2.4.1 Inhibition of Intracellular Enzymes and Proteins
23.2.4.2 Targeting Nucleic Acids
23.2.4.3 Modulating the Intracellular Environment
23.3 Antimicrobial Target Sites and Action Mechanisms in Bacterial Spores
23.4 Antimicrobial Target Sites and Action Mechanisms in Fungi
23.4.1 Mechanisms of Cell Wall and Membrane-Targeting Antifungals
23.4.2 Mechanisms of Intracellular Targeting Antifungals
23.5 Mechanism of Action Investigations
23.5.1 Identifying the Target Site
23.5.1.1 Initial Antimicrobial Activity Assessment
23.5.1.2 Cell Wall as the Target Site
23.5.1.3 Cell Membrane as the Target Site
23.5.1.4 Intracellular Target Site
23.5.1.5 Biosynthesis or Metabolic Pathways as Targets
23.5.2 Identifying the Mechanism of Action
23.5.2.1 Elucidating Membrane-Destabilizing Action Mechanisms
23.5.2.2 Intracellular Action Mechanisms
23.5.2.3 Molecular Mechanism of Action
23.5.3 Guide for Conducting a Mechanism of Action Study
23.6 Resistance and Stress Adaptation
23.6.1 Definition of Terms
23.6.2 Intrinsic Resistance and Stress Adaptation Mechanisms
23.6.2.1 LPS Modifications
23.6.2.2 Peptidoglycan
23.6.2.3 Cytoplasmic Membrane Modifications
23.6.2.4 Efflux Pumps
23.6.2.5 Proteome
23.6.2.6 The Stringent Response
23.6.2.7 Biofilm
23.6.2.8 Antimicrobial Inactivation
23.6.3 Acquired Resistance Mechanisms
References
Index

Citation preview

Antimicrobials in Food

Antimicrobials in Food Fourth Edition

Edited by

P. Michael Davidson T. Matthew Taylor Jairus R. D. David

Fourth edition published 2021 by CRC Press 6000 Broken Sound Parkway NW, Suite 300, Boca Raton, FL 33487-2742 and by CRC Press 2 Park Square, Milton Park, Abingdon, Oxon, OX14 4RN © 2021 Taylor & Francis Group, LLC First edition published by Marcel Dekker 1983 Third edition published by CRC Press 2005 CRC Press is an imprint of Taylor & Francis Group, LLC Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, access www​.copyright​.com or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. For works that are not available on CCC please contact mpkbookspermissions​@tandf​.co​​.uk Trademark notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe.

Library of Congress Cataloging-in-Publication Data Names: Davidson, P. Michael, 1950- editor. | Taylor, T. M. (T. Matthew), editor. | David, Jairus R. D., editor. Title: Antimicrobials in food / edited by P. Michael Davidson, T. Matthew Taylor, Jairus R. D. David. Description: Fourth edition. | Boca Raton : CRC Press, 2020. | Includes bibliographical references and index. Identifiers: LCCN 2020020320 | ISBN 9780367178789 (hardback) | ISBN 9780429058196 (ebook) Subjects: LCSH: Food additives. | Anti-infective agents. Classification: LCC TX553.A3 A57 2020 | DDC 664/.06--dc23 LC record available at https://lccn.loc.gov/2020020320 ISBN: 978-0-367-17878-9 (hbk) ISBN: 978-0-429-05819-6 (ebk) Typeset in Times by Deanta Global Publishing Services, Chennai, India

Contents Preface...................................................................................................................................................... vii Acknowledgments...................................................................................................................................... ix Editors ....................................................................................................................................................... xi List of Contributors..................................................................................................................................xiii 1. Food Antimicrobials – An Introduction......................................................................................... 1 T. Matthew Taylor, P. Michael Davidson, and Jairus R. D. David 2. Methods for Activity Assay and Evaluation of Results .............................................................. 13 Aurelio López-Malo, Emma Mani-López, P. Michael Davidson and Enrique Palou 3. Sodium Benzoate and Benzoic Acid...............................................................................................41 John R. Chipley 4. Sorbic Acid and Sorbates................................................................................................................ 89 Jarret Stopforth and Travis Kudron 5. Organic Acids.................................................................................................................................133 T. M. Taylor and Stephanie X. Doores 6. Sulfur Dioxide and Sulfites............................................................................................................191 Lilian Were and Leandra Filiaci 7. Nitrite...............................................................................................................................................219 R. Bruce Tompkin, Wendy Bedale, Andrew Milkowski, Kathleen Glass, and Jeffrey J. Sindelar 8. Nisin................................................................................................................................................ 309 Noushin Eghbal, Nour-Eddine Chihib, and Adem Gharsallaoui 9. Natamycin...................................................................................................................................... 339 P. Michael Davidson and Craig Doan 10. Lauric Arginate Ethyl Ester ........................................................................................................ 357 Cangliang Shen and T. Matthew Taylor 11. Medium-Chain Fatty Acids (>C8) and Monoesters.................................................................. 373 P. Michael Davidson, Jon J. Kabara and Douglas L. Marshall 12. Parabens......................................................................................................................................... 405 P. Michael Davidson 13. Dimethyl Dicarbonate and Diethyl Dicarbonate.........................................................................421 Randy W. Worobo, Rebecca M. Cheng, and Cornelius S. Ough 14. Lysozyme........................................................................................................................................ 445 T. Matthew Taylor, Eric A. Johnson, and Ann E. Larson v

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15. Bacteriocins and Their Applications in Foods............................................................................475 Pushpinder Kaur Litt, Dallas G. Hoover and Haiqiang Chen 16. Bacteriophages................................................................................................................................513 Pushpinder Kaur Litt, Kalmia E. Kniel, and Manan Sharma 17. Naturally Occurring Compounds – Plant Sources.................................................................... 527 Aurelio Lopez-Malo, Stella M. Alzamora, María J. Paris, Leonor Lastra-Vargas, María Bernarda Coronel, Paula L. Gómez, and Enrique Palou 18. Naturally Occurring Compounds – Animal Sources................................................................ 595 Jarret Stopforth and Travis Kudron 19. Use of Antimicrobials as Processing Aids in Food Processing................................................. 647 Emefa A. Monu and Jairus R. D. David 20. Antimicrobial Delivery Systems.................................................................................................. 665 Adriano Brandelli, Cicero C. Pola, and Carmen L. Gomes 21. Hurdle Technology – or Is It? Multifactorial Food Preservation for the Twenty-First Century................................................................................................................... 695 J. David Legan and Jairus R. D. David 22. Applications of Antimicrobials to Foods – A Food Industry Perspective.................................715 Jairus R. D. David and Peter Taormina 23. Mechanisms of Action, Resistance, and Stress Adaptation...................................................... 735 M. Hyldgaard Index....................................................................................................................................................... 785

Preface For the sake of maintaining the health and wellbeing of all, protection of the microbiological quality and safety of food is of utmost importance. The presence of contamination, and the unchecked growth of microbes in foods, is well-known to open the door both to the transmission of human pathogens to consumers and potential disease, and the loss of a food’s usefulness as a nutrient via spoilage. Foods may be prevented from becoming vehicles of disease transmission and protected from spoilage by application of one or more physical processing measures, such as heat, dehydration, or high-pressure processing, or by the use of chemical antimicrobial preservatives. Chemical antimicrobials, which act by a wide variety of mechanisms to inhibit microbial growth or inactivate microbes, may be derived from both natural and synthetic sources. Their activity is variable as to types of microorganisms affected, and their activity is often highly influenced by the environment and the physical and chemical composition of foods. They are subject to a diverse and complex set of regulatory requirements from nations around the globe based upon their usefulness and toxicology. It has been nearly 15 years since the publication of the 3rd edition of Antimicrobials in Food, an excellent reference for the food microbiologist, food safety specialist, industrial researcher, and student alike. This 4th edition comes about in a time of great change within the food industry, driven in large part by consumer purchasing trends demanding foods with fewer traditional additives, including antimicrobials, as well as foods processed so as to retain a greater fresh-like character during the food’s shelf-life. Compared to 2005, consumer purchase trends are apparently more influenced by numerous considerations, including budgetary restrictions, labeling claims regarding whether a food is “Clean,” “Natural,” “Organic,” “Raised Without Antibiotics,” or any of a number of others. The popularity of differing social media applications and networks also contributes to a consumer’s decision on whether to purchase and consume a particular food product or brand versus another. The 4th edition of Antimicrobials in Food addresses these and a number of other concerns in the following ways. First, a comprehensive revision has been completed of many chapters that appeared in previous editions of the text. Recent research has been integrated with important historical work that is still relevant. Highlights include chapters on individual or groups of compounds that generally detail current knowledge on the spectrum of activity of the antimicrobials, their mechanisms of activity, influences of food process measures on antimicrobial activity, regulatory status, and toxicology, among others. Traditional antimicrobials are covered, but new chapters have been added on lauric arginate, bacteriophages, and delivery systems for antimicrobials. A new comprehensive revision of Chapter 1 provides discussion on the key market trends which have influenced the use of food antimicrobials in recent years, including the clean label trend, the voice of the consumer and its ever-evolving role in the manufacture of foods, and the greater awareness of the need to reduce food insecurity through enhanced preservation technologies. There are new chapters on processing aides (Chapter 19), an evaluation of the hurdle concept (Chapter 21), and applications of antimicrobial food preservatives, including a discussion of the challenges and opportunities for industry members (Chapter 22). The final chapter (Chapter 23) is a comprehensive discussion of the mechanisms of action of antimicrobial preservatives as well as resistance development. Last, while P. M. Davidson, the heart and soul of this text, has returned as editor for this 4th edition, the current text brings on new editors from both academia and industry. This change from the last edition brings a broader perspective by blending both academic research and knowledge and experience from food industry applications. As with past editions, the editors are delighted to report a diverse and highly qualified set of contributing authors, assembled from universities and corporations around the U.S. and a host of other nations. An outline of content, albeit slightly different from chapter to chapter, has been retained from previous editions in an effort to improve the reader’s ability to utilize individual chapters for easy reference, wherein an antimicrobial’s spectrum of activity, uses within foods, mechanisms of action, regulations vii

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surrounding its use, and its toxicology are described. Finally, the editors wish to express their deep gratitude to the contributing authors whose efforts at assembling, reviewing, and integrating a large amount of recently published refereed research into existing knowledge, or the development of wholly new content, made this book possible. We do hope this book will be a useful reference and resource for those needing insight as to the use of antimicrobial food preservatives for protecting the quality and safety of foods from microorganisms. T. Matthew Taylor College Station, Texas P. Michael Davidson Coeur d’Alene, Idaho Jairus R. D. David Omaha, Nebraska

Acknowledgments To my family, friends, and co-workers at Texas A&M University whose desire for my success is constant – thank you. To Mike Davidson, to whom, like my fellow editor Dr. Jairus David, I must say congratulations for his contributions to the field of food safety microbiology and the use of food preservatives. In addition, I thank Dr. Davidson for his mentorship during my time at University of Tennessee. I thank also my students, both past and present, who have shouldered many burdens I’ve laid on them, because without them my research is not completed, my classes are not taught as effectively, and my time spent on this book would not be possible. Thank you all. T. Matthew Taylor First, I extend many thanks to the authors of the chapters of this 4th edition of Antimicrobials in Foods. I believe this to be the best of all the editions based on its breadth, depth, and appeal to academic, industrial, and governmental food microbiologists alike. Thanks too to my fellow co-editors, Drs. Matt Taylor and Jairus David, for, as Jairus often calls it, the “heavy lifting,” and to my previous co-editors, Dr. A. Larry Branen and Dr. John N. Sofos, both titans in the food science field, whom I both admire and respect. Few people advance science without support from others. If others judge that I have contributed positively to the field of food microbiology, I could not have done so without the cooperation and collaboration of outstanding professional colleagues, post-docs, and graduate and undergraduate students. Thus, I must thank them all for the opportunity to have interacted with them in an area that I have truly enjoyed. And finally I would like to dedicate the book to my spouse, Linda, for her encouragement and support during what is supposed to be our retirement, and my daughters, Laurel and Holly, and their families for making our life fun and livable. P. Michael Davidson I would like to express my thanks to the following: my wife, Shelley Zylstra-David, for her loving encouragement of this my work, and to our three millennial children, Adriana, Brennan, and Blake, for “daring” me to write “another book.” Dr. P. Michael Davidson for his pioneering contributions to the advances in antimicrobials and food safety, and to my professional development in the field covered. Dr. Larry Steenson, John Wyatt, Jerry Erdmann (Danisco); Dr. Athula Ekanayake (P&G); Weylan Bosse, Saurabh Kumar (Corbion-Purac); Michael Matthews, Emma Cahill (Kerry); Dr. Kathy Glass, Dr. Chuck Czuprynski, Max Golden, Dr. Wendy Bedale (Food Research Institute, University of Wisconsin); Dr. Emilia Rico (BCN Mold Research Labs) for introducing me to the world of natural and clean label antimicrobials, delivery systems, and microbial challenge studies. Dr. Al Bolles, Senior Executive Vice President, R&D, Conagra Brands, Dr. Richard McArdle, Vice President, Innovation, and Dr. Corey Berends, Vice President, R&D for their visionary leadership and entrusting me to lead, scout, and manage a program on the use of natural and clean label antimicrobials and novel processing technologies to provide competitive advantage and food protection. And to my peers – Kevin Gatlin, Susan Janssen, John Hinchik, Kathy Landis, Christie Hancock, Amy Kerby, Denise Becker, Edith Zambrana, Dr. Fred Cook, Dr. Lalit Bohra, Dr. Shecoya White, Dr. Lenora Howard, Larry Quint, Cort Ballard, Indarpal Singh, Dr. Eric Brown, Dr. Ric Gonzalez, Dr. Brian Degner, Dr. Curtis Stowe, Dr. Rodney Green, and Dr. Andrew Wassinger. My fellow editors – Dr. T. Matthew Taylor and Dr. P. Michael Davidson – for their visionary leadership and this our collaboration, and my learning. Jairus R. D. David

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Editors Jairus R. D. David – Biographical Sketch Jairus R. D. David, Ph.D., is a thought leader in the agro-food industry and an authority on food preservation science, food safety, and quality. Jairus has worked with the food industry on the use of natural and clean-label ingredients, especially antimicrobial preservatives for designing premium foods, and in developing microbiology and thermal-processing food safety objectives and compliance for over 35 years, balancing applied research with process and quality optimization, and the launch of innovative products. His industrial experience spans working for family-owned small companies and large corporations, on highly regulated baby foods, infant formula, and consumer-packaged goods. Jairus earned his Ph.D. in microbiology with emphasis in thermal processing from the University of California, Davis, under the tutelage of Dr. Larry Merson. Earlier, he earned his M.Sc. (Food Technology) from FAO Central Food Technological Institute (CFTRI), Mysore, India, and his B.Sc. (Agriculture) from the University of Agricultural Sciences, Bangalore, India. He is a Certified Quality Manager (CQM) and Certified Quality Engineer (CQE), from American Society for Quality. He has participated in the leadership program at the Kellogg School of Management, Center for Creative Leadership, Stephen Covey Leadership Forum, and the Massachusetts Institute of Technology. A Fellow of the Institute of Food Technologists (IFT), he is the recipient of IFT’s prestigious Industrial Scientist Award (2006). He is recognized for developing and influencing public health food safety policy on the use of honey in cereals and bakery products for the prevention of infant botulism in infants under 12 months of age. Currently, all honey and honey-containing food products in commerce carry a warning label: “Do not feed honey to infants less than one year of age.” Jairus has authored or coauthored several refereed papers, book chapters, books on aseptic technology, abstracts, and patents. Jairus is currently Principal Consultant at JRD Food Technology Consulting, LLC, Omaha, Nebraska. www​.JRDFoodTech​.com. Open for both domestic and international assignments, able and willing to do pro bono consulting for projects in developing “Third-World” and “Fourth-World” countries, rural communities, and non-profit organizations. P. Michael Davidson – Biographical Sketch Dr. P. Michael Davidson is a University of Tennessee (UT) Institute of Agriculture Chancellor’s Professor Emeritus and former Head (2005–2013) of the Department of Food Science & Technology at UT. Prior to retirement in 2016, he served on the faculty at UT for 30 years and was Professor in Food Science and Toxicology at the University of Idaho for eight years. He earned a Ph.D. in food science at Washington State University in 1979, an M.S. in food science from the University of Minnesota, and a B.S. in microbiology from the University of Idaho. Dr. Davidson’s research program involves microbiological food safety. His primary research area in food safety has been characterizing regulatory-approved and naturally occurring antimicrobial food preservatives. He is co-editor of the book Antimicrobials in Foods, 3rd Edition along with John Sofos and Larry Branen. A secondary research area has been the development and characterization of thermal and novel non-thermal processes to control pathogenic and spoilage microorganisms in foods. Dr. Davidson has authored or co-authored over 200 refereed journal articles, book chapters, and books and given over 300 scientific presentations at national and international meetings, industry workshops, and universities. He previously served as a Co-Scientific Editor for the Journal of Food Protection and on the Board of Directors of the Institute of Food Technologists (IFT). Davidson was presented with Honorary Life Membership in the International Association of Food Protection (IAFP) in 2018, the inaugural IFT Gerhardt Haas Award in 2017 for outstanding contribution to food safety, the Frozen Food Research Award from the Frozen Food Foundation in 2016, the IFT IAFP President’s Recognition Award in 2005, and the IFT Food Microbiology Division Distinguished Service Award in 2000. He was elected Chair of the IFT Food Microbiology Division in 1996 and Chair of the Food Microbiology Division of the American Society for Microbiology in 1993. For his contributions xi

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to microbiology, food science and technology, and food safety, Dr. Davidson was elected a Fellow of the American Academy of Microbiology, the Institute of Food Technologists, and the International Association of Food Protection, respectively. T. Matthew Taylor – Biographical Sketch Dr. Matthew Taylor is an associate professor of food microbiology in the Department of Animal Science at Texas A&M University, College Station, TX. He received the B.S. in Food Science and B.A. in Sociology in 2000 from North Carolina State University. He then obtained the M.S. degree in Food Science from North Carolina State University in 2003, and earned his Ph.D. in Food Science and Technology from the University of Tennessee, Knoxville in 2006. He joined Texas A&M University in June 2007. Dr. Taylor’s primary research interests are in the utilization and mechanisms of food antimicrobials to inhibit bacterial foodborne pathogens. Specifically, research is conducted to investigate and determine the manner by which food antimicrobials inhibit microbial pathogens. Additionally, research is conducted that seeks to overcome obstacles to the use of food antimicrobials in some product by the encapsulation of food antimicrobials. Dr. Taylor functions as lead instructor for undergraduate and graduate courses elaborating the microbiology of human foods. Dr. Taylor also regularly provides guest lectures for Texas A&M University courses, as well as teaching in food safety microbiology for members of the food processing industry and regulatory agency officers. Dr. Taylor is an active member of the Institute of Food Technologists and was previously recognized for excellence in service to the IFT Food Microbiology Division. He currently chairs and co-chairs various committees for the International Association for Food Protection, Phi Tau Sigma Honorary Food Science Society, and Gamma Sigma Delta Society. He sits on the editorial boards of multiple refereed journals. He also provides expert reviews in food safety microbiology for multiple journals publishing refereed food microbiology research. He has participated in over $33 million dollars in competitively funded and contracted research throughout his career.

List of Contributors Stella M. Alzamora Departamento de Industrias Ciudad Autónoma de Buenos Aires Buenos Aires, Argentina Wendy Bedale Food Research Institute University of Wisconsin Madison, Wisconsin Adriano Brandelli Department of Food Science Federal University of Rio Grande do Sul Porto Alegre, Brazil Haiqiang Chen Department of Animal and Food Sciences University of Delaware Newark, Delaware Rebecca M. Cheng Department of Food Science Cornell University Geneva, New York Nour-Eddine Chihib UMET-PIHM Laboratory University of Lille Villeneuve d’Ascq, France John R. Chipley Food Microbiologist (Retired) Watkinsville, Georgia María B. Coronel Departamento de Industrias Ciudad Autónoma de Buenos Aires Buenos Aires, Argentina Jairus R. D. David JRD Food Technology Consulting, LLC Omaha, Nebraska P. Michael Davidson Department of Food Science (Retired) University of Tennessee

Knoxville, Tennessee Current Address: Coeur d’Alene, Idaho Craig Doan Impact Washington Bothell, Washington Stephanie Doores Department of Food Science (Retired) Pennsylvania State University University Park, Pennsylvania Noushin Eghbal Department of Food Science, Engineering and Technology University of Tehran Karaj, Iran Leandra Filiaci Schmid College of Science and Technology Chapman University Orange, California Adem Gharsallaoui LAGEPP Laboratory University of Lyon Villeurbanne, France Kathleen Glass Food Research Institute University of Wisconsin Madison, Wisconsin Carmen L. Gomes Department of Mechanical Engineering Iowa State University Ames, Iowa Paula L. Gómez Departamento de Industrias Ciudad Autónoma de Buenos Aires Buenos Aires, Argentina Dallas G. Hoover Department of Animal and Food Sciences University of Delaware Newark, Delaware xiii

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Morten Hyldgaard DuPont Nutrition and Biosciences Brabrand, Denmark

Douglas Marshall Eurofins Scientific Inc. Fort Collins, CO

Eric A. Johnson Food Research Institute University of Wisconsin Madison, Wisconsin

Andrew Milkowski Department of Animal Sciences University of Wisconsin Madison, Wisconsin

Jon J. Kabara (Deceased) Med-Chem Labs Longboat Key, Florida

Emefa A. Monu Department of Poultry Science Auburn University Auburn, Alabama

Kalmia E. Kniel Department of Animal and Food Sciences University of Delaware Newark, Delaware Travis Kudron Nobilus, LLC Seattle, Washington Ann E. Larson Department of Environment, Health, and Safety University of Wisconsin Madison, Wisconsin Leonor Lastra-Vargas Departamento de Ingenieria Química y Alimentos Universidad de las Américas Puebla Cholula, Puebla, Mexico J. David Legan Eurofins Microbiology Laboratories Madison, Wisconsin Pushpinder K. Litt Department of Animal and Food Sciences University of Delaware Newark, Delaware Aurelio López-Malo Departamento de Ingenieria Química y Alimentos Universidad de las Américas Puebla Cholula, Puebla, Mexico Emma Mani-López Departamento de Ingenieria Química y Alimentos Universidad de las Américas Puebla Cholula, Puebla, Mexico

Cornelius S. Ough (Deceased) University of California Davis, California Enrique Palou Departamento de Ingenieria Química y Alimentos Universidad de las Américas Puebla Cholula, Puebla, Mexico Maria J. Paris Departamento de Ingenieria Química y Alimentos Universidad de las Américas Puebla Cholula, Puebla, Mexico Cicero C. Pola Department of Mechanical Engineering Iowa State University Ames, Iowa Manan Sharma Agricultural Research Service U.S. Department of Agriculture Beltsville, Maryland Cangliang Shen Davis College of Agriculture, Natural Resources and Design West Virginia University Morgantown, West Virginia Jeffrey J. Sindelar Department of Animal Sciences University of Wisconsin Madison, Wisconsin Jarret Stopforth Nobilus, LLC Seattle, Washington

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List of Contributors Peter J. Taormina Etna Consulting Group Jacksonville, Florida T. Matthew Taylor Department of Animal Science Texas A&M University College Station, Texas R. Bruce Tompkin ConAgra Refrigerated Foods (Retired) Downers Grove, Illinois

Lilian Were Schmid College of Science and Technology Chapman University Orange, California Randy W. Worobo Department of Food Science Cornell University Geneva, New York

1 Food Antimicrobials – An Introduction T. Matthew Taylor, P. Michael Davidson, and Jairus R. D. David CONTENTS 1.1 Role of Additives in Food................................................................................................................. 1 1.2 Definition and Function of Chemical Food Preservatives................................................................ 1 1.3 Selection of Antimicrobials.............................................................................................................. 3 1.3.1 Antimicrobial Spectrum....................................................................................................... 4 1.3.2 Physico-Chemical Properties of the Antimicrobial............................................................. 5 1.3.3 Food-Related Factors............................................................................................................ 5 1.3.4 Process Factors..................................................................................................................... 6 1.3.5 Resistance Development....................................................................................................... 6 1.4 Considerations in the Applications of Food Antimicrobials............................................................ 6 1.4.1 Toxicological Safety............................................................................................................. 7 1.4.2 Labeling with Respect to Antimicrobials............................................................................ 7 1.5 Future of Antimicrobials................................................................................................................. 10 References..................................................................................................................................................11

1.1 Role of Additives in Food Food additives may be classified by one of six primary functions they serve: preservation, improvement in nutritional value, addition or replacement of color, addition or replacement of flavor, improvement in texture, or processing aids (Branen and Haggerty, 2002). The focus of this book is food additives that contribute to preservation, although some food antimicrobials also contribute to color stability (sulfites, nitrites) or flavor (nitrites and certain organic acids). Despite the recognized requirement for food additives, their toxicological safety continues to be evaluated and is often questioned. Few new additives have been approved by regulatory agencies in recent years, and it is doubtful that many additional ones will be approved in the future. Although there is no question that the risk from an additive must be minimal, it is apparent that such risk must be balanced against the benefits of the use of such additives. For example, reduction in the number of foodborne illness cases or reduced food waste via spoilage losses are benefits provided by antimicrobial additives. Balancing the risks vs. the benefits is not easy and requires extensive research about the usefulness and toxicological safety of the additives in question. In very few cases are additives totally devoid of risk, and thus an assessment of the degree of acceptable risk is often required. Responsibility for determining if risks outweigh benefits for any particular additive is with scientists, legislators, regulatory personnel, food processors, and consumers. It is essential that all involved in the decision process be acutely aware of the risks and benefits of all additives.

1.2 Definition and Function of Chemical Food Preservatives Drying, cold, fermentation, and heating have always been the primary methods used to prolong the shelf-life of food products. However, since prehistoric times, some chemicals, such as salt, have been

1

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Antimicrobials in Food

added to preserve freshly harvested foods for later use. While certain chemical food preservatives, such as salt, nitrites, and sulfites, have been in use for many years, most others have seen extensive use only recently. One of the reasons for increased use of chemical preservatives has been the change in the ways foods are produced and marketed. Today, consumers expect foods to be readily available year-round, to be “free” of foodborne pathogens, and to have a reasonably long shelf-life. While many improvements have been made using packaging and processing systems to preserve foods, antimicrobials play a significant role in protecting consumers from premature food loss and potential episodes of foodborne illness. Because of changes in the marketing for foods to a more global system, products are seldom grown and sold locally as in the past. Today, foods produced in one geographic area are often shipped to another area for processing and to several other areas for distribution. Several days, months, or even years may elapse from the time food is produced until it is consumed. To accomplish the longterm shelf-life necessary for such a system, multiple effective means of preservation are often required including the use of antimicrobials. It is important to note that food antimicrobials are not designed to be able to conceal spoilage of a food product. Rather they actually prevent or slow the growth of foodborne microorganisms which means the food remains wholesome during its extended shelf-life. Also, because most food antimicrobials are generally only bacteriostatic or fungistatic, they will not preserve a food indefinitely. Regulatory agencies generally classify food antimicrobials as “preservatives.” Chemical preservatives are defined by the U.S. Food and Drug Administration (FDA; 21CFR 101.22(a)(5)) as any chemical that, when added to food, tends to prevent or retard deterioration thereof, but does not include common salt, sugars, vinegars, spices, or oils extracted from spices, substances added to food by direct exposure thereof to wood smoke, or chemicals applied for their insecticidal or herbicidal properties.

Therefore, preservatives are used to prevent or retard both chemical and biological deterioration of foods. Those preservatives used to prevent or delay chemical deterioration include antioxidants used against autoxidation of pigments, flavors, lipids, and vitamins, anti-browning compounds used against enzymatic and nonenzymatic browning, and anti-staling compounds used against deleterious texture changes. Those additives used to prevent or delay biological deterioration are termed “antimicrobials.” FDA defines antimicrobial agents (21CFR 170.3(o)(2)) as “substances used to preserve food by preventing growth of microorganisms and subsequent spoilage, including fungistats, mold and rope inhibitors.” The European Union definition for “preservatives” is the same as the US definition of antimicrobials, i.e., those compounds that “extend the shelf-life of foods by protecting against loss of quality caused by microorganisms or protecting against the growth of illness-causing microorganisms” (European Union, 2019). Historically, the primary function of food antimicrobials has been to prolong shelf-life and preserve quality through the inhibition of spoilage microorganisms. Surprisingly, few food antimicrobials have been used exclusively to control the growth of specific foodborne pathogens. The one example of longterm usage is that of nitrite to inhibit Clostridium botulinum in cured meats. More recently, a number of compounds have been used against pathogens including organic acids as spray sanitizers against pathogens on beef carcasses, nisin and lysozyme against Clostridium botulinum in pasteurized process cheese, and lactate and diacetate to inactivate Listeria monocytogenes in processed meats. Thus, antimicrobials are being increasingly looked at as primary interventions for the inhibition or inactivation of pathogenic microorganisms in foods. Antimicrobials may be classified as traditional or naturally occurring (Davidson et al., 2013a). The former is approved for use in foods by many international regulatory agencies (Tables 1.1 and 1.2). Naturally occurring antimicrobials include compounds from microbial, plant, and animal sources. They are, for the most part, only proposed for use in foods. A few, such as nisin, natamycin, lactoferrin, lauric arginate, and lysozyme, are approved by regulatory agencies in some countries for application to foods (Tables 1.1 and 1.2). Extensive reviews on natural antimicrobials may be found in Dillon and Board (1994), Sofos et al. (1998), Naidu (2000), Burt (2004), Davidson et al. (2013a,b), and Davidson et al. (2014).

3

Food Antimicrobials – An Introduction TABLE 1.1 Traditional or Regulatory-Approved (U.S. Food and Drug Administration, 2018) Food Antimicrobials Compound(s)

Microbial Target

Primary Food Applications

Acetic acid, acetates, diacetates, dehydroacetic acid

Yeasts, bacteria

Baked goods, condiments, confections, dairy products, fats/oils, meats,a sauces

Benzoic acid, benzoates

Yeasts, molds

Dimethyl dicarbonate Lactic acid, lactates

Yeasts Bacteria

Beverages, fruit products, margarine Beverages Meats,a fermented foods

Lactoferrin

Bacteria

Meatsa

Lauric arginate Lysozyme

Bacteria Clostridium botulinum, other bacteria

Natamycin Nisin

Molds Clostridium botulinum, other bacteria

Nitrite, nitrate

Clostridium botulinum

Meatsa Cheese, casings for frankfurters,a cooked meat, and poultry products Cheese Cheese, casings for frankfurters, cooked meat, and poultry productsa Cured meatsa

Parabens (alkyl esters (propyl, methyl, heptyl) of p-hydroxybenzoic acid) Propionic acid, propionates

Yeasts, molds, bacteria (Gram-positive)

Beverages, baked goods, syrups, dry sausage

Molds

Sorbic acid, sorbates

Yeasts, molds, bacteria

Bakery products, dairy products Most foods, beverages, wines

Sulfites

Yeasts, molds

a

Fruits, fruit products, potato products, wines

21 CFR1 Section 172.130, 182.6197, 184.1005, 184.1185, 184.1721, 184.1754 184.1021, 184.1733 172.133 184.1061, 184.1207, 184,1639, 184.1768 GRAS Notices 000067 and 000130 USDA-FSIS GRAS Notice No. 000064

172.155 184.1538

172.160, 172.170, 172.175, 172.177 172.145, 184.1490, 184.1670 184.1081, 184.1221, 184.1784 182.3089, 182.3225, 182.3640, 182.3795 182.3616, 182.3637, 182.3739, 182.3766, 182.3798, 182.3862

For meat products, antimicrobials permitted by USDA Food Safety and Inspection Service are listed in USDA-FSIS (2020).

1.3 Selection of Antimicrobials It is not an easy process to select the appropriate preservation system for a particular food product. The target pathogen or spoilage microorganisms must be identified and then the possible preservation systems must be evaluated via model studies and studies in the food product in question. Generally, a combination of chemical preservatives and other preservation methods is needed (Leistner, 2000). Selection of the proper antimicrobial is dependent upon several primary factors, including the spectrum of antimicrobial activity, chemical properties of the antimicrobial, physicochemical properties and composition of the food product in question, and the type of preservation or processing and storage systems utilized. The individual chapters in this book examine these factors for all of the various natural and synthetic food antimicrobials. This discussion gives a brief overview of those factors and their importance.

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Antimicrobials in Food TABLE 1.2 Regulatory-Approved Food Antimicrobial Designations in the European Union (E Numbers) and in Codex Alimentarius (INS Numbers) Compound

E or INS Number

Compound

E or INS Number

200 202 203 210 211 212 213 214 215 218 219 220 221 222 223 224 226 227 228 234 235

DMDC Na lauroyl arginate K nitrite Na nitrite Na nitrate K nitrate Acetic acid K acetate Na acetate Ca acetate Lactic acid Propionic acid Na propionate Ca propionate K propionate Boric acid Na tetraborate Na lactate K lactate Ca lactate

242 243 249 250 251 252 260 261 262 263 270 280 281 282 283 284 285 325 326 327

Sorbic acid K sorbate Ca sorbate Benzoic acid Na benzoate K benzoate Ca benzoate Ethyl paraben Na ethyl paraben Methyl paraben Na methyl paraben Sulfur dioxide Na sulfite Na hydrogen sulfite Na metabisulfite K metabisulfite Ca sulfite Ca hydrogen sulfite K hydrogen sulfite Nisin Natamycin

a

Source: European Union, 2008. a Dimethyl Dicarbonate.

1.3.1 Antimicrobial Spectrum The initial selection of the antimicrobial is normally based on an assessment of the overall microbial spectrum of the chemical in question. Antimicrobial spectrum may be defined as the efficacy of a compound against various types of microorganisms (e.g., bacteria, yeasts, molds) and forms of those microorganisms (vegetative cells vs. spores). Even species, strain, and Gram reaction (positive vs. negative) can have dramatic influences on apparent activity. For example, the activity of very hydrophobic antimicrobials may be limited against Gram-negative bacteria which have an outer membrane lipopolysaccharide layer which helps to screen out certain antimicrobials. Quite often, a broad spectrum of activity, although desired, is not easy to achieve. Few chemicals have the ability to inhibit several different types, species, or strains of microorganism. The antimicrobial spectrum of a compound is generally determined by following the growth of organisms in the presence of various concentrations of the antimicrobial. Appropriate methods for in vitro evaluation of the activity of food antimicrobials are described in Chapter 2, “Methods for Activity Assay and Evaluation of Results.” Seldom, however, does growth in a synthetic microbiological medium parallel that in a food product. Thus, one must be wary of an antimicrobial spectrum determined only in synthetic medium. The final confirmatory test for the antimicrobial spectrum and activity must be carried out in a food product since food components and properties can dramatically alter the overall spectrum and activity of the antimicrobial. As a rule, it is important to know the conditions under which any antimicrobial spectrum was determined before projections are made regarding the usefulness of an antimicrobial. Probably the best method for determining what type of food antimicrobial to use would be based upon its mechanism of action and/or target in the cell. However, the exact mechanisms through which

Food Antimicrobials – An Introduction

5

antimicrobials affect microbial growth are complex and difficult to determine. Mechanisms of action of food antimicrobials generally are classified as reaction with the cell membrane causing permeability changes or interference with uptake and transport, inactivation of essential enzymes, interference with genetic mechanisms, or inhibition of protein synthesis. These are discussed within individual chapters and collectively in Chapter 23, “Mechanisms of Action, Resistance, and Stress Adaptation.” If the mechanism of the compound is known, combinations of antimicrobials with different mechanisms could be utilized against the microorganisms in the food product (Branen and Davidson, 2004; Davidson et al., 2013a).

1.3.2 Physico-Chemical Properties of the Antimicrobial The overall microbial spectrum, the mode of action, and the efficacy of compounds are largely dependent on the chemical and physical properties of the antimicrobial. The polarity of a compound is probably the most important physical property. Water solubility or hydrophilic properties appear to be necessary to assure that the antimicrobial is soluble in the water phase, where microbial growth occurs. At the same time, however, antimicrobials acting on the hydrophobic cell membrane most likely require some lipophilic properties. Thus, the most effective food antimicrobials are amphiphilic and require a specific hydrophile–lipophile balance for optimal activity. The balance needed in a synthetic medium, however, may differ significantly from that needed for a food product owing to polarity of the food components.

1.3.3 Food-Related Factors The chemical reactivity of the antimicrobial with other food components can significantly affect activity. Chemical reactions, in addition to decreasing antimicrobial activity, can also result in the formation of off-flavors, -odors, and -colors. Sorbic acid, for example, can be degraded by certain Penicillium species isolated from cheese to produce 1,3 pentadiene which has a kerosene off-odor. A sensory evaluation is often needed to assure that antimicrobials do not directly or indirectly through chemical reaction alter the color, flavor, or texture of a food product. Another primary factor leading to reduced effectiveness among food antimicrobials is food component interactions (Davidson et al., 2013a). As stated above, since most food antimicrobials are amphiphilic, they can solubilize in or be bound by lipids or hydrophobic proteins in foods, making them less available to inhibit microorganisms in the food product. Interaction with lipids probably results in the greatest interference with antimicrobial activity. Highly active antimicrobial compounds that are hydrophobic tend to partition into the lipid areas of the food and away from the water phase, where microbial growth occurs (Rico-Muñoz Davidson, 1983). The pH of a food can alter the effectiveness of an antimicrobial. For example, organic acids are most effective in their undissociated form. The concentration of the undissociated acid is dictated by the food pH and pKa of the acid. In high-acid foods (generally less than pH 4.5–4.6), this is not a problem because all regulatory-approved organic acids have pKa values less than 5.0. For food products with a pH of 5.5 or greater, there are very few organic acid-based compounds that are effective at low concentrations. In the undissociated form, organic acids can penetrate the cell membrane lipid bilayer more easily. Once inside the cell, the acid dissociates because the cell interior has a higher pH than the exterior (Davidson, 2013b). Bacteria maintain internal pH near neutrality to prevent conformational changes to the cell structural proteins, enzymes, nucleic acids, and phospholipids. Protons generated from intracellular dissociation of the organic acid acidify the cytoplasm and must be extruded to the exterior. Since protons generated by the organic acid inside the cell must be extruded using energy in the form of ATP, the constant influx of these protons will eventually deplete cellular energy. Chelating compounds such as ethylenediaminetetraacetic acid (EDTA) or its salts have a potentiating effect on some antimicrobials. They expand the activity of certain antimicrobials (e.g., nisin, lysozyme) to include Gram-negative bacteria which are not normally inhibited by the compounds alone (Cutter and Siragusa, 1995; Branen and Davidson, 2004). In addition, some Gram-positive bacteria are more susceptible to certain antimicrobials in the presence of chelators including EDTA. It is theorized that chelators may destabilize the LPS layer of the outer cell membrane of the Gram-negative bacteria allowing the antimicrobials access to the inner cell membrane.

6

Antimicrobials in Food

1.3.4 Process Factors The type of preservation process used in conjunction with antimicrobials has a significant influence on the type and level of antimicrobial needed. Certain preservation processes may result in the need to control spore-formers that have the ability to survive the heating process. A lowering of the water activity can select for those organisms that have the ability to survive and/or grow at lower water activity. Generally, molds survive and yeasts can grow at a lower water activity than bacteria, thus indicating the need for a different antimicrobial. Refrigeration procedures generally select for psychrotrophic Gramnegative microorganisms, thus requiring an antimicrobial capable of limiting the growth and activity of these organisms. Packaging can directly alter the environment of the food and thus influence the overall growth pattern and type of organisms in the food. Vacuum or modified-atmosphere packaging results in low oxygen tension which inhibits the growth of molds and several bacteria but allows certain facultative anaerobic microorganisms such as lactic acid bacteria to grow. Much research has been done on the addition or incorporation of food antimicrobials to packaging materials, which themselves may be antimicrobials (e.g., chitosan). Also, active packaging systems which incorporate compounds that continuously alter the atmosphere are available. This could allow for inhibition of spoilage or pathogenic microorganisms in the packaged product. Food antimicrobial compounds are often primary contributors to a combination of inhibitors and inhibitory conditions (e.g., low pH, low temperature). This is sometimes termed “hurdle technology” (Leistner and Gorris, 1995; Leistner, 2000). The concept of the use of multiple factors is discussed in detail in Chapter 21, “Hurdle Technology – or Is It? Multifactorial Food Preservation for the TwentyFirst Century.”

1.3.5 Resistance Development Since the activity spectra are often different for each antimicrobial, the microflora contaminating a food product significantly influences the choice of the antimicrobial needed. One should be cautious, however, not to select an antimicrobial solely according to its ability to control the predominant microorganism present. Because of their specificity, selecting antimicrobials that control some genera but not others may result in selecting for and creating favorable conditions for the growth of other organisms. For example, phenolic compounds may inhibit certain Gram-positive food-poisoning bacteria, but because of the reduced activity against Gram-negative bacteria, favorable conditions can be created for growth and spoilage by these latter organisms. Potential food antimicrobials should not contribute to the development of resistant strains nor alter the environment of the food in such a way that growth of another pathogen is selected. This subject is discussed in detail in Chapter 23, “Mechanisms of Action, Resistance, and Stress Adaptation.” Microorganisms exposed to a stress factors (e.g., heat, cold, starvation, low pH/organic acids) may become more resistant and have enhanced survival to subsequent stresses. For example, it has been demonstrated that some bacterial pathogens may develop a tolerance or adaptation to organic acids following prior exposure to low pH. While this increased resistance may be a problem in the application of organic acids for controlling pathogens, it has not been shown to occur in an actual food-processing system (Davidson and Harrison, 2002).

1.4 Considerations in the Applications of Food Antimicrobials Detailed considerations in the application of food antimicrobials may be found in Chapter 22, “Applications of Antimicrobials to Foods – A Food Industry Perspective.” Subjects covered include research and development perspectives, how to determine efficacy with single and multiple antimicrobials, and the steps used in the food industry for identifying and determining the feasibility for application of antimicrobials to a food product. The technology development steps involve determinations of efficacy, cost-in-use, capital expenditure, minimum order quantity, patents, regulatory aspects, challenge

7

Food Antimicrobials – An Introduction

studies, sensory and shelf-life studies, and production impacts. Scale-up and commercialization for various types of food products are also detailed. Other considerations for application include toxicological safety, and labeling.

1.4.1 Toxicological Safety Perhaps the most important aspect of any compound proposed for use as a food additive would be its toxicological characteristics. It is obviously essential that an additive for use as a food antimicrobial be safe for human consumption. The safety of some food antimicrobials has been questioned which has led to limitations on their use. The stringent requirements for toxicological safety testing can result in costs in the millions of dollars before approval for a new additive can be obtained. Ultimately, of course, an antimicrobial must be nontoxic to test-animals and humans, based on several studies. It is also important that the antimicrobial be metabolized and excreted by the body. The compound or its breakdown products should also not result in buildup of residues in body tissues. Valid assay methods for the antimicrobial are also a necessity so that levels can be easily followed. The majority of the antimicrobials now being used in food products have been extensively tested for toxicological safety, and although questions will continue to be asked, an evaluation of the risks vs. the benefits indicates that these antimicrobials are acceptable. A possible total or partial shift to naturally occurring antimicrobials is being investigated by many researchers at the present time. Because they occur in nature, it is often thought that naturally occurring antimicrobials are less toxic than synthetic compounds. Obviously, this is not always true. A naturally occurring antimicrobial must be shown to be non-toxic either by animal testing or by its continuous consumption by consumers as a food over a long period of time. The latter may be problematic even for some common potential natural antimicrobials such as spice extracts. This is because, while spices have been consumed for centuries, they are not normally consumed in the concentrations necessary to achieve antimicrobial activity. In addition to lack of toxicity, naturally occurring compounds must be able to be metabolized and excreted so as to not lead to residue build-up. In addition, they should be non-allergenic and should not bind or destroy important nutrients in a food product (Harlander, 1993).

1.4.2 Labeling with Respect to Antimicrobials One of the alleged attractions of naturally occurring antimicrobials is their reduced negative impact on the labeling of foods. Consumers are reportedly concerned about the presence of synthetic chemicals in their foods and would prefer natural compounds. This has led to the so-called “clean-label” movement which has its roots in the health and wellness movement of the early 2000s. Clean labels are not regulatory body-driven but rather appear to be consumer-driven (Table 1.3). To date there is not an established, objective, and common definition of what a clean label is, but rather several definitions or interpretations,

TABLE 1.3 Regulatory Definitions for Use of Food Antimicrobials Processing Aids Preservatives Antimicrobials: -FDA -USDA FSIS Natural Antimicrobials: -FDA -USDA Clean Label: FDA and USDA

21 CFR 178 Indirect Food Additives 21 CFR 101.22 (a) (5) 21 CFR 170.3 (o) (2) 9 CFR 430.1 No definition currently, proposed rule and comment period 21 CFR 101.22 (1982) – Vinegar and Lemon Juice Not defined

8

Antimicrobials in Food TABLE 1.4 Clean Label “Interpretation”—Based on Consumer, Retail, and Restaurant Perspective (David, 2016) Consumer Requirements for “Clean Label” Fewer ingredients (30–35 mm in diameter, intermediate with a zone of 20–30 mm, or resistant with a zone log 6 CFU/ml because selection of resistant mutants may occur. A tube or flask without added antimicrobial should be prepared as a control. The medium is incubated at the optimum temperature of the test microorganism for up to 48 h. The medium is sampled at appropriate times (e.g., 0, 2, 4, 8, 12, 24 and 48 h) and the number of viable microorganisms determined by the spread or pour plate method. Several responses of the test microorganism may be encountered in this type of analysis, including stationary-phase growth level suppression, lag-phase increase, decrease in the growth rate during log phase and lethality (Davidson and Parish, 1989). In practice a distinction is frequently drawn between a fungistatic or bactericidal action and a fungicidal or bactericidal action. This difference may not be completely justified since the former two differ from the latter in the death rate of the microorganisms. In the long term, the effect of added antimicrobial in food is either to inactivate (kill) the microorganisms or to eventually allow them to recover and grow. The governing factor here is the dosage of antimicrobial (Figure 2.7). Inhibition curves can be used to evaluate the antimicrobial action of several agents against bacteria, yeast and molds. However, for molds their radial growth rate as well as the germination time (time to form a visible colony) are often used as measures of antimicrobial effectiveness (López-Malo et al., 1995, 1997, 1998; Lahlali et al., 2005; Baert et al., 2007; Gómez-Ramírez et al., 2013; Mani-López et al., 2018). The inhibition curves methodology is the only technique that demonstrates lethality. This method is versatile but has some disadvantages, including that no single statistic is produced to compare such treatments as MIC. It is also labor-intensive and expensive. Modeling the growth kinetics of microorganisms (Zwietering et al., 1990, 1991; McMeekin et al., 1993; Whiting, 1993, 1995, 1997; Velázquez-Nuñez et al., 2013; Ozcakmak and Gul, 2017; Mani-López et al., 2018; Ro et al., 2018) allows improved statistical analysis of growth-inhibition curves in the presence of food antimicrobials. Secondary models also can describe the individual or combined effects of antimicrobials under specific growth conditions (pH, aw, temperature) using parameters estimated from primary models (germination time, radial growth rate, lag phase duration and decimal reduction rate). In addition, secondary empirical models describe only the effect of antimicrobials under specific tested conditions on microorganism growth. Zhang et al. (2015) described the antimicrobial interaction effects of apple skin polyphenols, acetic acid, oregano essential oil and carvacrol on Salmonella in Without antimicrobial action Insufficient antimicrobial action

microbiostasis

Sufficient antimicrobial action Disinfection Time FIGURE 2.7  Effect of antimicrobial agent dose on microbial response. (Adapted from Lück and Jager, 1997.)

27

Methods for Activity Assay and Evaluation of Results

sliced cooked ham through a secondary empirical model. A secondary model obtained from Gompertz parameters was used to describe the growth rate of Aspergillus flavus under different values of cinnamon essential oil and pH, aw, casein or fat, to identify the effects of their interactions on the mold growth (Kosegarten et al., 2017). Probabilistic modeling has been used to determine the growth/no-growth interface. Probabilistic models based on logistic regression analysis provide a useful way to describe the growth/no growth boundary (Ratkowsky and Ross, 1995; Presser et al., 1998; López-Malo, et al., 2000; Gómez-Ramírez et al., 2013; Jamshidi et al., 2014; Kosegarten et al., 2017), including the effect of selected antimicrobial concentrations. This methodology allows the definition of the antimicrobial concentration range necessary to inhibit growth under selected environmental conditions (Alzamora and López-Malo, 2002; Gómez-Ramírez et al., 2013; Jamshidi et al., 2014; Kosegarten et al., 2017). A second method for determining antimicrobial effectiveness over time is to measure turbidity increases with a spectrophotometer. A major disadvantage of this type of analysis is the sensitivity of the instrument. Spectrophotometers generally require log 6.0–7.0 CFU/ml for detection (Brock et al., 1984; Piddock, 1990). This may create a situation in which no growth (i.e., no absorbance increase) is observed when, in fact, undetectable growth is occurring at levels below log 5.0 CFU/ml. An erroneous interpretation of “lethality” could result (Davidson and Parish, 1989). A method of using turbidimetric analysis to determine infinite inhibitory concentrations (IIC, analogous to MIC) was developed by Marwan and Nagel (1986). In their procedure, the time for a microbial population to reach a specific turbidity in the presence of an antimicrobial was divided by the time for the same population to reach the turbidity in the absence of the antimicrobial. This was termed the relative effectiveness (RE). When 1/RE was plotted versus concentration of the inhibitor, the response was linear. Marwan and Nagel determined that at 1/RE = 0 (IIC), the concentration of antimicrobial would totally inhibit the test microorganism. As an example, the inverse of lag time of Penicillium glabrum inoculated in PDA formulated with selected concentrations of vanillin and potassium sorbate is shown in Figure 2.8. The response was linear and the calculated inhibitory concentrations, infinite lag time, correspond to MIC determined using the agar dilution method. Lambert and Pearson (2000) described a test to obtain MIC using optical density (OD) determinations. The basis of the technique is the comparison of the area under the optical density versus the time curve of the control (microorganism without antimicrobial) with the areas of the microbial curve with the test antimicrobial in several dilutions. The authors use 100-well plates and an OD automatic reader capable of giving an output signal based on a microbial growth criterion. As the amount of antimicrobial agent in a well increases, the effect on the growth is noticed by a reduction in the area under the curve. In general, plotting the inhibitor concentration on a logarithmic scale gives a characteristic sigmoid-shaped curve. The curve can be split into three principle regions: a region where the presence of the antimicrobial has no effect on the organisms relative to the control growth, a region where there is increasing inhibition of growth and a region where there is no measurable growth relative to the 1/Lag 0.014

Penicillium glabrum

0.012 0.01

Vanillin

0.008 0.006 0.004

Potassium sorbate

0.002 0

0

200

400

600

800 1000 1200 1400 1600

Antimicrobial concentration (ppm) FIGURE 2.8  Relative effectiveness (inverse of lag time) of selected concentrations of vanillin and potassium sorbate on Penicillium glabrum inoculated in PDA.

28

Antimicrobials in Food

control. This approach permits the calculation of the MIC as the concentration above which no growth is observed relative to the control. To summarize the use of in vitro methods, they should be used together, one end-point and one descriptive method. The end-point method helps to determine the approximate effective concentration, and the descriptive method evaluates the effect of a compound on growth over time.

2.6 Combined Antimicrobial Systems Traditionally, it was common to use only one chemical antimicrobial agent in a food product for preservation purposes (Busta and Foegeding, 1983); however, in recent years, the use of combined agents in a single food system has become more frequent. The use of combined antimicrobial agents theoretically provides a greater spectrum of activity, with increased antimicrobial action against the pathogenic or spoilage organisms. It is thought that combined agents act on different species of a mixed microbiota or act on different metabolic elements within similar species or strains. This theoretically results in improved microbial control over the use of one antimicrobial agent alone; however, actual proof of improved efficacy requires subjective interpretations. Although testing of combined antibiotics for clinical use is well-studied and relatively well-standardized (Barry, 1976; Krogstad and Moellering, 1986; Eliopoulos and Moellering, 1991), application of such methodology to antimicrobials in food is much less so. However, such systems have gained attention more recently because of the desire to minimize the sensory effects of high concentrations of certain antimicrobials or to potentiate the antimicrobial effect of single antimicrobials. There have been several studies on the antimicrobial combinations to preserve or extend the shelf life of foods including nisin and bovine lactoferrin (Turkish-style meatball, Colak et al., 2008), reuterin and nisin (milk, Arqués et al., 2011), cumin essential oil and nisin (commercial barley soup, Pajohi et al., 2011), chitosan and oregano essential oil (red porgy, Vatavali et al., 2013), dimethyl decarbonate and nisin (litchi juice, Yu et al., 2013), clove and mustard essential oil both in vapor phase (strawberries, Aguilar-González et al., 2015), Chinese cinnamon and cinnamon essential oils (readyto-cook meat; Ghabraie et al., 2016), oregano and rosemary essential oils (iceberg lettuce and chard, de Medeiros Barbosa et al., 2016), Blepharis cuspidata and Boswella agadensis essential oils (Gadisa et al., 2019) and thyme and Mexican oregano essential oils in vapor phase (Reyes-Jurado et al., 2019a). Methods of testing combined antimicrobials usually involve agar diffusion, agar or broth dilution, time-kill curves and vapor-phase methods. Agar diffusion is a simple qualitative method that often requires subjective interpretation of inhibition zone shapes to determine the efficacy of combined antimicrobials (Barry, 1976). This method provides a measure of microbial growth inhibition but does not address biocidal activity. Dilution and vapor-phase methods yield quantitative data and are most often conducted with various combined concentrations of two antimicrobials arranged in a “checkerboard” array. The checkerboard method is the technique used most frequently to assess antimicrobial combinations in vitro, presumable because (a) its rationale is easy to understand, (b) the mathematics necessary to calculate and interpret the results are simple, (c) it can readily be performed in the laboratory using microdilution systems and (d) it has been the technique most frequently used in studies that have suggested an advantage of synergistic interactions of antibiotics in clinical treatments (Eliopoulos and Moellering, 1991). The term checkerboard refers to the pattern (of tubes or microtiter wells or closed systems) formed by multiple dilutions of the two antimicrobials being tested in concentrations equal to, above and below their MIC (Table 2.3). Traditional clinical dilution testing uses two-fold dilutions of test compounds, but testing of food antimicrobials is often conducted with alternate dilution schemes (Rehm, 1959; Davidson and Parish, 1989; Eliopoulos and Moellering, 1991). Inhibition or time-kill curves can also be used for the evaluation of combined antimicrobials (Krogstad and Moellering, 1986; Eliopoulos and Moellering, 1991). The advantage of this test is that rates of lethality can be determined; however, repetitive sampling limits the number of combinations that can be assayed at any one time. Combined studies are conducted to determine if specific types of interactions occur between the two combined antimicrobials. Traditionally, the terms “additive”, “antagonistic” and “synergistic” were used to describe possible antimicrobial interactions. Garreti (1958) suggested the terms “additivity” and

29

Methods for Activity Assay and Evaluation of Results TABLE 2.3

Checkerboard Array in Which Serial Dilutions of Two Antimicrobials Are Performed Using Concentrations Proportional to the MICs of the Antimicrobials Being Tested Antimicrobial B

2.0 1.0 0.50 0.25 0.12 0.06 0

2.0/0 1.0/0 0.50/0 0.25/0 0.12/0 0.06/0 0/0 0

2.0/0.06 1/0.06 0.50/0.06 0.25/0.06 0.12/0.06 0.06/0 0/0.06 0.06

2.0/0.12 2.0/0.25 1.0/0.12 1.0/0.25 0.50/0.12 0.50/0.25 0.25/0.12 0.25/0.25 0.12/0.12 0.12/0.25 0.06/0.12 0.06/0.25 0/0.12 0/0.25 0.12 0.25 Antimicrobial A

2.0/0.50 1.0/0.50 0.50/0.50 0.25/0.50 0.12/0.50 0.06/0.50 0/0.50 0.50

2.0/1.0 1.0/1.0 0.50/1.0 0.25/1.0 0.12/1.0 0.06/1.0 0/1.0 1.0

2.0/0 1.0/0 0.50/0 0.25/0 0.12/0 0.06/0 0/2.0 2.0

“equivalence” to describe combination interactions; however, our discussion is limited to the more traditional descriptors. Additivity occurs when two combined antimicrobials give results that are equivalent to the sum of each antimicrobial acting independently. There is no enhancement or reduction in overall efficacy for the combined antimicrobials compared to the individual results. This is also sometimes referred to as “indifference” (Krogstand and Moellering, 1986; Eliopoulos and Moellering, 1991). Antagonism refers to reduced efficacy of the combined agents compared to the sum of the individual results. Synergism is an increase or enhancement of overall antimicrobial activity when two agents are combined compared to the sum of individual results. A conclusion of synergism must be approached with caution since it implies that a reduction in the overall antimicrobial concentrations might be achieved in a food system without a reduction in efficacy. Gardner (1977) stated that true synergism is quite rare in relation to combined antibiotics. Other concerns about the misuse of the term “synergism” in relation to antimicrobials have been previously cited (Davidson and Parish, 1989; Garrett, 1958). Most commonly, additive interactions are misidentified as synergistic. A case in which an increase in antimicrobial activity is observed upon the addition of a second compound to a food system does not necessarily constitute synergy. A conclusion of synergism requires that the overall efficacy of the combination be significantly greater than the sum of the efficacies of the individual compounds. From a practical standpoint, processors must beware of the temptation to utilize combined antimicrobials without thorough scientific scrutiny. This is mainly because of possible antagonistic interactions that can lower overall antimicrobial efficacy compared to the individual compounds used singly. Some food-related antimicrobial combinations that have been interpreted as antagonistic include sulfite and butylated hydroxyanisole, ethanol and butyl paraben, SO2 and butyl paraben, SO2 and benzoate and SO2 and sorbate (Parish and Carroll, 1988b; Davidson and Parish 1989).

2.7 Application Studies The primary purpose of in vitro methods is to determine the concentration of an antimicrobial necessary to inhibit a microorganism in a laboratory medium. However, once it has been determined that an antimicrobial is effective using in vitro methods, it is still necessary to apply the compound to a food system. Often, an antimicrobial that performed well in a microbiological medium is shown to have little or no effect in a food. This is the result of many interacting factors in a food, such as proteins, lipids, cations, binding to food components, inactivation by other additives, pH effects on antimicrobial stability and activity, uneven distribution in the food matrix and poor solubility, among others. Before actual application of an antimicrobial to a food, it may be useful to evaluate the effect of some components in a food system that may influence the effectiveness of the compounds, such as lipids, proteins, additives and divalent cations. Lipids may case a decrease in activity of lipophilic compounds,

30

Antimicrobials in Food

and since many food antimicrobials have a hydrophobic character, there is invariably some reduction (Rico-Muñoz and Davidson, 1983; García-Díez et al., 2017; Kosegarten et al., 2017). Proteins may cause binding of some compounds and reduce activity (Rico-Muñoz and Davidson, 1983) or not influence the antimicrobial effect (Kosegarten et al., 2017; García-Díez et al., 2017). López-Malo et al. (1995) prepared fruit-based agars containing mango, papaya, pineapple, apple and banana with up to 2000 ppm vanillin to assess the effect of fruit composition on the MIC to inhibit A. flavus, A. niger, A. parasiticus and A. ochraceus. The small differences in protein and lipid composition between banana or mango and the other fruits increased vanillin MIC; the effect is attributed to binding of the phenolic vanillin by protein. Divalent cations may affect the activity of some compounds by affecting the microorganism itself or by interacting with the antimicrobial (Rico-Muñoz and Davidson, 1984). Further information can be gained by using model systems that include a percentage of a food in a buffer or microbiological medium. For example, Dje et al. (1989) evaluated antimicrobials in model systems containing a 10% (wt/vol) suspension of raw chicken or frankfurters. They prepared the suspensions by homogenizing 25 g product with 225 ml 0.1% peptone diluent. A homogeneous suspension of frankfurter was facilitated by adding 0.25% Tween 80 (sorbitan monooleate polyoxyethylene). In addition to determining the effect of food components on activity, this procedure allows better control of environmental conditions and natural microbiota. In both preceding model systems, a modified broth dilution assay could be used to determine an MIC (end-point method), or periodic sampling (descriptive method) could be run. In applying an antimicrobial to a food, the microorganisms should include natural contaminants (bioburden) and the pathogen of interest. The incubation conditions should reflect normal storage and abuse. The success of the test may be through increased shelf life or reduced growth rate of a food-borne pathogen. Few food antimicrobials actually cause lethality to food-borne microorganisms at normal use concentrations. No standardized methods are available or application methods; however, a modification of the end-point method or time-kill curve is possible. Although it is impractical to design uniform procedures for all products, as is done for screening methods, a good approach is to develop a uniform inhouse procedure based upon standardized procedures. Microbial challenge testing can be used to study the fate of pertinent microorganisms in foods formulated with antimicrobial agents as a way to assess product stability and safety (Tapia et al., 1995). The microbial challenge test has become an established technique within the food industry in order to simulate what can happen to a product during processing, distribution and subsequent handling, following inoculation with relevant microorganisms and further holding under controlled conditions (Notermans et al., 1993). This procedure can be used to verify antimicrobial agent effectiveness in foods. For foods, the common application method of antimicrobial agents in vapor-phase is through the vapor-phase assay in which the system is scaled to a closed “big box” instead of a Petri dish. However, for certain foods such as bread slices, Petri dishes can be utilized; the antimicrobial agent is loaded in a paper disc placed in the plate lid (Krisch et al., 2013). Hermetic glass jars, hermetic polypropylene boxes or polyethylene terephthalate (PET) jars are commonly utilized to create an atmosphere of vapors of the antimicrobial agent which are thus in contact with the tested food (Aguilar-González et al., 2015; ManiLópez et al., 2018). A known concentration of antimicrobial agent is placed inside the container separately from food. If the amount to be tested is small (μl range), the compound may be added to a piece of paper, or if a larger amount is necessary (ml range) a glass container may be used (see Figures 2.9 and 2.10). The jars (or boxes) are kept at a controlled temperature under standard conditions depending on the target microorganism and observed after the corresponding incubation period to determine the antimicrobial affect. The MIC is defined as the lower concentration in which microbial growth is inhibited and is expressed as μl (or ml) of antimicrobial agent per liter of air.

2.7.1 Interpretation of Results The most difficult aspect of antimicrobial testing is appropriate data analysis and interpretation. Erroneous research conclusions from improper data interpretation can ultimately lead to industrial losses when practiced commercially. It is therefore necessary for data analysis to be conducted with great care. Agar diffusion techniques produce quantitative zone diameter data. These results indicate relative microbial inhibition among a group of antimicrobials or test microorganisms. In antibiotic disk testing,

31

Methods for Activity Assay and Evaluation of Results Paper loaded with the antimicrobial agent (essential oil)

apple strawberry

FIGURE 2.9  Vapor-phase test application for apple or strawberry in polyethylene terephthalate (PET) jars.

Glass container of antimicrobial agent (essential oil) inside the glass jars. Paper loaded with the antimicrobial agent (essential oil) inside the glass jar.

FIGURE 2.10  Vapor-phase tests application for bread (left) or alfalfa seeds (right) in glass jars.

the inhibition zone diameter of a specific test organism and antibiotic is compared to zone sizes produced with control organisms (Figure 2.1). The method correlates inhibition zone size with the concentration of the antimicrobial agent (Conte and Barriere, 1992). It must be noted that inhibition zone sizes are influenced by a number of factors, including inoculum density, agar depth, agar medium composition, incubation time and temperature (Barry, 1986). Combined antimicrobial testing using the agar diffusion method produces inhibition zones that are interpreted depending upon their shape. This technique makes use of paper strips or disks impregnated with a specific quantity of antimicrobial gent and placed on an agar surface seeded with the test microorganism. This method is more qualitative than dilution testing but is useful as an end-point method for large numbers of preservative combinations. As previously discussed, dilution testing results in an MIC. This reflects the minimum concentration of a compound necessary to inhibit the growth of a microorganism under specific conditions including time and temperature. It should be noted that MIC could vary depending on factors mentioned earlier. Interpretation of dilution results for combined testing is usually conducted with isobolograms. An isobologram may be thought of as an array of differing concentrations of two compounds, where one compound ranges from lowest to highest concentration on the x-axis and the other on the y-axis (Figure 2.11). All possible permutations of combined concentrations are reflected within the array. If those concentrations that inhibit the growth of the test organism fall on an approximately straight line that connects the

32

Antimicrobials in Food

Compound B

Antagonism Additive

Synergism

Compound A FIGURE 2.11  Interpretation of time-kill curves for combined antimicrobials.

individual MIC on the x- and y-axes, the combined effect is additive. Deviation of linearity to the left or right of the additive line is interpreted as synergism or antagonism, respectively. Isobologram construction can be simplified using fractional inhibitory concentrations (FIC), which are MIC normalized to unit. The FIC is the concentration of a compound needed to inhibit growth (expressed as a fraction of its MIC) when combined with a known amount of a second antimicrobial compound. It is calculated as the ratio of the MIC of a compound when combined with a second compound divided by the MIC of the first compound alone. Additive, synergistic or antagonistic interactions are interpreted as with an MIC isobologram. For example, if the MIC for sorbate (under specific test conditions) is determined to be 200 ppm and the MIC for sorbate when combined with x ppm of propylparaben is 120 ppm, then the FIC for sorbate in the presence of x ppm propylparaben is 120/200 = 0.6. A modified method for determining the fractional inhibitory concentration of two or more compounds in combination was described by Techathuvanan et al. (2014). The method essentially selects combinations of antimicrobials to be tested that lie in the three potential areas of an isobologram, antagonism, additive or synergism. By starting with MIC and using fractions thereof for each antimicrobial, one can interpret responses of growth or no growth as antagonism, additive or synergism (Table 2.4). This technique has potential to be used to rapidly screen multiple combinations only for synergistic effects if desired. The FIC of two compounds in an inhibitory combination may be added to give a total FICindex. If the FIC of propylparaben in the previous example was determined to be 0.5, then FICindex would be FIC (sorbate) + FIC (propylparaben) = 0.6 + 0.5 = 1.1. An FICindex near 1 indicates additivity, whereas 1 indicates antagonism (Table 2.5). The degree to which a result must be less than or greater than 1 to indicate synergism or antagonism is a matter of interpretation. Squires and Cleeland (1985) proposed that additive results are indicated by FICindex between 0.5 and 2.0 for antibiotic testing. Synergism and antagonism are indicated by results 2.0, respectively. Research is needed to provide a database for proper interpretation of FIC and of the FICindex in relation to food antimicrobial systems. Data interpretation must be conducted conditionally and will depend upon a number of variables, such as specific test conditions, microbial strain and target food system. It should be noted that interpretations might also vary depending upon the specific concentrations of each antimicrobial used in combination. Parish and Carroll (1988a) observed additivity between SO2 and either sorbate or butylparaben when the inhibitory concentration contained less than 0.25 FIC of SO2; however, the same combination at higher SO2 FIC indicated antagonistic results. Rehm (1959) observed similar anomalies when sodium sulfite was combined with formate or borate. Fractional lethal concentrations (FLC) can be determined when a broth dilution assay is used. This is the concentration of single or combined agents that kills the population of test organism to a non-recoverable level under the test conditions. The FLC is determined by inoculating broth media without antimicrobial agents with aliquots from those samples in the MIC test that inhibited growth. The concentrations

33

Methods for Activity Assay and Evaluation of Results TABLE 2.4

Use of Modified Fractional Antimicrobial Concentrations for Antimicrobial Combination Testing of Two or More Antimicrobials Antimicrobial (AM) (x MIC) AM1a

AM 2b

AM3c

Fractional Inhibitory Concentration Index

Binary antimicrobial combinations 0 0 NAd 1.0 0 NA 0 1.0 NA 0.75 0.25 NA

0 1 1 1

0.50

0.50

NA

1

0.25

0.75

NA

1

0.25

0.25

NA

0.5

0.75

0.75

NA

1.5

Triple antimicrobial combinations 0 0 0 1 0 0 0 1 0 0 0 1 0.33 0.33 0.33

0 1 1 1 1

0.50

0.25

0.25

1

0.25

0.50

0.25

1

0.25

0.25

0.50

1

0.167

0.167

0.167

0.5

0.50

0.50

0.50

1.5

Bacterial Growth

Descriptor of Results

Growth No growth No growth Growth No growth Growth No growth Growth No growth Growth No growth Growth No growth

Control Minimum Inhibitory Conc. Minimum Inhibitory Conc. Additivee or antagonistic Additive effect

Growth No growth No growth No growth Growth No growth Growth No growth Growth No growth Growth No growth Growth No growth Growth No growth

Control Minimum Inhibitory Conc. Minimum Inhibitory Conc. Minimum Inhibitory Conc. Additive or antagonistic Additive effect Additive or antagonistic Additive effect Additive or antagonistic Additive effect Additive or antagonistic Additive effect Additive or antagonistic Synergistic effect Antagonistic effect Additive or synergistic

Additive or antagonistic Additive effect Additive or antagonistic Additive effect Additive or antagonistic Synergistic effect Antagonistic effect Additive or synergistic

AM1 = first antimicrobial, bAM2 = second antimicrobial, cAM3 = third antimicrobial. NA = not applicable. e Synergistic, additive and antagonistic effects of combined antimicrobials correspond to FICI of 1.5, respectively. Adapted from: Techathuvanan et al., 2016. a

d

that had a lethal effect remain negative for growth after appropriate incubation, but growth is observed in the concentrations that had only an inhibitory effect. Inhibition or time-kill curves (log CFU/ml versus time) can be graphically interpreted for combination interaction (Barry, 1976; Krogstad and Moellering, 1986; Eliopoulos and Moellering, 1991). Since this technique uses a fixed concentration of each antimicrobial and must be repeated when a concentration is altered, it is logistically difficult and is most often used to enhance results observed with dilution methods. Survivor curves are generated with four treatments: (1) compound A alone, (2) compound B alone, (3) compound A and B together and (4) control A without A or B (Figure 2.12). Additivity is observed when the time-kill curve for the combined antimicrobials is similar to the curve for the more active

34

Antimicrobials in Food

TABLE 2.5 Quantitative Definitions of Results with Antimicrobial Combinations Additivity   The result with two antimicrobials is equal to the   sum of the results for each of the antimicrobials   used separately Antagonism   The result with two antimicrobials is significantly   less than the additive response Synergism   The result with two antimicrobials is significantly greater   than the additive response

FICA + FICB = 1.0

FICA + FICB > 1.0

FICA + FICB < 1.0

FICA and FICB are the fractional inhibitory concentrations of antimicrobial A and B, respectively.

Log CFU/ml

control

(a)

control

control

A

A

A

B

B

B

A+B

A+B A+B Time

(b)

Time

(c)

Time

FIGURE 2.12  An isobologram than enables interpretation of dilution results when testing combined antimicrobials: (a) additive, (b) synergistic, or (c) antagonistic results.

compound alone. Antagonism results if the combined effect is significantly less than that observed for either compound alone. If the time-kill curve for the combined compounds is significantly reduced from both individual curves, the result is interpreted as synergistic.

2.8 Acknowledgments Authors López-Malo, Mani-López and Palou gratefully acknowledge financial support from the National Council for Science and Technology (CONACyT) of Mexico (Project CB-2016-01-283636) and Universidad de las Américas Puebla (UDLAP Projects 2409 and 3555).

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Parish, M. E. and Carroll, D. E. 1988a. Minimum inhibitory concentration studies of antimicrobial combination against Saccharomyces cerevisae in a model broth system. J. Food Sci. 53:237–239. Parish, M. E. and Carroll, D. E. 1988b. Effects of combined antimicrobial agents on fermentation initiation by Saccharomyces cerevisae in a model broth system. J. Food Sci. 53:240–242. Paton, J. H., Holt, H. A. and Bywater, M. J. 1990. Measurement of MICs of antibacterial agents by spiral gradient endpoint compared with conventional dilution method. Int. J. Exp. Clin. Chemotherapy. 3:31–38. Pereira, E., Santos, A., Reis, F., Tavares, R. M., Baptista, P., Lino-Neto, T. and Almeida-Aguilar, C. 2013. A new effective assay to detect antimicrobial activity of filamentous fungi. Microbiol. Res. 168:1–5. Piddock, L. J. 1990. Techniques used for determination of antimicrobial resistance and sensitivity in bacteria. J. Appl. Bacteriol. 68:307–318. Presser, K. A., Ross, T. and Ratkowsky, D. A. 1998. Modelling the growth limits (growth/no growth interface) of Escherichia coli as a function of temperature, pH, lactic acid concentration and water activity. Appl. Environ. Microbiol. 64:1773–1779. Ratkowsky, D. A. and Ross, T. 1995. Modeling the bacterial growth/no growth interface. Lett. Appl. Microbiol. 20:29–33. Rehm, H. 1959. Untersuchung zur Wirkung von Konservierungsmittel kombinationen I. Z. Lebesm. Untersuch. Forsch. 110:283–293. Reyes-Jurado, F., Cervantes-Rincón, T., Bach, H., López-Malo, A. and Palou, E. 2019a. Antimicrobial activity of Mexicano regano (Lippia berlandieri), thyme (Thymus vulgaris) and mustard (Brassica nigra) essential oils in gaseous phase. Ind. Crops Prod. 131:90–95. Reyes-Jurado, F., Navarro-Cruz, A. R., Ochoa-Velasco, C. E., Palou, E., López-Malo, A. and Ávila-Sosa, R. 2019b. Essential oils in vapor phase as alternative antimicrobials: A review. Crit. Rev. Food Sci. Nutr. 18:1–10. Rico-Muñoz, E. and Davidson, P. M. 1983. Effect of corn oil and casein on the antimicrobial activity of phenolic antioxidants. J. Food Sci. 48:1284–1288. Rico-Muñoz, E. and Davidson, P. M. 1984. Effect of calcium and magnesium on the antibacterial activity of phenolic antioxidants against Staphylococcus aureus A100. J. Food Sci. 49:282–283. Ro, E. Y., Kim, G. S., Kwon, D. Y., Park, Y. M., Cho, S. W., Lee, S. Y., Yeo, I. H. and Yoon. K. S. 2018. Effect of natural antimicrobials with modified atmosphere packaging on the growth kinetics of Listeria monocytogenes in ravioli at various temperatures. J. Food Safety 38:e12392 (8 pp). Rodrigues, L. B., dos Santos, L. R., Rizzo, N. N., Ferreira, D., de Oliveira, A. P., Levandowski, R., Webber, B. and do Nascimento, V. P. 2018. ATP-bioluminescence and conventional microbiology for hygiene evaluation of cutting room surfaces in poultry slaughterhouse. Acta Sci. Vet. 46:1534 (6 pp). Schalkowsky, S. 1986. Plating systems. In: Food Borne Microorganisms and Their Toxins: Developing Methodology, edited by M. Pierson and N. Stern, Marcel Dekker, New York. Schoenknecht, F. D., Sabath, L. D. and Thornsberry, C. 1985. Susceptibility tests: Special test. p. 1000. In: Manual of Clinical Microbiology, 4th ed., edited by E. Lennette, American Society for Microbiology, Washington, DC. Seo, H. S., Beuchat, L. R., Kim, H. and Ryu, J. H. 2015. Development of an experimental apparatus and protocol for determining antimicrobial activities of gaseous plant essential oils. Int. J. Food Microbiol. 215:95–100. Spiral Biothec, Inc. 2001. SGE, Spiral Gradient Endpoint, User Guide, Spiral Biothec, Norwood, MA. Squires, E. and Cleeland, R. 1985. Methods of Testing Combinations of Antimicrobial Agents, HoffmanLaRoche, Inc., Nutley, NJ. Sun, W., Weingarten, R. A., Xu, M., Southall, N., Dai, S., Shinn, P., Sanderson, P. E., Williamson, P. R., Frank, K. M. and Zheng, W. 2016. Rapid antimicrobial susceptibility test for identification of new therapeutics and drug combinations against multidrug-resistant bacteria. Emerg. Microbes Infect. 5:11 p. Synbiosis, A Division of Synoptics Ltd. 2019. Protocol 3 – Automatic colony counting and zone measuring, Product description, Cambridge, UK. Tapia de Daza, M. S., Argaiz, A., López-Malo, A. and Díaz, R. V. 1995. Microbial stability assessment in high and intermediate moisture foods: special emphasis on fruit products. p. 575. In: Food Preservation by Moisture Control, Fundamentals and Applications, edited by G. V. Barbosa-Cánovas and J. WeltiChanes, Technomic Publishing, Inc, Lancaster, PA.

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3 Sodium Benzoate and Benzoic Acid John R. Chipley CONTENTS 3.1 3.2 3.3

Introduction and Historical Background........................................................................................41 Physical and Chemical Properties and Natural Occurrence......................................................... 42 Antimicrobial Activity.................................................................................................................. 44 3.3.1 Spectrum of Action........................................................................................................... 44 3.3.2 Influence of Other Chemicals and Physical Environment................................................ 48 3.3.3 Mechanism of Action........................................................................................................ 52 3.3.4 Acquired Resistance to Benzoic Acid.............................................................................. 55 3.4 Microbial Metabolism of Benzoic Acid........................................................................................ 59 3.5 Reaction with Food Constituents...................................................................................................61 3.6 Regulatory Status.......................................................................................................................... 62 3.7 Applications................................................................................................................................... 63 3.7.1 Use in Various Food Systems........................................................................................... 63 3.7.2 Use as a Postharvest Fungicide......................................................................................... 66 3.7.3 Other Applications............................................................................................................ 66 3.7.4 Storage and Handling....................................................................................................... 67 3.8 Toxicology..................................................................................................................................... 68 3.9 Assay............................................................................................................................................. 72 3.10 Acknowledgments......................................................................................................................... 73 References................................................................................................................................................. 73

3.1 Introduction and Historical Background Benzoic acid is one of the oldest chemical preservatives used in the cosmetic, drug, and food industries. Sodium benzoate was the first chemical preservative approved for use in foods by the U.S. Food and Drug Administration (FDA) (Jay, 2000). Its preservative action appears to have been first described in 1875 when researchers discovered that sodium benzoate had strong antifungal properties, and was relatively tasteless in foods, easy to make, and inexpensive. It was not introduced for commercial food preservation until around 1900 because benzoic acid could not be produced synthetically in large quantities (Lueck, 1980). An informative historical narrative published recently (Blum, 2018) details the dangers of consuming processed food in the U.S. at the end of the nineteenth century. In most cases, substandard dairy and meat products, alcoholic beverages, and canned vegetables which should have been discarded were knowingly sold to customers for consumption after the addition of chemicals, often at lethal concentrations. For example, milk adulterated with formaldehyde was estimated to have killed thousands of children in New York City alone. In the production of ketchup, tomato trimmings were “thickened” with various non-tomato byproducts, processed under non-sanitary conditions, and a large dose of preservative added to extend shelf life. The preservative of choice was sodium benzoate. Food companies found it profitable to use rotten tomato waste and scraps in their production ketchups and supplement them with up to four times the proposed government standard of 0.10% for this preservative (Blum, 2019). 41

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Currently, the advantages of low cost, ease of incorporation into products, lack of color, and relatively low toxicity have subsequently caused benzoic acid (and its salts) to become one of the most widely used preservatives in the world (Davidson, 1997) when used at legally prescribed levels. During the last 15 years, several articles and references concerning various aspects of food additives and preservatives (e.g., benzoic acid) have been published. General evaluations of the use of preservatives in foods (Vogel, 1992), in prevention of microbial spoilage (Giese, 1994), in meat products (Gerhardt, 1995), in beverage manufacture (Giese, 1995), and in consumer attitudes toward the use of preservatives (Jager, 1994) serve as examples. In addition, references detailing characteristics of food spoilage yeasts (Deak, 2007), food preservation (Rahman, 2007), fungal food spoilage (Pitt and Hocking, 2009), and chemical preservatives and natural antimicrobial compounds (Davidson and Taylor, 2007; Davidson et al., 2013) provide excellent background information. Recently, an extensive review of benzoic acid as a naturally occurring compound in foods and as an antimicrobial additive was published by del Olmo et al. (2017). Detailed coverage of its uses in foods and non-food products, toxicology in humans and animals, current controversy involving long-term exposure, and analytical methods of detection were also presented.

3.2 Physical and Chemical Properties and Natural Occurrence Benzoic acid (C6H5COOH) and sodium benzoate (C6H5COONa) have the structural formulas shown in Figure 3.1. Benzoic acid (molecular weight 122.1), also called phenylformic acid or benzenecarboxylic acid, occurs in pure form as colorless or white needles or leaflets. It is soluble to a limited extent in water (0.18, 0.27, and 2.2 g dissolves in 100 ml water at 4°C, 18°C, and 75°C, respectively.) Sodium benzoate (molecular weight 144.1) is a white granular or crystalline powder. It is much more soluble in water than benzoic acid (62.8, 66.0, and 74.2 g dissolves in 100 ml water at 0°C, 20°C, and 100°C, respectively). For this reason, it is preferred for use in many cases. Potassium benzoate and calcium benzoate have also been approved for use, although their solubility in water is less than that of the sodium salt. Benzoic acid occurs naturally in several foods and commodities (Table 3.1). It accounted for approximately 16% of the growth inhibition of Saccharomyces bayanus and Pseudomonas fluorescens from ethanolic extracts of cranberries (Marwan and Nagel, 1986a). Benzoic acid has also been identified as a major constituent in extracts of blackberries (Humpf and Schreier, 1991); of mushrooms, depending on the variety (Abdullah et al., 1994); and of fresh tomatoes (Marlatt et al., 1992). Yogurts have been found to contain natural levels of benzoic acid (Stijve and Hischenhuber, 1984; Teuber, 1995). In an extensive review, Sieber et al. (1995) surveyed and analyzed many types of cultured dairy products and cheeses for the natural occurrence of benzoic acid (Table 3.2). Its presence appears to occur as a by-product of the microbial degradation of either hippuric acid or phenylalanine in these products (Figure 3.2). Oxidation of benzaldehyde may also contribute to benzoic acid generation. A fourth reaction involves hydrolysis of hippuric acid to yield benzoic acid and glycine, e.g., during the fermentation of goat’s and sheep’s milk (Hornickova et al., 2014). Both of these acids were detected at higher levels in milk from sheep (Table 3.3). This may be of significance in countries like Switzerland, where benzoic acid has not been approved as a food additive (Sieber et al., 1989, 1990). Benzoic acid has also been identified as a natural by-product in culture filtrates of Lactobacillus plantarum (Niku-Paavola et al., 1999). Several new types of low-molecular-mass compounds were

FIGURE 3.1  Structures of benzoic acid and sodium benzoate.

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Sodium Benzoate and Benzoic Acid TABLE 3.1 Natural Occurrence of Benzoic Acid Category Fruits and berries

Fermented products

Spices and flavors

Others

Product Apples Apricots Berries (i.e., blackberries) Blueberries Cherries Cranberries Grapes Plums, greengage Prunes Strawberries Tomatoes Beers Dairy, cultured Teas, black Wines Anise Cinnamon Chocolate Cloves, ripe Licorice Vanilla Brewer’s yeast Coffee beans Honey Milk Mushrooms Teas, green Tobacco

Note: Levels of benzoic acid vary (10 to 1000 mg/kg) depending on the food. Source: Adapted from Fenaroli (2002); Chipley (2005); Qi et al. (2009).

TABLE 3.2 Benzoic Acid Content of Cultured Dairy Products Product

Range of Concentration (mg/kg)

Yogurts Yogurt, fruit Sour cream Buttermilk Cottage cheese Cheese Cheese, nonsmear, ripened Cheese, smear, ripened

9–56 5–39 10–18 10–19 2–18 0–200 0–41 0–622

Source: Adapted from Sieber et al. (1995).

also identified, and a mixture of these, along with benzoic acid, was found to have antimicrobial properties. Naturally occurring non-flavonoid phenolic compounds, including derivatives of benzoic acid, were used to characterize commercial fruit juices (Fernandez de Simon et al., 1992). Orange, apple, pineapple, peach, apricot, pear, and grape juices were analyzed to establish phenolic profiles unique to each type of juice. In four of the seven juices, p-hydroxybenzoic acid was detected at 0.2 to 2.6 mg/l.

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FIGURE 3.2  Biosynthesis of benzoic acid by lactic acid bacteria in fermented dairy products. [Source: Adapted from Sieber et al. (1995).]

TABLE 3.3 Content of Hippuric Acid in Raw Milk and Benzoic Acid in Fermented Milk (A) Hippuric Acid (mg/kg) Average Range (B) Benzoic Acid (mg/kg) Average Range

Raw Goat’s Milk 15.5 ± 8.3 5.8–25.1

Raw Sheep’s Milk 43.3 ± 12.3 20.3–55.6

Ferm. Goat’s Milk 20.3 ± 13.9 5.0–52.8

Ferm. Sheep’s Milk 29.5 ± 16.1 4.9–77.8

Source: Adapted from Hornickova et al. (2014).

3.3 Antimicrobial Activity 3.3.1 Spectrum of Action Because the quantity of undissociated acid decreases with increasing pH (Table 3.4), the use of benzoic acid or sodium benzoate as a food preservative has been limited to those products that are acid in nature. TABLE 3.4 Effect of pH on the Dissociation of Benzoic Acid pH 3 4 5 6 7 pK

Undissociated Acid (%) 93.5 59.3 12.8 1.44 0.144 4.19

Source: From Baird-Parker (1980).

Sodium Benzoate and Benzoic Acid

45

Currently, these compounds are used primarily as antimycotic agents, and most yeasts and fungi are inhibited by 0.05% to 0.1% of the undissociated acid. Food-poisoning and spore-forming bacteria are generally inhibited by 0.01% to 0.02% undissociated acid, but many spoilage bacteria are much more resistant. Therefore, benzoic acid cannot be relied on to effectively preserve foods capable of supporting bacterial growth (Chichester and Tanner, 1972; Baird-Parker, 1980). MICs for some of the bacteria, yeasts, and fungi involved in food poisoning and food spoilage are given in Tables 3.5–3.6. Several factors interact to determine the MIC, including pH, temperature, genus and species of the microorganism in question, composition of the growth medium, prior exposure to the preservative, and environment from which the microorganism was originally isolated. The sensitivity of 42 yeast cultures to sorbic and benzoic acids, potassium sorbate, and sodium benzoate was determined (Manganelli and Casolari, 1983). The scattering of MIC values was lower with potassium sorbate and benzoic acid and higher with sorbic acid and sodium benzoate. The addition of benzoate to chemostat cultures of S. cerevisiae decreased the biomass and increased the specific oxygen uptake rate of cells (Verduyn et al., 1992). The effects of several preservatives and antimicrobial agents on aflatoxin B1 production by A. flavus have been reported (Bauer et al., 1981). In liquid media, all treated cultures produced measurable levels of toxin 3 to 7 days later than controls. In benzoic acid-supplemented cultures, aflatoxin B1 production was higher than in controls. The authors concluded that subinhibitory concentrations of these compounds may stimulate toxin production in some cases. Sodium benzoate was found to control both growth and aflatoxin production by Aspergillus parasiticus in liquid media (El-Gazzar and Marth, 1987). Increasing the concentration of sodium benzoate increased the percentage of inhibition at the end of incubation (10 days). The average accumulation of mycelial dry weight was greater in the presence of benzoate than in its absence, however, with the greatest increase occurring when the medium contained 0.3% sodium benzoate. Different concentrations of benzoic acid were tested to determine the effective levels capable of reducing the mycelial growth of six Fusarium and eight Penicillium species by 50% (Thompson, 1997). In general, Fusarium species were more sensitive to benzoic acid (210 to 420 µg/ml) than were Penicillium species (250 to 3000 µg/ml). Growth and aflatoxin production by toxigenic strains of Aspergillus were partially or completely inhibited by the undissociated form of six organic acid preservatives, including benzoic (Rusul and Marth, 1988). Salts, such as sodium and potassium chlorides and sodium nitrate, enhanced aflatoxin production when present at low levels but became inhibitory at higher levels. Garri, produced from fermented cassava roots, spoils quickly due to the germination and growth of Aspergillus. Effects of sodium benzoate on survival, growth, and aflatoxin production in packaged garri have been reported (Ogiehor and Ikenebomeh, 2004). Three Aspergillus species were inoculated into garri with or without benzoate and stored at ambient temperature. Decreased growth and no aflatoxin production occurred in benzoate-treated samples at the end of storage. Neosartorya fischeri is one of the most frequently isolated heat-resistant fungi causing spoilage of fruit juices and other heat-processed fruit-based products (Nielsen et al., 1989). Growth of this fungus was accompanied by production of fumitremorgin mycotoxins. Fungal growth was reduced by lowering the pH of laboratory media from 7.0 to 2.5; selected organic acids promoted growth and toxin production when added to the media. Small amounts (75 mg/L) of potassium sorbate or sodium benzoate completely inhibited the germination of ascospores and subsequent outgrowth. Both fungistatic and fungicidal properties have been attributed to benzoic acid, according to the results of a study involving several strains of Trichophyton and Microsporum (Pelayo, 1979). Sodium benzoate has been suggested as an inhibitor of cellulose-decomposing bacteria and fungi (Sauer, 1977). Under appropriate conditions, bacteriostatic and bactericidal properties of benzoic acid can also be demonstrated. Beuchat (1980) reported that sodium benzoate (300 µg/ml) inhibited the growth of Vibrio parahaemolyticus in laboratory media and enhanced the rate of thermal inactivation of this organism at slightly higher concentrations. Ten generally regarded as safe (GRAS) substances, including benzoic acid, were tested against both the opaque and translucent morphotypes of Vibrio vulnificus (Sun and Oliver, 1995). Eight of these had a lethal effect on both morphotypes of this bacterium.

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TABLE 3.5 Antimicrobial Spectrum of Benzoic Acid against Selected Bacteria, Yeasts, and Fungi Microorganisms Bacteria Bacillus cereus Escherichia coli Escherichia coli Klebsiella pneumoniae Lactobacillus sp. Listeria monocytogenes Micrococcus sp. Pseudomonas sp. Pseudomonas aeruginosa Staphylococcus aureus Streptococcus sp. Yeasts Sporogenic yeasts Asporogenic yeasts Candida krusei Debaryomyces hansenii Hansenula sp. Hansenula subpelliculosa Oospora lactis Pichia membranefaciens P. pastori Rhodotorula sp. Saccharomyces bayanus S. cerevisiae Torulopsis sp. Zygosaccharomyces bailii Z. lentus Z. rouxii Fungi Alternaria solani Aspergillus sp. Aspergillus parasiticus Aspergillus niger Byssochlamys nivea Chaetomonium globosum Cladosporium herbarum Mucor racemosus Penicillium sp. Penicillium citrinum Penicillium glaucum Rhizopus nigricans

pH

MIC*

6.3 5.2–5.6 6.0 6.0 4.3–6.0 5.6 (21°C) 5.6 (4°C) 5.5–5.6 6.0 6.0 6.0 5.2–5.6

500 50–120 100–200 100–200 300–1800 3000 2000 50–100 200–480 200–500 50–100 200–400

2.6–4.5 4.0–5.0

4.8 4.0 4.0 4.8

20–200 70–150 300–700 500 180 200–300 300 700 300 100–200 330 600 200–500 4500 1200 500–1100 1000

5.0 3.0–5.0 5.5 5.0 3.3 5.0 5.1 5.0 2.6--5.0 5.0 5.0 5.0

1500 20–300 >4000 2000 500 1000 100 30–120 30–280 2000 400–500 30–120

4.8 4.0

4.0 4.0

* Minimum inhibitory concentration in µg/ml (ppm). Source: Adapted from Chipley (1983, 1993, 2005); Davidson and Juneja (1990); Russell (1991); and Steels et al. (1999).

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Sodium Benzoate and Benzoic Acid TABLE 3.6 Minimum Inhibitory Concentration of Benzoic Acid for Food Spoilage Yeasts Isolate* Kluveromyces fragilis Kloeckera apiculata Pichia ohmeri Hansenula anomala Saccharomyces cerevisiae Zygosaccharomyces rouxii Zygosaccharomyces bisporus Candida krusei Saccharomycodes ludwigii Schizosaccharomyces pombe Zygosaccharomyces bailii

MIC (ppm) 173 188 200 223 170–450 242–330 200–350 440 500–600 500–567 600–1300

* A total of 23 isolates were tested. Most were isolated from spoiled foods that had contained preservative. Isolates were grown at 25°C in yeast extract medium containing 5% glucose (pH 3.5) without addition of benzoic acid. Source: Adapted from Warth (1989c).

In a series of studies involving L. monocytogenes (El-Shenawy and Marth, 1988; Yousef et al., 1989), it was found that benzoic acid at concentrations of approximately 1000 to 3000 ppm had strong bacteriostatic, but relatively modest bactericidal, activities against cells in a liquid minimal medium. Incubation of cells in minimal media caused injury that depended on the temperature of incubation but not on the presence of benzoic acid. The authors questioned the suitability of benzoic acid alone to control this pathogen in foods. This organism was isolated from milk, and its survival and growth were determined in media supplemented with organic acids and sodium benzoate (El-Shenawy and Marth, 1989). In general, inactivation or inhibition of growth occurred in inoculated media when a lower incubation temperature (13°C) and pH (5.0) were used and required lower levels of sodium benzoate. Combining glycerol monolaurate with benzoic acid gave greater inhibition of L. monocytogenes than when each preservative was tested alone (Oh and Marshall, 1994). The effects of temperature, pH, sodium chloride, and three preservatives on the growth of three foodborne bacterial pathogens were studied using gradient gel plates (Thomas et al., 1993). Potassium sorbate was completely effective against Verocytotoxigenic E. coli at all temperature/pH/NaCl combinations. It was also the most effective against B1 cereus. At C8) and Monoesters

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Skřivanová, E., Pražáková, Š., Benada, O., Hovorková, P. and Marounek, M. 2014. Susceptibility of Escherichia coli and Clostridium perfringens to sucrose monoesters of capric and lauric acid. Czech J. Anim. Sci. 59:374–380. Smith, J.L. and Palumbo, S.A. 1980. Inhibition of aerobic and anaerobic growth of Staphylococcus aureus in a model sausage system. J. Food Safety 2:221–233. Stecchini, M.L., Di Luch, R., Bortolussi, G. and Del Torre, M. 1996. Evaluation of lactic acid and monolaurin to control Listeria monocytogenes on Stracchino cheese. Food Microbiol. 13:483–488. Stillmunkes, A.A., Prabhu, G.A., Sebranek, J.G. and Molins, R.A. 1993. Microbiological safety of cooked beef roasts treated with lactate, monolaurin or gluconate. J. Food Sci. 58:953–958. Takano, M., Simbol, A.B., Yasin, M. and Shibasaki, I. 1979. Bactericidal effect of freezing with chemical agents. J. Food Sci. 44:112–115. Tetsumoto, S. 1933a. Sterilizing action of acids. II: Sterilizing action of saturated monobasic fatty acids. J. Agric. Chem. Soc. Jpn. 9:388–397. Tetsumoto, S. 1933b. Sterilizing action of acids. III: Sterilizing action of saturated monobasic fatty acids. J. Agric. Chem. Soc. Jpn. 9:563–567. Thomas, L.V., Davies, E.A., Delves-Broughton, J. and Wimpenny, J.W.T. 1998. Synergist effect of sucrose fatty acid esters on nisin inhibition of Gram-positive bacteria. J. Appl. Microbiol. 85:1013–1022. Thormar, H., Isaacs, E.C., Brown, H.R., Barshatzky, M.R. and Pessolano, T. 1987. Inactivation of enveloped viruses and killing of cells by fatty acids and monoglycerides. Antimicrob. Agents Chemother. 31:27–31. Thornton, R.H. 1963. Antifungal activity of fatty acids to Pithomyces chartatum and other fungi. N.Z. J. Agric. Res. 6:469–483. Tokarskyy, O. and Marshall, D.L. 2008. Mechanism of synergistic inhibition of Listeria monocytogenes growth by lactic acid, monolaurin, and nisin. Appl. Environ. Microbiol. 74:7126–7129. Trotter, T.M. and Marshall, D.L. 2003. Influence of pH and NaCl on monolaurin inactivation of Streptococcus iniae. Food Microbiol. 20:187–192. Tsuchido, T. and Takano, M. 1988. Sensitization by heat treatment of Escherichia coli K-12 cells to hydrophobic antibacterial compounds. Antimicrob. Agents Chemother. 32:1680–1683. Tsuchido, T., Saeki, T. and Shibasaki, I. 1981. Death kinetics of Escherichia coli in a combined treatment of heat and monolaurin. J. Food Safety 3:57–68. USDA FSIS. 2019. FSIS Directive. Safe and suitable ingredients used in the production of meat, poultry, and egg products. 7120.1 Revision 15. USDA Food Safety and Inspection Service, Washington, DC. Unda, J.R., Molins, R.A. and Walker, H.W. 1991. Clostridium sporogenes and Listeria monocytogenes: Survival and inhibition in microwave-ready beef roasts containing selected antimicrobials. J. Food Sci. 56:198–205, 219. Vasseur, C., Rigaud, N., Hebraud, M. and Labadie, J. 2001. Combined effects of NaCl, NaOH, and biocides (monolaurin or lauric acid) on inactivation of Listeria monocytogenes and Pseudomonas spp. J. Food Prot. 64:1442–1445. Verhaegh, E.G.A., Marshall, D.L. and Oh, D.H. 1996. Effect of monolaurin and lactic acid on Listeria monocytogenes attached to catfish fillets. Intl. J. Food Microbiol. 29:403–410. Venkitanarayanan, K.S., Zhao, T. and Doyle, M.P. 1999. Inactivation of Escherichia coli O157:H7 by combinations of GRAS chemicals and temperatures. Food Microbiol. 16:75–82. Walker, J.E. 1924. Germicidal properties of chemically pure soaps. J. Infect. Dis. 35:557–566. Wang, L.L. and Johnson, E.A. 1992. Inhibition of Listeria monocytogenes by fatty acids and monoglycerides. Appl. Environ. Microbiol. 58:624–629. Wang, L.L. and Johnson, E.A. 1997. Control of Listeria monocytogenes by monoglycerides in foods. J. Food Prot. 60:131–138. Welsh, J.K., Arsenakis, M., Coelen, R.J. and May, J.T. 1979. Effect of antiviral lipids, heat, freezing on the activity of viruses in human milk. J. Infect. Dis. 140:322–328. Winslow, C.E.A. and Lochridge, E.E. 1906. The toxic effect of certain acids upon typhoid and colon bacilli in relation to the degree of their dissociation. J. Infect. Dis 3:547–571. Woolford, M.K. 1975. Microbiological screening of the straight chain fatty acids (C1-C12) as potential silage additives. J. Sci. Food Agric. 28:219. Wyss, O., Ludwig, B.J. and Joiner, R.R. 1945. Fungistatic and fungicidal action of fatty acids and related compounds. Arch. Biochem. 7:415–425.

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Yamazaki, K., Yamamoto, T., Kawai, Y. and Inoue, N. 2004. Enhancement of antilisterial activity of essential oil constituents by nisin and diglycerol fatty acid ester. Food Microbiol. 21:283–289. Yang, C.-M. Luedecke, L.O., Swanson, B.G. and Davidson, P.M. 2003. Inhibition of microorganisms in salad dressing by sucrose and methylglucose fatty acid monoesters. J. Food Proc. Preserv. 27:285–298. Yuda, M., Matsuura, H., Takeuchi, A. and Iwasaki, T. 1977. Preservatives for starch-containing foods. Japan Kokai JP 52/21317 (77/21317), February 17, 1977. Zhang, H., Wei, H., Cui, Y. Zhao, G. and Feng, F. 2009. Antibacterial interactions of monolaurin with commonly used antimicrobials and food components. J. Food Sci. 74:M418–M421. Zhang, S., Xiong, J., Lou, W., Ning, Z., Zhang, D. and Yang, J. 2019. Antimicrobial activity and action mechanism of triglycerol monolaurate on common foodborne pathogens. Food Control 98:113–119.

12 Parabens P. Michael Davidson CONTENTS 12.1 Chemical and Physical Properties............................................................................................... 405 12.2 Antimicrobial Activity................................................................................................................ 405 12.2.1 Bacteria......................................................................................................................... 407 12.2.2 Fungi............................................................................................................................. 409 12.3 Mechanism of Action...................................................................................................................411 12.4 Applications..................................................................................................................................412 12.5 Regulatory Status.........................................................................................................................413 12.6 Toxicology....................................................................................................................................414 12.7 Assay............................................................................................................................................415 References................................................................................................................................................415 The methyl and propyl esters of p-hydroxybenzoic acid, are allowed to be directly added to foods as antimicrobials in many countries. The following sections will review various characteristics of these compounds along with the ethyl, butyl, and heptyl esters which are approved for use in foods by selected countries.

12.1 Chemical and Physical Properties Parabens have the general structure shown in Figure 12.1. The molecular weights of the various esters are as follows: methyl (Chemical Abstracts Service (C.A.S.) 99-76-3), 152.15; ethyl (C.A.S. 120-47-8), 166.18; propyl (C.A.S. 94-13-3), 180.21; butyl (C.A.S. 94-26-8), 194.23; and heptyl, 236.21. Solubility data for these compounds are shown in Table 12.1. Water solubility is inversely related to alkyl chain length. Parabens are stable in air and are resistant to cold and heat, including steam sterilization. Aalto et al. (1953) detected no hydrolysis in solutions of parabens buffered to pH 3.0 and 6.0 and heated at 120°C for 30 min. At pH 8.0, 6% hydrolysis was detected under the same conditions. Solutions of the parabens buffered to pH 3.0, 6.0, and 8.0 remained unchanged during storage at 25°C for 6 weeks (Aalto et al., 1953). The compounds are all odorless, except for methyl paraben, which reportedly has a faint characteristic odor (Aalto et al., 1953, Chichester and Tanner, 1972).

12.2 Antimicrobial Activity The first reports on the antimicrobial activity of the parabens came from Sabalitschka and co-workers in the early 1920s (Prindle, 1983). Esterification of the carboxyl group of benzoic acid allows the molecule to remain undissociated up to pH 8.5 versus the pKa of benzoic acid which is 4.20. Thus, while the pH optimum for antimicrobial activity of benzoic acid is 2.5–4.0, parabens are effective at pH 3–8 (Aalto et al., 1953; Chichester and Tanner, 1972). Propyl paraben has been demonstrated to be two to eight times 405

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FIGURE 12.1  Structure of the methyl and propyl esters of p-hydroxybenzoic acid.

TABLE 12.1 Solubility of the Parabens in Various Solvents Solubility (g/100 g) Solvent Water

Ethanol

Propylene glycol

Olive oil Peanut oil

Temperaturea

Methyl

Ethyl

Propyl

Butyl

Heptyl

25°C 10°C 80°C 25°C 50% (25°C) 10% (25°C) 25°C 50% (25°C) 10% (25°C) 25°C 25°C

0.25 0.20 2.0 52.0 18.0 0.5 22.0 2.7 0.3 2.9 0.5

0.17 0.07 0.86 70.0 25.0 3.0 1.0

0.05 0.025 0.30 95.0 18.0 0.1 26.0 0.9 0.06 5.2 1.4

0.02 0.005 0.15 210.0 110.0 9.9 5.0

1.5 mg -b -

Solubilities were determined at the temperatures specified; all solvents were pure, except where percentages are shown, which indicates percentage in water. b Not reported. Sources: Aalto et al. (1953); Lück and Jager (1997). a

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more effective at inhibiting bacterial growth than sodium benzoate or sorbate at pH 6.8–7.0 (Aalto et al., 1953; Jurd et al., 1971). Additionally, Eklund (1985a) showed that, while the antimicrobial activity of parabens was related to pH, this dependence was not related to dissociation of the compounds.

12.2.1 Bacteria The antimicrobial activity of parabens has been evaluated against a wide variety of Gram-negative and Gram-positive food-related bacteria (Table 12.2). It should be noted that many inhibition studies were done with different strains of bacteria, incubation conditions (pH, time, temperature), media, assay techniques, and data analyses. Because of these differences, it is difficult to compare results of different studies, except in relative terms. From the results of the minimum inhibitory concentration studies shown in Table 12.2, it can be seen that, as the alkyl chain length of the parabens increases, inhibitory activity generally increases. Increasing TABLE 12.2 Concentration Ranges of Esters of p-Hydroxybenzoic Acid Necessary for Total Growth Inhibitiona of Growth of Various Bacteria (pH, Incubation Temperature, and Time Vary) Concentration (μg/ml) Microorganism Gram-positive Bacillus cereus Bacillus megaterium Bacillus subtilis Clostridium botulinum Clostridium perfringens Lactococcus lactis Listeria monocytogenes Micrococcus sp. Micrococcus (Sarcina) lutea Staphylococcus aureus Streptococcus faecalis Gram-negative Aeromonas hydrophila Enterobacter aerogenes Escherichia coli Klebsiella pneumoniae Pseudomonas aeruginosa Pseudomonas fluorescens Pseudomonas fragi Pseudomonas putida Pseudomonas stutzeri Salmonella Salmonella Typhimurium Vibrio parahaemolyticus Yersinia enterocolitica

Methyl

Ethyl

Propyl

Butyl

Heptyl

1000–2000 1000 1980–2130 1000–1200 500 (3:1 methyl and propyl) 1430–1600 4000 1670–4000 -

830–1000 1000–1330 800–1000

125–400 320 250–450 200–400

63–400 100 63–115 200

12 -

60–110 1000 1000–2500 130

400 512 10–100 400–500 350–540 40

125 120–200 -

12 12 12 -

550 2000 1200–2000 1000 4000 1310 450 500–750 2000 350

1000 1000–2000 500 1000–4000 400–500 1000 -

100 1000 400–1000 250 8000 670 4000 250–300 1000 180–300 50–100 -

4000 1000 125 8000 100 1000 -

-

-

-

Sources: Aalto et al. (1953), Bargiota et al. (1987), Dymicky and Huhtanen (1979), Eklund et al. (1981), Eklund (1985a), Jurd et al. (1971), Juneja and Davidson (1993), Kato and Shibasaki (1975), Klindworth et al. (1979), Lee (1973), Lewis and Jurd (1972), Lück and Jager (1997), Moir and Eyles (1992), Moustafa and Collins (1969), Payne et al. (1989), Pierson et al. (1980), Reddy et al. (1982), Robach and Pierson (1978), Sokol (1952), Tattawasart et al. (1999), Venugopal et al. (1984). a As defined by the authors of the references cited.

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activity with decreasing polarity is more evident against Gram-positive than against Gram-negative bacteria. Gram-positive bacteria are generally more susceptible to non-polar phenolic compounds than Gram-negative bacteria. Both Eklund (1980) and Freese et al. (1973) stated that Gram-negative bacteria were, most likely, resistant to the parabens owing to a screening effect by the cell wall lipopolysaccharide layer. Fukahori et al. (1996) studied the relationship between uptake and the antimicrobial activity of the methyl, ethyl, propyl, and butyl esters of p-hydroxybenzoic acid using Escherichia coli. They reported that the uptake of the parabens was logarithmically proportional to the alkyl chain length from methyl to butyl. However, free energy change measurements indicated that the transfer of the compounds also involved hydrophilic interactions. In addition, they found that the concentration of parabens necessary for antimicrobial activity decreased in logarithmic relationship to alkyl chain length. They concluded that the antibacterial activity of parabens is dependent upon alkyl chain length both for uptake and for concentration at the cellular target. In addition to total inhibition achieved in studies shown in Table 12.2, others have reported variable results with partial or no inhibition. For example, Martin et al. (1972) found no inhibition of Alcaligenes viscolactis in skim milk with up to 600 μg/ml propyl paraben. Moustafa and Collins (1969) showed that, while 4000 μg/ml propyl paraben inhibited the growth of Pseudomonas fragi, 2000 μg/ml actually stimulated growth. At 200 μg/ml, propyl paraben inhibited protease secretion by Aeromonas hydrophila (Venugopal et al., 1984). Ahmedy et al. (1999) found no difference in susceptibility of pathogenic and non-pathogenic strains of Yersinia to methyl paraben despite the fact that the same strains showed differences in resistance to certain antibiotics and cationic biocides. Darwish and Bloomfield (1997) evaluated the effect of co-solvents ethanol, propylene glycol, and glycerol on the activity of methyl and propyl parabens against Staphylococcus aureus and Pseudomonas aeruginosa. Interestingly, the antimicrobial activity of the parabens increased with increasing hydrophobicity of the co-solvent, being greatest with the most hydrophobic co-solvent, ethanol. In contrast, the uptake of parabens was apparently determined by the hydrophilicity of the co-solvent, with glycerol causing the greatest uptake by the cells. Uptake however, was not correlated with inhibition. It was concluded that inhibition was a combination of the action of the solvent and the paraben on the integrity of the outer (P. aeruginosa) and cytoplasmic (both genera) membranes. Moir and Eyles (1992) compared the effectiveness of methyl paraben and potassium sorbate on the growth of four psychrotrophic foodborne bacteria, A. hydrophila, Listeria monocytogenes, Pseudomonas putida, and Yersinia enterocolitica. At pH 5, little difference was found between minimum inhibitory concentrations (MIC) of methyl paraben and potassium sorbate at 5°C or 30°C. At pH 6 however, methyl paraben was effective at a lower concentration than potassium sorbate for all pathogens except A. hydrophila, where the two were equal. Little or no adaptation was found to occur when cells were exposed to sub-inhibitory concentrations of antimicrobials. At 5°C in the presence of 1000 μg/ml methyl paraben, A. hydrophila survived for 1–2 days, P. putida and Y. enterocolitica for 1–2 weeks, and L. monocytogenes for over 4 months. Injury occurred with A. hydrophila and L. monocytogenes but was variable for Y. enterocolitica and did not occur for P. putida. Razavilar and Genigeorgis (1998) studied the influence of temperature, time, and inoculum on the efficacy of methyl paraben against growth of Listeria monocytogenes, L. innocua, L. ivanovii, and L. seeligeri. Methyl paraben at 1000 μg/ml allowed growth of all species of Listeria at pH 6.0–6.2 in brain heart infusion (BHI) broth at 20 and 30°C. In contrast, no growth occurred with any species at 4 or 8°C and 1000 μg/ml methyl paraben. At 1500 μg/ ml and 20°C, methyl paraben had inhibitory effects (e.g., increased lag time, decreased final growth level) on the Listeria species but did not completely inhibit growth of any species except L. ivanovii. Fyfe et al. (1998) evaluated the antimicrobial activity of 1000 μg/ml methyl paraben or benzoic acid with plant oil extracts (fennel, anise, or basil) against Listeria monocytogenes and Salmonella Enteritidis. Methyl paraben, by itself, did not inhibit the growth of either microorganism under the test conditions. However, when combined with 0.2% anise, fennel, or basil oil, L. monocytogenes was reduced by 5.1, 5.7, and >8 logs compared to the control after 24 h, respectively. The combinations were even more effective against Salmonella Enteritidis, reducing viable cells by >8 logs compared to the control with all combinations at 24 and 48 h. All combinations containing methyl paraben were more inhibitory than those containing benzoic acid. This was not surprising since the tests were done in a microbiological medium with a pH around 7.0. Er et al. (2014) investigated the inhibitory concentrations of methyl and propyl

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409

parabens, among others, against Salmonella spp. isolated from chicken as both planktonic and biofilm cells. The MIC (as defined as no growth in 24 h) for the planktonic cells ranged from 60 to 70 μg/ml for methyl and 50–120 for propyl. Biofilm cells were significantly more resistant than the planktonic cells, requiring up to around 2000 μg/ml for methyl and propyl (although one strain required ca. 8000 μg/ml for propyl paraben at pH 6.0). They demonstrated the reduced susceptibility of Salmonella in biofilms. Propyl paraben was evaluated against Listeria monocytogenes Scott A in 10% chicken and hot dog suspensions (Dje et al., 1989). The effectiveness of propyl paraben was dependent upon the product. In chicken, L. monocytogenes was inhibited 99.9% compared to the control after 24 h incubation at 35°C. In contrast, little inhibition of L. monocytogenes growth was demonstrated by propyl paraben in the hot dog suspension. The difference in effectiveness was theorized to be due to a higher fat content in the hot dogs. Dje et al. (1990) evaluated the effect of 1000 μg/ml propyl paraben and 1000 μg/ml methyl:propyl paraben in salt brine (13%) on L. monocytogenes suspended in the brine or inoculated on hot dogs which were dipped in brine. Propyl paraben alone had little or no effect on L. monocytogenes survival in salt brine at 4°C or on the surface of hot dogs which were dipped in brine for 5 min and incubated at 24°C. In contrast, 1000 μg/ml methyl:propyl paraben caused a 2–3 log decrease in viable L. monocytogenes in the salt brine at 4°C. On hot dogs, the antimicrobial combination was less effective, delaying growth for around 4 h at 24°C of one of two strains tested. In a similar study by Blom et al. (1997), propyl paraben added to vacuum-packaged sliced ham or sausage inoculated with Listeria monocytogenes and stored at 4 or 9°C for 5 weeks was ineffective in controlling the microorganism. The efficacy of a blend of benzalkonium chloride, acetic acid, and methyl paraben was evaluated as a wash to reduce Listeria monocytogenes, Salmonella, and Escherichia coli O157:H7 on fresh-cut romaine and iceberg lettuce (Sevart et al., 2017). While up to a 3% solution reduced the numbers of test microorganisms in wash water, it did not significantly reduce cells on the lettuce itself. Robach and Pierson (1978) investigated the effect of methyl and propyl paraben on toxin production of Clostridium botulinum NCTC 2021. At 100 μg/ml of methyl and 100 μg/ml of propyl paraben, toxin formation was prevented while 1200 μg/ml methyl and 200 μg/ml propyl were necessary for growth inhibition. Reddy and Pierson (1982) and Reddy et al. (1982) determined the effect of methyl, ethyl, propyl, and butyl parabens on the growth and toxin production of ten Clostridium botulinum strains (5 Type A, 5 Type B). In TYG medium at pH 7.0 and 37°C, 1000 μg/ml methyl paraben blocked growth and toxin formation for only 1 day. Ethyl and propyl paraben, at the same concentration, prevented growth and toxin production for the maximum incubation time of 7 d. Butyl paraben, as might be expected, was most effective and prevented growth and toxin production for 7 d at 200 μg/ml. Reddy and Pierson (1982) also evaluated ethyl, propyl, and butyl parabens in TYG medium in the presence of 0.05 M phosphate buffer at pH 7.0 and 6.0. Ethyl paraben prevented growth and toxin production by C. botulinum at 37°C for 7 d at 1000 μg/ml (pH 7.0) and 800 μg/ml (pH 6.0). Propyl paraben at 800 μg/ml and 400 μg/ml, and butyl paraben at 200 μg/ml and 100 μg/ml, were similarly effective against C. botulinum at pH 7.0 and 6.0, respectively. Draughon et al. (1982) demonstrated effective growth inhibition of C. botulinum by 1000 μg/ml of all esters of the parabens. Ethyl paraben at 1000 μg/ml was also effective in inhibiting toxin formation in canned comminuted pork. However, inhibition of C. botulinum by the parabens in many food systems has been reported to be much less than in laboratory media (Sofos and Busta, 1980). Little research on the activity of the n-heptyl ester in foods has been published. Chan et al. (1975) did show that this compound was very effective in inhibiting bacteria involved in the malolactic fermentation of wines.

12.2.2 Fungi The antifungal effectiveness of the parabens has been evaluated against several food-related fungi (Table 12.3). In comparison to bacteria, fungi are much more susceptible to parabens. As with bacteria, inhibition of fungi increases as the alkyl chain length of the parabens increases. Thompson (1994) evaluated butyl, propyl, ethyl, and methyl parabens, alone and in combination, against strains of mycotoxigenic Aspergillus, Penicillium, and Fusarium. The most effective parabens were the propyl and butyl esters with minimum inhibitory concentrations of 1.0–2.0 mM in potato dextrose agar. Combinations of the various parabens were reported to have synergistic activity against the

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TABLE 12.3 Concentration Ranges of Esters of p-Hydroxybenzoic Acid Necessary for Total Growth Inhibitiona of Various Fungi (pH, Incubation Temperature, and Time Vary) Concentration (μg/ml) Fungi Alternaria sp. Aspergillus flavus Aspergillus niger Byssochlamys fulva Candida albicans Debaryomyces hansenii Penicillium digitatum Penicillium chrysogenum Rhizopus nigricans Saccharomyces bayanus Saccharomyces cerevisiae Torula utilis Torulaspora delbrueckii Zygosaccharomyces bailii Zygosaccharomyces bisporus Zygosaccharomyces rouxii

Methyl

Ethyl

Propyl

Butyl

Heptyl

1000 1000 500 500 500 930 1000 -

400–500 500–1000 400 250 250 250 500 700 900 400 700

100 200 200–250 200 125–250 63 125–200 125 220 125–200 200 -

125–200 125 8000 mg/kg body weight. The sodium salts of methyl, ethyl, propyl, and butyl paraben elicited oral LD50 values of 2000, 2500, 3700, and 950 mg/kg, respectively. The chief toxic effect in dogs was acute myocardial depression with hypotension, but the effects were transient and non-cumulative. In subchronic testing, 500 mg/kg of methyl paraben caused no ill effects to rabbits over a 6-day period, while 3000 mg was toxic to the animals (Lück and Jager, 1997). Lück and Jager (1997) also reported that propyl paraben showed similar subchronic toxicity responses. Prindle (1983) reported that feeding 2–20 mg/kg per day of the “lower” esters of parabens to rabbits, guinea pigs, or rats caused no harmful effects after 120 days. Rats fed up to 60 mg/kg per day for 30 days also showed no effects. For chronic toxicity testing, white rats were fed diets containing 2% (0.9–1.2 g/kg per day) and 8% (5.5–5.9 g/kg per day) each of methyl and propyl paraben (Matthews et al., 1956). After 96 weeks, the animals at the 2% level had no depressed weight gains or any histological changes in internal organs. At 8% of the diet however, a mild growth retardation was observed. The same researchers found that mongrel dogs could tolerate daily doses of 1 g/kg of the methyl and propyl esters for 1 year with no ill effects. Tissue samples from these animals were normal. Matthews et al. (1956) reported that none of the esters produced skin irritation in humans at concentrations of 5%. Epstein (1968) however, reported that parabens in foods were associated with dermatitis. Contact dermatitis involving parabens has been reported, although it is associated with topical use (Reitschel and Fowler, 2001). The concentrations required to elicit such a response are generally high and no mechanism is known for the sensitivity (Soni et al., 2001). Similarly, allergic-type reactions have been reported with parabens but evidence of allergenicity of the compounds is lacking (Soni et al., 2001).

Parabens

415

In the early 2000s, the parabens family of preservatives came under intense public scrutiny because of concerns over their potential estrogenicity or endocrine disruption capabilities and due to possible links of paraben-containing cosmetics to breast cancer (Darbre et al., 2004). While this issue attracted a great deal of public and global media attention, no objective evidence has demonstrated any significant risk from current cosmetic or food usage of parabens, and various scientific and regulatory bodies have consistently reported that parabens do not cause cancer (CIR, 2008; US FDA, 2018). Like many other food ingredients, parabens have safe dietary intake amounts that have been established by scientific and government regulatory agencies (U.S. Code of Federal Regulations, 2019; 21 CFR 184.1490, 21 CFR 184.1670). Bio-monitoring studies on levels of parabens found in human urine reflect the cumulative exposures across all product types, including foods, cosmetics, and drugs (Calafat et al., 2010). These monitoring studies on levels of parabens provide physicians and public health officials with reference values that facilitate ongoing research on exposure and health effects. While parabens have been proven safe over decades of research and scrutiny, consumers have shown a growing trend and desire for products that are “natural,” “naturally derived,” or “nature-inspired.” This trend has provoked a strong push by the foods and cosmetics industries to search for natural ingredients that have similar antimicrobial efficacy and can ultimately replace traditional preservatives like parabens. The irony behind this public perception driving the consumer trend away from traditional preservatives is that PHBA is the building block of parabens and is found in nature as a truly natural preservative (Baardseth and Russwurm, 1978).

12.7 Assay Several methods are available for the qualitative or quantitative determination of the parabens. According to the Joint FAO/WHO Expert Committee on Food Additives (2000), 2 g of a dried sample to be assayed for parabens should be weighed to the nearest mg and transferred to a flask. Forty milliliters of 1 N sodium hydroxide are added and the sides of the flask rinsed with water. The flask is covered with a watch glass, and the solution boiled gently for 1 h and cooled. Five drops of bromothymol blue TS are added and the mixture titrated with 1 N sulfuric acid, comparing the color with a buffer solution (pH 6.5) containing the same proportion of indicator. Blank determinations with the reagents are performed to make any necessary corrections. Each ml of 1 N sodium hydroxide is equivalent to 152.2 mg of (methyl) C8H8O3, 166.18 mg of (ethyl) C9H10O3, or 180.2 mg of (propyl) C10H12O3. With the increased interest in the toxicological aspects of parabens has come with a plethora of newly developed methods for analysis. These involve more sophisticated extraction procedures and automated analytical techniques. In an article describing the development of a method for screening of foods using high-performance liquid chromatography (HPLC) and photodiode array detection, Maher et al. (2018) reviewed some of the methods that had been developed to analyze for parabens in foods. Those methods include gas chromatographic analysis following derivatization flame ionization detector (Jain et al., 2013), capillary electrophoresis (Alshana et al., 2015; Bottoli et al., 2011), HPLC with UV detection (Saad et al., 2005; Yang et al., 2014) and mass spectrometry detection (Cao et al., 2013; Yin et al., 2018; Molognoni et al., 2018; Song et al., 2017), and ultraperformance liquid chromatography with an electrochemical detector (Chuto et al., 2013), among others.

REFERENCES Aalto, R.R., Firman, M.C., and Rigler, N.E. 1953. p-Hydroxybenzoic acid esters as preservatives. I. Uses, antibacterial and antifungal studies, properties and determination. J. Am. Pharm. Assoc. 42:449. Ahmedy, A.M., Hussein, M.M.M., El-Falaha, B.M.A., and Megid, R.M.A. 1999. Antimicrobial susceptibility and surface hydrophobicity of Yersinia species in relation to genetic elements. Al-Azhar J. Microbiol. 45:125. Alshana, U., Ertaş, N., and Göğer, N.G. 2015. Determination of parabens in human milk and other food samples by capillary electrophoresis after dispersive liquid–liquid microextraction with back-extraction. Food Chem 181:1–8.

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Anonymous. 2019. Eur-Lex Access to European Union Law. https​:/​/eu​​r​-lex​​.euro​​pa​.eu​​/eli/​​reg​/2​​0 08​/1​​333​​/2​​019​ -0​​6 ​-18 Baardseth, P., and Russwurm, H. 1978. Content of some organic acids in cloudberry (Rubus chamaemorus). Food Chem. 3:43. Barberis, C.L., Astoreca, A.L., Dalcero, A.M., and Magnoli, C.E. 2010. Food-grade antioxidants and antimicrobials to control growth and ochratoxin a production by Aspergillus Section Nigri on peanut kernels. J. Food Prot. 73:1493–1501. Bargiota, E.E., Rico-Munoz, E., and Davidson, P.M. 1987. Lethal effect of methyl and propyl parabens as related to Staphylococcus aureus lipid composition. Intl. J. Food Microbiol. 4:257. Blom, H., Nerbrink, E., Dainty, R., Hagtvedt, T., Borch, E., Nissen, H., and Nesbakken, T. 1997. Addition of 2.5% lactate and 0.25% acetate controls growth of Listeria monocytogenes in vacuum-packed, sensory acceptable servelat sausage and cooked ham stored at 4°C. Intl. J. Food Microbiol. 38:71–76. Bottoli, C.B.G., Gutiérrez-Ponce, M.D.S., Aguiar, V.S., and de Aquino W.M. 2011. Determination of parabens in sweeteners by capillary electrochromatography Braz. J. Pharm. Sci. 47: 779–785. Bredin, J., Davin-Regli, A., and Pages, J.-M. 2005. Propyl paraben induces potassium efflux in Escherichia coli. J. Antimicrob. Chemother. 55:1013–1015. Calafat, A.M., Ye, X. Wong, L-Y., Bishop, A.M. and Needham, L.L. 2010. Urinary concentrations of four parabens in the U.S. Population: NHANES 2005–2006. Environ. Health Perspectives 118:679–685. Cao, S., Liu, Z., Zhang, L., Xi, C., Li, X., Wang, G., Yuana, R., and Mu, Z. 2013. Development of an HPLC–MS/ MS method for the simultaneous analysis of six kinds of parabens. Food Anal. Methods 5:1016–1023. Chan, L., Weaver, R., and Ough, C.S. 1975. Microbial inhibition caused by p-hydroxybenzoate esters in wines. Am. J. Enol. Vitic. 26:201. Chichester, F., and Tanner, F.W. 1972. Antimicrobial food additives. In Handbook of Food Additives, 2nd Ed., edited by T.E. Furia. CRC Press, Cleveland, OH, p. 115. Chung, D., Papadakis, S.E., and Yam, K.L. 2001a. Release of propyl paraben from a polymer coating into water and food simulating solvents for antimicrobial packaging applications. J. Food Proc. Preserv. 25:71. Chung, D., Chikindas, M.L., and Yam, K.L. 2001b. Inhibition of Saccharomyces cerevisiae by slow release of propyl paraben from a polymer coating. J. Food Prot. 64:1420. Chuto, M., Chaiyo, S., Siangproh, W., and Chailapakul, O. 2013. Rapid separation and highly determination of paraben species by ultra-performance liquid chromatography—electrochemical detection. Detection 1:21–29. Code of Federal Regulations. 2019. Title 21: Food and Drugs, Parts 170–199. Off. Federal Regulations, National Archives Records Serv., General Service Admin., Washington, DC. Accessed at https​:/​/ww​​w​ .ecf​​r​.gov​​/cgi-​​bin​/t​​ext​-i​​dx​?gp​=​&SID​​= d8a3​​1c956​​01598​​0bae9​​6c405​​675b1​​226​&m​​c​=tru​​e​&tpl​=​/ecf​​rbrow​​​ se​/ Ti​​tle21​​/21ta​​b​_02.​​tpl Cosmetic Ingredient Review Expert Panel (CIR). 2008. Final amended report on the safety assessment of methylparaben, ethylparaben, propylparaben, isopropylparaben, butylparaben, isobutylparaben, and benzylparaben as used in cosmetic products. Intl. J. Toxicol. 27 (Suppl 4):1–82. Dan, V., Raducan, A., Stoicescu, A., and Nicolau, A. 1996. Effect of esterparabens upon biotechnological processes in brewing. Roum. Biotechnol. Lett. 1:181. Darbre, P.D., Aljarrah, A., Miller, W.R., Coldham, N.G., Sauer, M.J., and Pope, G.S. 2004. Concentrations of parabens in human breast tumours. J. Appl. Tox. 24:5. Darwish, R.M., and Bloomfield, S.F. 1997. Effect of ethanol, propylene glycol and glycerol on the interaction of methyl and propyl p-hydroxybenzoate with Staphylococcus aureus and Pseudomonas aeruginosa. Intl. J. Pharma. 147:51. Dje, Y., Payne, K.D., and Davidson, P.M. 1989. Inhibition of Listeria monocytogenes Scott A by propyl paraben in non-fat milk and chicken and hot dog suspensions. Ann. Mtg. Inst. Food Technol., Chicago, IL. Dje, Y., Davidson, P.M., and Rico, E. 1990. Effect of antimicrobials against Listeria monocytogenes on hot dogs and in brine. Ann. Mtg. Inst. Food Technol., Anaheim, CA. Dobias, J., Chudackova, K., Voldrich, M., and Marek, M. 2000. Properties of polyethylene films with incorporated benzoic anhydride and/or ethyl and propyl esters of 4-hydroxybenzoic acid and their suitability for food packaging. Food Addit. Contam. 17:1047. Draughon, F.A., Sung, S.C., Mount, J.R., and Davidson, P.M. 1982. Effect of the parabens with and without nitrite on Clostridium botulinum toxin production in canned pork slurry. J. Food Sci. 47:1635.

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Dymicky, M., and Huhtanen, C.N. 1979. Inhibition of Clostridium botulinum by p-hydroxybenzoic acid n-alkyl esters. Antimicrob. Agents Chemother. 15:798. Eklund, T. 1980. Inhibition of growth and uptake processes in bacteria by some chemical food preservatives. J. Appl. Bact. 48:423. Eklund, T., Nes, I.F., and Skjelkväle, R. 1981. Control of Salmonella at different temperatures by propyl paraben and butylated hydroxyanisole. In Psychrotrophic Microorganisms in Spoilage and Pathogenicity, edited by T.A. Roberts, G. Hobbs, J.H.B. Christian and N. Skovgaard. Academic Press, London, UK. Eklund, T. 1985a. Inhibition of microbial growth at different pH levels by benzoic and propionic acids and esters of p-hydroxybenzoic acid. Intl. J. Food Microbiol. 2:159. Eklund, T. 1985b. The effect of sorbic acid and esters of p-hydroxybenzoic acid on the protonmotive force in Escherichia coli membrane vesicles. J. Gen. Microbiol. 131:73. Epstein, S. 1968. Paraben sensitivity: Subtle trouble. Ann. Alergy 26:185. Er, B., Demirhan, B., Onurdag, F., Özgacar, S., and Öktem, A. 2014. Antimicrobial and antibiofilm effects of selected food preservatives against Salmonella spp. isolated from chicken samples. Poultry Sci. 93:695–701. Fagundes, C., Perez-Gago, M.B., Monteiro, A.R., and Palou, L. 2013. Antifungal activity of food additives in vitro and as ingredients of hydroxypropyl methylcellulose-lipid edible coatings against Botrytis cinerea and Alternaria alternata on cherry tomato fruit. Intl. J. Food Microbiol. 166:391–398. FAO/WHO. 2000. Joint FAO/WHO Expert Committee on Food Additives. FAO Food and Nutrition Papers 55th Meeting. Freese, E., Sheu, C.W., and Galliers, E. 1973. Function of lipophilic acids as antimicrobial food additives. Nature 241:321. Fukahori, M., Akatsu, S., Sato, H., and Yotsuyanagi, T. 1996. Relationship between uptake of p-hydroxybenzoic acid esters by Escherichia coli and antibacterial activity. Chem. & Pharm. Bull. 44:1567. Furr, J.R., and Russell, A.D. 1972. Some factors influencing the activity of esters of p-hydroxybenzoic acid against Serratia marcescens. Microbios 5:189. Fyfe, L., Armstrong, F., and Stewart, J. 1998. Inhibition of Listeria monocytogenes and Salmonella enteritidis by combinations of plant oils and derivatives of benzoic acid: The development of synergistic antimicrobial combinations. Intl. J. Antimicrob. Agents 9:195. Government of Canada. 2019. List of Permitted Preservatives (Lists of Permitted Food Additives), Accessed 5 May 2019 at https​:/​/ww​​w​.can​​ada​.c​​a ​/en/ ​​healt​​h​-can​​ada ​/s​​ervic​​es​/fo​​od​-nu​​triti​​on ​/fo​​od​-sa​​fety/​​food-​​addit​​ iv​es/ ​​lists​​-perm​​itted​​.html​ Jain, R., Mudiam, M.K.R., Chauhan, A., Ratnasekhar, Ch., Murthy, R.C., and Khan, H.A. 2013. Simultaneous derivatisation and preconcentration of parabens in food and other matrices by isobutyl chloroformate and dispersive liquid–liquid microextraction followed by gas chromatographic analysis. Food Chem. 141:436–443. Japan External Trade Association. 2011. Specifications and Standards for Foods, Food Additives, etc. Under the Food Sanitation Act (Abstract) 2010. Accessed at https​:/​/ww​​w​.jet​​ro​.go​​.jp​/e​​xt​_im​​ages/​​en​/re​​ports​​/ regu​​latio​​ns​/pd​​f​​/foo​​dext2​​010e.​​pdf Jermini, M.F.G., and Schmidt-Lorenz, W. 1987. Activity of Na-benzoate and ethyl-paraben against osmotolerant yeasts at different water activity values. J. Food Prot. 50:920–927. Jones, P.S., Thigpen, D., Morrison, J.L., and Richardson, A.P. 1956. p-Hydroxybenzoic acid esters as preservatives. III. The physiological disposition of p-hydroxybenzoic acid and its esters. J. Am. Pharm. Assoc. 45:268. Juneja, V.K., and P.M. Davidson. 1993. Influence of altered fatty acid composition on resistance of Listeria monocytogenes to antimicrobials. J. Food Prot. 56:302–305. Jurd, L., King, A.D., Mihara, K., and Stanely, W.L. 1971. Antimicrobial properties of natural phenols and related compounds. I. Obtusastyrene. Appl. Microbiol. 21:507. Kato, A., and Shibasaki, I. 1975. Combined effect of different drugs on the antibacterial activity of fatty acids and their esters. J. Antibact. Antifung. Agents (Japan) 8:355. Klindworth, K.J., Davidson, P.M., Brekke, C.J., and Branen, A.L. 1979. Inhibition of Clostridium perfringens by butylated hydroxyanisole. J. Food Sci. 44:564. Lee, J.S. 1973. What seafood processors should know about Vibrio parahaemolyticus. J. Milk Food Technol. 36:405.

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Lewis, J.C., and Jurd, L. 1972. Sporostatic action of cinnamylphenols and related compounds on Bacillus megaterium. In Spores V, edited by H.O. Halvorson, R. Hewson, and L.L. Campbell. Amer. Soc. Microbiol., Washington, DC. Liao, C. Liu, F., and Kurunthachalam, K. 2013. Occurrence of and dietary exposure to parabens in foodstuffs from the United States. Environ. Sci. & Technol. 47:3918–3925. Lück, E., and Jager, M. 1997. Antimicrobial Food Additives. Characteristics, Uses, Effects, 2nd Ed. SpringerVerlag, Berlin. Maher, H.M., Alzoman, N.Z., Almeshal, M.A., Alotaibi, H.A., Alotaibi, N.N., and Al-Showiman, H. 2018. Quantitative screening of parabens in ready-to-eat foodstuffs available in the Saudi market using high performance liquid chromatography with photodiode array detection. Arabian J. Chem. 13:2897–2911. https​:/​/do​​i​.org​​/10​.1​​016​/j​​.arab​​jc​.20​​​18​.07​​.019 Martin, J.H., Chung, K.S.S., and Ogrosky, L. 1972. Inhibition of growth of Alcaligenes viscolactis by some common food preservatives. J. Dairy Sci. 55:1179. Marwan, A.G., and Nagel, C.W. 1986. Quantitative determination of infinite inhibition concentrations of antimicrobial agents. Appl. Env. Microbiol. 51:559. Matthews, C., Davidson, J., Bauer, E., Morrison, J.L., and Richardson, A.P. 1956. p-Hydroxybenzoic acid esters as preservatives. II. Acute and chronic toxicity in dogs, rats, and mice. J. Am. Pharm. Assoc. 45:260. Moir, C.J., and Eyles, M.J. 1992. Inhibition, injury and inactivation of four psychrotrophic foodborne bacteria by the preservatives methyl p-hydroxybenzoate and potassium sorbate. J. Food Prot. 55:360. Molognoni, L., Daguer, H., de Sá Ploêncio, L.A., and De Dea Lindner, J. 2018. A multi-purpose tool for food inspection: Simultaneous determination of various classes of preservatives and biogenic amines in meat and fish products by LC-MS. Talanta 178:1053–1066. Moustafa, H.H., and Collins, E.B. 1969. Effects of selected food additives on growth of Pseudomonas fragi. J. Dairy Sci. 52:335. Murata, W., Yamaguchi, Y., Fujita, K.-I., Yamauchi, K., Tanaka, T., and Ogita, A. 2019. Enhancement of paraben-fungicidal activity by sulforaphane, a cruciferous vegetable-derived isothiocyanate, via membrane structural damage in Saccharomyces cerevisiae. Lett. Appl. Microbiol. 69​:doi​.org​/10​.1111​/ lam​​.13230 Nesci, A., Rodriguez, M., and Etcheverry, M. 2003. Control of Aspergillus growth and aflatoxin production using antioxidants at different conditions of water activity and pH. J. Appl. Microbiol. 95:279. Nguyen, T., Clare, B., Guo, W., and Martinac, B. 2005. The effects of parabens on the mechanosensitive channels of E. coli. Eur. Biophys. J. 34:389–395. Oka, S. 1960. Studies on the transfer of antiseptics to microbes and their toxic effect. Part III. Adsorption of esters of p-hydroxybenzoic acid on yeast cell and their toxic effect. Bull. Agric. Chem. Soc. Jpn. 24:412. Passone, M.A., Resnik, S.L., and Etcheverry, M. 2005. In vitro effect of phenolic antioxidants on germination, growth and aflatoxin B1 accumulation by peanut Aspergillus section Flavi. J. Appl. Microbiol. 99:682–691. Payne, K.D., Rico-Munoz, E., and Davidson, P.M. 1989. The antimicrobial activity of phenolic compounds against Listeria monocytogenes and their effectiveness in a model milk system. J. Food Prot. 52:151–153. Pierson, M.D., Smoot, L.A., and Vantassell, K.R. 1980. Inhibition of Salmonella typhimurium and Staphylococcus aureus by butylated hydroxyanisole and the propyl ester of p-hydroxybenzoic acid. J. Food Prot. 43:191. Prindle, R.F. 1983. Phenolic compounds. In Disinfection, Sterilization, and Preservation, 3rd Ed., edited by S.S. Block. Lea and Febiger, Philadelphia, p. 197. Razavilar, V., and Genigeorgis, C. 1998. Prediction of Listeria spp. growth as affected by various levels of chemicals, pH, temperature and storage time in a model broth system. Intl. J. Food Microbiol. 40:149. Reddy, N.R., and Pierson, M.D. 1982. Influence of pH and phosphate buffer on inhibition of Clostridium botulinum by antioxidants and related phenolic compounds. J. Food Prot. 45:925. Reddy, N.R., Pierson, M.D., and Lechowich, R.V. 1982. Inhibition of Clostridium botulinum by antioxidants, phenols and related compounds. Appl. Environ. Microbiol. 43:835. Reitschel, R.L., and Fowler, J.F. 2001. Fisher’s Contact Dermatitis, 5th Ed. Lippincott, Williams and Wilkins, Philadelphia. Rico-Munoz, E., Bargiota, E.E., and Davidson, P.M. 1987. Effect of selected phenolic compounds on the membrane-bound adenosine triphosphatase of Staphylococcus aureus. Food Microbiol. 4:239. Robach, M.C., and Pierson, M.D. 1978. Influence of para-hydroxybenzoic acid esters on the growth and toxin production of Clostridium botulinum 10755A. J. Food Sci. 43:787.

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Saad, B., Bari, M.F., Saleh, M.I., Ahmad, K., and Talib, M.K. 2005. Simultaneous determination of preservatives (benzoic acid, sorbic acid, methylparaben and propylparaben) in foodstuffs using high-performance liquid chromatography. J. Chromatogr. A 1073:393–397. Sethi, V. 1994. Efficacy of various preservatives for preserving whole tomato concentrate. Ind. Food Packer 48:11. Sevart, N., Gragg, S., Krug, M., Jenott, J., and Phebus, R. 2017. A Blend of benzalkonium chloride, acetic acid, and methyl paraben effectively reduces Escherichia coli O157:H7, Salmonella spp., and Listeria monocytogenes in produce wash water, but lacks consistent efficacy on romaine and iceberg lettuce. International Association of Food Protection P3-60, Tampa, FL. Sofos, J.N., and Busta, F.F. 1980. Alternatives to the use of nitrite as an antibotulinal agent. Food Technol. 34:244. Sokol, H. 1952. Recent developments in the preservation of pharmaceuticals. Drug Stand. 20:89. Song, S., Zhang, Z., Zou, N., Chen, R., Han, L., Pan, C., and Sapozhnikova, Y. 2017. Determination of six paraben residues in fresh-cut vegetables using QuEChERS with multi-walled carbon nanotubes and highperformance liquid chromatography–tandem mass spectrometry. Food Anal. Methods 10:3972–3979. Soni, M.G., Burdock, G.A., Taylor, S.L., and Greenberg, N.A. 2001. Safety assessment of propyl paraben: A review of the published literature. Food & Chem. Toxicol. 39:513. Soni, M.G., Taylor, S.L., Greenberg, N.A., and Burdock, G.A. 2002. Evaluation of the health aspects of methyl paraben: A review of the published literature. Food & Chem. Toxicol. 40:1335. Tattawasart, U., Maillard, J.-Y., Furr, J.R., and Russell, A.D. 1999. Comparative responses of Pseudomonas stutzeri and Pseudomonas aeruginosa to antibacterial agents. J. Appl. Microbiol. 87:323. Thompson, D.P. 1994. Minimum inhibitory concentration of esters of p-hydroxybenzoic acid (paraben) combinations against toxigenic fungi. J. Food Prot. 57:133–135. Tomaszewski, M. 1960. The occurrence of p-hydroxybenzoic acid and some other simple phenols in vascular plants. Bull. Acad. Pol. Sci. 8:61. Torres, A.M., Ramirez, M.L., Arroyo, M. Chulze, S.N., and Magan, N. 2003. Potential use of antioxidants for control of growth and fumonisin production by Fusarium verticillioides and Fusarium proliferatum on whole maize grain. Intl. J. Food Microbiol. 83:319. US Food and Drug Administration. 2018. Parabens. Accessed at http:​/​/www​​.fda.​​gov​/c​​osmet​​ics​/p​​roduc​​tsing​​ redie​​nts​/i​​ngred​​ients​​​/ucm1​​28042​​.htm USDA Foreign Agricultural Service Global Agricultural Information Network. 2015. Chinese Standards for Food Additives - GB2760-2015. https​:/​/ga​​in​.fa​​s​.usd​​a​.gov​​/ Rece​​nt​%20​​GAIN%​​20Pub​​licat​​ions/​​Stand​​ ard​%2​​0for%​​20Foo​​d​%20A​​dditi​​ve​%20​​Use​%2​​0 -​%20​​GB276​​0 ​-201​​5​_ Bei​​jing_​​China​​%20-%​​20Peo​​p​les%​​ 20Rep​​ublic​​%20of​​_4​-28​​-2015​​.pdf USDA Foreign Agricultural Service Global Agricultural Information Network. 2019. Russian Federation Food and Agricultural Import Regulations and Standards Report RS1838. Accessed at http:​/​/agr​​iexch​​ ange.​​apeda​​.gov.​​in​/ IR ​​_ Stan​​dards​​/ Impo​​r t​_ Re​​g ulat​​ion​/ F​​oodan​​d Agri​​cultu​​ralIm​​portR​​egula​​tions​​a ndSt​​ andar​​dsRep​​ortMo​​scowR​​uss​ia​​nFede​​ratio​​n2132​​019​.p​​df Valkova, N., Lepine, F., Bollet, C., Dupont, M., and Villemur, R. 2002. prbA, a gene coding for an esterase hydrolyzing parabens in Enterobacter cloacae and Enterobacter gergoviae strains. J. Bacteriol. 184:5011. Valkova, N., Lepine, F., Labrie, L., Dupont, M., and Beaudet, R. 2003. Purification and characterization of PrbA, a new esterase from Enterobacter cloacae hydrolyzing the esters of 4-hydroxybenzoic acid (parabens). J. Biol. Chem. 278:12779. Venugopal, V., Pansare, A.C., and Lewis, N.F. 1984. Inhibitory effect of food preservatives on protease secretion by Aeromonas hydrophila. J. Food Sci. 49:1078. Wang, Y., Oxer, D., and Hekimi, S. 2015. Mitochondrial function and lifespan of mice with controlled ubiquinone biosynthesis. Nat Commun. 6:6393. doi: 10.1038/ncomms7393 Yang, J., Li, Y., Gong, W., Wang, C., Liu, B., and Sun, C. 2014. Simultaneous determination of six parabens in foods by matrix liquid-phase dispersion extraction combined with high-performance liquid chromatography. Food Anal. Methods 7: 1693–1702. Yin, Q., Zhu, Y., and Yang, Y. 2018. Dispersive liquid–liquid microextraction followed by magnetic solidphase extraction for determination of four parabens in beverage samples by ultra-performance liquid chromatography tandem mass spectrometry Food Anal. Methods 11:797–807.

13 Dimethyl Dicarbonate and Diethyl Dicarbonate Randy W. Worobo, Rebecca M. Cheng, and Cornelius S. Ough CONTENTS 13.1 Introduction..................................................................................................................................421 13.2 Chemistry.................................................................................................................................... 422 13.2.1 Description.................................................................................................................... 422 13.2.2 Ethyl Carbamate Formation......................................................................................... 422 13.2.3 Synthesis....................................................................................................................... 423 13.2.4 Reactions....................................................................................................................... 423 13.3 Antimicrobial Activity................................................................................................................ 424 13.3.1 Yeast.............................................................................................................................. 424 13.3.2 Bacteria......................................................................................................................... 426 13.3.3 Molds............................................................................................................................ 427 13.3.4 Physical and Chemical Effects on Activity.................................................................. 427 13.3.4.1 Temperature................................................................................................. 427 13.3.4.2 pH................................................................................................................. 428 13.3.4.3 Ethanol and Other Constituents................................................................... 429 13.4 Mechanism of Action.................................................................................................................. 429 13.5 Applications..................................................................................................................................431 13.5.1 Fruit Juices.....................................................................................................................431 13.5.2 Grape Juice and Wines..................................................................................................432 13.5.3 Soft Drinks....................................................................................................................433 13.5.4 Foods..............................................................................................................................433 13.5.5 Beer............................................................................................................................... 434 13.6 Regulatory Status........................................................................................................................ 434 13.7 Toxicology....................................................................................................................................435 13.7.1 Foods..............................................................................................................................435 13.7.2 Direct Exposure............................................................................................................ 436 13.7.3 Indirect Exposure......................................................................................................... 436 13.8 Analysis....................................................................................................................................... 436 13.8.1 Direct............................................................................................................................ 436 13.8.2 Indirect.......................................................................................................................... 437 References............................................................................................................................................... 438

13.1 Introduction Diethyl dicarbonate (DEDC), also known as diethyl pyrocarbonate (DEPC), has been mentioned in the Russian literature as a cause for effervescence in sparkling wine (Parfentjev and Kovalenko, 1951a, b; Merzhanian, 1951). Kozenko (1952), in further discussing DEDC, mentioned that in 1933 a Russian investigator expressed the opinion that the neutral esters of carbonic acid were involved in sparkling wine effervescence. Although these works were erroneous in their conclusions, they stimulated interest in these types of compounds. 421

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Boehm and Mehta (1938) first isolated and properly identified the alkyl esters of pyrocarbonic acid. Bayer AG first introduced DEDC into the market for trial tests about 1959, at which time they also applied for a U.S. patent (Bernhard et al., 1959). Hennig (1959) reported on the effectiveness of DEDC as a fungicide in wine and also reported that it caused no off-aromas or off-flavors. Before the introduction into the market, Bayer AG tested a number of analogs of DEDC. The choice of DEDC over the equally effective fungicide dimethyl dicarbonate (DMDC), also known as dimethyl pyrocarbonate, was made, according to Genth (1972), because of the innocuous hydrolysis products, ethanol and carbon dioxide. However, a 1972 ban on the use of DEDC in the United States resulted in greater focus on DMDC.

13.2 Chemistry 13.2.1 Description DEDC is a colorless liquid with a faint fruit-like odor. The boiling point at 760 mm Hg is 155°C, at which temperature it decomposes. The density of DEDC is 1.12 g/cm3 at 20°C, the refractive index is 1.397 at 25°C, and the viscosity is 1.97 cP at 20°C. It is only slightly soluble in water, with a solubility of about 0.6 g per 100 g at 18°C. The compound also hydrolyzes rapidly in water. In ethanol, solubility is greater (50 g per 100 g) and hydrolysis is slower. DEDC is a lacrimator and causes skin irritation. DMDC is a colorless, fruity smelling liquid, and it has a melting point of 15.2°C, a boiling point of 123–149°C, with decomposition, and a density of 1.26. It is also only slightly soluble in water and more soluble in organic solvents. Skin and eye contact should be avoided.

13.2.2 Ethyl Carbamate Formation One source of this compound is the reaction of carbamoyl phosphate with ethanol. This ubiquitous compound, of the ornithine cycle, is found in all living cells. The reaction is:





(13.1)

It has been conclusively shown by Ough et al. (1988b, 1990) and Monteiro et al. (1989) that fermenting yeasts metabolize arginine and excrete urea into the medium. This is the primary source in wine.





(13.2)

Dimethyl Dicarbonate and Diethyl Dicarbonate



423

(13.3)

13.2.3 Synthesis The synthesis of dialkyl dicarbonate esters was described by Boehm and Mehta (1938). Their method involved the heating of the chlorocarbonic ethyl esters of emetine (6', 7', 10, 11 - tetromethoxyemetan), an alkaloid, with dilute potassium hydroxide. DEDC could be formed, as well as some other dialkyl dicarbonates, but DMDC could not be recovered. Kovalenko (1952) devised a more direct method and was able to synthesize DMDC. The reaction used was:

CH 3OCOOCl + NaOCOOCH 3 ® CH 3OCOOCOOCH 3 + NaCl (13.4)

13.2.4 Reactions The diethyl and dimethyl dicarbonates are extremely reactive substances. Some of the reactions that can take place are as follows:

R-dicarbonate = DMDC or DEDC (13.5)

(R = either two methyl groups or two ethyl groups).

R1 -dicarbonate + H 2O ® 2R1OH + 2CO2 (13.6a)



R1 -dicarbonate + R 2OH ® R1OCOOR 2 + R1OH + CO2 (13.6b)

(R2 = any alkyl or aromatic group).

R1 -dicarbonate + R 2 NH 2 ® R1OCONHR 2 + R1OH + CO2 (13.6c)

(R2 = alkyl group, alkyl-C-–, aromatic, or H).

R1 -dicarbonate + R 2COOH ® R1OCOOCOOR 2 + R1OH + CO2 (13.6d)

(R2 = any alkyl or aromatic group).

R1 -dicarbonate + R 2SH ® R1OCOSR 2 + R1OH + CO2 (13.6e)

(R2 = any alkyl or aromatic group). The hydrolysis reaction (6a) was recognized by Kovalenko (1952). The alcoholysis (6b) was documented by Kielho ̈fer and Wu r̈ dig (1963a). Larrouque r̀ e (1963) reacted alkyl and aromatic amines with DEDC to produce reaction (6c). Thoukis et al. (1962) reported reactions of DEDC with –NH2 groups and noted carbethyoxy compounds could be formed with amino acids. The mixed esters formed by reaction of DEDC with the desired alkyl acid (6d) were indicated by Thoma and Rinke (1959). Mu ̈hlra d́ et al.

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FIGURE 13.1  Hydrolysis of DEDC and DMDC at different temperatures. Half-life refers to the time for half of the DEDC or DMDC to hydrolyze: DMDC in water (circles); DMDC in 14.6% vol/vol ethanol (squares); DEDC in water (triangles).

(1967) stated that the sulfydryl groups of cysteine were carbethoxylated by DEPC (6e). Larrouque r̀ e (1965) also investigated the reaction of DEDC with thiol groups. Similar reactions [(4) through (6)] for DMDC have been reported by Peterson and Ough (1979), Ough and Langbehn (1976), and Stafford and Ough (1976). Other reactions of DEDC that have been investigated include those with phenols and phenol glucosides (Paulus and Lorke, 1967), those with vitamins (Fischer, 1970), and those with malic acid (Schelenz and Fischer, 1971). The report of Duhm et al. (1966) indicated that a large range of food chemicals reacted with DEDC. The hydrolysis rate of DEDC was considered by Pauli and Genth (1966) and further by Schelenz and Fischer (1970). The kinetics of hydrolysis of DMDC have been reported by Peterson (1978) and Genth (1979). Figure 13.1 gives the half-life of DMDC and DEDC in water and that of DMDC in 14% vol/vol ethanol at a variety of temperatures.

13.3 Antimicrobial Activity After the initial publications of the effectiveness of DEDC against wine yeast, a number of investigators further examined its usefulness as a bactericide and fungicide. Because the dicarbonates rapidly hydrolyze to the corresponding alcohol (methyl or ethyl) and carbon dioxide, the challenge is to destroy target microorganisms quickly, before hydrolysis is completed. As such, the dicarbonates cannot be relied upon for long-term protection against recontamination or later outgrowth of surviving organisms.

13.3.1 Yeast Reviews by Pauli and Genth (1966) and Genth (1964) included numerous reports on the fungicidal action of DEDC on yeasts. Their fungicidal data plus those of Alimukhamedova (1975), Grospicova et al. (1969), and Todor et al. (1967) are shown in Table 13.1. Many of the differences in antifungal effectiveness of DEDC are due to the conditions under which each test was made. Test conditions as well as tolerance differences between genera, species, or even strains could account for most of the variation.

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Dimethyl Dicarbonate and Diethyl Dicarbonate TABLE 13.1 Fungicidal Activity of Diethyl Dicarbonatea

Yeast Saccharomyces acidifaciens S. apiculatus S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae S. cerevisiae var. ellipsoides Burgundy S. cerevisiae var. ellipsoides Champagne S. carlsbergensis S. chevalieri S. globosus S. heterogenicus S. ludwigii S. oviformis S. pastorianus S. pastorianus S. pastorianus S. rouxii S. uvarum S. uvarum Schizosaccharomyces pombe Brettanomyces bruxellensis B. vini B. claussenii B. species B. species Pichia alcoholophila P. farinosa P. farinosa P. membranefaciens P. membranefaciens Torula utilis Torulopsis candida T. colliculosa T. colliculosa T. versatilis Rhodotorula glutinosa R. mucilaginosa R. rubra R. rubra Candida krusei

Starting Cell Concentration per mL 5.05 × 102 1.0 × 105 5.0 × 102–1.0 × 104 5.0 × 104 1.50 × 102 5.00 × 102 4.0 × 103 1.0 × 103 3.6 × 106 1.4 × 104 5.0 × 104 6.0 × 105 4.0 × 103 5.6 × 102 8.0 × 102 5.2 × 102 5.0 × 104 5.0 × 104 4.0 × 105 4.0 × 102 4.0 × 105 1.0 × 105 5.0 × 104 2.0 × 103 5.0 × 102 1.0 × 105 1.0 × 104 5.0 × 102 5.0 × 104 4.6 × 103 1.0 × 103 4.0 × 105 4.0 × 103 5.0 × 102 2.0 × 104 1.0 × 105 1.0 × 105 5.6 × 102 5.0 × 102 5.0 × 102 5.6 × 102 1.0 × 105 2.0 × 103 5.0 × 103 1.0 × 105

DEDC Concentration (mg/L) Sufficient to Be Fungicidal (100% kill) 200 60–90 30–80 50 50–100 50 500–1000 500 50 12 100 100 250–1000 30 30 30 25 50 100 100 100 100–500 25 200 400 250 25 200 25 100 100 100 100 25 14 250 100 30 200 200 30 200 200 300 60–200 (Continued)

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TABLE 13.1 (CONTINUED) Fungicidal Activity of Diethyl Dicarbonatea

Yeast C. krusei C. krusei C. lipolytica C. mycoderma C. mycoderma C. parapsilosis C. pseudotropicalis C. pulcherima Cryptococcus diffluens Kloeckera apiculata Debaryomyces kloeckeri Hanseniaspora apiculta Hansenula anomala H. anomala H. anomala Fabospora macedoniensis Zygosaccharomyces eupagicus Zygosaccharomyces sp. a

Starting Cell Concentration per mL

DEDC Concentration (mg/L) Sufficient to Be Fungicidal (100% kill)

5.0 × 102 5.0 × 104 1.0 × 105 5.0 × 102 2.5 × 103 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 104 1.0 × 104 5.0 × 102 1.0 × 104 4.0 × 103 5.0 × 102 2.0 × 103 4.0 × 102 6.3 × 102 1.0 × 105

200 200 90–500 200 100 200 200 200 75 200 200 25 250–500 200 100 30 30 80

Reports of various tests done under varied environmental conditions.

Genth (1979) reported the effectiveness of DMDC as a fungicide when used in alcohol-free beverages (Table 13.2). Daudt and Ough (1980) reported its use as a sterilant against yeast in wine (Table 13.3). DMDC has proven to be equal or superior to DEDC as a fungicide in both instances. Pauli and Genth (1966) showed that, for every ten-fold increase in cell count, the amount of DEDC required for fungicidal action was doubled. Daudt and Ough (1980) showed that 50 mg/L of DMDC effectively sterilized a wine with 10% vol/vol ethanol containing 2 × 104 cells. Viable cells remained, however, if only 25 mg/L of DMDC was used, with as few as 50 cells/mL in the wine initially. The rate of kill is proportional to the concentration of DEDC (Ough and Ingraham, 1961). Over 2 hours were required for complete kill of 50 yeast cells/mL with the addition of 40 mg/L of DEDC; less than half an hour was required with 120 mg/L of DEDC. Similar results were found using DMDC (Daudt and Ough, 1980). A delay in the lethal effect at lower DEDC or DMDC concentrations could be due to the capability of the yeast cells to reproduce for short periods before permanent damage and death of the organism. Ough et al. (1988c) investigated the interaction of sulfur dioxide and DMDC. They found additive effects between the two inhibitors when testing a very resistant yeast. For a lethal dose of DMDC for this yeast, a much larger than normal amount was required.

13.3.2 Bacteria A comparison of the antibacterial effectiveness of DEDC on a number of different genera has been summarized in Table 13.4 (Pauli and Genth, 1966). Similar tests using DMDC (Genth, 1979, 1980) are shown in Table 13.5. Results on the antimicrobial activity of these compounds indicated that they had approximately equivalent bactericidal activity (Genth, 1979, 1980). Murata (1974) found that 1300 mg/L of DEDC was capable of destroying a Lactobacillus caseii bacteriophage. Ough et al. (1988c) found Oenococcus oeni relatively insensitive to DMDC at normal wine treatment levels. It had a slight additive effect on the bacteria when used with sulfur dioxide.

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Dimethyl Dicarbonate and Diethyl Dicarbonate TABLE 13.2 Effective Fungicidal Concentration of DMDC in Alcohol-Free Beverages a Yeast

DMDC Concentration (mg/L) Sufficient to Be Fungicidal (100% kill)

Saccharomyces carlsbergensis S. carlsbergensis S. diastaticus S. oviformis S. bailii S. cerevisiae S. uvarum S. pastorianus S. apiculatus S. globosus Zygosaccharomyces priorianus Rhodotorula mucilaginosa R. glutinosa R. rubra Candida krusei Pichia membranefaciens P. farinosa Torulopis candida T. versatilis T. stallata Torula utilis Endomyces lactis Kloeckera apiculata Hansenula anomala a

100 70 200 100 120 30 20 100 70 40 90 50 30 200 200 30 100 100 100 75 250 70 30 50

The cell counts were about 500 cells/mL, temperature 28°C, and pH between 2.8 and 4.7.

13.3.3 Molds For activity against molds, higher levels of DEDC were required than for either yeast or bacteria. Table 13.6 shows the results summarized by Pauli and Genth (1966). Genth (1979) reported that 500 spores per milliliter of Penicillium glaucum, Byssochlamys fulva, Botrytis cinerea, Mucor racemosus, and Fusarium oxysporum required 200, 100, 100, 500, and 100 mg/L of DMDC, respectively, for effective killing. Vegetative cells and mold conidia are relatively sensitive to both dicarbonates. However, heat-resistant structures such as ascospores of B. fulva are quite resistant and, thus, likely to survive exposure to up to 200 ppm DMDC (Splittstoesser and Wilkinson, 1973; Van der Riet et al., 1989).

13.3.4 Physical and Chemical Effects on Activity It has been recognized that DEDC activity could be altered by some of the physical or chemical variables existing during handling of products. Such variables are temperature, pH, chemical makeup of product, and the amount of other reactive substances in the treated material.

13.3.4.1 Temperature The hydrolysis rate of DEDC or DMDC is dependent on temperature—the higher the temperature the more rapid the rate. The half-life values for both DEDC and DMDC are given in Figure 13.1. Despite the

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TABLE 13.3 Antimicrobial Effectiveness of DMDC in a Standard Wine Containing 10% Ethanol DMDC Added (mg/L) Microorganism Saccharomyces cerevisiae Montrachet S. bayanus Champagne Schizosaccharomyces pombe Saccharomyces cerevisiae Steinberg S. fermantati Florb Brettanomyces spp. Saccharomyces cerevisiae Distillers S. cerevisiae Geisenhiem S. cerevisiae Burgundy S. cerevisiae Rankine 350 S. cerevisiae Eperney Rankine 729 S. cerevisiae Tokayb Champagne Epernay (Geisenheim Reinzucthefen) Saccharomyces bayanus (formerly oviformis) S. oviformis (S. bayanus) Inst. Pasteur Schizosaccharomyces pombe Moscovinb Geisenheim 1949 (Geisenheim Reinzucthefen) Dekkera intermedia FS&T 71-12 Rhodotorula rubra a b

c

0

6.25

12.5

25

50

100

58 2500 42 96 203 10,000 78 88 2000 89 2500 887 5000

16 419 14 50 53 2000 0 0 310 64 2000 129 3000

0 729 4 43 9 100 0 0 0 38 844 144 350

0 0 0 0 0 0 0 0 0 0 0 0 0

0 0 0 0 0 0 0 0 0 0 0 0 0

0 0 0 0 0 0 0 0 0 0 0 0 0

17 214 2200 1200 2,000 1,000

19 346 529 800 600 800

0c 200 2000 640 70 720

0 11 4 5 0 650

0 0 0 0 0 0

0 0 0 0 0 0

a

Values are the number of viable cells remaining in wine after 38 h. Yeasts did not grow well in wines with 10% ethanol; wines of 8% ethanol were used instead. S. fermentati Flor needs to stay at least 4 days in the incubator (at 37°C) to grow on the plates; S. cerevisiae Tokay and S. pombe Moscovin need 3 days. S. fermentati Flor also grew better in 8% ethanol wines, but the data presented are for 10% ethanol wines. Although nothing grew on the plates, growth developed later in the bottles, indicating some viable cells remaining.

more rapid hydrolysis at higher temperatures, Turtura (1966) found that increased temperature enhanced the effectiveness of DEDC as a sterilant. He noted that DEDC was twice as effective at 27°C as at 0°C. Grospicova et al. (1969) also studied the effect of treatment temperature and found a two-fold increase in effectiveness from 6 to 20°C on two Lactobacillus species. Shibasaki et al. (1969) noted the temperature coefficient for molds in distilled water and in phosphate buffer was 2.5 and 2.3, respectively, but the temperature coefficient for the microbiocidal activity was 5 and 10 for the same two solutions. Their work with Penicillium thomii showed that at pH 4.0, about a four-fold increase in the antiseptic effectiveness of DEDC was evident as the temperature was increased from 10 to 20°C, and another ten-fold increase occurred at 20–30°C. Splittstoesser and Wilkinson (1973) reported a 100-fold increase in effectiveness of DEDC against S. cerevisiae and Lactobacillus plantarum by raising the temperature from 20 to 40°C. Goto et al. (1970) found no temperature effect; however, they used only 250 mg/L of DEDC against high numbers of yeast cells. At the higher temperature (35°C), the rapid growth rate could have overcome the sterilization effects in the rich medium used. They also found no effect of ethanol or sugars on the kill rates (Goto et al., 1970).

13.3.4.2 pH Reaction of R–NH groups with DEDC and DMDC (one of the proposed modes of action) is dependent on the degree of ionization of the –NH group. In the pH range 3.0–4.0, this relation is demonstrated by the

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Dimethyl Dicarbonate and Diethyl Dicarbonate TABLE 13.4 Antibacterial Effectiveness of Diethyl Dicarbonatea

Bacteria Acetobacter pasteurianus A. xylinum Bacterium coli Escherichia coli Bacterium aceticum Staphylococcus aureus Lactobacillus arabinosus L. helveticus L. pastorianus Lactobacillus buchneri Bacillus subtilis Salmonella Typhimurium Bacillus mediosporum B. megatherium Salmonella Thompson Pseudomonas cocciformis Pseudomonas, pigmented Pseudomonas, unpigmented P. erithrogloem P. flavum P. mildenbergii Micrococcus annulatus M. lacteus M. lacteus Sarcina flavescens a

Starting Cell Concentration per mL

DEDC Concentration (mg/L) Sufficient to Be Bactericidal (100% kill)

5.0 × 102 6.0 × 103 1.0 × 105 1.0 × 105 5.0 × 104 1.0 × 103 1.0 × 106 1.0 × 107 6.0 × 102 4.8 × 102 4.0 × 109 1.0 × 105 4.8 × 102 4.8 × 102 1.0 × 105 8.0 × 102 1.0 × 105 1.0 × 105 7.2 × 102 4.8 × 102 6.4 × 102 4.8 × 102 6.4 × 102 6.4 × 102 8.0 × 102

80 300 400 1500 500 70–100 250 100–300 300 30 3000 2000 30 30 2000 30 200 500 30 30 30 30 30 30 30

Determined under various environmental conditions.

reaction of DEDC and DMDC with ammonia to form urethanes (Figure 13.2). As the ammonia becomes ionized at the lower pH, the reaction is lessened (Ough, 1976b; Ough and Langbehn, 1976).

13.3.4.3 Ethanol and Other Constituents The rate of hydrolysis of DEDC or DMDC can be significantly decreased by increasing the ethanol concentration. Other water-soluble organic solvents also have similar effect. The reactivity of DEDC or DMDC is also such that any proteinaceous material suspended in the solution reacts and decreases the antimicrobial effectiveness of the treatment. Porter and Ough (1982) found that ethanol increased the effectiveness of DMDC and that 20°C was the optimum use temperature.

13.4 Mechanism of Action The inactivation of microorganisms by DEDC or DMDC appears strongly related to the inactivation of the enzymes of the organism. Protein modification, through reaction of nucleophilic groups, such as imidazoles, amines, or thiols, can readily occur with the dicarbonate (Osterman-Golkar et al., 1974; Ehrenberg et al., 1976). The point of attack is one of the central carbon atoms. The residual portion of the dicarbonate is unstable and decomposes rapidly. Reactions with protein imidazole (7) and amine

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TABLE 13.5 Antibacterial Effectiveness of Dimethyl Dicarbonatea

Bacteria Acetobacter pasteurianus Escherichia coli Pseudomonas aeruginosa Staphylococcus aureus Lactobacillus buchneri Lactobacillus pastorianus Lactobacillus brevis Pediococcus cerevisiae a

Starting Cell Concentration per mL 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 102 5.0 × 102

DEDC Concentration (mg/L) Sufficient to Be Bactericidal (100% kill) 80 400 100 100 30 300 200 300

Temperature 28°C and pH between 2.8 and 4.7 in an artificial medium.

TABLE 13.6 Fungicidal (Mold) Effectiveness of Diethyl Dicarbonate

Mold Aspergillus flavus A. niger A. niger A. terreus Penicillium digitatum P. expansum P. glaucum P. italicum P. leteum Mycoderma species Trichoderma viride Botrytis cinerea Oidium lactis Fusarium oxysporum Fusarium orthoceras Neurospora sitophila Mucor racemosus Rhizophus nigricans Pullularia pullulans Cladosporium herbarum Paecilomyces spp. a

Starting Spore Concentration per mL

DEDC Concentration (mg/L) Sufficient to Be Fungicidal (100% kill)

5.0 × 102 1.0 × 104 2.9 × 102 3.0 × 102 3.0 × 102 1.0 × 102 3.0 × 102 3.0 × 102 3.0 × 102 4.0 × 103 1.0 × 103 1.0 × 103 3.0 × 102 1.0 × 103 1.0 × 103 3.0 × 102 3.0 × 102 3.0 × 102 3.0 × 102 5.0 × 102 1.3 × 102

1000 150 750 250–400 120–250 100 250 100 300 500–100 500 100 300–700 100 100 50–200 300–700 300 100–200 100 1000

Tested under various environmental conditions.

groups are of particular importance owing to the ease and rapidity with which they occur (Means and Feeney, 1971).



(13.7)

Dimethyl Dicarbonate and Diethyl Dicarbonate

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FIGURE 13.2  Influence of ammonia, pH, and DEDC on the amounts of ethyl urethane formed, mg/L of DEDC: 50 (triangles); 100 (hexagons); 200 (squares); 400 (circles).

Mühlrád et al. (1967) investigated the reactions of DEDC with a number of amino acids. The reaction with histidine is similar to that of the imidazole reaction, i.e., a reaction with the imino nitrogen. Holbrook and Ingram (1973) found that the enzyme lactate dehydrogenase was completely inactivated by reaction of DEDC with the histidyl groups of the enzyme. Many other reports (Huc et al., 1971; Dann and Briton, 1974; Choong et al., 1977) with other enzymes have shown that DEDC reacts primarily with the histidyl moeity to form carbethoxy-histidine. Enzyme inhibition results from active site blocking and conformational changes. For example, the histidyl group in alcohol dehydrogenase has been proposed (Ringold, 1966) to be involved in the oxidation-reduction process. Blocking of either nitrogen in the histidine ring would inactivate the enzyme. DEDC has been used routinely to inhibit nuclease in the extradition of nucleic acids. Any single key enzyme inactivated could eventually cause the death of an organism. A report by Ehrenberg et al. (1976) reviews the many reactions that can take place with DEDC.

13.5 Applications Pauli (1984) reviewed the nature of food additives and their possible hazardous reactions. DEDC was one of concern. The usefulness of DMDC as a beverage preservative, its antimicrobial activity, and some of its chemistry were reviewed by Thoukis (1983). Because of its regulatory status, DMDC is approved only for use in various beverages. Furthermore, its application to beverages requires use of special equipment for metering and mixing DMDC.

13.5.1 Fruit Juices Martienssen (1961) found DEDC could be used to stabilize fruit juice and prevent fermentation for several days. The authors suggested that syrups with over 50% sugar would delay the hydrolysis of the ester. Mehlitz and Gierschner (1964), who investigated DEDC sterilization failures, noted that possible mold contamination points, such as corks, had to be sterilized before treatment of juices with DEDC. Wucherpfenning (1966) pointed out that the DEDC reacts with many components in the juices and therefore the juice can no longer be considered “natural.” Attempts to use DEDC to prevent malolactic

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fermentation in apple juice have met with little success (Hara and Otsuka, 1967). Genth (1969) has reviewed the use of DEDC for cold sterilization of fruit juices. Fisher and Golden (1998) reported that Escherichia coli O157:H7 populations were reduced from about 7.0 log CFU/mL to undetectable levels within three days in apple cider containing 250 ppm DMDC and stored at 4°C; increasing the storage temperature to 25°C enhanced DMDC effectiveness, and E. coli O157:H7 was undetectable within two days of storage. Basaran-Akgul et al. (2009) treated 8 different apple ciders inoculated with different strains of E. coli O157:H7 and found that 250 ppm of DMDC was able to achieve greater than a 5.0-log reduction after 6 hours at room temperature. DMDC has also been found to be effective at reducing Salmonella enterica in orange juice at concentrations of 172 and 200 ppm, resulting in a 5-log reduction after 24 hours when held at 4°C (Cheng et al., 2018). DMDC has also been used with other non-thermal processes, such as high-pressure processing, ultraviolet (UV) radiation, or additional preservatives to increase DMDC effectiveness. Combination of DMDC with sodium benzoate or potassium sorbate offers enhanced protection by providing a secondary barrier against surviving spoilage organisms (Golden, 2002; Worobo, 2002). E. coli O157:H7 populations were reduced from 7.0 log CFU/mL to undetectable levels at 4°C within 72 hours in apple cider and 48 hours in orange juice containing 250 ppm DMDC (Lakins, 2002). A combination of 250 ppm DMDC and 450 ppm sodium benzoate rendered E. coli O157:H7 undetectable within 48 hours in apple cider and 24 hours in orange juice. Lakins (2002) reported that inoculated Salmonella (7.0 log CFU/mL initial) was undetectable in apple cider containing 250 ppm DMDC within 48 hours; the same result was achieved within 24 hours in cider containing a combination of 250 ppm DMDC and 450 ppm sodium benzoate. Williams et al. (2006) reported that a combination of ozone, DMDC, and 24-hour storage at 4°C was effective at reducing populations of E. coli O157:H7 in apple cider by 5.7 log CFU/mL and Salmonella spp. in apple cider and orange juice by 5.8 and 5.6 log CFU/mL, respectively. DMDC has also been used in conjunction with non-thermal processes, such as high pressures or UV radiation to achieve microbial inactivation. Combination treatments of 400 MPa with 62.5 ppm DMDC and 550 MPa with 125 ppm DMDC were able to achieve greater than 5.0 log reductions of Salmonella Agona in orange juice and E. coli O157:H7 in apple juice, respectively (Whitney et al., 2008). After 24-hour storage at 4°C, E. coli O157:H7 was undetectable in apple juice treated at 550 MPa with both 62.5 and 125 ppm DMDC. Gouma et al. (2015) combined DMDC with UV radiation and mild heat (55°C) to inactivate E. coli in apple juice. Combination UV and heat treatments using 25 mg/L, 50 mg/L, and 75 mg/L of DMDC resulted in a greater than 6.0-log reduction after 3.58 min, with the addition of DMDC reducing UV dose and processing time (Gouma et al., 2015). Worobo (2002) reported that strain variation and differences in apple varietals used in cider-making affect the sensitivity of E. coli O157:H7 in apple cider. Using 250 ppm DMDC, E. coli O157:H7 D-values ranged from a low of 0.24 hours in apple made from McIntosh apples to a high of 1.02 hours in cider made from Red Delicious apples. B. fulva ascospores were resistant to DMDC up to 1000 mg/L concentration in apple juice, while only 25–75 mg/L was required to kill the vegetative cells. Increased effectiveness was noted at higher treatment temperatures (Van der Riet et al., 1989).

13.5.2 Grape Juice and Wines The effectiveness of DEDC as a sterilant, particularly in grape juice and wines, has been reviewed by Rankine (1964), Pauli and Genth (1966), Pauli (1969), Genth (1971), Barnick (1973), and Ough (1978). Beuchat (1976) investigated the ability of DEDC to inhibit the growth of Byssochlamys nivea ascospores in grape juice. This microorganism is known to produce the toxin patulin. It took 500 mg/L of DEDC to inhibit growth of the spores. The initial work outside the Bayer AG laboratories was reported by Hennig (1959, 1960) and indicated that DEDC was a satisfactory inhibitor and had active cidal properties. He indicated that the inhibitory effect required about one-third the amount of DEDC, as did the killing effect. Hennig (1961) also developed the atomizing pump system that came into general use for application of the compound. Mayer and Luthi (1960) also did some of the initial testing on wine in Switzerland and suggested 50–100 mg/L

Dimethyl Dicarbonate and Diethyl Dicarbonate

433

of DEDC as an effective sterilizing level. Ough and Ingraham (1961) confirmed that 50–100 mg/L of DEDC was a safe sterilizing level in wine and also noted no identifiable sensory problems with DEDC. A sensory threshold for DEDC was found at about 600 mg/L (Van Zyl, 1962; Blouin and Barthe, 1963). Van Zyl (1962) also noted the synergistic effect of DEDC with sulfur dioxide. Italian studies by Fontana and Colagrande (1964) indicated that levels of 150–200 mg/L DEDC were required for effective sterilization. This was further verified for both wine and grape juice by Garoglio and Stella (1964) and Minarik (1964) in Czechoslavakia, Adams (1965) in Canada, Kalugina (1965) in Russia, Todor et al. (1967) in Romania, Ivanov et al. (1967) in Bulgaria, Prillinger (1964) in Austria, and Rankine and Pilone (1974) in Australia. All these researchers tested DEDC in a practical manner, and most determined it to be a very effective antimicrobial agent for use in bottling wine or grape juice in conjunction with the use of good standard sanitary practices. These are only a representative group of the published reports. Relatively large amounts of ethyl urethane (up to 600 mg/L) were found in a number of commercial Japanese sakes (Ough, 1978). No diethyl carbonate was found in these samples, indicating that no ethanol was present when the DEDC was used. It appears that this method was used commercially in Japan to sterilize the koji before the alcoholic fermentation (Hara et al., 1970). Further studies (Ough et al., 1988a) showed that heating of ethyl alcohol and urea in water solutions resulted in the formation of ethyl carbamate. In wine, Ough et al. (1988b) found that urea when formed during fermentation could remain in the wine. If the wine was heated ethyl carbamate formed. Freshly made sake has no more urea than any other similar ferment; however, in the processing of sake it is heated. This results in significant amounts of ethyl carbamate. DMDC has been reviewed as an agent for inhibiting yeasts and fermentation in wine (Delfini et al., 2002; Costa et al., 2008; Zuehlke et al., 2015). Porter and Ough (1982) investigated the use of DMDC as an alternative sterilant to DEDC for alcoholic beverages, especially in wine. DMDC at 100 mg/L in 8% v/v ethanol at 30°C effectively reduced S. cerevisiae Montrachet to undetectable levels after 20 minutes. For the same concentration of DMDC in 10% v/v ethanol at 30°C, the same result was achieved within 10 minutes, suggesting DMDC is more effective at higher ethanol concentrations. Divol et al. (2005) found that DMDC used in concentrations of 200 mg/L was more effective than SO2 at stopping fermentation in pure cultures of S. cerevisiae, Candida stellata, and Zygosaccharomyces bailii and in botrytized must. Fermentative spoilage of grape juice at 21°C was prevented by addition of 0.8 mM DMDC when juice was inoculated with S. cerevisiae Montrachet at levels of 2 or 200 CFU/mL (Terrell at al., 1993). These researchers also noted that DMDC was generally more effective than sulfur dioxide and sorbic acid, and that the effectiveness of DMDC, but not the other chemicals, was enhanced when the storage temperature was increased to 31°C. Similarly, Morris et al. (1996) demonstrated that 0.8 mM DMDC effectively prevented spoilage of grape juice held at 31°C, while sorbic acid, potassium metabisulfite, and combinations of these two delayed but did not prevent spoilage.

13.5.3 Soft Drinks DEDC appears to have some value in inhibiting growth of osmophilic yeasts in natural fruit juice soft drinks (Pozsonyi, 1972). A maximum of 300 mg/L of DEDC was suggested for use in sterilizing the drink against yeast or bacterial growth (Pozsonyi, 1972). Molds were not found to be sufficiently killed or inhibited. Pátkay et al. (1973) investigated the use of DEDC with various diluted and carbonated commercial syrups and found that a level of 200 mg/L of DEDC was sufficient to protect the drinks. DMDC has been suggested for commercial use in Germany as a sterilant for artificial drinks (Genth, 1980). The advantage of using DMDC is that no reactions occur with sugar, sugar alcohols, or artificial sweeteners, such as saccharin or cyclamate.

13.5.4 Foods Rash (2003) reported that soaking cantaloupes for 3 minutes in a 10,000-ppm solution of DMDC reduced the population of Salmonella from 5.01 log cfu/cm2 to undetectable levels; Salmonella populations were detected only by enrichment after a 3-minute treatment with 5000 ppm DMDC. Mount et al. (1999)

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Antimicrobials in Food

demonstrated that shelf life of fresh salsa could be extended by several weeks by a combination of 100 ppm DMDC and 1% potassium sorbate. They reported that the aerobic plate count and yeast/mold count of fresh salsa decreased from 6 and 2 log CFU/g, respectively, to undetectable levels within 8 days of storage at 4°C. Chen et al. (2013) dipped Chinese cabbage parts in a 200 mg/L DMDC solution and found that the solution reduced total aerobic plate counts and total yeast and mold count of the leaf and stalk by at least 4.0 log CFU/g and 3.0 log CFU/g, respectively. The DMDC solution was also able to inhibit polyphenol oxidase and peroxidase activity, which is linked to enzymatic browning reactions, thus preserving the quality of Chinese cabbage (Chen et al., 2013). Wang et al. (2013) performed similar analyses using a 200 mg/L DMDC solution as a disinfectant for fresh cut carrots, reporting that total aerobic plate count and total yeast and mold counts were reduced by more than 3.0 log CFU/g. Molin et al. (1963) summarized the possible uses of DEDC in foods. They found that the growth of B. cinerea on strawberries was delayed by dipping the fruit into 100 and 1000 mg/L solutions of DEDC. No adverse sensory effects of the DEDC were noted. In testing apple sauce, they noted that up to 1000 mg/L was required to inhibit mold growth and was not very effective in preventing fermentation of apple sauce. The storage life of freshly slaughtered chicken carcasses dipped into a solution of DEDC by SchmidtLorentz (1962) was extended by 65%. Surface bacteria showed a 10- to 20-fold reduction. Hara and Otsuka (1966a) studied the use of DEDC to sterilize koji. They determined the most satisfactory approach was to mix the koji in water and then add DEDC. The wild yeast and some bacteria present were killed by 100–200 mg/L, but the lactic bacteria and Aspergillus oryzae required more than 600 mg/L of DEDC.

13.5.5 Beer Mönch (1961) appears to have been the first to test DEDC in beer sterilization. His studies indicated levels between 250 and 1000 mg/L were required for effective action against spoilage by yeast. He noted a taste threshold in beer of 1000 mg/L of added DEDC. Molin et al. (1963) indicated that 0.01% DEDC doubled the shelf life of a beer. A system was suggested by Kozulis et al. (1971) in which DEDC was used in conjunction with p-hydroxybenzoate esters for beer sterilization.

13.6 Regulatory Status DEDC may not be used for treatments involving food or beverages in the United States. In 1972, based upon a report by Löfroth and Gejvall (1971), the U.S. Food and Drug Administration (FDA) banned the use of DEDC (FDA, 1972). Until that time, it was legally used in a number of beverages. The first approved use for DMDC was for addition to wines as a yeast inhibitor at a level not to exceed 200 ppm (FDA, 1988). The FDA, in 2001, amended the food additive regulations to provide for a more descriptive term, “microbial control agent,” in place of “inhibitor of yeast,” for the safe use of DMDC based upon a food additive petition filed by Bayer Corp. (FDA, 2001a). DMDC presently is approved for use as a direct food additive to be used as a microbial control agent in certain beverages in which the microbial population has been reduced to 500 microorganisms per mL or less by current good manufacturing practices. Beverages in which DMDC is approved for use are: (1) wine, dealcoholized wine, and low-alcohol wine, at a limit of 200 ppm; (2) ready-to-drink teas, at a limit of 250 ppm; (3) carbonated or non-carbonated, nonjuice-containing (≤1% juice), flavored or unflavored beverages containing added electrolytes (5–20 meq/L Na+ and 3–7 meq/L K+), at a limit of 250 ppm; and (4) carbonated, dilute beverages containing juice, fruit flavor, or both, in which the juice content does not exceed 50%, at a limit of 250 ppm (FDA, 2001a). In 2000, Bayer Corp. filed an effective notification of food-contact substance (FCN 0035) with the FDA, thereby allowing for use of DMDC as a microbial control agent in non-carbonated juice beverages containing up to and including 100% juice (FDA, 2000). A food-contact substance is defined as one intended for use as a component of material used in manufacturing, packaging, transporting, or holding food where the substance is not intended to have a technical effect on the food (FDA, 2002). Limitations for use of DMDC as a food-contact substance are similar to those specified for its use as a food additive,

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i.e., the microbial load of the juice must be reduced by current technologies (e.g., heat) prior to the addition of DMDC at a limit of 250 ppm (FDA, 2003). DMDC breaks down into carbon dioxide and methanol at very low concentrations, making it an incidental additive, a processing aid that breaks down into constituents that are either normally present in the food or found at insignificant levels (FDA, 2001b). Incidental additives are exempt from label reporting. DMDC is marketed and sold for application to foods under the registered trademark, Velcorin®. The basis for the ban on DEDC was that a small amount of ethyl carbamate, a carcinogen, was formed by the reaction of ammonia with the DEDC. Despite extensive proof of the error of that report (Fischer, 1972; Ough, 1976a), the ban was upheld, based on the Delaney Clause of the Food Additive Amendment. Interestingly, all fermented products tested in one study, including yogurt, beer, and soy sauce, had small but detectable amounts of ethyl carbamate (Ough, 1976a). As it turns out, the amount of ethyl carbamate added to the food chain by the use of DEDC would be minor. The natural formation of precursors in fermented foods and beverages and in drugs using alcohol as either a carrier or as an antiseptic far exceeds any formed from the use of DEDC. Nevertheless, this has not altered the FDA’s position on this compound.

13.7 Toxicology Since the dicarbonates are expected to be decomposed before consumption, the reaction products are of major concern when considering their use in foods. Direct consumption or contact with the material, however, must not be ignored.

13.7.1 Foods Since DMDC is hydrolyzed to carbon dioxide and methanol almost immediately after addition to beverages, the FDA determined that there would be virtually no exposure of consumers to the additive itself when used within the limits of 200 ppm in beverages (FDA, 1988). Methanol is the principal hydrolysis product of concern resulting from addition of DMDC to wine. Theoretically, complete hydrolysis of DMDC would yield two moles of methanol from one mole of DMDC added to wine. Using a worstcase scenario, the FDA determined that consumption of wine containing 200 ppm DMDC at a rate of 232 grams per person per day would result in a daily intake of not more than 22 milligrams per person per day, well within safe limits (FDA, 1987). An adult human can metabolize up to 1500 milligrams of methanol per hour with no adverse symptoms or effects (Lehman, 1963). A fairly extensive report of the safe use of DMDC in wine, including reference to breakdown and reaction products, is described in the FDA response to the original DMDC food additive petition (FDA, 1988). Hecht (1961) found no significant toxic effects in rats consuming DEDC-treated grape juice for a 59-day period. This observation was confirmed by Bornmann and Loeser (1961). Although DEDCtreated milk caused some weight loss, no other symptoms were detected by these workers. Bornmann and Loeser (1966) did long-range studies using the reaction product of DEDC and ethanol (diethyl carbonate, DEC). No harmful effects were found when DEC was given to rats at 0.3% in drinking water for 100 weeks or by force feeding it to dogs at 600 mg/kg per day for 3 months. Sharratt et al. (1972) repeated the short-term tests using wine, beer, orange juice, and blackcurrant juice. Treatments with DEDC caused no adverse effects from the reaction products on the rats being tested. Studies by Lang et al. (1966) showed that most of the reaction products were hydrolyzed enzymatically to the original compound plus carbon dioxide and ethanol. The exception was the carbethyoxy ascorbic acid. This product hydrolyzed into the original ascorbic acid, dehydroascorbic acid, diketogluconic acid, and furfural after a few hours. Paulus and Lorke (1967) tested these products in short-term studies on rats and found no adverse effects. Wolf et al. (1969) found no toxic effects in five human volunteers, who over a 40-day period ingested 250 mL wine per day that was treated with 150 mg/L of DEDC. A single treatment of 1 liter of the treated wine caused no toxic symptoms in the group. Zaitsev et al. (1970) suggested that DEDC not be used in foods because of its toxicity. Their fear was that the compound would not completely hydrolyze.

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Santini et al. (1985) tested the mutagenicity of DEDC on two strains of Salmonella Typhimurium (His–) and on Bacillus subtilis 170 (Trp –, Ura–) and found no reversion to prototrophy. They also found no effect of ethyl carbamate on the mutagenicity of B. subtilis. There was some transformation reduction. The Joint Food and Agriculture Organization/World Health Organization Expert Committee on Food Additives (JECFA) (1972) studied the use of DEDC and concluded beverages over pH 4.0 with a significant content of ammonia, amino acids, or proteins (for example, milk) should not be treated with DEDC. Treatment of beer and fruit juices is not technically justified, nor should wine be treated with DEDC. The acceptable level of treatment for soft drinks, carbonated or not, is 250 mg/L. Under no circumstances should the human daily intake of ethyl carbamate exceed 100 μμg/day. Short-term toxicological studies reported by Genth (1980) indicated the acute toxicity for rats of DMDC was 330–900 mg/kg body weight. The Ames test for mutagenicity was negative. No adverse effects were found after feeding fruit juice and alcoholic beverages treated with 4000 mg/L of DMDC for 3 months. The amounts of methanol produced were well below toxicological levels. Methyl urethane, another reaction product, was reported not to be a carcinogen (Pound, 1967). The JECFA (1991) has also reviewed studies regarding short- and long-term DMDC toxicity studies in rats and dogs, where no evidence of toxic effects was found from consumption of DMDC-treated beverages. The JECFA has ruled that since the concentrations of methanol resulting from DMDC use are similar to naturally occurring amounts in fruit and alcoholic beverages, DMDC is acceptable to use as a cold sterilization agent up to a concentration of 250 mg/L and in accordance with good manufacturing practices (GMPs).

13.7.2 Direct Exposure The median lethal dose (LD50) values of DEDC for mice, rats, cats, rabbits, and dogs were reported to be 1558, 850, 100–250, 500–750, and greater than 500 mg/kg body weight, respectively, by the oral route (Joint FAO/WHO Expert Committee on Food Additives, 1972). Tests with rabbits, guinea pigs, rats, and mice (Hecht, 1961) showed 1 mg/L exposure in air caused chronic respiratory symptoms and 10 mg/L was lethal. It is a strong skin and eye irritant.

13.7.3 Indirect Exposure The carcinogenicity of ethyl carbamate has been well-documented in animals (Allen et al., 1986; Miller and Miller, 1983; Mirvish, 1968; Schmähl et al., 1977; Woo, 1983). Uzvolgyi et al. (1983) found that DEDC and ammonia incubated in vitro in rat gastric juice and then administered to mice orally caused lung cancer in the mice. If DEDC and ammonia were administered by probe in vivo to mice, the same effect was found. Later Uzvolgyi (1986) reported contrary results when testing lactating mice and their offspring.

13.8 Analysis Both DEDC and DMDC are very reactive, and if any water or other proton donor substance is present, they break down autocatalytically. Two types of analyses are available: one to measure the pure or nearly pure substance and the other to measure the residual amount of a specified reactant in the treated material and calculate the amount of dicarbonate added. The first is used primarily to determine the purity of the dicarbonate or to study the kinetics of its reaction. The other method is used at some later date to determine the amounts added originally.

13.8.1 Direct The dicarbonates, either diethyl or dimethyl, can be determined. DEDC was measured with a relative standard deviation of ±0.3% by Cuzner et al. (1971). A measured amount of dicarbonate was combined with an excess but known amount of morpholine. The morpholine reacted with the dicarbonate.

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The remaining morpholine was titrated with standardized methanolic HCI solution to a pH end point of 4.0. Hara and Otsuka (1966b) tested three other amines besides morpholine and determined that n-butylamine and isobutylamine gave the sharpest end point. They extracted dicarbonate from sake with organic solvents before analysis. Moncelsi (1970) has suggested a spectrophotometric method using 4-aminoantipyrine, which requires an extraction step. Peterson (1978) used the method suggested by Berger (1975) for his studies on hydrolysis rate. This spectrophotometric method lends itself to multi-sample analysis. The reactions involved in this method are as follows:



(13.8)

The colored compound 5-thio-2 nitrobenzoic acid is destroyed by the dicarbonate. Pseudo-first-order kinetics describe the reaction. The rate of the color disappearance is proportional to the initial amount of dicarbonate present.

13.8.2 Indirect The reactions of DEDC with alcohols were postulated earlier. Kielhöfer and Würdig (1963a, b) were first to use the formation of diethyl carbonate to measure the original amount of DEDC added to an alcoholic beverage. They found no DEC in untreated wine. The wines treated with DEDC formed DEC in amounts proportional to the amount of ethanol present in the wine. The method consisted of extraction of the wine with carbon disulfide or pentane and determining the DEC by gas chromatography. The amounts found are in the range of 5–10 mg/L of DEC in most wines treated with the usual amount of DEDC. Prillinger (1964) verified the earlier work. Garschagen (1967) found that DEC could be measured by gas chromatography directly without extraction. A very sensitive headspace method that could detect DEC to 0.1 mg/L was suggested by Kunitake (1969). Wunderlich (1972) reported a collaborative study for the Association of Official Analytical Chemists. The method involved extracting the DEC and determining the amount with quantitative gas chromatography. McCalla et al. (1977) questioned the use of the 15% Carbowax 20 M column used for the separations. They found an unknown peak at the same retention time as DEC. Using headspace sampling and chromatography coupled to a mass spectrometer, Van Lierop and Nootenboom (1979) were able to measure as little as 1 g/L of DEC. The specificity of measurement was enhanced when the mass spectrometer was selectively tuned to direct M/E 63 and 91, the main fragments of DEC. The official method (Horwitz, 1980) is extraction with carbon disulfide, separation on the gas chromatographic column, and comparison of the DEC peak area with nonalcoholic foods or beverages. The DMDC reaction product with ethanol is ethyl methyl carbonate (EMC). Stafford and Ough (1976) reported a method to detect the amount of EMC formed and to relate it to the amount of DEDC added to an alcoholic beverage. They used a gas chromatographic method similar to that of Wunderlich (1972) for DEC measurement but used an internal standard (DEC) to allow more accurate determination. Bandion et al. (1979) reported a similar determination using glass capillary columns for better separation.

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Molin N., Satmark, L., and Thorell, M. 1963. Pyrocarbonic acid diethyl ester as a potential food preservative. Food Technol. 17:797. Moncelsi, E. 1970. Determinazione spettrofotometrica del dietile pircocarbonato con 4-amminoantipirina. Chem. Ind. (Milan) 52:637. Mönch, G. 1961. Pyrokohlens ure-Di thylester, ein neues Konservierungsmittel f r Bier? Brawissenshaft 14:257. Monteiro, F. F., Trousdale, E. K., and Bisson, L. F. 1989. Ethyl carbamate formation in wine: Use of radioactively labeled precursor to demonstrate the involvement of urea. Am. J. Enol. Vitic. 40:1. Morris, J. R., Main, G., and Threlfall, R. 1996. Fermentations: Problems, solutions, and prevention. Vitic. Enol. Sci. 51(3):210. Mount, J. R., Kasrai, S., and Draughon, F. A. 1999. Dimethyl dicarbonate and potassium sorbate effects on microbial growth in fresh salsa. Ann. Mtg. Inst. Food Technol., Chicago, IL, July 24–28, No.22D-9, p. 49. Mühlrád, A., Hegyi, G., and Tóth, G. 1967. Effect of diethylprocarbonate on proteins. I. Reaction of diethylpyrocarbonate with amino acids. Acta Biochim. Biophys. Acad. Sci. Hung. 2:19. Murata, A. 1974. Growth inhibition of Lactobacillus casei phage J1 by diethyl dicarbonate. J. Gen. Appl. Microbiol. 20:71. Osterman-Golkar, A., Ehrenberg, L., and Solymosy, F. 1974. Reaction of diethyl pyrocarbonate with nucleophiles. Acta Chem. Scand. 28B:215. Ough, C. S. 1976a. Ehthylcarbonate in fermented beverages and foods. I: Naturally occurring ethylcarbamate. J. Agric. Food Chem. 24:323. Ough, C. S. 1976b. Ethylcarbamate in feremented beverages and foods. II: Possible formation of ethylcarbamate from diethyl dicarbonate addition to wine. J. Agric. Food Chem. 24:328. Ough, C. S. 1978. A Davis scientist reviews and excellent fungicide, DEDC. Wine Vines 59(4): 30. Ough, C. S., and Ingraham, J. L. 1961. The diethylester of pyrocarbonic acid as a bottled-wine sterilizing agent. Am. J. Enol. Vitic. 12:149. Ough, C. S., and Langbehn, L. 1976. Measurement of methylcarbamate formed by the addition of dimethyl dicarbonate to model solutions and to wines. J. Agric. Food Chem. 24:428. Ough, C. S., Crowell, E. A., and Gutlove, B. R. 1988a. Carbamyl compound reactions with ethanol. Am. J. Enol. Vitic. 39:239. Ough, C. S. Crowell, E. A., and Mooney, L. A. 1988b. Ethyl carbamate precursors during grape juice fermentation. I. Addition of amino acids, urea and ammonia. Am. J. Enol. Vitic. 39:243. Ough, C. S., Kunkee, R. E., Vilas, M. R., Bordeu, E., and Huang, M. C. 1988c. The interaction of sulfur dioxide, pH and dimethyl dicarbonate on the growth of Saccharomyces cerevisiae Montrachet and Leuconostoc oenos. Am. J. Enol. Vitic. 39:279. Ough, C. S., Stevens, D., Sendovski, T., Huang, Z., and An, D. 1990. Factors contributing to urea formation in commercially fermented wines. Am. J. Enol. Vitic. 41:68. Parfentjev, L. N. and Kovaklenko, V. I. 1951a. The possible participation of pyrocarbon esters in forming champagne properties in sparkling wines. Vinodel. Vinograd. SSSR 11(3):16. Parfentjev, L. N., and Kovaklenko, V. I. 1951b. The theory of champagne fermentation. Vinodel. Vinograd. SSSR 113:17. Pátkay, G., Hergár , E., and Gyns, L. 1973. Szénsavas, Üdititalok tartósitása piroszénsavdietiléeszterrel. K. Paprikaipar. 38:95. Pauli, O., and Genth, H. 1966. Diethyl pyrocarbonate. I: Properties, effects, and analysis. Zeitschrift fuer Lebensmittel-Untersuchung und -Forschung 132:216–227. Pauli, O. 1969. Le pyrocarbonate d’ethyle, un nouveau produit pour la sterilisation des boissons. Spectre d’efficacit , utilisation, analyse, toxicologie et legislation. Anal. Falsif. Expert. Chim. 61:377. Pauli, G. H. 1984. Chemistry of food additives: Direct and indirect effects. J. Chem. Educ. 61:332. Paulus, W., and Lorke, D. 1967. Diethyl pyrocarbonate. III. Preparation and toxicological testing of the representative reaction products of diethyl pyrocarbonate. Zeitschrift fuer Lebensmittel-Untersuchung und -Forschung 132:325–333. Peterson T., and Ough, C. S. 1979. Dimethyldicarbonate reaction with higher alcohols. Am. J. Enol. Vitic. 30:119. Peterson, T. W. 1978. Reaction of dimethyldicarbonate with higher alcohols and water, M.S. thesis, University of California, Davis.

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Porter, L. J., and Ough, C. S. 1982. The effects of ethanol, temperature and dimethyl dicarbonate on viability of Saccharomyces cerevisiae Montrachet No. 522 in Wine. Am. J. Enol. Vitic. 33:222. Pound, A. W. 1967. Initiation of skin tumours in mice by homologous and N-substituted derivatives of ethylcarbamate. Aust. J. Exp. Biol. Med. Sci. 45:507. Pozsonyi, F. 1972. A dietilpirokarbon t alkalmaz s nak el yer az dit ital gy rt sban. Szeszipar 20:28. Prillinger, F. 1964. ber den Nachweis und die Bestimmung von Di thylkarbonat in mit Baycovin behandelten Weinen. Mitt. Rebe Wein Klosterneuburg 14A:29. Rankine, B. C. 1964. Di-ethyl pyro-carbonate. Int. Bottler Packer, December, p. 68. Rankine, B. C., and Pilone, D. A. 1974. Yeast spoilage in bottle table wine and its prevention. Aust. Wine Brew. Spirit Rev. 92(11):36, 38, 40. Rash, V. A. 2003. Physical and chemical treatments for control of Salmonella on cantaloupe rinds. M.S. thesis. University of Tennessee, Knoxville. Ringold, H. J. 1966. Proposed catalytic role for histidine in pyridine nucleotide-linked alcohol dehydrogenases. Nature 210:535. Santini, P., Moretton, J., and D’Aquino, M. 1985. Detection of the genetic toxicity of diethyl pyrocarbonate using bacterial systems. Rev. Latinoam. Microbiol. 27:157 [CA 103(19):159244z]. Schelenz, R., and Fischer, E. 1970. Untersuchungen mit 14C-markeitem Pyrokohlens ure-Di thylester. III. Zur hydrolyse and Alkoholyse. Z. Lebensm. Unters. Forsch. 145:279. Schelenz, R., and Fischer, E. 1971. Untersuchung mit 14C-markiertiem Pyrokohlens ure-Di thylester. IV. Reaktion mit dl-Apfels ure. Z. Lebensm. Unters. Forsch. 147:145. Schmahl, D., Port, R., and Wahrendorf, J. 1977. A dose response study of urethane carcinogenesis in rats and mice. Intl. J. Cancer 19:77. Schmidt-Lorenz, W. 1962. Uber die Anwendlbarkeit von Pyrokohlens ure-di thylester als Konzervierungsmittel f r Schlachtgefl gel. Z. Lebensm. Unters. Forsch. 117:231. Sharratt, M., Gaunt, I. F., Grasso, P., Kiss, I. S. Hooson, J., Wright, M. G., and Ganolli, S. D. 1972. Short-term toxicity studies on diethyl pyrocarbonate-treated beverages in the rat. Food Cosmet. Toxicol. 10:743. Shibasaki, I., Matsumoto, K., and Horie, H. 1969. Factors affecting the microbicidal activity of diethylpyrocarbonate. Nippon Shokuhin Kogyo Gakkai-Shi 16:405. Splittstoesser, D. F., and Wilkinson, M. 1973. Some factors affecting the activity of diethylpyrocarbonate as a sterilant. Appl. Microbiol. 25:853. Stafford, P. A., and Ough, C. S. 1976. Formation of methanol and ethylmethyl carbonate by dimethyl dicarbonate in wine and model solutions. Am. J. Enol. Vitic. 27:7. Terrell, F. R., Morris, J. R., Johnson, M. G., Gbur, E. E., and Makus, D. J. 1993. Yeast inhibition in grape juice containing sulfur dioxide, sorbic acid, and dimethyldicarbonate. J. Food Sci. 58(5):1132. Thoma, W., and Rinke, H. 1959. Synthesen mit Pyrokohlens ure-estern. Liebigs Ann. Chem. 624:30. Thoukis, G. 1983. Dimethyl dicarbonate as a beverage conservative. In Instrumental Analysis of Foods, edited by G. Charalamvous, Academic Press, New York, pp. 455–462. Thoukis, G. R. Bouthilet, J., Ueda, M., and Caputi, A., Jr. 1962. Fate of diethyl pyrocarbonate in wine. Am. J. Enol. Vitic. 13:105. Todor, T., Trifon, I., and Georgi, B. 1967. Effect of pyrocarbonic acid diethyl ester on some microorganisms in grape juice. Lozar. Vinar 15(3):36. Turtura, G. S. 1966. Ricerche sul potere microbicida dell’estere dietilico dell’acido pirocarbonico. Ric. Sci. 36:638. Uzvolgyi, E., Bojan, F., and Arany, I. 1983. In vitro and in vivo formation of carcinogens from diethyl pyrocarbonate in the presence of ammonia. Magy. Onkol. 27:165 [CA 99(25):211314n]. Uzvolgyi, E. 1986. Another possibility of development of carcinogenic substances from noncarcinogenic precursors occurring in foods. Egeszsegtudomany 30(1):32 [CA 105(3):2321w]. Van der Riet, W. B., Botha, A., and Pinches, S. E. 1989. The effect of dimethyldicarbonate on the vegetative growth and ascospores of Byssochlamys fulva suspended in apple juice and strawberry nectar. Intl. J. Food Microbiol. 8:95. Van Lierop, B. H., and Nootenboom, H. 1979. Gas-liquid chromatographic-mass fragmentographic determination of low levels of diethylcarbonate in beverages. J. Assoc. Off. Anal. Chem. 62:253. Van Zyl, J. A. 1962. The microbiology of South African wine making II. The preservation of musts and wines with pyrocarbonic acid diethyl ester. S. Afr. J. Agric. Sci. 5:293.

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Wang, C., Chen, Y., Xu, Y., Wu, J., Xiao, G., Zhang, Y., and Liu, Z. 2013. Effect of dimethyl dicarbonate as disinfectant on the quality of fresh-cut carrot (Daucus carota L.): Dimethyl dicarbonate used on freshcut carrot. J. Food Process Preserv. 37:751–758. Whitney, B. M., Williams, R. C., Eifert, J., and Marcy, J. 2008. High pressures in combination with antimicrobials to reduce Escherichia coli O157:H7 and Salmonella Agona in apple juice and orange juice. J. Food Prot. 71:820–824. Williams, R. C., Sumner, S. S., and Golden, D. A. 2006. Inactivation of Escherichia coli O157:H7 and Salmonella in apple cider and orange juice treated with combinations of ozone, dimethyl dicarbonate, and hydrogen peroxide. J Food Sci 70:M197–M201. Wolf, A., Hrivnak, D., and Machovcova. 1969. Beitrag zur Toxicit t des Dikohlens uredi thylesters. Z. Lebensm. Unters. Forsch. 139:287. Woo, Y. 1983. Carcinogenicity, mutagenicity and teratogenicity of carbamates, thiocarbamates and related compounds: An overview of structure-activity relationships and environmental concerns. Environ. Sci. Health C1:97. Worobo, R. W. 2002. Personal communication. Wucherpfenning, K. 1966. Moeglichkeiten des Einsatzes von Pyrokohlens uredi thylester zur haltbarmachung von Fruschs ften und seive Beurteilung hinsichtlich de Erhaltung der Naturreinkeit. Int. Fruchs ftunion Wiss. Tech. Komm. 7:49. Wunderlich, H. 1972. Collaborative study of a gas-liquid chromatographic method for the determination of diethylcarbonate in wine. J. Assoc. Off. Anal. Chem. 55:557. Zaitsev, A. N., Rakhamanina, N. L., and Dyubyuk, N. E. 1970. Hygenic study of PIREF and its possible use in the food industry. Vopr. Pitan. 29:14. Zuehlke, J. M., Glawe, D. A., and Edwards, C. G. 2015. Efficacy of dimethyl dicarbonate against yeasts associated with Washington State grapes and wines: Use of DMDC in winemaking. J. Food Process Preserv. 39:1016–1026.

14 Lysozyme T. Matthew Taylor, Eric A. Johnson, and Ann E. Larson CONTENTS 14.1 Background and General Properties of Lysozyme................................................................... 445 14.2 Properties.................................................................................................................................. 446 14.2.1 Chromatographic Elution Characteristics................................................................. 446 14.2.2 Basic Enzymatic and Hydrolytic Properties............................................................. 447 14.2.3 Common Properties amongst Differing Lysozyme Types........................................ 448 14.3 Uses and Stability in Foods....................................................................................................... 450 14.3.1 Thermal Stability in Solution and in Food Products during Heat-Processing......... 450 14.3.2 Lysozyme Stability to Non-Thermal Processing and Food Preservatives.................451 14.4 Antimicrobial Spectrum of Activity..........................................................................................452 14.4.1 Gram-Positive and Gram-Negative Bacteria Sensitivity to Lysozyme......................452 14.4.2 Lysozyme Sensitivity as a Function of Microbial Physiological Status....................453 14.5 Activity against Food-Related Microorganisms....................................................................... 454 14.5.1 Antimicrobial Activity of Lysozyme against Gram-Positive Rods in Foods........... 454 14.5.2 Antifungal Activity against Foodborne Yeasts and Spoilage Bacteria in Foods.......455 14.6 Effect on Heat Resistance of Bacterial Spores...........................................................................455 14.7 Enhancement of Activity by Other Chemical Agents............................................................... 456 14.7.1 Enhancement of Lysozyme by Chelators against Foodborne Bacteria..................... 457 14.7.2 Lysozyme Use in Combination with Antimicrobial Polypeptides and Bacteriocins.....458 14.8 Enhancement by Physical Processes......................................................................................... 459 14.9 Food Applications..................................................................................................................... 459 14.9.1 Functionality in Ready-to-Eat Further Processed Foods.......................................... 459 14.9.2 Lysozyme Utility on Animal Carcass and Derived Product Surfaces...................... 460 14.9.3 Lysozyme Applications for Food Contact Surfaces Disinfection............................. 461 14.10 Non-Enzymatic Antimicrobial Activity of Lysozyme and Lysozyme-Peptides....................... 461 14.10.1 Bactericidal Characteristics of Lysozyme................................................................. 461 14.10.2 Lysozyme-Derived Antimicrobial Peptides.............................................................. 461 14.11 Recombinant Lysozymes for Use as Food Antimicrobials....................................................... 462 14.12 Regulatory Status and Toxicology............................................................................................ 462 14.13 Summary and Perspectives....................................................................................................... 463 References............................................................................................................................................... 464

14.1 Background and General Properties of Lysozyme Lysozyme was discovered in 1921 by Alexander Fleming (1881–1955), who described it as a “remarkable bacteriolytic principle” (Fleming, 1922). His subsequent discovery of penicillin occurred in the late 1920s. Fleming’s lysozyme research emerged from his demonstration that chemical antiseptics were ineffective in treating infections. He showed that his own nasal mucus inhibited the growth of a Micrococcus spp., a fortuitous discovery since micrococci are among the most sensitive organisms to lysozyme. He first thought that the inhibitory factor was a bacteriophage, but later showed that it was 445

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an enzyme that lysed bacterial cells (Fleming, 1922). He established that lysozyme was an endogenous (innate) antimicrobial in the body, and his work supported the concept that an effective way to treat infections was to enhance the body’s own immune responses. Fleming found that lysozyme was present in nasal mucus and tears, and that hen egg white had a particularly high level of the protein. It is noteworthy that Laschenko (1909) and Rettger and Sperry (1912) showed that egg white was capable of causing lysis of certain strains of bacteria and spores. It was Fleming, however, who actually isolated the lytic agent and showed that it occurred in human secretions. Lysozyme was the first enzyme whose primary amino acid sequence was determined, and was also the first enzyme whose structure was determined by X-ray crystallography (Blake et al., 1965; Phillips, 1966). It has an ellipsoid structure with dimensions of approximately of 4.5 × 3.0 × 3.0 nm. Its catalytic activity was demonstrated in 1966 by John Rupley, who showed that lysozyme cleaved purified oligosaccharides of N-acetylglucosamine containing a β-(1-4) glycosidic bond (Rupley, 1967). Oligosaccharides with more than three sugar units were hydrolyzed, and the hexasaccharide was shown to be the optimum length substrate. Several families of lysozyme have been found, but they all share the characteristic property of cleaving a β-glycosidic bond between the C-1 of N-acetylmuramic acid and the C-4 on N-acetylglucosamine of bacterial peptidogylcan. Although traditionally associated with eggs of birds, especially those of domestic hens, lysozyme is widespread in nature and is found in many sources including certain vegetables, insects, plants, and fungi (Masschalck et al., 2001; Tranter, 1994; Jolles and Jolles, 1984). Lysozyme is present in human colostrum (Mathur et al., 1990) and mammalian tissues and fluids such as milk, saliva, mucus, blood, and tears. It is also present in high concentrations in macrophages, leukocytes, monocytes, and neutrophilic granulocytes. Lysozyme has important roles in the immune response of organisms in reaction to infections and inflammation (Osserman et al., 1974; Sava, 1996). The major families of naturally occurring lysozyme that differ markedly in amino acid primary structure and secondary structure: “C” for chicken or classical, “G” for goose, bacterial lysozyme (autolysins), phage lysozyme, and plant lysozyme (Jolles, 1996), but only the C enzyme from the hen egg whites is currently permitted for use in food preservation. Approximately 3.5% of the total protein content in hen egg white is lysozyme (Alderton et al., 1945; Sofos et al., 1998; Yamamoto et al., 1997). Lysozyme is present at concentrations of about 0.1, 0.13, and 0.25 µg/ml in milk from sheep, cows, and goats, respectively (Chandan et al., 1968), while human milk contains 0.4 µg/ml lysozyme (Reiter, 1978). Lysozyme is considered to be one of the most important factors of nonspecific immunity in human breast milk (Hennart et al., 1991). It also occurs at much higher concentrations in colostrum than in milk, and may have positive attributes on the intestinal flora of nursing infants (Barbara and Pellegrini, 1976). Lysozyme has been proposed for use in various clinical applications, including antibacterial, antiviral, and anti-inflammatory treatments in humans and animal species (Sava, 1996; Proctor and Cunningham, 1988). As described in detail in the following sections, lysozyme is also used to control microbial growth in foods such as cheese and wine, and has potential uses as a preservative in other food systems. Lysozyme serves as a good model of an ideal food preservative in many respects, since it is an innate component of the human immune system and thus would be expected to have low toxicity, it is an enzyme that acts catalytically and can be used at low concentrations in foods, it is specific for bacterial peptidoglycan and does not react with human tissue, and it has certain desirable resistance properties to heat, reduced pH, and other intrinsic and extrinsic factors of foods.

14.2 Properties 14.2.1 Chromatographic Elution Characteristics Early researchers described various methods for the purification and assay of lysozyme from egg white (Alderton et al., 1945; Meyer et al., 1936). Purified commercial lysozyme for food use is currently produced from hen egg whites in high yield by cation exchange resins. Its high affinity to cation resins is due to the basic character of the enzyme, its isoelectric point of 10.5–11, and its monomeric structure and molecular weight of ~14,400 Da (Cunningham et al., 1991). Lysozyme is eluted from the resins by NaCl,

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and is recovered as the hydrochloride salt. In addition to removal of lysozyme from egg albumin, the process also removes more than 90% of the avidin from eggs. The extracted egg whites are not nutritionally or functionally changed, and are approved for food uses. Lysozyme is generally used as the hydrochloride salt, which is freely soluble in water and buffers. Lysozyme hydrochloride appears as a white, fine powder and has a slightly sweet taste. A 2% lysozyme hydrochloride solution has a pH of about 3.3. It is insoluble in most organic salts and concentrated salt solutions, but regains activity when transferred back to an aqueous solution. Depending on the pH conditions, lysozyme can polymerize (Sophianopoulos and Van Holde, 1964). Basic properties of lysozyme are described in Table 14.1.

14.2.2 Basic Enzymatic and Hydrolytic Properties Lysozyme (E.C. 3.2.1.17) is classified as a mucopeptide N-acetylmuramyl hydrolase by the Commission on Enzymes (Connor, 1993) and is commonly referred to as muramidase (Proctor and Cunningham, 1988). The primary amino acid sequence and structure as determined by X-ray crystallography of hen egg-white lysozyme (HEWL) has been described in detail (Blake et al., 1965; Phillips, 1966; Jolles et al., 1963; Rees and Offord, 1972). Egg-white lysozyme is a single polypeptide chain of 129 amino acids cross-linked by four disulfide bridges (Jolles et al., 1963; Jolles, 1996; Phillips, 1966) with a molecular weight of approximately 14,400 Da (Barbara and Pellegrini, 1976; Jolles and Jolles, 1984). Enzymatic activity of the molecule is lost if at least two of the disulfide bridges are not intact, or if all disulfide bonds are reduced (Proctor and Cunningham, 1988; Wang and Shelef, 1992). Lysozyme has a hydrophobic core with hydrophilic amino acid side chains toward the surface, giving the molecule stability (Proctor and Cunningham, 1988). The lytic activity of lysozyme has traditionally been measured by observing lysis of UV-killed and lyophilized Micrococcus luteus (M. lysodeikticus) cells based on the early observations of Fleming, and this assay has been optimized by Shugar (Fleming, 1922; Parry et al., 1969; Shugar, 1952). Numerous variations for assay of lysozyme by turbidimetry have been reported (detailed in Proctor and Cunningham, 1988). One unit of lysozyme is defined as the smallest quantity leading to complete lysis of Micrococcus luteus in a serial dilution test (Alderton et al., 1945). Specifically, one Shugar unit is that quantity of enzyme in 1 ml of a suspension of Micrococcus luteus inactivated cells at pH 7.0, with an initial absorbance of 0.750 at 450 nm in a pathlength of 1 cm, which causes the absorbance to decrease at a rate of 0.001 per minute. The maximal specific activity of lysozyme at 25°C using M. luteus as the substrate generally is ~50,000 units/mg, but can be much less (~20,000 units/mg) depending on the TABLE 14.1 Properties and Specifications of Food-Grade Lysozyme Hydrochloride for Use in the United Statesa a. GRAS substance: Egg-white lysozyme (CAS Reg. No. 9001-62-2) obtained by extraction of egg whites. b. Enzyme property: Peptidioglycan N-acetylmuramoylhydrolase (EC No. 3.2.1.17) that catalyzes the hydrolysis of peptidoglycan in the cell walls of certain bacteria. The ingredient is used as an enzyme as defined in §170(o)(9) in 21 CFR §184. c. Primary food application: Cheeses as defined in 21 CFR, §51, Sec. 170.3 (n) (5), in accordance with §184.1(b)(3). The primary enzyme target is germinating spores of Clostridium tyrobutyricum, an organism responsible for late blowing of certain cheese varieties. d. Use level: Levels not to exceed current good manufacturing practices in cheese. e. Label requirement: Bulk and packaged foods that contain cheese manufactured using egg-white lysozyme shall include the usual name “egg-white lysozyme” on their ingredient labels. f. Commercial ingredient: Food-grade lysozyme hydrochloride. g. Food use properties: Freely soluble in water; pH 3.3; electrophoretically pure. h. Limit specifications: Satisfies limit tests and regulations for heavy metals, arsenic, and microbial limit tests. According to IDF (1987) salmonellae absent in 25 g; Staphylococcus aureus, Pseudomonas aeruginosa, E. coli, sulfite-reducing clostridia absent in 1 g. Coliforms max 30/g. a

Sources include: International Dairy Federation 1987; U.S. FDA 1998; www​.fordras​.com; www​.inovatech​.com. For regulatory information for other countries, the manufacturer is advised to contact the major manufacturers of lysozyme, whose websites are listed above.

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lysozyme preparation. From a practical perspective, it is often desirable to use a wavelength of 650 nm to reduce absorption by colored substances in food extracts. Certain compounds can inhibit the lysozyme assay including surfactants such as sodium dodecyl sulfate (SDS), iodine, and fatty acids and alcohols of C12 or higher (Smith and Stroker, 1949). Several alternate methods of assay have been developed including chromatographic separations, agar plate lysis, radioimmunoassays, and fluorescence systems, but these methods are generally only used for special purposes and the M. luteus lysis method is the standard (Proctor and Cunningham, 1988; Jolles and Jolles, 1984). A sensitive reverse-phase HPLC detection method for lysozyme was recently developed for use in milk and cheese samples (Pellegrino and Tirelli, 2000). The optimum pH for lysis of Micrococcus lysodeikticus in vitro for egg-white lysozyme is ~6.6 (Barbara and Pellegrini, 1976). Enzymatic activity is observed over the pH range of 3.5–7.0 (Wang and Shelef, 1992), although lower activity has been observed at pH values 7.0 (Smolelis and Hartsell, 1952; Yang and Cunningham, 1993). The optimum pH for activity is strongly dependent on salt concentration (Chang and Carr, 1971; Davies et al., 1969). At pH values 10 krad gamma-irradiation at ambient temperature in phosphate buffer at pH 6.2, although inactivation was concentration-dependent (Eitenmiller et al., 1971). Inactivation of lysozymes by gamma-irradiation was due in part to hydroxyl radicals (Eitenmiller et al., 1971). Kume et al. (1973) observed a 37% inactivation of lysozyme activity in aqueous solution by 0.35 Mrad gamma-irradiation, although only slight inactivation of lysozyme occurred in irradiated egg white treated under the same conditions, possibly due to protection by high protein levels in the egg white. Results of another study showed that both purified lysozyme in buffer and endogenous lysozyme in egg white were not significantly affected by 0.195 kGray gamma-irradiation at 60°C (Schaffner et al., 1989). Gamma-irradiation at 20 kGy polymerized lysozyme through covalent bonds, especially in the presence of peroxidizing lipids and at higher water activity levels (Kanner and Karel, 1976). Lysozyme activity can be increased by physical processes used in many food preservation systems. Freezing and thawing of Escherichia coli caused sensitivity to lysozyme (Kohn, 1960; Ray et al., 1984), although the organism regained resistance after incubation at 37°C or exposure to calcium (Ray et al., 1984). Transmission electron microscopy demonstrated protoplast formation and cell wall damage in unfrozen or frozen/thawed L. monocytogenes treated with lysozyme (El-Kest and Marth, 1992). The sensitization of Gram-negative cells to lysozyme by high pressure is dependent on factors such as pressure, temperature, pH, medium, growth stage, and bacterial species (Masschalck et al., 2001). High pressure has been shown to sensitize E. coli to lysozyme (Garcia-Graells et al., 1999; Masschalck et al., 2000), although the sensitization is transient (Hauben et al., 1996; Masschalck et al., 2001). In addition, four of six Gram-negative bacteria tested were sensitive to lysozyme during treatment by high hydrostatic pressure (Masschalck et al., 2001). EDTA increased the activity of lysozyme against E. coli during high-pressure treatment (Hauben et al., 1996). However, exposure of E. coli to lysozyme or lysozymeEDTA immediately after high-pressure treatment did not decrease the number of viable cells (Hauben et al., 1996). Pulses of high hydrostatic pressure may sensitize some bacterial strains more effectively to lysozyme than treatment with continuous pressure (Masschalck et al., 2001). Sublethal injury by high pressure might be a useful hurdle along with lysozyme or lysozyme-EDTA in certain foods. Lower pressures required to inactivate certain bacteria in the presence of lysozyme (Masschalck et al., 2001) may lead to new combination treatments for the non-thermal preservation of foods. Similarly, pulsed electric field (PEF) was enhanced in its anti-S. aureus efficacy in skim milk when milk was first treated with 5.0 IU/mL nisin and 3000 IU/mL lysozyme in milk adjusted to pH 5.0, yielding a 4.8 log10-cycle reduction as compared to other treatments. PEF application time was also a key factor in determining S. aureus reductions observed (Sobrino-López and Martín-Belloso, 2008). Lysozyme retains activity in the presence of many common components and compounds inherently present in or added to foods. Activity of lysozyme was not affected by the presence of certain antimicrobial compounds used in foods, including sodium nitrite, ethanol, butylated hydroxyanisole, calcium propionate, potassium sorbate, and propyl paraben (Yang and Cunningham, 1993). Potassium dichromate had no effect on lysozyme activity in mastitic bovine milk (Weaver and Kroger, 1978). Some organic solvents used in foods, such as ethanol, do not denature lysozyme (Sofos et al., 1998). Certain chemicals which may be present in foods can have a detrimental effect on lysozyme activity. Lysozyme can bind to certain food components, leading to a loss in activity (Cunningham and

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Lineweaver, 1967; Sofos et al., 1998). Lysozyme may be less heat-stable in proteinaceous foods than in vitro, due to the formation of mixed disulfide-linked molecules or other interactions with proteins or other constituents in the food (Johnson, 1994). Purified lysozyme in buffer at pH 6.2 was inactivated by egg yolk (Cunningham and Cotterill, 1971), possibly due to electrostatic interaction between lysozyme and substances in the yolk (Proctor and Cunningham, 1988). Activity was lost in the presence of high concentrations of lactic acid, acetic acid, and chlorine (Yang and Cunningham, 1993). Lysozyme can also be inhibited by substances such as alkyl sulfates, aliphatic long chain alcohols, and fatty acids and their sodium salts (Smith and Stroker, 1949). Polysaccharides with carboxylic acid and sulfonic acid groups, such as pectin and alginate, also reduced lysozyme activity, although the addition of 1–6% salt minimized inactivation by these compounds (Yashitake and Shinichiro, 1977). High concentrations of polyvalent and divalent cations such as cobalt, manganese, mercury, and copper can inactivate lysozyme, as well as low concentrations of iodine (Proctor and Cunningham, 1988). Salt level plays a critical role in the enzymatic activity of lysozyme. Non-specific activation occurs at low salt concentrations (Tranter, 1994; Chang and Carr, 1971), although lysozyme activity is inhibited at high salt concentrations greater than 0.05–0.1 M (Chang and Carr, 1971; Davies et al., 1969; Smolelis and Hartsell, 1952), at least partially through loss of enzyme solubility and formation of crystalline structures and/or precipitation from aqueous solution (Galm et al., 2015). The addition of 3 mM NaCl improved the lysis of rapidly growing sensitive organisms by lysozyme (Vakil et al., 1969). Inhibition of lysozyme activity in high ionic strength solutions was stronger at alkaline pH values (Davies et al., 1969). The presence of sodium nitrate (180 mg/L) or sodium chloride (270 mg/L) did not inhibit the activity of lysozyme (450 mg/L) against several Gram-positive and Gram-negative bacteria under non-growthrestricted conditions (Gill and Holley, 2002).

14.4 Antimicrobial Spectrum of Activity Lysozyme exhibits antimicrobial activity against vegetative cells of a wide variety of organisms, including numerous foodborne pathogens and spoilage organisms. Egg-white lysozyme is generally most active against certain Gram-positive organisms, and ineffective against dormant bacterial spores and Gram-negative bacteria. Antimicrobial activity of the enzyme can vary widely within groups of similar organisms, and its lytic activity is dependent on growth medium, test conditions, age of cells, presence of interfering substances, and many other parameters.

14.4.1 Gram-Positive and Gram-Negative Bacteria Sensitivity to Lysozyme Gram-positive bacteria vary markedly in their sensitivity to lysozyme depending on the strain, species, and conditions of microbe handling and testing. Spoilage of foods by Gram-positive organisms including the heterofermentative LAB, staphylococci, Brochothrix, and certain other Gram-positives is a major problem in the food industry, but these groups of organisms are mostly resistant to the enzyme. Species of the Gram-positive genera Micrococcus and Sarcina are generally more sensitive to lysozyme than are Lactobacillus and Bacillus species (Connor, 1993). Ashton et al. (1975) demonstrated that the prevention of growth of Geobacillus. stearothermophilus and Paenibacillus coagulans by egg albumin was due to the presence of lysozyme. Human lysozyme inhibited S. aureus (Maga et al., 1998), but the egg white enzyme is generally inactive by itself against S. aureus. Lysozyme susceptibility among species of Gram-positive bacteria may decrease by derivatization of the amino groups and may vary due to differences in the relative proportions of N-acetylamino sugars in the cell walls (Salton and Pavlik, 1960). Specifically, higher proportions of β-1-6 and β-1-3 linkages between N-acetylmuramic acid and N-acetylglucosamine are found in the peptidoglycan of some Gram-positive bacterial species, which are more resistant to lysozyme than β-1-4 linkages. These differences in cell surface structure contribute to lysozyme susceptibility amongst the Gram-positives (Vakil et al., 1969). Bacterial spores can be sensitized to lysozyme by strong reducing agents or oxidizing agents to rupture disulfide bonds (Gould and Hitchins, 1963), but these methods are not compatible with food use (Gould, 2002). Recent research, nevertheless, has demonstrated spores of the gastrointestinal pathogen Clostridium difficile

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to be sensitive to combined application of egg-white lysozyme and nisin in synergistic fashion (Chai et al., 2016). Gram-negative bacteria are generally less sensitive than Gram-positive bacteria to lysozyme due mainly to protection of the cell wall by the outer membrane. Resistant organisms include the important pathogens Salmonella spp. and E. coli O157:H7. However, certain physiological conditions, physical treatments, or chemicals that disrupt or alter the outer membrane can enhance the activity of lysozyme against Gram-negatives. At pH 3.5, most Gram-negative bacteria tested in one study exhibited some sensitivity to lysis by lysozyme, especially Salmonella and Brucella species (Peterson and Hartsell, 1955). Ibrahim (1998) reported enhanced antimicrobial activity of heat-denatured lysozyme against Gram-negative bacteria. In another study, human and bovine milk lysozymes displayed activity against five Gram-negative bacteria, including some strains of Gram-negative bacteria that are resistant to eggwhite lysozyme (Vakil et al., 1969), demonstrating variability in antimicrobial activity between lysozymes isolated from different sources. The susceptibility of various species of food-related organisms, summarized in Table 14.2, represents the authors’ assessment of the available literature. However, as discussed throughout this chapter, resistant organisms can often be made sensitive by physical treatments or chemical agents that disrupt outer membrane layers and capsules, allowing penetration of lysozyme and consequent lytic activity. Thus, in evaluating the potential for lysozyme function as a preservative in foods, a systems-based approach is often desirable whereby combinations of physical treatment or chemical agents are used in combination with lysozyme. In recent years, researchers have explored opportunities for increasing the spectrum of antimicrobial activity of lysozyme against Gram-negative genera through multiple means, including thermal or chemical modification of lysozyme, or incorporating lysozyme into some form of carrier system. For example, a lysozyme–dextran conjugate produced a consistent 2.0 log10-cycle decrease in E. coli numbers in a buffer compared to native, non-modified lysozyme when tested at 100, 250, or 400 µg/mL. When applied in cheese curd aged 39 days at 4°C, E. coli numbers treated by the dextran–lysozyme (400 µg/mL lysozyme) conjugate were >3.0 log10 CFU/g less than non-treated controls at day 39, and ~2.0 log10 CFU/g less than numbers of the same organism treated by native lysozyme (Amiri et al., 2008). Similar findings have been reported for lysozyme conjugated with polysaccharides such as xanthan and guar gum, wherein conjugates demonstrated enhanced antimicrobial activity against Gram-negative bacteria, including E. coli and Salmonella, versus native egg-white lysozyme (Hamdani et al., 2018; Hashemi et al., 2014). Thermo-chemical modified lysozyme (egg white), treated with 1 or 2% hydrogen peroxide and heating at 70°C produced significant improvement in the inhibition of Pseudomonas fluorescens versus nonmodified lysozyme during 24 h of incubation at 37°C (Cegielska-Radziejewska et al., 2010). However, populations of the organism recovered to non-differing numbers at 48 h of incubation. Exposure to conjugate egg-white lysozyme and the antibacterial phenolic triclosan caused a 6.0 to almost 10.0 log10 CFU/ mL decrease in Pseudomonas aeruginosa, Salmonella Typhimurium, Klebsiella pneumoniae, or E. coli depending upon the organism tested (Hoq et al., 2008).

14.4.2 Lysozyme Sensitivity as a Function of Microbial Physiological Status Sensitivity of organisms to lysozyme may be dependent on physiological state. Most studies examining the effectiveness of lysozyme have tested cells in nutrient-deficient environments. Cell wall synthesis in rapidly growing cultures may exceed the rate of degradation by lysozyme (Hughey and Johnson, 1987). Lysozyme–EDTA combinations were less effective against rapidly growing Gram-negative organisms in broth media than in buffer (Gill and Holley, 2002). The antimicrobial effect of lysozyme and nisin against lactic acid bacteria was sometimes enhanced when the growth media was diluted (Chung and Hancock, 2000). Inhibition of growth of L. monocytogenes in culture media by lysozyme at 5°C and 25°C increased when the pH was lowered from 7.2 to 5.5, although lysis of non-growing cells in buffer by lysozyme was not increased by a reduction in pH (Johansen et al., 1994). In that study, lower pH may have inhibited growth of the organism enough to allow the rate of hydrolysis by lysozyme to exceed the growth rate (Johansen et al., 1994). Heat-treated cells of Yersinia enterocolitica grown at 4°C were resistant to lysozyme, but heat-treated cells grown at 37°C were sensitive (Pagan et al., 1999). However, the activity of lysozyme against Gram-positive organisms was enhanced by EDTA under

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non-growth-restricted conditions (Gill and Holley, 2002). During evaluation of the antimicrobial efficacy of lysozyme and lysozyme–antimicrobial combinations, testing of both growing and non-growing cells may be advantageous.

14.5 Activity against Food-Related Microorganisms In the following section, selected studies of the effects of lysozyme on food-related organisms are described. Salmonella Senftenberg was not detected on inoculated turkey drumsticks treated with 0.1% lysozyme for 3 hours at 22°C, or 0.05% lysozyme for 1 minute at 63°C (Teotia and Miller, 1975). The heat resistances of Salmonella serovars Enteritidis and Typhimurium, E. coli O157:H7, and S. aureus were significantly reduced in buffer in the presence of eggshell membranes containing lysozyme (Poland and Sheldon, 2001). In this system, components of the albumin may have enhanced the activity of enzyme against these organisms.

14.5.1 Antimicrobial Activity of Lysozyme against Gram-Positive Rods in Foods The antimicrobial activity of lysozyme has been extensively tested against L. monocytogenes, a Grampositive pathogen of great concern to the food industry. Lysozyme (>2 mg/ml) prevented growth of L. monocytogenes in culture media at pH 7.0, and 2 mg/ml had listericidal effects at pH 9.0 (Wang and Shelef, 1991). Lysozyme was bactericidal to L. monocytogenes in milk, mozzarella, taleggio, and grana cheeses, frankfurters, and on the surface of chickens (Dell’Acqua et al., 1989). Poor survival of the organism in egg white (Sofos et al., 1998; Erickson and Jenkins, 1992) is probably due to the presence of lysozyme. Lysozyme was bactericidal to L. monocytogenes in egg white-containing cholesterol-free mayonnaise (Erickson and Jenkins, 1991) and may be partially responsible for the rapid inactivation of E. coli O157:H7 in commercial mayonnaise made with whole eggs (Raghubeer et al., 1995). Pretreatment of raw cod-fish fillets with 3 mg/ml lysozyme delayed growth of L. monocytogenes at abuse temperatures, and was listericidal at refrigeration temperatures, although it did not affect growth of natural microflora in the product (Wang and Shelef, 1992). Egg-white lysozyme alone (100 ppm) killed or prevented growth of L. monocytogenes in several foods, particularly vegetables (Hughey et al., 1989). Lysozyme in Camembert cheese initially was bactericidal to L. monocytogenes, although the organism eventually grew in the cheese after 35 days (Hughey et al., 1989). Listeria monocytogenes cells grown at low temperatures were more sensitive to lysozyme than cells grown at ambient temperature (Johnson, 1994) or 37°C (Smith et al., 1991), indicating lysozyme may be useful for controlling the organism in refrigerated foods. Milk pasteurization in combination with added lysozyme may eliminate L. monocytogenes in certain cheeses prepared from milk contaminated with the organism. Numbers of heat-stressed L. monocytogenes significantly decreased in Cheddar and Camembert-type cheeses containing lysozyme (Johnson, 1994). Conversely, growth of Listeria monocytogenes was not inhibited on frankfurters formulated with 0.01% lysozyme (Bedie et al., 2001). Mangalassary et al. (2008) treated turkey bologna surfaces with 0.5 U lysozyme/cm2 prior to a thermal intervention, which reduced numbers of L. monocytogenes on the deli meat surfaces to non-detectable counts during subsequent refrigeration storage. Lysozyme had limited effectiveness against L. monocytogenes in pork sausage (Hughey et al., 1989) and pork, beef, or turkey frankfurters (Johnson, 1994). L. monocytogenes was resistant to hen egg-white lysozyme in milk (Hughey et al., 1989; Kihm et al., 1994), but sensitive to the enzyme in media or buffer, or in mineralized milk during heating (Kihm et al., 1994). Mineral or mineral-associated components in milk may protect the organisms from inactivation by lysozyme and heat in milk, probably by increasing the stability of the cell surface (Kihm et al., 1994). Egg-white lysozyme was antimicrobial to C. botulinum in foods such as turkey, pork sausage, salmon, asparagus, potatoes, tomatoes, and mushrooms (Johnson, 1994; Sofos et al., 1998). Lysozyme delayed botulinum toxin production in bratwurst and salmon, as well as cod suspensions (Johnson, 1994). Lysis of C. botulinum by lysozyme in culture medium was enhanced at lower temperatures and pH (Johnson, 1994).

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The effect of lysozyme against clostridial species has been well-documented. Spores of C. tyrobutyricum survive milk pasteurization and cause “late blowing” of cheeses such as Edam and Gouda during ripening, which results in cracks or fissures in the finished cheeses. Lysozyme lyses C. tyrobutyricum cells as they outgrow from germinated spores and can prevent blowing of cheeses through lactate utilization by the organism (Carini and Lodi, 1982). Late blowing by sporeformers in Edam cheese prepared with low-quality milk (i.e., high sore load) was inhibited by 500 units/ml lysozyme (Wasserfall et al., 1976). Growth of C. tyrobutyricum in Italian cheeses was also inhibited by 50 ppm lysozyme or nontoxic lysozyme salts added to butter or cheese (Ferrari and Dell’Acqua, 1979). Vegetative cells of C. tyrobutyricum can develop resistance to lysozyme (Wasserfall and Teuber, 1979). However, lysozyme prevented the growth of the organism in Edam cheese, even when spores from lysozyme-resistant vegetative cells were used to contaminate the cheese milk (Wasserfall and Prokopek, 1978). Lysozyme inhibition of LAB members, including many starter cultures used in foods as well as spoilage organisms, is variable among species. Lysozyme in milk is usually more inhibitory to pathogenic and spoilage organisms than LAB (Cunningham et al., 1991). In culture media, lysozyme had no effect on the growth of Gouda cheese starter cultures (Bester and Lombard, 1990). Another study demonstrated inhibition of one Streptococcus starter culture from semi-hard cheese (Wasserfall, 1978). Spoilage lactobacilli were inhibited by 20 ppm lysozyme in sweetened sake (Uchida et al., 1972). Lactobacillus delbrueckii subsp. lactis was sensitive to >0.5 mg/l lysozyme in culture media, though susceptibility of the organism to the bacteriocin acidocin was reduced in the presence of lysozyme (Chumchalova et al., 1998). The application of lysozyme did not affect growth of LAB or yeasts on ready-to-use pear cubes, demonstrating that the action of lysozyme may be affected by substrate (Pittia et al., 1999).

14.5.2 Antifungal Activity against Foodborne Yeasts and Spoilage Bacteria in Foods Chitinase activity exhibited by lysozyme is probably responsible for the weak activity of lysozyme against several pathogenic and spoilage yeasts, including Candida albicans, Cryptococcus neoformans, Saccharomyces cerevisiae, and Zygosaccharomyces rouxii (Johnson, 1994). Lysozyme inhibits growth of spoilage lactic acid bacteria, and thermo-modified lysozyme was shown by researchers to inhibit wine-spoiling acetic acid bacteria (AAB) members, including Acetobacter aceti and Gluconobacter oxydans at contents ranging from 0.01 to 0.09 mg/mL following thermo-modification (Carillo et al., 2014; Gerbaux et al., 1997).

14.6 Effect on Heat Resistance of Bacterial Spores The safety of minimally processed, extended shelf-life foods, including sous-vide and cook-chill products, is usually due to a minimal heat treatment to achieve a 6-log reduction in heat-sensitive nonproteolytic Clostridium botulinum spores, followed by refrigeration below 10°C (Gould, 1999). However, heat treatment guidelines at 65–95°C do not usually take into consideration the effect of lysozyme on spore heat resistance (Fernandez and Peck, 1999). Lysozyme, which is naturally present in many foods, is relatively heat-resistant and may be active in foods after mild heat processes such as pasteurization (Peck et al., 1992). At concentrations as low as 0.1 µg/ml (Peck et al., 1992) lysozyme increases the apparent heat resistance of non-proteolytic Clostridium botulinum spores (Alderton et al., 1974; Fernandez and Peck, 1999). Increased heat resistance of proteolytic type A C. botulinum spores has been shown, although the effect was minimal compared to the effect on type E nonproteolytic spores (Alderton et al., 1974). Lysozyme also increased the recovery of heat-treated spores of C. perfringens (Adams, 1974), and possibly Bacillus cereus (Blocher and Busta, 1982), but not Bacillus thuringiensis (Faille et al., 1999). Approximately 0.1– 20% of nonproteolytic C. botulinum spores are permeable to lysozyme, resulting in a biphasic survival curve upon heating (Peck et al., 1993). Lysozyme in the recovery medium is thought to compensate for a heat-damaged germination system (Broda et al., 1998) in lysozyme-permeable heat-sensitive bacterial spores (Peck et al., 1992), probably by degrading the spore cortex, allowing core hydration, and inducing spore germination (Gould, 1989).

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Vegetable juice extracts with measurable lytic activity when added to the recovery medium increased the measurable heat resistance of nonproteolytic C. botulinum (Stringer and Peck, 1996). In other research, heating in a meat medium resulted in a 6-log reduction in numbers of viable nonproteolytic C. botulinum in the absence of lysozyme, but not with lysozyme present (Peck and Fernandez, 1995). It is unknown whether the unusually high measured heat resistance of type E C. botulinum in Dungeness crab meat (Peterson et al., 1997) was due to the presence of lysozyme in the product. A combination of lysozyme with high-pressure processing increased the lethality against B. cereus spores in milk during post-process aging and cheese manufacture (López-Pedemonte et al., 2003). Polypeptides produced from egg-white lysozyme also were reported to reduce Bacillus subtilis spores to non-detectable counts at levels of 10 up to 100 μg/mL (Abdou et al., 2007). Predictive models for the inactivation of nonproteolytic C. botulinum in minimally processed refrigerated foods in the presence of lysozyme were developed by Fernandez and Peck (1999). Because inadequate heat treatments may lead to subsequent growth of the organism during refrigerated storage, additional research was needed to verify the ability of currently used heat treatments to adequately kill nonproteolytic C. botulinum in such foods containing lysozyme or similar lytic enzymes. Stochastic modeling of B. cereus growth inhibition in liquid medium at 16 or 25°C when supplemented with nisin (0.13 μM) and 0.66 mM lysozyme produced significant extensions in the lag phase of vegetative cells of the pathogen as compared to controls ranging anywhere from 3.25 to 13 h (Antolinos et al., 2011).

14.7 Enhancement of Activity by Other Chemical Agents The outer membrane of most Gram-negative bacteria protects the cell wall from lysozyme and may limit the effectiveness of lysozyme in food applications (Gill and Holley, 2002; Davidson and Harrison, 2002). The antimicrobial effect of lysozyme, particularly against Gram-negative bacteria, can often be enhanced by the presence of specific chemicals or physical treatments (Table 14.3). The sensitivity of TABLE 14.3 Physical Treatments and Chemical Agents Reported to Increase the Activity of Lysozyme against Selected Organisms Physical Treatments Heat Alkaline pH Osmotic shock Freeze-thawing High pressure Chemical Treatments EDTA, DTPA, other divalent metal chelators Trisodium phosphate Cysteine, proline Conalbumin Lactoferrin, other transferrins Lysolecithin, polyethylene glycol, other surfactants. Glycine Poly-L-lysine Nisin, possibly other bacteriocins Hydrogen peroxide Ascorbic acid Lactic acid Trypsin, proteinase K Lipase Carvacrol, nerolidol, certain other terpenoids

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Gram-negative species to lysozyme is increased by chelators, certain antibiotics, amino acids, alkaline pH, osmotic shock, drying, and freeze-thawing (Tranter, 1994; Proctor and Cunningham, 1988), as well as hydrogen peroxide and ascorbic acid (Miller, 1969). Lysozyme activity is also enhanced by food additives including EDTA, butyl paraben, and tripolyphosphate, as well as naturally occurring antimicrobials such as conalbumin, lactoferrin, nisin, and transferrin (Johnson, 1994). Other food components promoting lysozyme activity include lactic acid, trypsin, proteinase K, and lipase (Sofos et al., 1998).

14.7.1 Enhancement of Lysozyme by Chelators against Foodborne Bacteria Many chemical agents can permeabilize the outer membrane of Gram-negative bacteria, as classically shown by the metal chelator EDTA. Early studies demonstrated that Versene (EDTA) could increase the lytic activity of lysozyme against certain Gram-negative bacteria (Repaske, 1958). Chelators such as EDTA may remove stabilizing divalent cations (e.g., Mg+2, Ca+2) from lipopolysaccharide in the outer membrane, increasing permeability of the cells to antimicrobial agents, including lysozyme (Hancock, 1984; Lam et al., 2014). The combination EDTA-Tris-lysozyme was bactericidal to eight of ten Gram-negative bacterial pathogens tested in buffer, and two of six Gram-positive bacteria tested, including L. monocytogenes (Wooley and Blue, 1974). The presence of EDTA increased the activity of lysozyme against C. botulinum and L. monocytogenes (Hughey and Johnson, 1987) as well as B. subtilis and P. aeruginosa (Vakil et al., 1969). Lysis of coliforms implicated in bovine mastitis by bovine sera was markedly increased upon addition of lysozyme and EDTA, although lysozyme alone did not lyse the cells (Carroll, 1979). Growth of predominantly Gram-negative shrimp microflora in nutrient broth was slightly delayed by 150 µg/ml lysozyme, but strongly inhibited by >50 µg/ml lysozyme in the presence of 0.02% EDTA (Chander and Lewis, 1980). Lysozyme alone was shown to inhibit only 2 of 12 fungal species tested, although all but 1 species were inhibited by a lysozyme–EDTA combination (Razavi-Rohani and Griffiths, 1999). The potentiation of lysozyme activity by EDTA has also been demonstrated in food systems, and research has continued until the present. Somewhat contrary to others’ findings, Ko et al. (2008) did not report the addition of EDTA to be beneficial in the inhibition of L. monocytogenes growth in cooked hams, though a combination of ovotransferrin + lysozyme did produce a reduction in populations of the pathogen over post-lethality storage. Similar results were reported for the inhibition of E. coli O157:H7 populations on pork chops, though the blend of ovotransferrin with both EDTA and lysozyme proved best at reducing pathogen growth, with reductions of approximately 8.0 log10-cycles as compared to the use of only EDTA and lysozyme (Ko et al., 2009). Similarly, Pseudomonas numbers on pre-cooked chicken treated by 1.5% w/w of EDTA and lysozyme, stored under vacuum, were 3.0 log10-cycle lower compared to control samples not treated with antimicrobials and stored aerobically (Ntzimani et al., 2010). Ostrich meat patty counts of aerobic and lactic acid bacteria stored refrigerated aerobically or under vacuum were significantly reduced by the application of EDTA (20 mM), lysozyme, and nisin (250 ppm each), with lactic bacteria being reduced to a greater extent versus controls as compared to total aerobic loads (Mastromatteo et al., 2010). Bevilacqua et al. (2010) also reported 1.0 g/L lysozyme and 100 mM EDTA produced rapid inhibition of Hafnia alvei in medium. Most recently, pseudomonads were reported to be inhibited via application of silver-based nanoparticles bearing lysozyme and EDTA when applied in burrata-style cheese over 12 days’ storage under modified atmosphere storage (Costa et al., 2017). The effectiveness of lysozyme against C. botulinum can be increased using chelators such as EDTA, DTPA, and cysteine (Johnson, 1994). EDTA, nisin, butyl-paraben, and tripolyphosphate increased the delay of botulinal toxin production by lysozyme in bratwurst, salmon, and cod suspensions (Johnson, 1994). Toxin production by Clostridium botulinum was also delayed in turkey suspensions, potatoes, asparagus, and salmon by the addition of lysozyme, and the delay was increased in the presence of EDTA (Johnson and Dell’Acqua, 1995). Nonetheless, some food components, notably fat, have been reported to hamper/lower the antibotulinal effects of antimicrobial food preservatives, including lysozyme and EDTA (Glass and Johnson, 2004).

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14.7.2 Lysozyme Use in Combination with Antimicrobial Polypeptides and Bacteriocins Combinations of lysozyme and nisin, a food preservative effective against Gram-positive bacteria, are effective against numerous pathogenic and spoilage bacteria. Nisin concentrations of 100–500 units/g are usually sufficient to increase the antimicrobial effectiveness of lysozyme (Tranter, 1994). The increased efficacy of lysozyme–nisin combinations may be due to increased membrane damage, cell lysis, or the inhibition of energy-dependent processes that repair lysozyme/nisin damage to the cell (Chung and Hancock, 2000). The presence of lysozyme increased the depolarization of the cytoplasmic membranes of S. aureus by nisin (Chung and Hancock, 2000). Severe cell damage in the form of surface disruption was observed by scanning electron microscopy in Lactobacillus sakei treated with lysozyme–nisin combinations (Chung and Hancock, 2000). Lysozyme and nisin in combination was more effective than lysozyme alone against many LAB in culture media (Chung and Hancock, 2000), and can be bacteriostatic or bactericidal to L. monocytogenes under certain conditions (Monticello, 1990). Growth of L. monocytogenes was inhibited on sliced pork bologna by the use of a dip containing nisin and lysozyme, although lysozyme alone had no effect (Kain et al., 2001). Lysozyme–nisin combinations were generally more effective than either lysozyme or nisin alone to control the growth of the Gram-positive meat-spoilage bacteria Brochothrix thermosphacta and Carnobacterium spp. in culture media, meat juice extract, and on pork tissue (Nattress et al., 2001). The addition of 500 mg/kg lysozyme–nisin (1:3 mixture) and 500 mg/kg EDTA to ham or bologna sausage batter before cooking resulted in growth inhibition at 8°C of B. thermosphacta, L. monocytogenes, Lactobacillus curvatus, Lactobacillus mesenteroides on sausage, and E. coli O157:H7, L. mesenteroides, B. thermosphacta, and L. curvatus on ham (Gill and Holley, 2000a). The same combination was ineffective against L. sakei, Salmonella Typhimurium, Serratia grimesii, and Shewanella putrefaciens on both products (Gill and Holley, 2000a). Combinations of lysozyme, nisin, and hop acids were bactericidal to L. monocytogenes, B. cereus, and B. subtilis in culture media, and to L. monocytogenes on hot dogs or cooked hams when applied as a dip or spray, respectively (King and Ming, 2002). However, without nisin, hop resins did not increase the effectiveness of lysozyme against E. coli (Fukao et al., 2000). When added as a third preservative factor, lysozyme increased the bactericidal activity of nisin and carvacrol against B. cereus and L. monocytogenes in buffer (Pol and Smid, 1999). The bactericidal activity of pulsed electric field (PEF) against Salmonella Typhimurium in orange juice was increased in the presence of lysozyme or a combination of lysozyme and nisin (Liang et al., 2002). While nisin–lysozyme combinations are effective against many Gram-positive bacteria, lactoferrin may enhance the effectiveness of lysozyme against Gram-negative bacteria. Lactoferrin, a protein found in milk, appears to damage the outer membrane of Gram-negative bacteria, causing the release of LPS molecules and increasing bacterial susceptibility to lysozyme by increasing penetration of lysozyme through the outer membrane (Ellison and Giehl, 1991; Yamauchi et al., 1993). Lactoferrin has been shown to enhance the antimicrobial activity of lysozyme against E. coli (Yamauchi et al., 1993, Suzuki et al., 1989). The combination of apo-lactoferrin and lysozyme delayed growth of L. monocytogenes in UHT milk, but was not as effective against the organism as an EDTA–lysozyme combination (Payne et al., 1994). Similar delays in growth of L. innocua were reported in microbiological medium for a combination of lysozyme and lactoferrin delivered via carboxy methylcellulose (CMC)-derived films. The lag phase for untreated (control) cells was 1.86 h while for lysozyme alone it was 5.81 h, and for lysozyme + lactoferrin it was 6.5 h (Barbiroli et al., 2012). Egg-white lysozyme (0.08 mg/ml) increased the antimicrobial effect of bovine lactoferrin against Salmonella Enteritidis in buffer, though lysozyme alone was not bactericidal to the organism (Facon and Skura, 1996). The bactericidal effect of lysozyme–lactoferrin combinations is dose-dependent, is blocked by iron saturation of lactoferrin, and is inhibited by high calcium levels (Ellison and Giehl, 1991). Numerous other substances commonly found in foods have been shown to affect lysozyme activity. Glycine–lysozyme combinations are effective against Gram-positive and Gram-negative bacteria, and are used commercially in Japan in certain foods (Proctor and Cunningham, 1988). Lysozyme at 100 µg/g delayed botulinum toxin production by in aqueous turkey slurries, especially in the presence of cysteine, proline, and sodium lactate (Johnson, 1994). Pseudomonas aeruginosa, but not E. coli O157:H7

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or Salmonella Typhimurium, was sensitized to lysozyme by lactic acid or HCl disruption of the outer membrane (Alakomi et al., 2000). Oh et al. (2016) similarly reported synergistic inhibition of L. monocytogenes in liquid microbiological medium by the combined application of lysozyme with acetic, citric, lactic, and malic acid.

14.8 Enhancement by Physical Processes Lysozyme activity can be increased by physical processes used in many food preservation systems. Freezing and thawing of Escherichia coli causes sensitivity to lysozyme (Kohn, 1960; Ray et al., 1984), although the organism regained resistance after incubation at 37°C or exposure to calcium (Ray et al., 1984). Transmission electron microscopy demonstrated protoplast formation and cell wall damage in unfrozen or frozen/thawed L. monocytogenes treated with lysozyme (El-Kest and Marth, 1992). The sensitization of Gram-negative cells to lysozyme by high pressure is dependent on factors such as pressure, temperature, pH, medium, growth stage, and bacterial species (Masschalck et al., 2001). High pressure has been shown to sensitize E. coli to lysozyme (Garcia-Graells et al., 1999; Masschalck et al., 2000), although the sensitization is transient (Hauben et al., 1996; Masschalck et al., 2001). In addition, four of six Gram-negative bacteria tested were sensitive to lysozyme during treatment by high hydrostatic pressure (Masschalck et al., 2001). EDTA increased the activity of lysozyme against E. coli during high-pressure treatment (Hauben et al., 1996). However, exposure of E. coli to lysozyme or lysozyme– EDTA immediately after high-pressure treatment did not decrease the number of viable cells (Hauben et al., 1996). Pulses of high hydrostatic pressure may sensitize some bacterial strains more effectively to lysozyme than treatment with continuous pressure (Masschalck et al., 2001). Sublethal injury by high pressure might be a useful hurdle along with lysozyme or lysozyme–EDTA in certain foods. Lower pressures required to inactivate certain bacteria in the presence of lysozyme (Masschalck et al., 2001) may lead to new combination treatments for the non-thermal preservation of foods. Similarly, pulsed electric field (PEF) was enhanced in its anti-S. aureus efficacy in skim milk when milk was first treated with 5.0 IU/mL nisin and 3000 IU/mL lysozyme in milk adjusted to pH 5.0, yielding a 4.8 log10-cycle reduction as compared to other treatments. PEF application time was also a key factor in determining S. aureus reductions observed (Sobrino-López and Martín-Belloso, 2008).

14.9 Food Applications Egg-white lysozyme is desirable as a food preservative because of its ease of purification in economically feasible quantities from egg white, low toxicity, specific activity against target bacteria and fungi, low effective usage levels, and low impact on sensory qualities of foods (Sofos et al., 1998). Lysozyme is stable under many conditions typically found in foods, and is highly soluble in aqueous environments (Razavi-Rohani and Griffiths, 1996b). Lysozyme is typically used in foods at 20–400 ppm (Gould, 2002), and commercial preparations for food use are generally available in powdered or granular form. Possible limitations of lysozyme include cost, inactivation by endogenous food components, potential for allergenic response by some consumers, and limited antimicrobial spectrum (Holzapfel et al., 1995).

14.9.1 Functionality in Ready-to-Eat Further Processed Foods As described, lysozyme is currently used in foods such as cheese, frankfurters, cooked meat, and poultry products (Davidson and Harrison, 2002; Pina-Pérez et al., 2015; Taylor et al., 2019). A patented process in the United Kingdom applies to the addition of lysozyme to butter or milk used to make Italian cheeses in order to control undesirable organisms such as C. tyrobutyricum (Ferrari and Dell’Acqua, 1979). Lysozyme, used in combination with standard preservatives, reduced microbial spoilage of cooked, salami, and Vienna sausages (Akashi, 1969, 1970, 1971). Microbial lytic enzymes may also have applications as food preservatives. An N-acetylmuramidase purified from Streptomyces rutgerensis extended the shelf life of adzuki bean paste (Hayashi et al., 1989).

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Lysozyme may be used in lieu of nitrate to prevent late blowing in hard and semi-hard cheeses such as Gouda, Edam, Provolone, and Emmental in the U.S. as well as numerous other countries (International Dairy Federation, 1987). Late blowing is a ripening defect resulting from hydrogen and carbon dioxide gas production during the fermentation of lactate by C. tyrobutyricum, a heat-resistant spore-former commonly found in milk. Lysozyme is usually added to milk at 20–35 ppm, resulting in levels in cheese ranging from 200 to 400 ppm, which does not generally inhibit starter cultures (Sofos et al., 1998) or affect physical or organoleptic properties of the cheese, though its presence must be labeled in order to prevent potential food allergenicity-related food safety concerns in cheese consumers (EC Rule 2003/89/EC). Numerous Japanese patents cover the use of lysozyme as a preservative in foods such as fresh vegetables, meats, seafood such as oysters and shrimp, tofu, cheeses, butter, potato salad, noodles, custard, and infant milk formula (Tranter, 1994; Cunningham et al., 1991). Lysozyme has been investigated extensively as a preservative in vegetable, meat, and seafood products in Japan (Tranter, 1994), and has shown effectiveness as a preservative in products such as kimchi (fermented cabbage), rice sushi, noodles, and creamed custard (Cunningham et al., 1991). Other Japanese patents apply to the use of lysozyme applied as a coating on fruits, vegetables, meats, and seafoods (Cunningham et al., 1991; Sofos et al., 1998). Lysozyme used in a gelatin dip extended the shelf life of a type of seafood product made from lizard-fish (Akashi and Oono, 1972). Lysozyme has been used in Japan to extend the shelf life of wine and sake (Sofos et al., 1998, Tranter, 1994), and is utilized with increasing frequency to improve the quality of wines produced in a number of other countries. The addition of 250 mg/L lysozyme after malolactic fermentation increased the microbiological stability of red wine (Gerbaux et al., 1997). Such inhibition of spoilage bacteria in wine by lysozyme may contribute to the use of lower sulfite levels in some products. Numerous patents have been issued in the United States for lysozyme applications in food systems. These include patents for listericidal treatments utilizing lysozyme in dairy or meat products (Dell’Acqua et al., 1989), lysozyme and EDTA on vegetables (Johnson et al., 1991), and combinations of hop acids, nisin, and/or lysozyme as ingredients in or applied to the surface of solid foods (King and Ming, 2002). Another patent covers the use of lysozyme or nontoxic lysozyme salts along with chelators such as EDTA to control growth of Clostridium botulinum in animal or vegetable foods (Johnson and Dell’Acqua, 1995). Lysozyme added to infant formula or milk encourages the growth of Lactobacillus spp. (Schwimmer, 1981) and Bifidobacterium bifidus (Sawada et al., 1967; Nishihava and Isoda, 1967) in infant intestines, which may prevent growth of harmful bacteria. Daeschel and Kennedy (2006) reported a method to reduce lysozyme binding by wine tannins via tannin binding agents, thereby retaining greater antimicrobial activity of lysozyme against spoilage bacteria.

14.9.2 Lysozyme Utility on Animal Carcass and Derived Product Surfaces Lysozyme may also be effective in immersion or spray methods to decontaminate animal carcasses. Gram-negative organisms, including Salmonella, can become sensitive to lysozyme after a sudden decrease in osmolarity, possibly increasing the antimicrobial effectiveness of lysozyme used in the decontamination of poultry (Chatzolopou et al., 1993). Combinations of lysozyme with other antimicrobials may be useful when applied to the surface of meat products, either through a dip, application of an edible gel, or on packaging film (Gill and Holley, 2000a). The use of lysozyme–antimicrobial combinations as a surface application in packaging components would prevent inactivation of the enzyme during cooking (Gill and Holley, 2000b). A lysozyme– nisin gel coating (1:3 mixture; 25.5 mg/ml) with 25.5 mg/ml EDTA applied to the surface of cooked ham and bologna was bactericidal to B. thermosphacta, L. sakei, Leuconostoc mesenteroides, L. monocytogenes, and Salmonella Typhimurium, and inhibited growth of the organisms during storage at 8°C for 4 weeks (Gill and Holley, 2000b). Nisin and lysozyme incorporated together into whey-protein isolate film was antimicrobial to Salmonella enterica serotype Typhimurium colonies on agar plates (Rodrigues et al., 2002). Lysozyme incorporated into biodegradable packaging films was bactericidal to Lactobacillus plantarum and, in combination with EDTA, E. coli (Padgett et al., 1998). Incorporation of lysozyme into packaging films has been reported to allow direct contact of antimicrobials with pathogens present on the surface of foods (Barbiroli et al., 2012; Wang et al., 2018).

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14.9.3 Lysozyme Applications for Food Contact Surfaces Disinfection The use of lysozyme on food contact surfaces may prevent colonization by pathogens such as L. monocytogenes as well as other biofilm-forming bacteria (Bower et al., 1998). Hen egg-white lysozyme forms multilayers on hydrophobic metal surfaces (Schmidt et al., 1990). However, some loss of antimicrobial activity by adsorbed lysozyme has been demonstrated (Schmidt et al., 1990). Lysozyme treatment of stainless steel resulted in the attachment of increased numbers of vegetative Bacillus species, possibly due to removal of extracellular polysaccharides by the enzyme (Parkar et al., 2001). On the other hand, Sheffield et al. (2012) reported that 25–50 μg/mL lysozyme produced near-complete removal of K. pneumoniae biofilm material from steel coupons after a 1-h treatment, compared to only ~60% removal by dextranase at the same content under similar exposure conditions.

14.10 Non-Enzymatic Antimicrobial Activity of Lysozyme and Lysozyme-Peptides The antimicrobial activity of lysozyme resulting from its enzymatic muramidase activity on bacterial cell walls has been studied extensively. However, lysozyme has also been reported to have antimicrobial activity unrelated to its enzymatic activity (Chung and Hancock, 2000; Ibrahim, 1998; Ibrahim et al., 2001).

14.10.1 Bactericidal Characteristics of Lysozyme Cationic proteins often possess bactericidal properties (Pelligrini et al., 1992). The basic nature of the lysozyme molecule itself may result in antimicrobial activity against some organisms (Mayes and Takeballi, 1983; Salton, 1957). For instance, S. aureus will grow in egg white at pH 7.2, but not at pH 8.0, possibly due to negative charges on the cell leading to agglutination by positively charged molecules such as lysozyme (Ng and Garibaldi, 1975). Treatment of E. coli with egg-white lysozyme resulted in disintegration of the cell cytoplasm visible by electron microscopy, and lysozyme was detected within the cytoplasm of frozen cells by immunogold labeling (Pelligrini et al., 1992). Bactericidal properties of denatured or partially denatured lysozyme against Gram-negative and Gram-positive bacteria have been shown, with increased sensitivity of E. coli to heat-denatured lysozyme compared to native lysozyme (Ibrahim et al., 1996a, 1996b, 2001). Pelligrini et al. (1992) found that denaturation of lysozyme by dithiothreitol (DTT) to reduce disulfide bonds did not lead to a loss of antimicrobial activity against Gram-negative and Gram-positive bacteria, suggesting that the bactericidal efficacy of lysozyme is due not only to lytic muramidase activity, but also to cationic and hydrophobic properties. Native lysozyme and lysozyme denatured by DTT were equally bactericidal to oral streptococci (Liable and Germaine, 1985). The microbiocidal activity of T4 lysozyme may be due to membrane disruption by small peptide sequences rather than enzymatic cell wall degradation (During et al., 1999). Heat-denatured bacteriophage T4 lysozyme exhibited antimicrobial activity but no enzymatic activity (During et al., 1999).

14.10.2 Lysozyme-Derived Antimicrobial Peptides Antimicrobial cationic peptides with no enzymatic activity have been isolated from lysozyme (Pelligrini et al., 1997; During et al., 1999). Peptide digests of lysozyme without enzyme activity were strongly bactericidal to E. coli and S. aureus, with damage to cell membranes of both species evident by scanning electron microscopy (Mine et al., 2004). A 15-amino-acid sequence (amino acids 98–112) isolated from egg-white lysozyme and an identical synthesized peptide both exhibited antimicrobial activity without catalytic muramidase activity (Pelligrini et al., 1997). However, the peptide sequences were weaker against Gram-negative bacteria than parent lysozyme and were ineffective against some organisms that were lysed by the parent lysozyme, such as P. aeruginosa, M. luteus, and S. aureus (Pelligrini et al., 1997). Memarpoor-Yazdi et al. (2012) reported peptides, the products of enzymatic breakdown

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of HEWL, exhibited both antioxidant and antimicrobial activity against both Gram-positive and -negative bacterial organisms. Carrillo et al. (2018a, b) reported that heat-denatured hydrolysates exhibited greater antibacterial spectrum than native HEWL, producing significant structural damage in cell walls of Staphylococcus carnosus and E. coli, producing reductions in numbers of both pathogens at 0.5 to 2.0 mg/mL application. These researchers also previously reported inhibition of spoilage bacteria within the LAB and AAB groups relevant to wine spoilage (Carillo et al., 2014).

14.11 Recombinant Lysozymes for Use as Food Antimicrobials For several years, researchers have been interested in expressing recombinant (cloned) human lysozyme, egg-white lysozyme, or lysozymes from other sources. Recombinant lysozyme from humans or other sources may have reduced allergenicity compared to egg-white lysozyme. Directed mutations of the cloned lysozyme gene could result in lysozymes with enhanced resistance to intrinsic and extrinsic factors in foods, increased specificity for target molecules, and other desirable features. Lysozyme from human, bovine, and microbial sources has been expressed in organisms that are often compatible for use in foods (Sofos et al., 1998). For instance, active egg-white lysozyme was expressed by Saccharomyces cerevisiae (Oberto and Davison, 1985) and Aspergillus niger (Archer et al., 1990). Inactive egg-white lysozyme but active T4 and bacteriophage lysozymes were produced by transformed Lactococcus lactis subsp. lactis strains (van de Guchte et al., 1992). Human lysozyme has been expressed during growth in whey by the yeast Kluyveromyces lactis after transformation with the human lysozyme, potentially leading to use of the yeast as a lysozyme-producing starter culture in cheeses prone to late blowing caused by clostridial fermentation (Maullu et al., 1999). Similarly, human lysozyme secreted into the milk of transgenic mice retained its lytic activity, leading to potential applications in the dairy industry of lysozyme-containing milk produced by transgenic animals (Maga et al., 1995; 1998). Wu et al. (2015) demonstrated production of human lysozyme from transgenic chicken eggs, wherein nearly equivalent antibacterial activity was generated from the recombinant human lysozyme as compared to commercial sources of both hen and human lysozymes as determined from disc diffusion assay. Additionally, the pH and thermo-stability of the recombinant lysozyme were non-differing from commercial lysozymes as well. Genetic engineering resulted in a modified T4 lysozyme with a new disulfide bond, which increased thermostability by stabilizing the tertiary structure (Genentech and Genencor, 1985). Such modifications may increase applications of lysozyme in cheese-making and other food applications (Proctor and Cunningham, 1988). A hyperstable chicken lysozyme was isolated by directed mutations and expression (Shih and Kirsch, 1995). Increased numbers of recombinant enzymes with advantages over native are being introduced into the food industry, but their safety must be carefully evaluated prior to their use in commerce (Pariza and Johnson, 2001). Modification of pig-derived lysozyme with a short (six) multipeptide add-on resulted in inhibition of E. coli, K. pneumoniae, and Salmonella Enteritidis, where for native pig lysozyme no antibacterial activity was detectable (Zhu et al., 2017).

14.12 Regulatory Status and Toxicology Egg whites are well-known to be allergenic to sensitive individuals, particularly children (Poulsen et al., 2001), and lysozyme may constitute one of the allergens in egg albumin. However, the most allergenic proteins in eggs appear to be (in descending order) conalbumin (ovotransferrin), ovomucoid, ovalbumin, and lysozyme. Allergic reactions produced by egg-white lysozyme in animals and humans were fewer than reactions to other egg proteins such as ovalbumin and albumin (Bianchi, 1982), which have long been used as food ingredients. The acute toxicity of lysozyme in humans is unknown (Lück and Jager, 1997), although rodents and rabbits tolerated high intravenous doses (Barbara and Pellegrini, 1976; Bianchi, 1982). Several studies have indicated that lysozyme from egg white has negligible acute, subacute, and chronic toxicity in animals (Barbara and Pellegrini, 1976). In addition, lysozyme has a

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long history of safe consumption as an endogenous food ingredient in eggs, milk, and other foods. The presence of added egg-white lysozyme in foods at permitted levels is generally not considered to be a health concern. Lysozyme was previously considered as generally recognized as safe (GRAS) by the United States Food and Drug Administration for use in certain cheeses according to the tentative final rule (U.S. FDA, 1998). However, because of the potential for allergenicity of lysozyme, the FDA has tentatively concluded that bulk and packaged foods containing lysozyme must be labeled to indicate presence of “eggwhite lysozyme” (U.S. FDA, 1998). This is a condition of lysozyme use for GRAS status in such treated cheese. It is permitted for use in cheese at levels in accordance with good manufacturing practices. In addition, the FAO-WHO Joint Expert Committee on Food Additives concluded that the low additional intake of lysozyme in cheese made from treated milk was not a hazard to consumer health (JECFA, 1993). Use of hen egg-white lysozyme has been approved in many countries to control the growth of spoilage organisms in foods (Wang and Shelef, 1992). Lysozyme was approved as a preservative (E1105) under the European Additives Directive to prevent late blowing in ripened cheese. Lysozyme is also approved for use in cheese in Austria, Australia, Belgium, Canada, Denmark, Finland, France, Germany, Italy, the United Kingdom, and Spain, but as mentioned previously its use is to be labeled for purposes of food safety for consumers allergic to egg. The Organisation Internationale du Vin (OIV) approved the use of lysozyme in wine in 1997 to control undesirable bacterial growth, and its use in wines is awaiting approval in other countries. Applications for uses in other foods are being evaluated. The United States Department of Agriculture recently issued an acceptability determination regarding the use of a mixture of hops beta acids, egg-white lysozyme, and cultured skim milk in salad dressings used as a preservative in refrigerated meat and poultry deli salads (USDA, 2002). Interestingly, lysozyme has been removed in the current version of the U.S. Code of Federal Regulations, Title 21 Section 184, available at www​.ecfr​ .gov, where it was previously listed alongside other GRAS compounds, though authors are not aware of a reversal of GRAS status notice being promulgated by the FDA.

14.13 Summary and Perspectives In response to increased consumer perception and demand for more nutritious foods, the food industry is introducing foods into the marketplace that receive minimal processing and contain reduced levels of traditional preservatives such as salt, sugar, and acids, and few or no chemical preservatives, increasingly in favor of natural and/or “clean label” ingredients. To attain adequate shelf life for quality and to prevent microbial foodborne disease, there is significant interest by the industry in the use of natural preservation systems. A myriad of naturally occurring antimicrobials are being considered to develop safety systems usually based a multiple-hurdle system to inactivate or prevent the growth of pathogenic or spoilage organisms in foods (Sofos et al., 1998). In designing these systems, it is important to consider the interaction and multiple functions of antimicrobials as pioneered in the writings of Reiter (1978) and others. Naturally occurring antimicrobials, in conjunction with GMPs usage, can enhance the safety and quality of minimally processed refrigerated foods (Johnson, 1994). Egg-white lysozyme is a good example of a naturally occurring enzyme that has been employed by the industry to maintain product quality and reduce the incidence of spoilage. Combinations of natural antimicrobials and mild preservation techniques (i.e., hurdle processing) are an attractive approach to control pathogens and spoilage organisms in many foods. The addition of lysozyme to certain foods before heat processing may reduce the thermal requirements necessary to inactivate spores of some strains of thermophilic spore-forming spoilage bacteria that are particularly sensitive to lysozyme, such as G. stearothermophilus and Clostridium thermosaccharolyticum (Johnson, 1994). Lysozyme could play a key role in the development of antimicrobial systems for use in foods. Lysozyme is relatively inexpensive, readily available, and is stable under a wide variety of conditions. Lysozyme and lysozyme–antimicrobial combinations have many current and potential uses in the food industry to control pathogenic and spoilage organisms. Activity of lysozyme occurs over a wide range of temperatures and pHs. It is among the most stable of enzymes in foods and resists many food-processing procedures.

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These properties may lead to numerous applications for lysozyme minimally processed foods. The use of lysozyme in these foods for the control of pathogens and spoilage organisms may result in products with increased safety, enhanced quality, and long shelf life. Future research may consider the interaction of lysozyme with other antimicrobials in minimally processed or alternatively processed food products, the incorporation and delivery of lysozyme to foods via encapsulation technologies or edible films, and the potential utility of other forms of lysozyme (e.g., goose egg lysozyme, insect-derived lysozymes) or chemi-modified lysozyme derivatives as innovative antimicrobials for food safety protection and spoilage prevention.

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15 Bacteriocins and Their Applications in Foods Pushpinder Kaur Litt, Dallas G. Hoover and Haiqiang Chen CONTENTS 15.1 Introduction..................................................................................................................................475 15.2 Classification of Bacteriocins.......................................................................................................476 15.3 Genetics, Biosynthesis, and Mode of Action.............................................................................. 477 15.3.1 Organization of Gene Clusters..................................................................................... 477 15.3.2 Biosynthetic Pathway.....................................................................................................478 15.3.3 Post-Translational Modification, Activation, and Transport........................................ 479 15.3.4 Regulation of Biosynthesis........................................................................................... 480 15.3.5 Producer Immunity....................................................................................................... 481 15.3.6 Mode of Action............................................................................................................. 481 15.4 Production and Modeling............................................................................................................ 482 15.5 Resistance.................................................................................................................................... 482 15.6 Examples of Activity Spectra and Biochemical Properties........................................................ 483 15.6.1 Lactococcus.................................................................................................................. 483 15.6.2 Pediococcus.................................................................................................................. 485 15.6.3 Lactobacillus................................................................................................................ 489 15.6.4 Carnobacterium........................................................................................................... 493 15.6.5 Leuconostoc and Other Lactic Acid-Associated Bacteria........................................... 494 15.7 Applications in the Food Industry............................................................................................... 497 15.7.1 Dairy Products.............................................................................................................. 497 15.7.2 Meat Products............................................................................................................... 498 15.7.3 Fruits and Vegetables.................................................................................................... 499 15.8 Summation................................................................................................................................... 500 References............................................................................................................................................... 501

15.1 Introduction Interest in the development of bacteriocins as food preservatives and antimicrobial agents has robustly continued into the twenty-first century. A substantial portion of this effort has occurred in food technology laboratories working with bacteriocins produced by lactic acid bacteria (LAB). In the United States, the authorization by the U.S. Food and Drug Administration (FDA) in 1988 for the use of nisin as a food additive in pasteurized processed cheese spreads opened the door for the potential application of other bacteriocins as preservatives in foods. Covered earlier in this book (Chapter 8), nisin is an antibiotic-like, heat-stable peptide produced by Lactococcus lactis subsp. lactis that, among bacteriocins produced by LAB, is the most potent bacteriocin known. It is effective against an array of Gram-positive bacteria, most notably vegetative types and spores that are involved in foodborne illness and food spoilage. The effectiveness of nisin as a processing aid has been demonstrated in many products worldwide for over 50 years. An important contributory factor for the sustained interest in bacteriocins is the great concern that has grown in the food industry regarding Listeria monocytogenes, especially in American processed foods where there is zero tolerance for the pathogen. L. monocytogenes is a Gram-positive, 475

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psychrotrophic bacterium that is relatively common in raw foods and stubbornly resists normal means of control and elimination. Its human foodborne syndrome can sometimes be fatal, and the infective dose is highly variable among human subpopulations. The relevance is that nisin and other bacteriocins produced by some strains of LAB are specifically antagonistic against L. monocytogenes. Therefore, with the legal precedent set in the U.S. for use of nisin as an additive in foods, the potential exists for application of other bacteriocins as effective inhibitive agents against L. monocytogenes and other important Grampositive pathogens to enhance product safety. Recently, the nano-engineered antimicrobial peptides of nisin have shown to be effective against Gram-negative bacteria (Vukomanović et al., 2017). The production of bacteriocins from lactic cultures potentially shortens the regulatory process because most lactic acid bacteria have generally regarded as safe (GRAS) status, having been consumed in large numbers in acidified fermented foods by people for thousands of years with no ill effects. In this regard, bacteriocins from LAB have been described as “natural” inhibitors. This chapter focuses upon review of bacteriocins from LAB exclusive of nisin. Using current taxonomy, LAB associated with foods include members of the genera Lactococcus, Lactobacillus, Pediococcus, Leuconostoc, Carnobacterium, Enterococcus, Oenococcus, Streptococcus, Tetragenococcus, Vagococcus, and Weissella (Stiles and Holzapfel, 1997). We have not attempted to inclusively present the vast volume of information that now exists in this area. We have primarily addressed bacteriocins from the first six LAB genera listed above with inclusion of some specific examples of work on bacteriocins from Propionibacterium and Bifidobacterium, given the importance of these two genera in foods.

15.2 Classification of Bacteriocins The prototype bacteriocins were the colicins. First discovered by Gratia in 1925, “principe V” was produced by one strain of Escherichia coli against another culture of E. coli. The term “colicine” was coined by Gratia and Fredericq (1946); “bacteriocine” was used by Jacob et al. (1953) as a general term for highly specific antibacterial proteins. The term colicin now implies a bactericidal protein produced by varieties of E. coli and closely related Enterobacteriaceae (Cascales et al., 2007; Konisky, 1982). The original meaning of the term bacteriocin was thus greatly influenced by characteristics common to colicins. That is, bacteriocins were originally defined as bactericidal proteins characterized by lethal biosynthesis, a very narrow range of activity, and adsorption to specific cell envelope receptors (Jacob et al., 1953). A later amendment was the association of bacteriocin biosynthesis with plasmids. The description has expanded to recognize the differences between colicins and bacteriocins produced by Gram-positive bacteria (Yang et al., 2014). Bacteriocins from Gram-positive bacteria usually do not possess a specific receptor for adsorption, are commonly of lower molecular weight than colicins, can have different modes of killing, a wider range of effect, different modes of release and cell transport, and possess leader sequences cleaved during maturation (Reeves, 2012). Today, bactericidal peptides or proteins produced by bacteria are labeled as bacteriocins. Normally, to demonstrate the proteinaceous nature of a newly characterized bacteriocin, sensitivity to proteolytic enzymes such as trypsin, α-chymotrypsin, and pepsin is an expected demonstration. Usage in food preservation usually requires estimation of its heat resistance given the widespread use of thermal processing in food production. Bacteriocins are widely found throughout the bacterial world. These compounds are assumed to provide producing bacterial cells with a selective advantage over other bacteria. Synthesis of bacteriocins is found across all major groups of eubacteria and archaebacteria (Riley, 1998). There have been many valuable monographs written over the years that review colicins, bacteriocins, bacteriocins from LAB, and applications of specific bacteriocins (Yang et al., 2014; Chikindas et al., 2018; Field et al., 2018; Juturu and Wu, 2018; Alvarez- Sieiro et al., 2016). Normally, bacteriocins from LAB are cationic, and hydrophobic or amphiphilic molecules composed of 20 to 60 amino acid residues (Nes and Holo, 2000). Bacteriocins produced by Gram-positive bacteria are commonly classified into three classes (Rea et al., 2011; Cintas et al., 2001; Klaenhammer, 1993). Class I contains bacteriocins termed lantibiotics (from lanthionine-containing antibiotic). They are small (4 logs CFU/cm2 in cooked meat after 3 hours when compared to the control. An increase in C. jejuni populations was observed after 24 h at 24°C, which it was suggested could be due to moisture loss and restricted phage movement in the complex meat surface. Another study evaluated lytic bacteriophage application to eliminate E. coli O157:H7 on beef steak. The steak meat was cut into equal pieces (18 pieces) and spot-inoculated with a rifampin-resistant strain of E. coli O157:H7 (P1432) at 103 CFU/ml concentration and incubated at 37°C for 1 hour (O’Flynn et al. 2004). Afterwards, 1 ml of a phage cocktail at a titer of 108 PFU/ml was spot-inoculated onto the meat surfaces (9 pieces) and incubated for 1 h. In seven of the nine samples no E. coli O157:H7 was recorded after enrichment, and the other two pieces had less than 10 CFU/ml of viable E. coli O157:H7. Another study also showed that single phage (DT6), applied at a high MOI (103–104 MOI) to beef cuts (1 cm2), reduced non-O157 and O157 STEC populations by 0.33–0.77 log CFU/cm2 after 3 h, and 0.47–1.15 log CFU /cm2, respectively, and these reductions were significantly greater than control (non-bacteriophage) treatments (Tomat et al. 2013).

16.4.3 Food-Contact Surfaces Food-contact surfaces vary within food production and retail environments and are often areas of contamination and present risk of cross-contamination between food products. Lytic bacteriophages have been assessed for their effectiveness on several food-contact surfaces (Litt et al. 2017b, Viazis et al.

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2011a). The effect of a bacteriophage cocktail (BEC8) to control E. coli O157:H7 on food-contact surfaces was assessed (Viazis et al. 2011). Sterile stainless steel (SS), ceramic tile (CT), and high-density polyethylene (HDPE) were inoculated with E. coli O157:H7 (106, 105, 104 CFU/chip). Inoculated chips were treated with controls and BEC8 (106 PFU/chip) and incubated at 4, 12, 23, and 37°C and the surviving pathogen population was enumerated. In general, E. coli O157:H7 cells in low inoculum on chips were reduced by ca. 4 log CFU/chip quickly (10–60 min) at temperatures other than 4°C. E. coli O157:H7 populations (104, 106 CFU/chip) were reduced significantly compared to controls only after 24 hours at 37°C by ca. 5, 2.5, and 3 log CFU/chip on SS, CT, and HDPE, respectively (Viazis et al. 2011). Another study determined the effectiveness of a lytic bacteriophage (KH1) against E. coli O157:H7 attached to sterile stainless-steel coupons or which had formed biofilms on these coupons (Sharma et al. 2005). Sterile stainless-steel coupons were inoculated with E. coli O157:H7 (7–8 log CFU/coupon) and incubated at 22°C for 24 h, to encourage attachment with minimal biofilm formation, or 96 hours to encourage biofilm formation, before treatment with a single lytic phage specific to E. coli O157:H7 (7.7 logs PFU/ml) for 4 days (biofilm) or 1 day (attached cells). Phage treatment reduced attached bacterial cells by 1.2 logs CFU/coupon, after day 1, and maintained those levels of inactivation up to 4 days. In biofilms treated with KH1, E. coli O157:H7 ATCC 43895 populations did not differ significantly compared to populations in biofilms treated with controls (water), when treated with phage. By comparison, alkaline cleaners reduced both E. coli O157:H7 strain populations by 5–6 logs CFU per coupon. Overall, phage treatment was more effective against bacterial cells attached to stainless-steel coupon surfaces compared to cells in the biofilms (Sharma et al. 2005). Reduction of non-O157 and O157 shiga-toxigenic E. coli (STEC) strains on stainless-steel surfaces was also achieved by applying a two-phage cocktail on inoculated stainless-steel coupons at low (104) or high (106) MOI values. Both non-O157 and O157 STEC populations on stainless-steel surfaces were reduced more at 37°C (ca. 2–8 log CFU reductions) than at 4°C (ca. 2–4 log CFU reductions), and, in general, reductions were greater after 24 hours than after 1 or 3 hours (Tomat et al. 2014).

16.4.4 Pre-Harvest Control Some researchers have considered the application of bacteriophages in agricultural and food production systems to reduce pathogen contamination of foods (Atterbury et al. 2007). This may include the application of phages in crop or animal production, in agricultural waste management equipment, or oral administration in live food animals. A study was conducted to evaluate the efficacy of phage therapy to reduce C. jejuni colonization in broiler chickens (Carrillo et al. 2005). The 25-day-old broiler chickens were experimentally colonized with C. jejuni HPC5 and GIIC8 and treated with phages specific to C. jejuni (CP8 and CP34) through oral administration at different dosages. Phage-treated C. jejuni-colonized birds showed a reduction of C. jejuni populations ranging between 0.5 and 5 logs CFU/g of cecal contents compared to untreated controls after 5-day period. Reductions varied on the basis of the phage combination, the dose of phage given to the bird, and the time elapsed after administration. However, bacteriophage-insensitive mutant (BIM) C. jejuni cells were recovered from phage-treated chickens at a frequency of less than 4%. These BIM C. jejuni were compromised in their ability to subsequently colonize experimental chickens and rapidly reverted to a phage-sensitive phenotype in vivo. These results show the importance of using a cocktail of multiple phages specific to the target pathogen in the food animal (Carrillo et al. 2005). Another study looked at the application of phage at varying multiplicity of infection (MOI) in the treatment of dairy manure compost comprised of cow manure and sawdust, wasted feed, old hay, and vegetable wastes (Heringa et al. 2010). Compost (600 g) was spread and dried on trays in a biosafety hood at room temperature for 1 day. Salmonella Typhimurium strain 8243 (ca. 107 CFU/ml) was sprayed on the surface of compost and stored overnight at room temperature. A mixture of five lytic bacteriophages effective against several Salmonella serovars was sprayed at MOIs of 1, 10, and 50 onto 150 g of inoculated compost, mixed, and stored at room temperature (25°C) for 24 hours. Results showed significant reductions (~2.5 logs CFU/g) in S. Typhimurium populations compared to untreated controls following phage treatment at all MOI levels evaluated and at all compost moisture content levels (30, 40, 45, and 50%). Populations of S. Typhimurium were decreased by phage treatment within the first 4 h at all MOI

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levels evaluated, and these levels were maintained over 24 hours (Heringa et al. 2010). These results suggest that phage treatment can be applied during the compost process, even at higher moisture contents, to reduce foodborne pathogen counts.

16.4.5 Bacteriophage Insensitive Mutants Bacterial cells can become resistant to lytic phage infection by preventing phage adsorption via modifying the surface phage receptor structure, or by hindering the access of the phage to the receptor through the production of an excess of the extracellular matrix, or even by producing competitive inhibitors (Buckling and Rainey 2002). Phage-driven bacterial evolution increases its chances to adapt and survive in altered environmental conditions, ultimately becoming beneficial to bacteria (Pal et al. 2007). However, the coevolution of phage and bacteria in the natural environment has been shown, where phage-susceptible and -resistant bacterial populations intertwine with parent phages and evolved phages (mutant phages) that regain infectivity against the resistant bacteria (Buckling and Rainey 2002). Several studies have evaluated bacterial pathogens for BIM development after phage treatment in foods. BIM mutants of E. coli O157:H7 developed when meat was treated with individual lytic or a cocktail of lytic phages (O’Flynn et al. 2004), but most BIM colonies remained susceptible to at least of the one of lytic phages used in the cocktail. Another study showed that of eight presumptive BIM E. coli O157:H7 colonies isolated after phage treatment of beef samples, only two (25%) showed resistance to a single O157-specific phage infection (Tomat et al. 2013). A similar study examining S. Newport inactivation by lytic phages on whole cucumbers showed a relatively low rate (0.7%, 1/127 isolates) of BIM development (Sharma et al. 2017). The development of BIM may also be based on the storage temperature of foods treated with lytic bacteriophages, with higher temperatures potentially promoting higher growth rates and mutation rates of bacterial cells. However, BIM development for lytic phage applications to foods is unlikely, due to the presence of low levels of the pathogen in the environment, the use of multi-phage preparations in commercial lytic bacteriophage preparations which have different targets, and the lack/ slow growth of most bacterial pathogens at refrigeration or storage temperatures that slow the development of BIM mutant populations.

16.5 Where Does Phage Technology Stand Today? Several commercial phage products have been approved by regulatory agencies in the United States as food additives or to enhance food safety. The U.S. Department of Agriculture’s Food Safety and Inspection Service (USDA FSIS) has recorded of 12 entries for use of phage products which are listed as safe and suitable antimicrobials and could be utilized as processing aids in food production systems (FDA 2019). Most of these phage-based products do not have labeling requirements, except in cases where the product requires labeling of any additives. Phage products can be applied at any point including live animals/poultry pre-slaughter, poultry carcasses, raw poultry, beef parts and trims, and readyto-eat (RTE) meat and poultry. The Food and Drug Administration (FDA) has issued ten notices as generally recognized as safe (GRAS) for phage-based food safety products with ‘no questions’ notices and has not contested the manufacturer’s claim of GRAS (FDA 2019). Many companies have entered and are continuing to enter the bacteriophage production space. A few are mentioned here, and this discussion, showing the variety of materials approved and available, is not intended to be a complete discussion of all available technologies as this is a large area of research and development. A Dutch company, Micreos, PhageGuard, produces multiple FDA-approved phage preparations against foodborne pathogens, such as Listex P100 or PhageGuard Listex against L. monocytogenes, Salmonelex or PhageGuard S targeting Salmonella spp., and PhageGuard E for E. coli O157:H7 in raw and RTE foods (PhageGuard 2019a). PhageGuard Listex and S are approved for clean-label processing in the US, European Union (EU), Canada, Australia, New Zealand, Switzerland, and Israel (PhageGuard 2019a, 2019b). Intralytix is a United States-based biotech company in Baltimore, MD, producing phage-based products including ListShield targeting L. monocytogenes, EcoShield for E. coli O157: H7, SalmoFresh against Salmonella spp., and ShigaShield targeting Shigella spp. SalmoFresh has

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received GRAS status from the FDA and Health Canada, and is approved to be used as a processing aid in fresh and processed fruits and vegetables against Salmonella (Intralytix 2019a). Moreover, the National Food Service, Ministry of Health in Tel Aviv, Israel, approved the use of all FDA-approved phage preparations for similar applications on food (Intralytics 2019b). Another US phage company, Passport, has developed a phage product, Finalyse, to control E. coli O157:H7 and other STEC at preharvest level including animal hides and processing plants (Passport 2019).

16.6 Conclusion Bacteriophages have numerous applications for food safety, and can effectively reduce foodborne pathogens on various foods. Several commercial phage-based products have been approved for use in the food industry in a number of countries. Commercial phage products can be utilized for the reduction of specific foodborne pathogens at various timepoints in food production systems, such as in animal feed, sprayed on animal hides, animal contact surfaces or carcasses, and on raw poultry and beef trim, ready-to-eat (RTE) meat and poultry, dairy products, dip or spray wash produce, whole and fresh-cut fruits and vegetables, in fruit juices, or for surface-treatment of processing facilities. Lytic bacteriophage treatments seem to be most effective against bacterial foodborne pathogens when applied to foods in a way that delivers a high titer of phages to a large area, and then foods are stored at a temperature which prevents or slows the growth of foodborne pathogens after bacteriophage treatment. Bacteriophages can be an additional tool in a multi-hurdle approach to minimize the hazard of foodborne pathogens in foods with minimal processing and decrease the risk of bacterial infections to consumers, especially in fresh-cut produce (Abedon 2016, Mahony et al. 2011, Hagens and Loessner 2010). More understanding about the interactions of lytic bacteriophages with the overall microbiomes of foods can only improve the vast potential of lytic phages in foods to reduce the risk of foodborne illness.

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17 Naturally Occurring Compounds – Plant Sources Aurelio Lopez-Malo, Stella M. Alzamora, María J. Paris, Leonor Lastra-Vargas, María Bernarda Coronel, Paula L. Gómez, and Enrique Palou CONTENTS 17.1 Introduction............................................................................................................................... 527 17.2 Sources of Natural Antimicrobials from Plants........................................................................ 529 17.2.1 Natural Phenolic Compounds.......................................................................................531 17.2.2 Essential Oils............................................................................................................... 534 17.2.3 Phytoalexins..................................................................................................................537 17.2.4 Phytoanticipins............................................................................................................. 538 17.2.5 Antimicrobial Peptides (AMPs)....................................................................................539 17.2.6 Other Sources of Plant Antimicrobial Agents............................................................. 540 17.3 Testing the Efficacy of Antimicrobials..................................................................................... 540 17.3.1 In Vitro Testing.............................................................................................................541 17.3.2 Vapor Phase..................................................................................................................541 17.4 Mechanisms of Action.............................................................................................................. 544 17.5 Factors Affecting Activity......................................................................................................... 554 17.5.1 Plant Source Variation................................................................................................. 554 17.5.2 Extraction Methods.......................................................................................................555 17.5.3 Interaction with Food Matrix...................................................................................... 556 17.6 Increasing the Efficacy of Natural Antimicrobials from Plants............................................... 556 17.6.1 Application of Essential Oils in Gaseous Phase...........................................................557 17.6.2 Combination with Other Antimicrobials and/or Preservation Factors.........................558 17.6.3 Delivery Systems..........................................................................................................558 17.6.3.1 Emulsions....................................................................................................559 17.6.3.2 Nanosized Carriers/Coatings.......................................................................559 17.6.3.3 Packaging Films/Coatings.......................................................................... 560 17.7 Toxicity of Natural Antimicrobials and Its Evaluation and Regulatory Aspects..................... 562 17.8 Application in Food and Sensory Analysis............................................................................... 563 17.9 Final Remarks........................................................................................................................... 572 17.10 Acknowledgments..................................................................................................................... 573 References............................................................................................................................................... 573

17.1 Introduction Food market trends are changing, consumers more frequently demand high-quality foods with fresh-like attributes (Alzamora et al., 2016; Gould, 1995a, 1995b, 1996), and consequently less extreme treatments and/or additives are being required. To satisfy consumer demands, adjustments or reductions in conventionally used preservation techniques must be accomplished. Gould (2002) identified some food characteristics that must be attained in response to consumer demands; most of them occur within the minimal processing concept (Berdejo et al., 2019). To satisfy market requirements, the safety and quality of foods

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have to be based on substantial improvements in traditional preservation methods (Aguilar-González et al., 2015; Lorenzo-Leal et al., 2019a, 2019b; Mani-López et al., 2018; Reyes-Jurado et al., 2019b). Safe food may have different meanings to different people involved in the food chain; consumers for instance, relate a risk-free food as a safe one, and associate increased risk with the increased use of added substances such as antimicrobials, synthetic additives, and higher levels of sodium or fat, among others (Reyes-Jurado et al., 2019b). Scientists, public health officers, and international organizations define a safe food as one that provides maximum nutrition and quality while posing a minimal hazard to public health, and expect any risks that are present to be minimal (Shank and Carson, 1992). Consequences of quality loss caused by microorganisms include consumer hazards, due to the presence of pathogenic organisms or microbial toxins, as well as economic losses due to spoilage (Davidson, 2001). Inactivation, growth delay, or growth prevention of spoilage and pathogenic microorganisms are the main objectives of food preservation. Several factors influence microbial growth and survival; the appropriate modification and/or application of these factors is the base of preservation technologies. Food preservation technologies protect foods from the effects of microorganisms and inherent deterioration. Major food preservation technologies can be classified as those that act mainly by preventing or slowing down microbial growth (low temperature, reduced water activity, acidification, fermentation, modified atmosphere packaging, addition of antimicrobials, and compartmentalization in water-in-oil emulsions, among others), those that act by inactivating microorganisms (heat pasteurization and sterilization, microwave heating, ionizing radiation, high pressure, pulsed electric fields, and high-frequency ultrasound, among others), and those that prevent or minimize the entry of microorganisms into food or remove them (aseptic handling or packaging, centrifugation, and filtration, among many others) (Gould, 2002). In addition, techniques in combination, based on the “hurdle technology” concept (Alzamora et al., 2016; Leistner, 2000), may act by inhibiting or inactivating microorganisms, depending on the combination of hurdles applied to achieve food preservation (Berdejo et al., 2019; López-Malo et al., 2000). The use of chemical agents exhibiting antimicrobial activity (by inhibiting and/or reducing microbial growth or even by inactivating undesirable microorganisms) is one of the oldest and most traditional food preservation techniques. Antimicrobial agents are chemical compounds added to, or present in foods that retard microbial growth or cause microbial death. The antimicrobial activities of several plants used today as seasoning agents in foods and beverages have been recognized for centuries. The early Egyptians utilized plant extracts (spices and oils) as antimicrobials for the preservation of food as well as for embalming. Although ancient civilizations recognized the antiseptic or antimicrobial potential of many plant extracts, it was not until the eighteenth century that scientific information established the preservative effects of several plants. Antimicrobial agents are somewhat arbitrarily classified as traditional and naturally occurring (Davidson, 2001). Traditional antimicrobials are those that have been used for a long time, approved by many countries as antimicrobials in foods, or are produced by synthetic means or are inorganic. Antimicrobial agents may be either synthetic compounds intentionally added to foods or naturally occurring, biologically derived substances (the so-called naturally occurring antimicrobials), which may be used commercially as additives for food preservation besides exhibiting antimicrobial properties in the biological systems in which they are originally found (Sofos et al., 1998). However, as Davidson (2001) stated, ironically synthetic antimicrobial agents are found in nature, such as acetic, benzoic, and sorbic acids, among many others. Concerns about the use of antimicrobial agents in food products have been discussed for decades. Both the increasing demand for reduced-additive (including antimicrobial agents) and more “natural” foods, and the increasing demand for greater convenience have promoted the search for alternative antimicrobial agents or combinations to be used by the food industry (López-Malo et al., 2000). In this search, a wide range of natural systems from animals, plants, and microorganisms are being studied (Burt, 2004; Mani-López et al., 2018; Reyes-Jurado et al., 2019b). However, mainly economic aspects originating in the strict requirements to obtain approval and efforts to get the product onto the market restrict the spectrum of new chemical compounds that can help in the preservation of foods. The application of chemical antimicrobial agents in food preservation is regulated in the United States of America by the Food and Drug Administration (FDA) and in other countries by appropriate corresponding authorities. The Food

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and Agriculture Organization (FAO) and the World Health Organization (WHO) of the United Nations, testing and recommending usage and safety of chemicals in foods and acceptable daily intakes (ADI), regulate chemical antimicrobials internationally. In the US, chemicals in foods are examined according to the Food Additives Amendment of the Food, Drug and Cosmetic Act, specifying the procedures and conditions required for a chemical food additive to be approved. These obstacles have originated the search for emerging preservatives by examining compounds already utilized by the food industry, perhaps with other purposes, but with potential as antimicrobials, approved and not toxic in the levels used, many of them classified as generally recognized as safe (GRAS). Within these compounds are, for example, the so-called “green chemicals” present in plants that are utilized as flavor ingredients. Although many natural systems have potential to be used as antimicrobials, this chapter will focus mainly on natural antimicrobials from plants and their possible application in foods.

17.2 Sources of Natural Antimicrobials from Plants Plants, herbs, and spices as well as their derived essential oils and isolated compounds contain a large number of substances that are known to inhibit various metabolic activities of bacteria, yeast, and molds, although many of them are not yet completely exploited. The antimicrobial compounds in plant materials are commonly contained in the essential oil fraction of leaves (rosemary, sage), flowers and flower buds (clove), bulbs (garlic, onion), rhizomes (asafoetida), fruit (pepper, cardamom), or other parts of the plant (Nychas, 1995). Table 17.1 presents a list of some of the most highly recognized plants, herbs, and spices that have been reported as sources of natural antimicrobials. These compounds may be lethal to microbial cells or they may simply inhibit the production of a metabolite (e.g. mycotoxins) (Davidson, 2001). Zaika (1988) reviewed the literature reporting the antimicrobial activity of many spices and classified their inhibitory activities as strong, medium, or weak. According to this ranking, cinnamon and TABLE 17.1 Selected Plants and Their Major Antimicrobial Compounds Plant (Scientific Name) Allspice (Pimenta dioica) Basil (Ocimum basilicum) Black pepper (Pipper nigrum) Bay (Laurus nobilis) Caraway seed (Carum carvi) Celery seed (Apium graveolens) Cinnamon (Cinnamomum zeylanicum) Clove (Syzygium aromaticum) Coriander (Coriandum sativum) Cumin (Cuminum cyminum) Fennel (Foeniculum vulgare) Garlic (Allium sativum) Lemongrass (Cymbopogon citratus) citral Marjoram (Origanum majorana) Mustard (Brassica hirta, B. juncea, B. nigra) Onion (Allium cepa) Oregano (Origanum vulgare) Parsley (Petroselinum crispum) Rosemary (Rosmarinus officinalis) Sage (Salvia officinalis) Tarragon (Artemisia dracunculus) Thyme (Thymus vulgaris) Vanilla (Vanilla planifolia, V. pompona, V. tahilensis)

Major Components (in Descending Order) Eugenol, methyl ether cineol D-linalool, methyl chavicol, eugenol, cineol, geraniol Monoterpenes, sesquiterpenes Cineol, l-linalool, eugenol, geraniol Carvone, limonene D-limonene Cinnamic aldehyde, l-linalool, p-cymene, eugenol Eugenol, caryophyllene D-linalol, d-α-pinene, β-pinene Cuminaldehyde, p-cymene Anethole Diallyl disulfide, diethyl sulfide, diallyl trisulfide, allicin Geraniol Linalool, cineol, methyl chavicol, eugenol, terpinineol Allyl-isothiocyanate D-n-propyl disulfide, methyl-n-propyl disulfide Thymol, carvacrol, α-pinene, p-cymene Α-pinene, fenol-eter-apiol Borneol, cineol, camphor, α-pinene, bornyl acetate Thujone, cineol, borneol, thymol, eugenol Methyl chavicol, anethole Thymol, carvacrol, l-linalool, geraniol, p-cymene Vanillin, vanillic, p-hydroxybenzoic, p-coumaric acids

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clove were listed as exhibiting a strong inhibitory effect while allspice was classified with a medium inhibitory effect. Conner and Beuchat (1984a,b) screened 32 essential oils from plant sources for inhibitory effects on 13 food spoilage and industrial yeasts and identified cinnamon, allspice, and clove among the most inhibitory tested. Antimicrobial activity of cinnamon, allspice, and cloves is attributed to eugenol (2-methoxy-4-allyl phenol) and cinnamic aldehyde, which are major constituents of the volatile oils of these spices. Cinnamon contains 0.5 to 1.0% volatile oil, which contains 65 to 75% cinnamic aldehyde and 8% eugenol. Allspice contains up to 4.5% volatile oil, of which 80% is eugenol. Clove buds have an average essential oil content of 17% that is 93 to 95% eugenol (Bullerman et al.; 1977; Farrell, 1990). Major components with antimicrobial activity found in plants, herbs, and spices are phenolic compounds, terpenes (also called isoprenoids), terpenoids, aliphatic alcohols, aldehydes, ketones, acids, and isoflavonoids. Certain plant-based proteins and peptides, as well as some alkaloids, have also been reported as antimicrobials against foodborne bacteria, yeast, and molds (Savoia, 2012). As a rule, it has been reported that the antimicrobial activity of essential oils depends on the chemical structure of their components and on their concentration. Chemical structures of selected antimicrobial compounds from plant origin are presented in Figure 17.1.

FIGURE 17.1  Selected naturally occurring plant compounds with antimicrobial activity.

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FIGURE 17.1  Continued.

17.2.1 Natural Phenolic Compounds Derivatives of phenol, called phenolics, make up the second largest group of nutraceuticals after terpenoids and contain a phenol molecule with one or more hydroxyl groups (Gutierrez-del-Rio et al., 2018). This chemical alteration of phenol may increase its antimicrobial activity. Phenolic compounds have been used as antimicrobial agents since the early use of phenol as a sanitizer in 1867 (Davidson, 1993). As food antimicrobials, phenolic compounds can be classified, following Davidson (1993), as those currently approved (parabens), those approved for other uses (antioxidants), and those found in nature (polyphenolics, phenol). Naturally occurring phenolic compounds are widespread in plants and may be found in a great variety of food systems, and as phenol derivatives they may have antimicrobial activity (Chan et al., 2018; Gutierrez-del-Rio et al., 2018). These naturally occurring phenols and phenolic compounds may be classified into the following groups: simple phenols and phenolic acids (e.g. p-cresol, 3-ethylphenol, vanillic, gallic, ellagic, hydroquinone), hydroxycinnamic acid derivatives (e.g. p-coumaric, caffeic,

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ferulic, sinapic), flavonoids (e.g. catechins, proanthocyanins, anthocyanidins and flavons, flavonols and their glycosides), and “tannins” (e.g. plant polymeric phenolics with the ability to precipitate protein from aqueous solutions). These groups of phenolic compounds share the ability to inhibit microorganisms; therefore, they may have a common mode of action. Phenolics exert antimicrobial activity by injuring lipid-containing membranes, which results in changes in membrane structure and functionality, and in leakage of cellular contents (Freitas and Cattelan, 2018). They are extensively found in essential oils, as well as in many different plant extracts, and there are innumerous studies demonstrating their antimicrobial activity against fungi and bacteria. It is recognized that Gram-positive bacteria are generally more sensitive to these compounds, compared to Gram-negative ones. Flavonoids, a subgroup of phenolic compounds, can be found in fruits and vegetables, and their ability to work as antimicrobials is due to their capacity to form complexes with extracellular and soluble proteins, as well as with the bacterial membrane (Savoia, 2012). In a study developed by Yakoub et al. (2018), an ethanolic extract, rich in phenolic acids and flavonoid compounds, was obtained from Tossa jute. Phenolics acids such as quinic acid, 4-O-caffeoylquinic acid, 3,4-di-O-caffeoyquinic acid, protocatchuic acid, and caffeic acid, as well as flavonoids such as quercetin and cirsiliol, were among the major components detected. It was observed that increasing concentrations of phenolic extracts in the medium improved Micrococcus luteus, Bacillus cereus, Escherichia coli, Salmonella enterica, Salmonella typhi, and Enterobacter sp. inhibition (Yakoub et al., 2018); previous work had already suggested the efficacy of Tossa jute extract against bacteria and fungi. Vitamin P is another flavonoid that has been suggested as a potential antimicrobial; its addition to Chinese sausage resulted in Staphylococcus count reductions, although it allowed slight growth of certain beneficial and harmful bacteria (Tang et al., 2019). Unlike essential oils and their components that, as will be described later, are in part lipophilic, anthocyanins have the particularity of being water-soluble. This characteristic makes them interesting, given the aqueous nature of the majority of fresh food. Anthocyanins can be a potential choice of antimicrobials for direct application in food, while essential oils with considerable volatility are interesting for application in their vapor phase. Anthocyanins from Chinese wild blueberry extract destroyed the cell membrane of different pathogens, including Listeria monocytogenes, Staphylococcus aureus, Staphylococcus Enteritidis, and Vibrio parahaemolyticus; membrane destruction caused nucleic acid and protein leakage through the membrane (Sun et al., 2018). Sun et al. (2018) reported that anthocyanins could pass through the membrane and lower the activity of bacterial enzymes such as alkaline phosphatase, adenosine triphosphatase, and superoxide dismutase. Besides, they suggested that anthocyanins also weakened cellular respiration which, by affecting cellular energy supply, led to cell death. Apart from anthocyanins, both phenols and flavonoids with antimicrobial activity are found in Chinese wild blueberry extract (Sun et al., 2018). Epigallocatechin3-gallate (EGCG) from blueberry ash fruit and macadamia skin extracts was also effective against pathogenic and spoilage bacteria, as well as against fungi, when incorporated in active pea starchguar gum films; in this case, among the microorganisms studied, Salmonella Typhimurium and Rhizopus sp. were the most resistant to active films with the polyphenols, and in general, fungi resisted better than bacteria (Saberi et al., 2017). Grape seed is widely known for its polyphenol content. Grape seed extract is rich in cathechins, epicatechins, gallic acid, and proanthocyanidins; thus, it was used to mitigate L. monocytogenes growing in doner kebabs. The use of grape seed extract made the pathogen less resistant to heat and decreased the time needed for 4-log inactivation; the higher the concentration, the less time was needed for 4-log inactivation, and the less resistant the pathogen became. One percent grape seed extract was enough to cause immediate lethality to L. monocytogenes in doner kebabs at 65°C (Haskaraca et al., 2019). In a study ran by Zambrano et al. (2019), black grape extracts, also rich in phenolic compounds, showed antimicrobial activity against Bacillus subtilis and B. cereus. Pterostilbene, a polyphenol present in Xinjiang wine grape, had antifungal and antibacterial activity, damaging S. aureus and E. coli morphology, making cells irregular and wrinkled, and also causing cell lysis with nucleic acid and protein leakage; up-regulation of oxidative stress genes and down-regulation of genes involved in cell wall synthesis were also caused by the polyphenol (Ren et al., 2019). In another study, gallic acid grafting onto chitosan significantly inhibited E. coli and S. aureus growth, disrupting the cell membrane, which caused the

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release of cytoplasmic content and increased relative conductivity; cell morphology was also changed to an irregular shape, and cells gained a tendency to aggregate. The gallic acid–chitosan complex was able to cross the membrane and disrupt DNA synthesis in the nucleus (Li et al., 2019). Chitosan is, per se, recognized for its antimicrobial activity. However, the addition of gallic acid made the film more effective and more water-soluble, and made reactions with bacteria easier, compared to chitosan films without gallic acid. The higher the concentration of gallic acid, the more effect was observed. Gallic acid and chitosan were also loaded in glucomannan film, with remarkable antimicrobial activity against S. aureus and E. coli O157:H7 (Wu et al., 2019). Fang et al. (2018) developed a chitosan edible coating with gallic acid for fresh pork covering and, as in the last study mentioned, gallic acid improved chitosan film antimicrobial activity. Vanillin, a phenolic compound present in vanilla pods (Figure 17.1), also has antifungal activity (Cerrutti and Alzamora, 1996; López-Malo et al., 1995, 1997, 1998). Another phenolic compound present in plants with antimicrobial activity is oleuropein, obtained from green olive extracts (Beuchat and Golden, 1989; Nychas, 1995). The antimicrobial activity of naturally occurring phenols and phenolic compounds from olives, tea, and coffee has also been examined. Green and black tea, as well as coffee, are, in effect, known to be rich in polyphenols. Nychas (1995) reported that phenolic extracts of black and green tea and/or coffee could be bactericidal or bacteriostatic against Campylobacter jejuni, Campylobacter coli, Streptococcus mutans, Vibrio cholerae, Staphylococcus aureus, S. epiderimidis, Plesiomonas shigelloides, Salmonella typhi, S. Typhimurium, S. Enteritidis, Shigella flexneri, and S. dysenteriae. Moreover, Pseudomonas fragi, Lactobacillus plantarum, S. aureus, S. carnosus, Enterococcus faecalis, and Bacillus cereus were inhibited by ethyl acetate extracts from olives while B. subtilis, B. cereus, and S. aureus were inhibited by oleuropein. Coatings containing green tea extract, rich in polyphenols such as catechins, were used to protect strawberries and raspberries against viral attack (Falcó et al., 2019). Green tea extract owes its properties mainly to epigallocatechin-3-gallate (EGCG), the main catechin in green tea (Falcó et al., 2019). On the other hand, coffee waste containing gallic, chlorogenic, p-coumaric, sinapic acids, and caffeine inhibited Fusarium verticillioides, Fusarium sp., and Colletotrichum gloeosporioides when added to gellan gum films (Mirón-Mérida et al., 2019). Other extracts rich in phenolic compounds were obtained from Cynara cardunculus, an edible thistle; aqueous, ethanolic, and methanolic extracts of the leaves, containing chlorogenic acid, cynarin, luteolin-7-O-rutinoside, and cymaroside, had antimicrobial activity against foodborne Gram-negative bacteria, although they were not as effective against Gram-positive bacteria (Scavo et al., 2019). Also containing epicatechin, Hamelia patens extracts obtained by maceration, percolation, or Soxhlet extraction were successfully utilized as antimicrobials against S. aureus, E. coli, Salmonella Paratyphi, and S. Typhi (Paz et al., 2018). Pomegranate peel powder has a variety of polyphenols, including anthocyanins, gallotannins, ellagitannins, gallagyl esters, hydroxybenzoic acids, hydroxy cinnamic acids, and dihydroflavonol (Sharayei et al., 2019). There is evidence of E. coli O104:H4 inoculated in ground chicken becoming more sensitive to heat in the presence of the pomegranate powder extract (Juneja et al., 2016). Films prepared with fish gelatin and incorporated with pomegranate peel powder exhibited antibacterial activity against S. aureus, L. monocytogenes, and E. coli, with a more pronounced effect on Gram-positive bacteria rather than in E. coli (Hanani et al., 2019). When phenolic extracts from pomegranate peel, oregano, and clove were tested against Gram-positive and Gram-negative bacteria, the former were in general more sensitive to the extracts, as has been already mentioned. Foodborne pathogens such as Shigella flexneri and S. aureus were strongly inhibited by the extracts (Chan et al., 2018). In the same study, over-acidification was reported as one of the modes of action of phenolic acids, and as an outcome, lactic acid bacteria (LAB) were more resistant to the extracts compared to other foodborne bacteria, since they are naturally prepared to resist acidic environments. LAB were able to detoxify the phenolic acids through their metabolism, not being therefore inhibited by polyphenol-rich extracts (Chan et al., 2018). Pomegranate peel extract obtained by ultrasound-assisted extraction was also effective against Aspergillus niger; when pomegranate peel (3000 mg/L) and potassium sorbate (1000 mg/L) antifungal activities were compared, they were not significantly different in efficiency (Sharayei et al., 2019). Tannins, mainly catechin, catechin-3-gallate, gallocatechin, and gallocatechin-3-gallate, are present in persimmon fruit (Besada and Salvador, 2011). Persimmon tannins’ antibacterial activity is expressed

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through cell membrane hyperpolarization, which decreases membrane potential, leakage of intracellular ATP content, and cell protein defective synthesis and loss, as well as cell wall and membrane damage or partial disintegration. In fact, persimmon tannins were active against methicillin-resistant S. aureus isolated from pork, suppressing cell proliferation, changing cell morphology and destroying its integrity, decreasing membrane potential and intracellular APT concentration, and damaging the cell wall and membrane (Liu et al., 2019). Gingeron, zingerone, and capsaicin have been also found to be sporostatic for B. subtilis (Al-Khayat and Blank, 1985). When studying the relationship between structure and inhibitory actions, Katayama and Nagai (1960) assayed 32 pure phenol compounds. They found that 0.05% eugenol, carvacrol, isoborneol, thymol, vanillin, or salicyldehyde in agar were inhibitory against B. subtilis, S. Enteritidis, P. aeruginosa, P. morganii, and E. coli, concluding that the presence of a hydroxyl group enhanced the antimicrobial activity. López-Malo et al. (1995) demonstrated that vanillin concentration and the type of agar significantly (p < 0.05) affected the radial growth rate of Aspergilli. Also, differences among mold response were reported; the most resistant mold to the conditions assayed was A. niger, followed by A. parasiticus, A. flavus, and A. ochraceus. The presence of 1000 ppm of vanillin inhibited A. ochraceus growth for more than 2 months at 25°C in potato dextrose agar (PDA), while the growth of A. niger, A. flavus, and A. parasiticus was inhibited by 1500 ppm; results obtained in vitro were corroborated for A. flavus and A. parasiticus in apple-, banana-, mango-, papaya-, and pineapple-based agars and for A. ochraceus and A. niger in papaya-, pineapple-, and apple-containing agars. On the other hand, inhibitory concentrations of vanillin for A. niger and A. ochraceus in mango- and banana-based agars were greater than those found in PDA. Resnik et al. (1996) analyzed the effect of the concentration of vanillin on the growth rate and aflatoxin accumulation of A. parasiticus; the growth rate decreased abruptly in the presence of 250 ppm of vanillin, whereas 1500 ppm of vanillin inhibited mold growth during at least 37 days of storage at 28°C. However, 500 ppm of vanillin enhanced aflatoxins B1 and G1 accumulation, the toxin levels exceeding those of the control. Plant extracts, which may contain active compounds such as polyphenols or essential oils, can be used as a whole in microbial activity mitigation. Ginger, thyme, and mostly pomegranate and clove ethanolic extracts were effective against B. cereus, S. aureus, E. coli, and S. typhi (Mostafa et al., 2018). On the other hand, rosemary extract was effective against B. subtilis, mainly when nano-encapsulated in chitosan (Lee et al., 2019). Bullerman (1974) observed the inhibitory effect of cinnamon on an aflatoxinproducing mold, and reported that a 1% to 2% level of ground cinnamon in broth allowed some growth of A. parasiticus but eliminated approximately 99% of the production of aflatoxins.

17.2.2 Essential Oils Essential oils are one of the most studied groups of natural antimicrobials from plant sources. They contain a variety of compounds of different chemical classes, although natural phenolics have been reported to be the main antimicrobial compounds. Due to their low toxicity and easy degradation, they are considered environmentally friendly (Jafri et al., 2019), and as a matter of fact, most of the essential oils studied for their antimicrobial activity come from herbs or spices utilized as food (Freitas and Cattelan, 2018). Essential oils are volatile liquids, constituted by both polar and non-polar low-molecular-weight compounds, whose antimicrobial activity relies mostly on one or a few major components found in relatively high concentrations. Nevertheless, their effect on microbial cells can be due to synergy among different components (Gutierrez-del-Rio et al., 2018; Mani-López et al., 2018). Sometimes, their antimicrobial activity can be greater than conventional antimicrobials; such was the case of clove essential oil encapsulated in sodium alginate particles which showed stronger activity in situ against S. aureus compared to nitrite (Radunz et al., 2019). In general, the lipophilic and hydrophobic nature of an essential oil allows its passage through cell membranes, affecting the conformation of the polysaccharides, fatty acids, and phospholipids, and interacting with membrane proteins; as consequence, the leakage of ATP and other cellular content, as well as movement of proton flux outward from the cell, cause cell damage and death. The mitochondrial membrane is also damaged by essential oil components (Gutierrez-del-Rio et al., 2018; Mostafa et al., 2018; Dutra et al., 2019).

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The active compounds present in essential oils mainly belong to terpenes (monoterpenes, diterpenes, and sesquiterpenes) and terpenoids (monoterpenoids), although they can also be aldehydes, phenols, and methoxy derivatives (Gutierrez-del-Rio et al., 2018; Baldim et al., 2019; Jafri et al., 2019). Terpenes are hydrocarbons formed by jointed isoprene units and are the most diverse type of secondary metabolites with more than 55,000 constituents (Jafri et al., 2019). Thymol and carvacrol are, so far, considered the most active monoterpenes (Jafri et al., 2019) and together with p-cymene are the major volatile components of oregano, thyme, and savory, and likely account for their antimicrobial activity (Beuchat, 1976). The essential oil of oregano contains up to 50% thymol; thyme has 43% thymol and 36% p-cymene, while savory has 30–45% carvacrol and 30% p-cymene (Farag et al., 1989). On the other hand, terpenoids are terpene derivatives with additional oxygen or other elements, and highly active compounds with considerable antimicrobial activities; examples of terpenoids found in essential oils are linalool, linalyl acetate, citronellal, piperitone, menthol, and geraniol (Jafri et al., 2019). It has been reported that thymol, cinnamic aldehyde, eugenol, and other essential oil components have a wide antimicrobial spectrum (Wilkins and Board, 1989; Beuchat and Golden, 1989; Davidson, 1993). Thymol is the major component of thyme essential oil, as already suggested, and apart from interacting with periplasmatic protein and enzymes, it also induces the overexpression of outer-membrane proteins, causing unfolding of outer-membrane proteins and changing the membrane permeability. There is a loss of membrane potential, K+ leaking, and ATP losses (Jafri et al., 2019). Carvacrol, the main component of oregano essential oil, is believed to increase membrane fluidity and permeability due to the formation of channels in the membrane, allowing ion passage through the membrane (H+ inward and K+ outward), and leaking of nucleic acid and other intracellular materials (Dutra et al., 2019). It also affects ATP production and might even interfere in the bacterial quorum sensing signaling mechanism (D’Amato et al., 2018; Gutierrez-del-Rio et al., 2018; Dutra et al., 2019). Indeed, carvacrol was effective against E. coli strains O157, O26, O45, O103, O111, O121, O145, and O104, affecting their membrane permeability and also intracellular ATP levels (Stratakos et al., 2018). It has been reported that carvacrol and thymol had lesser inhibitory effects on mold growth than extracts from oregano and clove, suggesting that the antimicrobial activity may be due to several compounds from the extracts. These findings show that the phenolic extracts from plants may contain not only phenolics but also other compounds that could possess antimicrobial activities, and that, in effect, essential oil components can work synergistically. Paster et al. (1990), evaluating the antimycotic capacity of essential oil of oregano and clove against three strains of Aspergillus, indicated that the studied molds differed in their sensibility to tested extracts and found that A. flavus was sensitive to the essential oil of oregano. Zaika et al. (1983) found that little if any inhibition of growth and acid production by Lactobacillus plantarum and Pediococcus acidilactici was noted in the presence of 40 ppm oregano oil, whereas levels >200 ppm were bactericidal to both organisms; in addition, they reported that bacteria exposed to sub-lethal concentrations of oregano oil were able to overcome the inhibition and to develop resistance to the toxic effect of oregano oil or oregano. In a recent study, oregano essential oil was effective against bacteria of the Alicyclobacillus genera (Dutra et al., 2019). Apart from oregano and thyme, other essential oils such as cinnamon, clove, and rosemary essential oils are well-known as being very effective against microorganisms (Xu et al., 2018; Stojiljkovic et al., 2018). Cinnamon essential oil’s main component is cinnamaldehyde (or cinnamic aldehyde), a phenylpropene (Jafri et al., 2019; Chuesiang et al., 2019b), and was capable of affecting Alternaria alternata cell membrane integrity and permeability while changing the morphology of its hyphae (Xu et al., 2018). It is suggested that in fungi, cinnamaldehyde also inhibits cell division and disturbs the function of enzymes, including those involved in cell wall construction (Jafri et al., 2019). Cinnamaldehyde also has the ability to interfere with the activities of cytokinases, disrupting bacteria cell membranes and amino acid decarboxylases (D’Amato et al., 2018; Ji et al., 2019). Bullerman et al. (1977) demonstrated that the essential oil of cinnamon at a concentration of 200 ppm was inhibitory to growth and subsequent toxin production by A. parasiticus and that cinnamic aldehyde, the major component of cinnamon oil, was effectively inhibitory at a level of 150 ppm; however, they did not discard other minor constituents that may also have antimicrobial activity. In nanoemulsions formed by phase inversion temperature, cinnamon essential oil was able to disrupt the cell wall, leading to the expulsion of internal cellular material of E. coli, S. Typhimurium, S. aureus, and V. parahaemolyticus (Chuesiang et al., 2019a). Likewise, corn

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starch film with 2% cinnamon essential oil was effective against S. aureus, E. coli, and S. enterica for more than 120 days (Ke et al., 2019). Eugenol, a phenylpropene, can also be found in cinnamon essential oil (Ji et al., 2019), and it is known to affect the yeast cell membrane and wall, causing leakage of cellular content (Ji et al., 2019). Eugenol is also one of the main components of clove essential oil, together with carvacrol, thymol, and cinnamaldehyde (Chaieb et al., 2007). Eugenol has also exhibited antimicrobial activity, which is due to its capacity to permeate through the cell membrane causing K+ and ATP leakage, preventing cell enzyme activity, and interacting with proteins (D’Amato et al., 2018; Jafri et al., 2019). Eugenol has been reported by Al-Khayat and Blank (1985) as one of the most effective natural antimicrobials from plant origins acting as a sporostatic agent: significant reductions of viable Bacillus subtilis spores were obtained when exposed to 0.1–1.0 % eugenol for 8 days at 37°C. Clove essential oil had a bactericidal effect against S. aureus, E. coli, L. monocytogenes, and S. Typhimurium (Radunz et al., 2019). Rosemary essential oil is another effective antimicrobial and its main components are 1,8-cineole (33.08–38.50%) and camphor (13.55–18.13%) (Hcini et al., 2013; Stojiljkovic et al., 2018). The oil inhibited L. monocytogenes, E. coli, and S. enterica growth, and had an additive effect against Gram-positive and Gram-negative bacteria in combination with clove oil (Stojiljkovic et al., 2018). Ji et al. (2019) evaluated the essential oils vapors of 97 essential oils, among which garlic, cinnamon bark, mountain pepper, citronella, thyme, oregano, and spearmint displayed considerable activities against Penicillium corylophilum. The synergistic activity of cinnamon bark, citronella, and mountain pepper was also useful in the inhibition of P. corylophilum in beef jerky (Ji et al., 2019). α-terpineol and terpene-4-ol, the main components of tea tree essential oil, interfered with A. niger cell membrane permeability, hyphae morphology and mycelium growth, and spore morphology and germination, as well as with metabolic pathways (An et al., 2019). The main components, as well as the oil itself, showed strong antimicrobial activities against A. niger, in vitro and in grapes (An et al., 2019). Geraniol, nerol, and citronelol are examples of non-phenolic essential oil components with reported antimicrobial activity; Mahmoud (1994) found that 1000 ppm of geraniol, nerol, or citronelol inhibited Aspergillus flavus growth in nutritive broth (pH 5.5) during 15 days of incubation at 28°C, the MIC being 500 ppm for the three alcohols. López-Malo and Argaiz (1999) evaluated the effects of pH (6.5, 5.5, 4.5, or 3.5) and citral (3,7-dimethyl-2, 6-octadienal) concentration (0, 500, 1000, 1500, or 2000 ppm) on the growth of A. flavus, A. parasiticus, Penicillium digitatum, and P. italicum in PDA adjusted with sucrose to aw 0.97; the radial growth rate of molds increased as the pH increased and citral concentration decreased. Conversely, reducing pH and increasing citral concentration increased mold germination times; inhibitory citral concentration depended on pH and differed among molds. P. digitatum and P. italicum were inhibited with 500 ppm citral at pH 3.5 and 6.5, whereas 1000 ppm was required to inhibit A. flavus and A. parasiticus, while at pH 4.5 and 5.5 inhibitory concentrations varied from 1000 ppm for Penicillium to 2000 ppm for Aspergillus. The evaluation of essential oil antimicrobial activity against foodborne microorganisms is no longer confined to traditional in vitro methods; they have been tested in vapor phase, in foods, in foodpackaging models, combined with other technologies such as high-pressure and modified atmosphere, or even utilized in nets for crop protection against pests (Loziene et al., 2018; Black-Solis et al., 2019; Park et al., 2019; Reyes-Jurado et al., 2019b). Essential oil incorporation in carrier polymers has been widely considered recently, aiming at their holding and preservation, controlled release, improved activity, or to diminish the effect of their strong odors (Baldim et al., 2019; Mazarei and Rafati, 2019; Radunz et al., 2019). Studies have been implemented in which the antimicrobial activity of encapsulated or entrapped essential oil has been tested, reporting liposomes (Hammoud et al., 2019), starch films or bionanocomposites (Garrido-Miranda et al., 2018; Ke et al., 2019), and micro- and nanoemulsions (Mazarei and Rafati, 2019), among other means to entrap the essential oil. On this basis, lipid nanosystems were used for the encapsulation of Lippia sidoides essential oil, which were effective against Candida albicans (Baldim et al., 2019), while encapsulation using randomly methylated β cyclodextrin (RAMEB) improved thyme essential oil antimicrobial activity against Schizosaccharomyces pombe, S. aureus, and E. coli by 2 to 4 fold (Das et al., 2019). Nanoemulsions of carvacrol and Satureja khuzestancia essential oils showed antimicrobial activity against S. enterica and S. aureus (Mazarei and Rafati, 2019) while solid lipid micro-particles loaded with cinnamon oleoresin were effective against Candida

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pseudointermedia and Penicillium paneum (Procopio et al., 2018). Other nanostructures that were useful against fungi were zinc oxide nanoparticles, which combined with carvacrol showed bactericidal activity against Campylobacter jejuni; when carvacrol was used on its own, the effect was merely bacteriostatic while carvacrol induced cell membrane damage, and zinc oxide nanoparticles induced cell leakage (Windiasti et al., 2019). Regarding essential oil-loaded films with antimicrobial activity, corn-starch-based film containing cinnamon essential oil was effective against S. aureus, E. coli, and S. enterica (Ke et al., 2019). Tests have also been performed in foods, with successful results; coatings prepared with quinoa protein and chitosan, and filled with thymol nanoparticles, were able to reduce Botrytis cinerea growth in tomato (Robledo et al., 2018). In another study, the use of chitosan/montmorillonite bio-nanocomposites with rosemary essential oil for fresh poultry meat packaging reduced Bacillus cereus and Salmonella enterica counts in the meat (Souza et al., 2019). Likewise, carvacrol and thymol were effective in reducing microbial growth in marinated fresh chicken stored in vacuum packaging for 21 days at 4°C (Karam et al., 2019). Although some of the above-mentioned tests prove the efficacy of essential oils in protecting foods against foodborne microorganisms, essential oil efficacy in foods may differ from their efficacy in vitro. Essential oil antimicrobial activity is affected by different factors such as pH, temperature, and water activity, with increasing stability and effectiveness at low temperatures and pHs (Mani-López et al., 2018). Food composition may, indeed, increase or decrease essential oil activity (Freitas and Cattelan, 2018) and hinder or favor microbial growth. When microorganisms grow in food, they may find a friendlier environment, with more nutrients available, compared to growing medium, which help them fight the antimicrobial damage (Freitas and Cattelan, 2018). Besides, essential oil antimicrobial activity can decrease due to interaction with protein, fat, or enzymes present in a particular food, and because of such interactions, results are difficult to compare to other foods (Guitiérrez-del-Rio., 2018). If, in foods, higher essential oil concentrations are needed to inhibit microbial growth compared to growing media, food sensory properties might, in some cases, be affected (Goni et al., 2009). Requena et al. (2019) found carvacrol, eugenol, oregano, and clove essential oils more effective against E. coli and Listeria innocua in vitro, compared to their inoculation in foods such as cheese, chicken breast, pumpkin, and melon. The essential oils and their components had different effect on bacterial growth depending on the tested food, which highlighted the importance of food composition in foodborne microorganism nutrition and survival (Requena et al., 2019). Other spices and oils have also been shown to have some antimicrobial activity; Marth (1966) reviewed studies that reported antimicrobial activity by spice oils and extracts of laurel, peppers (chilies), coriander, anise, carvone, peppermint, caraway, cardamom, cumin, fennel, celery, dill, and mustard. Many of these preparations seemed to be active against only one microbial strain, and some were even active in one study but not in another. The normal inhibitory concentrations used in these studies were very high, indicating little activity. Other spices, including rosemary, sage, and turmeric, also possess antimicrobial activities. Curcumin, found in turmeric, for instance, was used for Aspergillus flavus spore inactivation (Temba et al., 2016). Hydrosols, the aqueous phase obtained from plant-containing essential oil hydro-distillation, hold small amounts of dispersed essential oil and can be used as antimicrobials, both in vitro and in foods, against bacteria and fungi. The essential oil volatile components present in the hydrosol, mainly phenolics, will affect the bacterial membrane structure and function through similar mechanisms known for pure essential oils (D’Amato et al., 2018).

17.2.3 Phytoalexins Phytoalexins are normally low-molecular-weight compounds (Ahuja et al., 2012; Kuc and Rush, 1985) elicited de novo by a plant when exposed to a hostile biotic or abiotic agent (Luis et al., 1997; Ejike et al., 2013). This occurs at the infection site, as a way to surpass the aggression. Biotic elicitation can be caused by bacteria, fungi, viruses, nematodes, and insects (Kuc and Rush, 1985), while abiotic elicitation is normally triggered by certain environmental condition or compounds such as ethylene and fungicides (Kuc and Rush, 1985). Their effectiveness against plant pathogens such as fungi and bacteria (Kuc and

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Rush, 1985; Ahuja et al., 2012) suggests their potential as foodborne pathogen antimicrobials, and it is, in fact, known that they are broad-spectrum antimicrobial agents. More than 200 phytoalexin compounds have been isolated from more than 20 families of plants, normally belonging to different classes of compounds such as terpenes, terpenoinds, and isoflavonoids and, to a lesser extent, proteins (chitinases, thionins, zeamatins, thaumatins, etc.) (Kuc and Rush, 1985; Sofos et al., 1998; González-Lamothe et al., 2009; Savoia, 2012). Their action against pathogens is thought to be multisite, and so affects different physiological and biochemical pathways. Phytoalexins with terpene, terpenoid, and isoflavonoid structures are believed to disrupt the cell membrane, a statement that could be supported by the phytoalexins’ lipophilic nature (Guest, 2017). Indeed, it was suggested that the mechanism through which glyceollins, a class of phytoalexins, express their antibacterial activity is by affecting cell membranes (Arora and Strange, 1991). On the other hand, there have been suggestions of dead causative oxidative phosphorylation in fungal cells due to phytoalexins, for which free phenolic groups seem to impart an important contribution (Guest, 2017). As a matter of fact, phytoalexins are mainly active against fungi, although they also have an effect on bacteria (Sofos et al., 1998; Smid and Gorris, 1999; González-Lamothe et al., 2009). There are reports of phytoalexin development in different food-based plants due to fungal attack, which is related, in a broad way, to phytoalexin antifungal capacity. Tsibulins 1d and 2d accumulated in onion bulbs, preferentially in necrotic tissue rather than in fresh tissue, as triggered by B. cinerea (Dmitriev et al., 1990). Likewise, Arora and Strange (1991) reported the development of phytoalexin in lettuce, as a response to the threat of B. cinerea, Bremia lactuacae, and Pseudomonas syringae. Three years later Bennett et al. (1994) concluded that lettucenin A, found in lettuce, had antimicrobial activity against B. cinerea, B. lactuacae, and P. syringae. Indeed, a correlation was found between lettucenin A accumulation and the inhibition of microbial growth, and it was also suggested that lettucenin A concentrations obtained in vivo could inhibit bacteria and fungi in vitro (Bennett et al., 1994). Phytoalexins have also been identified in food products such as banana leaves and fruits, triggered by kanamycin or Mycosphaerella fijiensis (Arora and Strange, 1991). Phytoalexins have also been found in legumes and cereals triggered either by fungal activity or by other factors. Phytoalexins occur in peanuts as a result of elicitation with resveratrol (Arora and Strange, 1991), and in sorghum the flavonoid 3-deoxyanthocyanidin can be found. In corn the diterpernoid kauralexin and the sesquiterpenoid zealexins are triggered after infection with fungi (Dmitriev et al., 1990). Phytoalexins are also produced in rice, normally diterpenoids, triggered by substances such as chitin oligosaccharides, proteins such as xylanase, fungus, or even UV light (Dmitriev et al., 1990). Phaseolin A occurs in bean pods, and it is triggered by monilicolin A from Monilinia fructicola, while glyceollins and glycinol are found in soybean cotyledons because of elicitation by Phytophthora megasperma cell wall glucan (Kuc and Rush, 1985). Finally, gluceollins, phytoalexins originated in soybean, showed antifungal activity against Diaporthe phaseolorum, Macrophomina phaseolina, Phytophthora sojae, Sclerotinia sclerotiorum, Cercospora sojina, Phialophora gregata, and Rhizoctonia solani and antibacterial activity against P. syringae, Xanthomonas campestris, Bacillus licheniformis, B. subtilis, S. aureus, and Corynebacterium flaccumfaciens (Arora and Strange, 1991). In many cases, the high concentrations needed to exert antimicrobial action in food matrices may explain the scarcity of examples of their actual use in foods (Sofos et al., 1998).

17.2.4 Phytoanticipins Phytoanticipins, compounds belonging to terpenoid derivatives, diterpenes, sesquiterpenes, flavonoid derivatives, and phenylpropanoid phenols, can also possess antimicrobial activity (Pedras and Yaya, 2015). Unlike phytoalexins, phytoanticipins are already present in plant tissue before it experiences unfavorable conditions. However, under hostile conditions, phytoanticipin concentration goes up (Ejike et al., 2013). Soledade et al. (1997) mentioned sinalexin as a phytoanticipin produced in white mustard, elicited by Alternaria brassicae (Soledade et al., 1997), that prevents white mustard from being attacked by Alternaria blackspot. This disease is caused by Alternaria brassicae, which is a hint of protection of some sort by the phytoanticipin (Soledade et al., 1997).

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Allicin, a phytoanticipin present in Allium genus plants, was described as a colorless oil, extremely pungent, that characterized the principal odor and taste of garlic and onion. It was reported that allicin in concentrations of 1:85,000 in broth was bactericidal to a wide variety of Gram-negative and Grampositive organisms. Investigations have shown that extracts from Allium bulbs inhibit the growth and respiration of pathogenic fungi and bacteria. Aqueous extracts from fresh garlic bulbs at levels of 3, 5, and 10% inhibited the growth of B. cereus on nutrient agar plates by 31.3, 58.2, and 100%, respectively (Saleem and Al-Delaimy, 1982). Similarly, allicin had a bactericidal effect against E. coli, proportional to the concentration of allicin used, and was effective against M. luteus in vapor phase. Other bacteria that were inhibited by allicin were Agrobacterium tumefaciens, Erwinia carotovora, P. syringae, and Xanthomonas campestris; some molds such as Alternaria brassicicola, B. cinerea, Magnaporthe grisea, and Plectosphaerella cucumerina were inhibited as well (Curtis et al., 2004). Many foodborne pathogenic bacteria are sensitive to onion and garlic extracts; S. aureus, B. cereus, Clostridium botulinum, S. Typhimurium, and E. coli have been adversely affected by garlic extracts (Beuchat and Golden, 1989). The antimicrobial effects reported for garlic and onion were attributed to the allicin concentrations and other sulfur compounds present in their essential oils, which might interfere with microorganism cell protein synthesis (Putnik et al., 2018). Mantis et al. (1978) studied the effect of garlic extracts on S. aureus in culture media; they reported that a 5% garlic extract concentration had a germicidal effect on S. aureus, whereas concentrations of garlic extracts equal to or greater than 2% had a clear inhibitory effect, while concentrations less than 1% were not considered inhibitory. An investigation of the effect of garlic and onion oils on toxin production by C. botulinum in meat slurry indicated that these oils, when used in the proportion of 1500 µg/g meat slurry, inhibited toxin production by C. botulinum type A; however, the inhibition was incomplete, and toxin production by C. botulinum type B and type E was not inhibited (DeWit et al., 1979). Yin and Tsao (1999) evaluated the antifungal activity of Allium plant (including garlic) extracts against Aspergillus niger, A. flavus, and A. fumigatus and found that garlic was effective as a fungicidal in concentrations between 35 and 104 ppm.

17.2.5 Antimicrobial Peptides (AMPs) Although not as well-known as phenolic compounds, plant proteins and peptides can also present antibiotic and antifungal activity, and be attractive for food applications. AMPs lack toxicity to mammalian cells (Fliss et al., 2015; Nordström and Malmsten, 2017); they are thermostable and specific to certain targets (Ben Said et al., 2019); these might be appealing characteristics. They normally have a molecular weight between 2 to 10 kDa, variable amino acid sequences and structure motifs (Fliss et al., 2015), are high in cysteine residues (4 to 12), and form disulfide bonds (Lipkin et al., 2005). AMPs are constituents of the plant immune defense mechanism, and as such can be isolated from different plant organs such as seeds, roots, stems, leaves, and flowers (Ben Said et al., 2019; Fliss et al., 2015; Tang et al., 2018). As part of plant defense, the specimen has increased resistance against the pathogen; this was observed in tomato plants expressing alpha-helical cationic peptide, which better resisted bacterial wilt caused by Ralstonia solanacearum (Morais et al., 2019). AMPs’ antimicrobial activity is based on their amphipathic structure, allowing attachment to both lipidic and phospholipidic groups, and the formation of ion channels in the membrane; these might destabilize and disintegrate the membrane, with irreversible damage to the cell (Cowan, 1999; González et al., 2017; Ben Said et al., 2019). They can also affect the formation of the cytoplasmatic membrane septum and cell wall or attach to outer-membrane AMP receptors, allowing the membrane to be permeable to metabolites (Ben Said et al., 2019; Tang et al., 2018). Some might instead interfere with intracellular functions, such as DNA replication and nucleic acid and protein synthesis; cellular enzymatic activity might also be altered, affecting microbial growth (Ben Said et al., 2019; Tang et al., 2018). When a proteolytic fraction from Vasconcellea cundinamarcencis was used against B. cinerea, the mold endured conidia germination and adhesion inhibition, as well as germ tube elongation; this mold also suffered from changes in the membrane integrity, and mycelium cells became hyper-sensible to cell wall-perturbing agents (Torres-Ossandón et al., 2019). B. cinerea was also inhibited by an AMP isolated from amaranth (Amaranthus retroflexus L.) seeds, along with other fungi such as Fusarium culmorum, Helminthosporium sativum, and Alternaria consortiale; the AMP, although not effective against

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Gram-negative bacteria, was effective against Gram-positive ones (Lipkin et al., 2005). As for Gramnegative bacteria, a 10 kDa AMP isolated from bitter melon seed extract was effective against E. coli, S. typhi, and Pseudomonas aeruginosa (Jabeen and Khanum, 2017). As natural antimicrobials regarded for food applications, AMP immobilization in suitable matrices has been suggested, aiming at improving their effectiveness (Ben Said et al., 2019; González et al., 2017). AMP isolation, however, is a challenging task that relies on elaborated techniques to be accomplished (Tang et al., 2018; Jabeen and Khanum, 2017).

17.2.6 Other Sources of Plant Antimicrobial Agents Organic acids are also known to hold antimicrobial properties, and their use in food preservation is not a recent practice. Organic acids such as citric, succinic, malic, and tartaric can be found in fresh fruits and vegetables. Others, such as lactic and propionic acids, can be obtained during fruit or vegetable fermentation (Gould, 2002). Another group of compounds that might sometimes present antimicrobial activity are alkaloids; those of Gymnema montanum and Tabernaemontana catharinensis extracts have antimicrobial properties and the ability to change membrane permeability (Savoia, 2012). Certain medium chain fatty acids such as caprylic, capric, and lauric acid are known as natural antimicrobials and were utilized in combination with essential oils for E. coli O157:H7 inhibition (Kim and Rhee, 2016). Likewise, a water-soluble polysaccharide labeled PLP1 was isolated from root barks of Periploca laevigata and exhibited antimicrobial activity against Gram-positive and Gram-negative bacteria; the polysaccharide was effective against E. coli but rather ineffective against S. Typhimurium (Hajji et al., 2019). Interestingly, Maillard reaction products obtained from the reaction of protein hydrolysate from sunflower meals, xylose, and cysteine (pH 7.4; heated at 120°C for 2 h) showed antimicrobial effect against S. aureus and E. coli; antimicrobial activity was attributed to melanoidins, which were suggested to destruct cell membrane through chelation of the stabilizing cation Mg2+ (Habinshuti et al., 2019).

17.3 Testing the Efficacy of Antimicrobials Essential oils of many plants possess useful biological and therapeutic activities and are extensively utilized for the preparation of pharmacologic drugs. They are commercially recovered from plant materials primarily by steam distillation, solvent extraction, or pressing, and their use in the food industry is influenced by the nature of their constituents. The antimicrobial activity of essential oils and extracts from plants, herbs, and spices depends on the extraction method but also on the initial quantity of essential oil in the plant. Within the same spice or plant, the levels of constituents and therefore active antimicrobial groups can substantially vary. Also, the geographic zone of cultivation may influence extract composition. The antimicrobial effectiveness of essential oils varies depending on the harvesting time within the year (Quesada et al., 2016). Therefore, there is a necessity to establish methods to fix or standardize essential oil purity or concentration of active components. Knowledge of the qualitative and quantitative composition of essential oils will allow valuable data to be obtained in systematic biological studies of plants as antimicrobial agents. Differences in the antimicrobial effectiveness of plant extract compounds from different sources, extraction methods, and geographic zones can be diminished if the active antimicrobial compounds from extracts or essential oils are identified. Gas chromatography is an interesting tool to measure the qualitative and quantitative composition of essential oils and its use has been widely reported in the literature (Velázquez-Nuñez et al., 2013; Hernández-Hernández et al., 2019; Requena et al., 2019). The adoption of plant or spice essential oils as alternatives to other preservatives will depend on antimicrobial uniformity and/or on the analytic methods available to normalize their antimicrobial potency (López-Malo et al., 2000). The concentration of antimicrobial compounds varies depending on the type of plant; among the richest are clove, nutmeg, laurel, and mustard, while the major constituents in most cases are phenolic compounds (Turgis et al., 2008; Aguila-González et al., 2015;

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Gómez-Heincke et al., 2016). Many publications about the antimicrobial activity of extracts, oils, spices, and herbs can be found in the food-related scientific literature. However, it is difficult to make quantitative comparisons of their effects, at least partially, because of the great variety of methods utilized to evaluate antimicrobial efficiency (Reyes-Jurado et al., 2019b; Freitas and Cattelan, 2018; Ncama et al., 2019). Parish and Davidson (1993) have critically discussed the numerous tests that have been used as well as the factors that are important to consider in determining the effectiveness of natural antimicrobials in foods. The screening and isolation of antimicrobial agents from plants require a multidisciplinary approach (Davidson and Naidu, 2000); in vitro or in vivo screening bioassays are useful determining factors for the successful isolation of active compounds. Solvent selection for extraction including solvent ability to extract components is an important factor determining operation efficiency. Dry plants could be extracted with a variety of solvents and sometimes sequentially from low to high polarity. Polar solvents such as ethyl acetate or methanol are often used. Ethyl acetate theoretically only extracts by leaching the sample while alcoholic solvents presumably rupture cell structures (membranes), extracting also intracellular materials. For fresh plant materials (with high water content) a solvent mixture of dichloro-methane-methanol gave better results during extraction. Methanol separation and sample partition followed by ethyl acetate and butanol extraction help to separate lipophilic compounds from water-soluble materials.

17.3.1  In Vitro Testing In vitro or explanatory (endpoint and descriptive methods) and applied (inhibition curves and endpoint methods) tests are the most utilized methods for antimicrobial efficiency evaluation of essential oils of plants and spices. A great number of these studies have been accomplished in vitro, and fewer have been accomplished in foods. Different methods, with different principles, are applied for the in vitro study of essential oils effectivity, namely dilution and diffusion methods. The former consists of diluting essential oil into the growing medium, while in the latter essential oil is loaded into a filter paper which then is added to the surface of a culture medium (Freitas and Cattelan, 2018). Ncama et al. (2019) distinguished between qualitative and quantitative methods. Some qualitative methods described are disc- or agar-diffusion, cylinder method, and thin-layer chromatography. In the first method a microbial culture is inoculated in agar, and a filter paper, dipped in the plant extract, is placed on the agar. In the second method, a specific volume of antimicrobial agent is placed into a cylinder containing the inoculated agar and mycelium growth is observed. In the third method a thin-layer chromatography plate with antimicrobial agents is dipped into broth containing the fungi and incubated in a humid atmosphere at a suitable temperature for fungal growth. The zones where inhibition occur can be visualized using reagents such as tetrazolium salts. A quantitative method described by Ncama et al. (2019) is known as the mycelia plug method and consists of a modified agar dilution method, in which agar is enriched with the antimicrobial agent at a specific concentration; inhibition is quantified through comparison between radial mycelia growth observed in medium treated with the antimicrobial agent and in the untreated control. Within the evaluation methods in model systems, descriptive (inhibition curves) methods have been commonly utilized for bacteria and yeasts, while endpoint tests have been used for bacteria, yeasts, and molds. Zaika (1988) concluded that many factors affect the antimicrobial activity of spices, extracts, and essential oils and mentioned that several aspects must be considered and reported; the observed inhibition depends on the evaluation method. Microorganisms differ in their resistance toward spices or herbs. Additionally, food components can increase (by the presence of acids, humectants, antimicrobials, etc.) or reduce (by partitioning of active components into the lipid phase, etc.) the antimicrobial capacity.

17.3.2 Vapor Phase Many of the studies regarding the application of essential oils against foodborne microorganisms entail the direct use of the oils in the medium or food. This practice carries some disadvantages when one considers the real application of essential oils in food products for human consumption. The most notorious drawback of direct application of essential oil in food is the effect on food sensory properties, which is

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not as pronounced when essential oils are used in the vapor phase, since essential oils are not in direct contact with food. On the other hand, when essential oils are used in the vapor phase, lower concentrations are needed to guarantee the same effectiveness, which also contributes to lowering the effect on sensory properties (Velázquez-Nuñez et al., 2013; Lorenzo-Leal et al., 2019). Some studies have, in fact, suggested that essential oils are more efficient in vapor phase compared to direct application in foods, and authors such as Aguilar-González et al. (2015), Songsamoe et al. (2017), and Chen et al. (2018b) provided recent results on essential oil effectiveness against bacteria and molds when applied in vapor phase. Essential oil antimicrobial components are mainly volatiles which supports their application in vapor phase for potentially improved results (Nedorostova et al., 2009). When essential oils are used in the vapor phase, volatiles act by destabilizing microbial cell membranes (Goni et al., 2009). Besides, essential oil antimicrobial components are known to be in the majority lipophilic, highly hydrophobic, and presenting low water solubility; therefore, due to their lack of affinity with water present in most food products, they form micelles in aqueous solutions, which hinders their activity as antimicrobials (ReyesJurado et al., 2019b). In direct application, essential oil activity is mainly due to the action of hydrophilic substances, which are normally not as effective as the hydrophobic ones. All of the abovementioned explains why essential oil application in vapor phase could be an alternative to reduce their impact on preserved food flavor (Lorenzo-Leal et al., 2019), and to reduce irritation while allowing the utilization of lower concentrations of an essential oil for the same antimicrobial activity (Reyes-Jurado et al., 2019b). Reyes-Jurado et al. (2019b) point out that since essential oils contain different volatile compounds with different volatilities, components will be released with different releasing times. If on one hand the antimicrobial activity of essential oils in vapor phase is directly related to their volatility, on the other hand in the direct addition of an essential oil to the medium, diffusivity and solubility are important for the distribution of the active compounds (Reyes-Jurado et al., 2019b). These factors might also suggest that the choice between applying essential oils in their vapor phase or directly to the medium depends on the food product to be preserved, the target microorganisms, and the solution sought. The vapor phase seems to be more suitable for microbial growth inhibition on food surfaces rather than in the inner parts of the food. Indeed, antimicrobial essential oil components with rather relevant volatilities can be well-distributed on the surface of growing medium or foods, and are effective against molds, which are very common on food surfaces. Reyes-Jurado et al. (2019b) emphasize the especial importance of using essential oils in vapor phase for mold growth inhibition due to easier airborne conidia inhibition, which are difficult to inactivate with the direct addition of antimicrobials. Another important application of essential oils in vapor phase consists in their incorporation into food package material for controlled release to the package headspace containing foods. Poly(vinyl alcohol) (PVA) films were incorporated with clove oil (1, 3, 5, 7, or 9% w/w) and used to cover minced Trichiurus haumela meat, without direct contact with the food; total viable counts in the fish meat preserved by the film lowered slightly with increasing essential oil content, and shelf life was extended for two days when films with at least 5% w/w essential oil were used (Chen et al., 2018b). Azadbakht et al. (2018) prepared chitosan films with Eucalyptus globulus essential oil, which reduced Listeria monocytogenes counts on sliced sausages, stored at 23°C. Although films with E. globulus essential oil showed increasing antibacterial activity with increasing essential oil concentration, the films were more efficient in direct contact with liquid medium than in vapor phase (Azadbakht et al., 2018). In another project, Michelia alba essential oil was used incorporated into an absorbent material based on the roots of water hyacinth for the inhibition of Aspergillus flavus in brown rice (Songsamoe et al., 2017). Several methods have been utilized to analyze essential oil effectiveness in vapor phase such as the inverted Petri dish method. In this method, a paper disc is impregnated with essential oil and introduced in the small headspace left by the Petri dish. Other methods comprise the use of a glass or plastic box in which the essential oil and agar (or food) inoculated with microorganism are introduced and kept apart, allowing contact of the essential oil vapor with the microorganisms (Reyes-Jurado et al., 2019b). In a study run by Tyagi and Malik (2011), E. globulus essential oil was tested in the vapor phase using the disc volatilization method and in direct addition using agar dilution and well diffusion methods; significantly higher antimicrobial activity was observed in vapor phase against Gram-positive (Bacilus subtilis and Staphylococcus aureus) and Gram-negative (Escherichia coli, Pseudomonas aeruginosa,

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and Pseudomonas fluorescens) bacteria, molds (Penicillium digitatum, Aspergillus flavus, Aspergillus niger, Mucor spp., Rhizopus nigricans, and Fusarium oxysporum), and yeasts (Candida albicans and Saccharomyces cerevisiae). Increasing essential oil concentration in the vapor phase increased microorganism inhibition zones, and these were higher in trials in the vapor phase compared to the ones with direct essential oil addition; microbial viability was also affected with 100% viability reductions for B. subtilis and C. albicans while 76.6, 72.3, and 82.9% viability reductions for P. aeruginosa, P. fluorescens, and E. coli, respectively, were observed after 12 h exposure. Ncama et al. (2019) cited a method consisting of putting the extract inside a well, and covering it with microtitration plates, on top of each of which the inoculated product is seated. Exposing the extracts to higher temperatures promoted evaporation, which allowed the vapors to be in contact with the product and avoid fungal spore or mycelial growth (Ncama et al., 2019). Additionally, time-saving methods have been suggested, using systems in which several microorganisms and essential oils can be tested at the same time. Kloucek et al. (2012) proposed a modified traditional disc volatilization method aiming at reducing labor and material needed, by using a foursection Petri dish (90 mm diameter), each with 5 mL of agar. Three compartments were inoculated while one was left un-inoculated for contamination control; a filter paper disc (85 mm diameter) was evenly impregnated with tested EO and laid on the walls dividing the compartments, allowing a 2 mm distance between the filter paper and growing media. Agar was poured into the lid as well, to function as a seal for the Petri dish. Once the Petri dish was closed with the agar-containing lid, it was incubated at selected conditions. There are several studies in the literature which report the synergistic effect of different essential oil components in the vapor phase; Ji et al. (2019) used an experimental set-up consisting of a modified checkerboard array to test the effect of combinations of garlic, cinnamon bark, may chang, citronella, oregano, and spearmint essential oils against Penicillium coryphilumon. A polycarbonate vial upper well containing agar inoculated with P. corylophilum was put into contact with a lower well containing a paper disc impregnated with different combinations of essential oils that eventually vaporized. After sealing the junction between the upper and lower wells with Parafilm, this system was incubated at 25°C for 120 hours, after which it was visualized for mold growth. Beef jerky inoculated with P. corylophilum was maintained in airtight container in the presence of essential oil to allow contact between essential oil vapor and food. Among the different combinations of essential oil tested, cinnamon bark combined with citronella essential oils’ vapors showed the highest synergistic antifungal activity against P. corylophilum, and in beef jerky this combination allowed a decrease in microbial population. The combination of cinnamon bark, citronella, and may chang essential oils was also very effective in vapor phase. In both cases, synergy between 1,8-cineole or eugenol with linalool was suggested. When each essential oil was tested separately, garlic essential oil showed the highest antimicrobial activity. Also using a checkerboard methodology, Netopilova et al. (2018) combined carvacrol and thymol in vapor phase, which generally generated a strong additive antimicrobial effect against 12 Staphylococcus aureus strains. When clove and cinnamon essential oils were combined in vapor phase, an antagonist effect was observed for E. coli, while against L. monocytogenes, B. cereus, and Yersinia enterocolitica the combination had a synergetic effect (Goni et al., 2009); the activity in vapor phase was better than with direct contact for the microorganisms mentioned, although this was not the case for P. aeruginosa. Solid-phase microextraction analysis identified eugenol as the major component found in the headspace; 1,8-cineole and camphor had a synergistic effect with eugenol when present in the headspace (Goni et al., 2009). According to the authors, the synergistic effect can be related to the sequential inhibition of certain biochemical pathways, the inhibition of protective enzymes, and the enhancing of antimicrobial uptake by the cell wall. On the other hand, an antagonist effect can be generated due to a decrease in components’ water solubility, making them less available to act on microorganisms (Goni et al., 2009). As previously mentioned, the effect of the application of M. alba essential oil in the vapor phase was tested against A. flavus on brown rice by Songsamoe et al. (2017), inhibiting mold spore germination and mycelium growth, and increasing brown rice shelf life up to four times; these authors also mentioned the advantage of essential oil components’ synergy, by pointing out the increasing antimicrobial effect of linalool, the major component of M. alba essential oil, by using only a small amount of caryophyllene, the second major component. Reyes-Jurado et al. (2019a) evaluated the effect of thyme, Mexican

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oregano, and mustard essential oils in vapor phase, against foodborne Gram-positive and -negative bacteria, as well as against fungi; the three studied oils were rather effective against tested fungi, among which are Aspergillus fumigatus, A. niger, Candida, and Penicillium expansum. They observed different sensitivities of the tested microorganisms to the essential oils, mustard vapor being the most effective in general. Oregano essential oil was not effective against the bacteria studied, including E. coli, P. aeruginosa, Salmonella Typhimurium, B. subtilis, and S. aureus. There also was a synergetic effect with the combination of thyme and Mexican oregano essential oils when these were used in proportions of either 25:25% or 25:50%. Aguilar-González et al. (2015) tested clove and mustard essential oils’ antimicrobial efficacy in vapor phase, by incorporating the essential oil in filter paper; tests were run against Botrytis cinerea in vitro and in strawberries. The two essential oils had synergistic effect, with lower minimal inhibitory concentrations compared to when the essential oils were used separately. Among a wide variety of essential oils tested by Hyun et al. (2015), thyme, oregano, and lemongrass essential oils showed strong activities in vapor phase against E. coli O157:H7, Salmonella Typhimurium, L. monocytogenes, S. aureus, and B. cereus, using a disc volatilization method. Lemongrass essential oil was effective in maintaining a reduced count of spoilage and pathogenic bacteria in cabbage stored at modified atmosphere with 100% CO2; SEM images of Candida albicans revealed shrunk and deformed surfaces in cells exposed to lemongrass essential oil in vapor phase, contrary to smooth surfaces in cells that did not undergo treatment. Although when the fungus was treated with direct application of lemongrass essential oil at its minimal inhibitory concentration the surface was also damaged, the harm was more pronounced in vapor phase. In general, lemongrass vapor was effective in a much lower concentration compared to the essential oil used directly, probably due to its better capacity to penetrate the exopolymeric matrix of Candida biofilms. While searching through the information available in the literature, it does not go unnoticed the number of studies regarding essential oil antimicrobial effectivity in vapor phase that have been performed using food products. An example is the application of garlic extract in zein film for the inhibition of P. expansum in bread (Heras-Mozos et al., 2019). Another example was the elaboration of polyvinyl alcohol microcapsules with oregano essential oil for microbial growth control in iceberg lettuce; a polypropylene-coated paper sachet was used to hold the spray-dried microcapsules, and these reduced the bacteria, mold, and yeast count in iceberg lettuce stored at 20°C and 85% RH (Chang et al., 2017). As has been widely observed throughout effectiveness studies in vapor phase, microorganism survival and growth differ depending on whether inoculation is made in growing media or foods. In the aforementioned study by Ji et al. (2019), P. corylophilum growth was easier in beef jerky than in agar DG18 due to beef’s lower carbohydrate content, which does not favor fungal growth. Lorenzo-Leal et al. (2019) found that allspice essential oil applied in vapor phase was more effective against L. monocytogenes and S. Typhimurium inoculated in alfalfa seeds, than in culture media. Phillips et al. (2012) also observed that microbial growth varied between culture medium and food; in culture medium, mycelial growth inhibition was observed, and spore germination was decreased for Penicillium chrysogenum, A. niger, and Alternaria alternata, using Citri-V™® (50:50 mix of orange:bergamot essential oils) in vapor phase. However, when Citri-V™® antifungal effectiveness against A. alternata was tested in tomatoes, there was no mold inhibition. In milling grain, on the other hand, the essential oils combination decreased A. niger and P. chrysogenum conidiophore growth (Phillips et al., 2012).

17.4 Mechanisms of Action The preceding reports indicate that some spice extracts (essential oils) have a broad spectrum of biological effects, whereas other extracts may be specific toward certain groups of microorganisms, such as Gram-positive or Gram-negative bacteria, or only bacteria, not yeasts or molds. Some essential oils, plant extracts, and oleoresins influence certain biochemical and/or metabolic functions, such as respiration or production of toxins or acids, indicating that the active components in various oils and oleoresins may have different specificity with regard to target sites, on or in microbial cells. The possible modes of action of phenolic compounds have been reported in different reviews; Swamy, Akhtar, and Sinniah (2016) explained the proposed mechanisms of action of essential oils against

Naturally Occurring Compounds – Plant Sources

545

bacterial and fungal pathogens, while Basak and Guha (2018) and Grande-Tovar et al. (2018) focused on the effects against fungi involved in fruit decay. Friedman (2017) and Marinelli, Di Stefano, and Cacciatore (2018) report findings related to the mode of action of cinnamaldehyde and carvacrol on bacteria respectively. A good summary of the experimental approaches used to identify target sites and modes of action of essential oils can be found in Hyldgaard et al. (2012). Table 17.2 gives an overview of the experimental approaches more recently taken in elucidating the mode of action of different plant antimicrobial components. The effect of phenolic compounds on microbial growth and toxin production could be the result of the ability of phenolic compounds to alter microbial cell permeability, permitting the loss of macromolecules from the interior. They could also interact with membrane proteins causing a disruption to the membrane structure and thus the functionality of embedded proteins (Reyes-Jurado et al., 2019b). Juven et al. (1994) found that increasing thyme essential oil, thymol, or carvacrol concentration was not reflected in a direct relationship with antimicrobial effects; however, they reported that after exceeding a certain (critical) concentration, a rapid and drastic reduction in viable cells of Salmonella Typhimurium was observed. Phenolic compounds could sensitize cellular membranes and when sites were saturated, a serious damage and a rapid collapse of cytoplasmatic membrane integrity could be presented, with the consequent loss of cytoplasmatic constituents. Ruiz-Barba et al. (1990), using scanning electron microscopy, observed that cells without treatment were smooth compared with those treated with phenols for 24 h, which appeared rugged and with irregular surfaces. Phenolic compounds could also denaturalize enzymes responsible for spore germination or interfere with amino acids necessary in germination processes (Nychas, 1995). So far, the mechanism of action proposed for the different plant antimicrobials tested have to do with one or several of the following aspects: induction of apoptosis, loss of membrane permeability, changes in DNA content, genetic expression, cell components, morphology and metabolism, and interaction with DNA. Ren et al. (2019) studied the effect of pterostilbene, a plant polyphenol, derived from Xinjiang wine grape on Staphylococcus aureus and Escherichia coli. The authors concluded that the antibacterial effect is explained by induction of bacterial cell wall and cell membrane damage and an interruption of cellular processes. These were evidenced by an increase detection of reactive oxygen species (ROS), a factor associated with apoptosis and an indicator of oxidative stress. These in turn explained the membrane depolarization, measured by an increase in membrane potential, and damage, as revealed by transmission electron microscopy (TEM) photographs in which cell lysis and leakage of intracellular substances could be observed. DNA content was also measured and a significant decrease in treated bacteria was observed. Leakage of intracellular nucleic acids and proteins revealed by significant changes in optical density values measured at 260 and 280 nm showed that pterostilbene increased cell membrane permeability with dose–effect relationships. Evident morphology changes were observed by scanning electron microscopy (SEM), in contrast to untreated cells presenting smooth and regular structures, S. aureus treated cells were described as irregular, shriveled, and adhered to each other while E. coli treated cells were found incomplete and with uneven shapes. Analysis on the expression of the genes TocL, DinF, Pal, MtgA, and NagA by qRT-PCR revealed changes at the molecular level. DinF was found to be significantly up-regulated which was explained as associated with oxidative stress. Pal, MtgA, and Nag were found to be down-regulated, and this was related to an inhibition of cell wall synthesis processes. TolC, the gene coding for tolerant colicin protein, an important component of the efflux pump of Gramnegative bacteria, was found to be up-regulated only in E. coli. Similar mechanisms involving the loss of membrane permeability, induction of apoptosis, and changes in morphology have been reported for different plant components against a variety of microorganisms (Table 17.2). The effects of clove oil on Listeria monocytogenes were studied by Cui et al. (2018); TEM analysis revealed a significantly deformed cell membrane and an incomplete and destroyed phospholipid bilayer. Evidence of damage to membrane integrity was found as a decrease in intracellular protein and ATP content as well as the detection of nucleic acid outside the cells. A decrease in DNA cell content was also found; this, and the decrease in cellular protein content, are explained by the authors both in terms of damage to membrane integrity as well as suppression of nucleic acid synthesis with a resultant negative effect on gene expression and protein synthesis. After treatment with clove oil, the activity of three types of ATPase (Na+K+-ATP, Ca2+-ATP, and Mg2+-ATP), as well as of the enzymes β-galactosidase and

Loss of membrane permeability

Induction of apoptosis

Mode of Action

Cell autolysis by measurement in absorbance changes at 600nm Reactive oxygen species (ROS) detection DNA fragmentation analysis by agarose gel electrophoresis Changes in optical density (OD) at 260 nm indicating leakage of 260 nm absorbing compounds such as nucleic acids

Experiment

Allium sativum essential oil

Taxus cuspidata stem essential oil

Eleutherococcus senticosus seed essential oil

Acantholippia seriphioides, Aloysia polystachya, Buddleja globose, Lippia turbinate, Minthostachys mollis, Schinus mole, and Solidago chilensis essential oils Helichrysum italicum oil Salvia sclarea essential oil Enteromorpha linza L. essential oil Chlorogenic acid Taxus cuspidata leaf essential oil

Zataria multiflora Boiss. essential oil Carvacrol

Pink pepper essential oil Anthocyanins

Bacillus subtilis

Phytic acid (2%), Syzygium cumini methanolic seed extract (2%), and sodium chloride (0.5%) Pterostilbene

E. coli and S. aureus E. coli and S. aureus E. coli ATCC 43890 S. aureus B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli O157:H7 ATCC 43889 B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli ATCC 43889

Gram-positive bacteria L. monocytogenes, S. aureus, S. Enteritidis, and V. parahaemolyticus Lactobacillus curvatus O157 and non-O157 Shiga-toxin producing E. coli Paenibacillus larvae

S. aureus and E. coli

S. aureus and Escherichia coli

Pterostilbene

Targeted Microorganisms Staphylococcus aureus

Inula helenium L. root essential oil

Agent

(Continued)

Sharma et al. (2013)

Bajpai et al. (2013b)

Bajpai et al. (2013a)

Cui et al. (2015a) Cui et al. (2015c) Patra et al. (2015) Li et al. (2014) Bajpai et al. (2014)

Pellegrini et al. (2017)

Ziaee et al. (2018) Stratakos et al. (2018)

Dannenberg et al. (2019) Sun et al. (2018)

Ren et al. (2019)

Yadav et al. (2018)

Stojanović-Radić et al. (2012) Ren et al. (2019)

References

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2

546 Antimicrobials in Food

Mode of Action

References

Li et al. (2015) Xueuan et al. (2018) Cui et al. (2018) Patra et al. (2015) Bajpai et al. (2014)

E. coli ATCC 43890 B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli O157:H7 ATCC 43889 Vibrio cholerae

Allium sativum extracts Carvacrol

Methyl gallate isolated from Acacia farnesiana

Eleutherococcus senticosus seed essential oil

Sánchez et al. (2013) (Continued)

Bajpai et al. (2013a)

Chen et al. (2018a) Stratakos et al. (2018)

Xiang et al. (2018) Sun et al. (2018)

S. aureus L. monocytogenes, S. aureus, S. Enteritidis, and V. parahaemolyticus Erwinia carotovora O157 and non-O157 Shiga-toxin producing E. coli Ralstonia solanacearum Botrytis cinerea L. monocytogenes

Artemisia argyi essential oil Anthocyanins

Inula helenium L. root essential oil S. aureus Alternatia alternata Botrytis cinerea Botrytis cinerea S. aureus and E. coli

Álvarez-Ordoñez et al. (2013) Stojanović-Radić et al. (2012) Guinoiseau et al. (2010) Xu et al. (2018) Xueuan et al. (2018) Shao et al. (2013a,b) Ren et al. (2019)

Inula graveolens and Santolina corsica essential oils Cinnamaldehyde Mint oil Tea tree oil Pterostilbene

Targeted Microorganisms Brachyspira hyodysenteriae, S. enterica, and E. coli S. aureus ATCC 6538

Agent BIOLL+®, a commercial citrus fruit extract

Macleaya cordata R. leaf essential oil Mint oil Measurement of intracellular protein Clove oil content Leakage of phosphate, potassium, Enteromorpha linza L. essential oil and other ions Taxus cuspidata leaf essential oil

Protein leakage by measuring changes in OD at 280 nm or at OD 595 after staining with Coomassie Brilliant Blue

Experiment

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 547

Mode of Action

Measuring dye intake into permeable cells

Measuring leakage of reducing sugars Changes in electrical conductivity

Experiment

Macleaya cordata R. leaf essential oil Enteromorpha linza L. essential oil Cinnamomum cassia bark essential oil Methyl gallate isolated from Acacia farnesiana α-terpineol and terpene-4-alcohol α-terpineol and terpene-4-ol Cinnamaldehyde Tea tree oi, terpinen-4-ol and 1,8-cineole Tea tree oil Zataria multiflora Boiss. essential oil Phytic acid (2%), Syzygium cumini methanolic seed extract (2%), and sodium chloride (0.5%) Pterostilbene Artemisia argyi essential oil

Pink pepper essential oil Allium sativum extracts Carvacrol

Macleaya cordata R. leaf essential oil

Inula helenium L. root essential oil

Cinnamomum verum bark essential oil Cistus ladanifer L. essential oil BIOLL+®, a commercial citrus fruit extract

Allium sativum essential oil

Taxus cuspidata stem essential oil

Agent

References

Ren et al. (2019) Xiang et al. (2018)

S. aureus and E. coli S. aureus

(Continued)

Li et al. (2015) Patra et al. (2015) Huang et al. (2014) Sánchez et al. (2013) Kong et al. (2019) An et al. (2019) Xu et al. (2018) Yu et al. (2015) Shao et al. (2013a,b) Ziaee et al. (2018) Yadav et al. (2018)

Dannenberg et al. (2019) Chen et al. (2018a) Stratakos et al. (2018)

Bouhdid et al. (2010) Upadhyay et al. (2018) Álvarez-Ordoñez et al. (2013) Stojanović-Radić et al. (2012) Li et al. (2015)

Sharma et al. (2013)

Bajpai et al. (2013b)

Gram-positive bacteria Erwinia carotovora O157 and non-O157 Shiga-toxin producing E. coli Ralstonia solanacearum E. coli ATCC 43890 S. aureus and E. coli Vibrio cholerae Aspergillus ochraceus Aspergillus niger Alternaria alternata Botrytis cinerea Botrytis cinerea Lactobacillus curvatus Bacillus subtilis

Ralstonia solanacearum

B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli ATCC 43889 S. aureus and P. aeruginosa Aspergillus flavus Brachyspira hyodysenteriae, S. enterica, and E. coli S. aureus ATCC 6538

Targeted Microorganisms

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

548 Antimicrobials in Food

Mode of Action

Intra-extracellular ATP

Measuring alkaline phosphatase (AKP), superoxide dismutase (SOD), and/or adenosine triphosphatase (ATPase) levels

Experiment

Methyl gallate isolated from Acacia farnesiana Taxus cuspidata stem essential oil

Helichrysum italicum oil Salvia sclarea essential oil Chlorogenic acid Taxus cuspidata leaf essential oil

Clove oil Tea tree oil Carvacrol

Cinnamomum verum bark essential oil ( E)-2-Hexenal Cinnamaldehyde Tea tree oi, terpinen-4-ol and 1,8-cineole Pterostilbene Artemisia argyi essential oil Chlorogenic acid Anthocyanins

Acantholippia seriphioides, Aloysia polystachya, Buddleja globose, Lippia turbinate, Minthostachys mollis, Schinus mole, and Solidago chilensis essential oils BIOLL+®, a commercial citrus fruit extract

Agent

L. monocytogenes Botrytis cinerea O157 and non-O157 Shiga-toxin producing E. coli E. coli and S. aureus E. coli and S. aureus S. aureus B. cereus ATCC 13061 and E. coli ATCC 43889 Vibrio cholerae B. cereus ATCC 13061 and E. coli ATCC 43889

(Continued)

Sánchez et al. (2013) Bajpai et al. (2013b)

Cui et al. (2015a) Cui et al. (2015c, 2016) Li et al. (2014) Bajpai et al. (2014)

Cui et al. (2018) Shao et al. (2013a,b) Stratakos et al. (2018)

Álvarez-Ordoñez et al. (2013) Bouhdid et al. (2010) Ma et al. (2019) Xu et al. (2018) Yu et al. (2015) Ren et al. (2019) Xiang et al. (2018) Li et al. (2014) Sun et al. (2018)

Brachyspira hyodysenteriae, S. enterica, and E. coli S. aureus and P. aeruginosa Aspergillus flavus Alternaria alternata Botrytis cinerea S. aureus and E. coli S. aureus S. aureus L. monocytogenes, S. aureus, S. Enteritidis, and V. parahaemolyticus

References Pellegrini et al. (2017)

Paenibacillus larvae

Targeted Microorganisms

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 549

Cytoplasmic pH Fatty acid composition

Experiment

Measurement of 4´, 6-diamidino-2phenylindole (DAPI) fluorescence. DAPI can penetrate bacterial cells and bind to DNA and RNA Random amplification of polymorphic bacterial DNA and resolution by gel electrophoresis Interaction with DNA Ultraviolet absorption spectroscopy Changes in morphology Scanning electron microscopy (SEM) and/or transmission electron microscopy (TEM) analysis

Changes in DNA content

Mode of Action

Targeted Microorganisms

Piper peltatum and Piper marginatum extracts

Artemisia argyi essential oil Phytic acid (2%), Syzygium cumini methanolic seed extract (2%), and sodium chloride (0.5%) Zataria multiflora Boiss. essential oil Anthocyanins

Lactobacillus curvatus L. monocytogenes, S. aureus, S. Enteritidis, and V. parahaemolyticus Alicyclobacillus acidoterrestris

L. monocytogenes E. coli and S. aureus E. coli, S. Typhimurium, and V. parahaemolyticus S. aureus Bacillus subtilis

pBR322 DNA S. aureus and E. coli

Eugenol Pterostilbene

Clove oil Helichrysum italicum oil Cinnamon oil nanoemulsions

S. aureus ATCC 29213 and E. coli OH17. H7 ATCC 25922

B. cereus ATCC 13061 and E. coli ATCC 43889 Vibrio cholerae Shewanella putrefaciens Alternaria alternata Aspergillus flavus L. monocytogenes

Ultrasonicated spinach leaf extracts

Methyl gallate isolated from Acacia farnesiana Cinnamon oil Cinnamaldehyde Cistus ladanifer L. essential oil Clove oil

Allium sativum essential oil

Agent

De Pascoli et al. (2018) (Continued)

Ziaee et al. (2018) Sun et al. (2018)

Xiang et al. (2018) Yadav et al. (2018)

Cui et al. (2018) Cui et al. (2015a) Chuesiang et al. (2019)

Cui et al. (2018) Ren et al. (2019)

Altemimi et al. (2017)

Arnon-Rips et al. (2019) Lyu et al. (2017) Xu et al. (2018) Upadhyay et al. (2018) Cui et al. (2018)

Sharma et al. (2013)

References

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

550 Antimicrobials in Food

Mode of Action

Experiment

Cold-pressed Valencia orange oil Inula graveolens and Santolina corsica essential oils Cinnamomum verum bark essential oil α-terpineol and terpene-4-alcohol ( E )-2-Hexenal α-terpineol and terpene-4-ol Cinnamaldehyde Mint oil Thymus vulgaris essential oil and thymol

Inula helenium L. root essential oil

Allium sativum essential oil

Taxus cuspidata stem essential oil

Eleutherococcus senticosus seed essential oil

Benzyl isothiocyanate Salvia sclarea essential oil Macleaya cordata R. leaf essential oil Enteromorpha linza L. essential oil Chlorogenic acid Cinnamomum cassia bark essential oil Taxus cuspidata leaf essential oil

Allium sativum extracts Ultrasonicated spinach leaf extracts

Agent

Methicillin-resistant S. aureus S. aureus S. aureus and P. aeruginosa Aspergillus ochraceus Aspergillus flavus Aspergillus niger Alternaria alternata Botrytis cinerea Alternatia alternata

Erwinia carotovora S. aureus ATCC 29213 and E. coli OH17. H7 ATCC 25922 E. coli E. coli and S. aureus Ralstonia solanacearum E. coli ATCC 43890 S. aureus S. aureus and E. coli B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli O157:H7 ATCC 43889 B. cereus ATCC 13061 and E. coli ATCC 43889 B. cereus ATCC 13061 and E. coli ATCC 43889 S. aureus ATCC 6538

Targeted Microorganisms

(Continued)

Stojanović-Radić et al. (2012) Muthaiyan et al. (2012) Guinoiseau et al. (2010) Bouhdid et al. (2010) Kong et al. (2019) Ma et al. (2019) An et al. (2019) Xu et al. (2018) Xueuan et al. (2018) Perina et al. (2015)

Sharma et al. (2013)

Bajpai et al. (2013b)

Bajpai et al. (2013a)

Clemente et al. (2017) Cui et al. (2015b) Li and Yu (2015) Patra et al. (2015) Li et al. (2014) Huang et al. (2014) Bajpai et al. (2014)

Chen et al. (2018a) Altemimi et al. (2017)

References

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 551

Changes in cell components

Several

Metabolic changes

Electron paramagnetic resonance spectroscopy Changes in membrane protein profile by microfluid chip technology

FT-IR spectroscope

Specific enzyme inhibition Tricarboxylic acid (TCA) pathway Raman spectroscopy

qRT-pCR, whole-genome DNA microarray approach, single-gene expression assessment

Experiment

Differential genetic expression

Mode of Action

Cinnamon bark and clove oil

Oregano essential oil

P. aeruginosa ATCC 27853 and P. aeruginosa ATCC 27857

Brachyspira hyodysenteriae, S. enterica, and E. coli L. monocytogenes ATCC 7644

Felso et al. (2013)

Álvarez-Ordoñez et al. (2013) Serio et al. (2010)

Kong et al. (2019) An et al. (2019) Ma et al. (2019) Cui et al. (2018) Clemente et al. (2017) Clemente et al. (2016)

Sánchez et al. (2013) Herman et al. (2013) Muthaiyan et al. (2012) Azizkhani et al. (2012)

Vibrio cholerae S. aureus ATCC 29213 Methicillin-resistant S. aureus S. aureus ATCC 29213 Aspergillus ochraceus Aspergillus niger Aspergillus flavus L. monocytogenes E. coli Aspergillus ochraceus

Siroli et al. (2018)

E. coli

References Yu et al. (2015) Shao et al. (2013) Tyagi and Malik (2010) Ren et al. (2019)

Thyme essential oil, carvacrol, 2-(E)-hexenal and citral Methyl gallate isolated from Acacia farnesiana Cinnamon and lavender oils Cold-pressed Valencia orange oil Zataria multiflora Boiss, Rosmarinus officinalis L., and Mentha longifolia L. essential oils α-terpineol and terpene-4-alcohol α-terpineol and terpene-4-ol ( E )-2-Hexenal Clove oil Benzyl isothiocyanate Antimicrobial packaging containing benzyl Isothiocyanate BIOLL+®, a commercial citrus fruit extract

Targeted Microorganisms Botrytis cinerea Botrytis cinerea Candida albicans S. aureus and E. coli

Tea tree oil, terpinen-4-ol, and 1,8-cineole Tea tree oil Lemon grass essential oil Pterostilbene

Agent

Experimental Approaches Recently Made in Elucidating the Mode of Action of Different Plant Antimicrobial Components and the Targeted Microorganisms

TABLE 17.2 (CONTINUED)

552 Antimicrobials in Food

Naturally Occurring Compounds – Plant Sources

553

alkaline phosphatase was significantly reduced. The study (Cui et al., 2018) also reported that clove oil could inhibit the physiological activity of L. monocytogenes by affecting metabolites and key regulatory enzymes of the tricarboxylic acid (TCA) pathway; the activity of the enzymes isocitrate dehydrogenase and α-ketoglutarate dehydrogenase decreased while the activity of citrate synthase increased or remained unchanged in treated cells. Intermediate compounds of the TCA cycle (citric, succinic, and L-malic acids) were significantly reduced after treatment; finally, eugenol, a major component of clove oil, was found to change the structure of DNA through formation of a eugenol-DNA chimera. An et al. (2019) studied the effect that tea tree oil and its components have on Aspergillus niger. An increase in relative electric conductivity, indicative of membrane permeability destruction, was reported as follows, from most to least damaging: α-terpineol > terpene-4-ol > tea tree oil > 3-carene. Amongst the morphological changes in external and internal structures described are a badly twisted, broken, wrinkling, and coarse mycelium with a flat-trip shape and absence of cytoplasmic content when cells were treated with α-terpineol; treatment with α-terpineol, terpene-4-ol, and tea tree oil clearly damaged the cell walls and structural clumps gathered from organelles, and abnormal vacuolations of varying size were observed. The damage to membranes was not restricted to the cytoplasmic membrane but included organelle membranes as well; metabolic changes reflected in a significant alteration of four metabolic pathways were also revealed. Changes in the histidine degradation pathway resulted in the formation of lower amounts of 10-formyltetrahydrofolate and L-glutamate, a detrimental consequence to the metabolism of A. niger. Levels of L-aspartate and oxaloacetate from the asparagine degradation pathway and 2-keto-3deoxy-6-phosphogluconate, glyceraldehyde-3-phosphate, and pyruvate from the 2-keto-3deoxy6-phosphogluconate cleavage pathway were also lowered; these led to lower availability of oxaloacetate and pyruvate to enter the TCA cycle and lower amounts of glyceraldehyde-3-phosphate entering the glycolytic pathway and participating in energy metabolism. All of these had an effect on compromising fungal life processes (An et al., 2019). Raman micro-spectroscopy is a spectroscopic technique that allows the researcher to obtain information on the chemical nature of the sample studied while information on the molecular composition, orientation, symmetry, and structure can be acquired (Clemente, Aznar and Nerín, 2016). Clemente, Aznar, and Nerín (2016) and Clemente et al. (2017) used this approach to investigate the effect of antimicrobial packaging containing benzyl isothiocyanate, an active agent found in plants of the mustard family (Brassicaceae), against Aspergillus ochraceus and E. coli, respectively. For A. ochraceus, it was concluded that the isothiocyanate group of benzyl isothiocyanate reacts with saccharides, amino acids, proteins, and lipids; possibly by the action of the highly electrophilic central carbon atom which can react readily with oxygen-, sulfur-, or nitrogen-centered nucleophiles, as reported by others (Zhang and Talalay, 1994). This in turn has impacts on cellular functions such as respiration, metabolism, and cell cycle; the effect was observed independently of the type of contact, liquid or vapor, although when spores were treated in vapor phase, some of them presented a reduced effect. In the case of E. coli, despite no cellular damage being observed, benzyl isothiocyanate accumulated in the cells and caused alterations to the previously mentioned essential cell components, saccharides, amino acids, proteins, and lipids (Clemente et al., 2016; Clemente et al., 2017). The major antimicrobial constituent of garlic and onion is allicin, along with several other sulfurcontaining compounds. Cavallito and Bailey (1944) were the first to isolate the major antimicrobial component from garlic bulbs by steam distillation of ethanol extracts. It was noted that allicin was extremely pungent and characterized the principal odor and taste of garlic and onion. The high antimicrobial activities of garlic and onion extracts are probably due to a high content of allicin and other sulfides in the essential oils of garlic and onion. Wills (1956) observed that the inhibition of sulfhydryl enzymes was associated with the presence of the –SO–S– grouping, not the –SO–, –S–S–, or –S– groups. Barone and Tansey (1977) reported that allicin disrupted microbial cell metabolism primarily by inactivation of –SH proteins by the oxidation of thiols to disulfides; competitive inhibition of the activity of sulfhydryl components, such as cysteine and glutathione, occurred by binding with them and/or by noncompetitive inhibition of enzyme functions by oxidation of the binding to –SH groups at allosteric sites. Chen et al. (2018a) reported that garlic extracts could alter cell membrane permeability and destroy the structural integrity of cell membranes of Erwinia carotovora, which they found by observing changes in membrane conductivity and leakage of proteins, respectively. SEM images showed different degrees of cell

554

Antimicrobials in Food

rupture and adherence in a time- and concentration-dependent manner; the authors concluded that the mechanism by which garlic extracts affected the microorganisms was by disruption of the cytoplasmic membrane. Siroli et al. (2018) studied the effect that different essential oils and their components had on E. coli K12 cells. Cells were exposed to sub-lethal concentrations of carvacrol, 2-(E)-hexenal, citral, and thyme essential oil for 1 hour and the composition of fatty acids and transcriptional response were analyzed; for all treatments, an increase in unsaturated and cyclic fatty acids was observed, where the greatest increase in unsaturated fatty acids was observed when cells were exposed to carvacrol. Exposure to citral, 2-(E)-hexenal, and thyme essential oil lead to an increase in mean chain length while citral treatment also caused an increase in branched fatty acids; authors reported that the increase in unsaturated fatty acids is related to bacterial response to different stresses – temperature, oxidative, and ethanol stresses or high pressures – and that the increase in chain length and cyclic fatty acids is then a response to counteract the fluidizing effect of unsaturated fatty acids. These findings were in agreement with what was observed in the transcriptomic analyses where the up-regulation of genes involved in fatty acid biosynthesis was evident. Despite clear differences between the numbers of genes being significantly upor down-regulated with each treatment, the effects observed were similar for all antimicrobials tested; overall results show that most of the affected genes are those involved in energy metabolism, protection against oxidative stress, purine/pyrimidine metabolism, fatty acid and phospholipid metabolism, and protein synthesis (Siroli et al., 2018).

17.5 Factors Affecting Activity 17.5.1 Plant Source Variation Several factors have a significant role in the composition/content of naturally occurring antimicrobial compounds: botanical source (species, variety, and genotype; part(s) of the plant, geographic origin, sampling time, and season and environmental conditions), growth and harvest conditions (wild or cultivated, soil composition and texture, good agricultural practices, site of collection, climate, time of harvest, stage of growth, storage conditions, and pre- and post-harvest phytosanitary treatments), steps of preparation, and extraction methods (Mukherjee et al., 2017). Essential oils are concentrated hydrophobic liquids containing more than 200 compounds; the volatile fraction (90–95 % w/w) contains monoterpene and sesquiterpene hydrocarbons and their oxygenated derivatives along with aliphatic aldehydes, alcohols, and esters, while the nonvolatile residue contains hydrocarbons, fatty acids, sterols, carotenoids, and flavonoids (Thakur et al., 2017). Their antimicrobial effect varies greatly due to differences in their chemical properties (Burt 2004). In a recent study, Semeniuc et al. (2018) compared and classified parsley, lovage, basil, and thyme essential oils based on their chemical composition, total phenolic content (TPC), and antibacterial activities against Salmonella Enteritidis and Listeria monocytogenes. Although all tested essential oils showed inhibitory and bactericidal effect on both bacteria, thyme essential oil showed the highest level of TPC and the stronger antibacterial activity; parsley essential oil was characterized by a low TPC and weak antibacterial activity against S. Enteritidis; lovage essential oil by a low TPC and moderate antibacterial activity; and basil essential oil by a low TPC and weak bacterial activity against L. monocytogenes. Moreover, plant bioactive compounds are secondary metabolites, which protect plants from predators and pathogens on the basis of their toxicity and repellency to herbivores and microbes, increasing plant’s ability to survive and overcome local challenges. These metabolites are often produced after growth, and their compositions vary due to genetic and environmental factors that influence genetic expression (Azmir et al., 2013; Thakur et al., 2017); some of them are preformed and some of them are induced by infection and adverse environmental factors (Taiz and Seiger 2002). As reviewed by Talhaoui et al. (2015), abiotic and biotic factors qualitatively and quantitatively affected the phenolic compounds composition in olive leaves. Oleuropein, the main phenolic compound of olive leaves, increased in response to water or cold stress while salinity stress associated with high sunlight enhanced the biosynthesis of other phenolic compounds, particularly flavonoids. Tyrosol, catechin, and

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oleuropein increased in leaves and other parts of the olive tree after fertilization with urea, nitrogen, copper, manganese, and zinc; changes in levels of phenolic compounds had been found to be involved in defensive reactions of olive leaves against fungi and bacteria (Talhaoui et al., 2015). Torres et al. (2018) demonstrated that mycorrhizal symbiosis induced the accumulation of flavonols and anthocyanins in the leaves of Tempranillo grapevines under stressful temperature conditions, although clonal diversity affected the secondary metabolism. The effect of the fertilizer regime (balanced or over-supply of macronutrients), locations (different soil type), cultivar, polyphenol content, and profile of polyphenols and their interactions in globe artichoke were examined by Lombardo et al. (2015); both the cultivar and the soil type modulate the plant’s response to fertilizer while the balanced fertilizer regime markedly favored the accumulation of polyphenols. The age of American cranberry plants significantly influenced secondary metabolites synthesis (total soluble phenolic compounds, flavonoids, catechins, procyanidins, and anthocyanins) (Berezina et al. 2017); catechins and flavonoids had the same accumulation dynamics, gradually decreasing their content as plants reached reproductive age. Phenolic compounds’ content in the berries was lower than in the leaves, the only exception being anthocyanins. This natural and induced variability in chemical composition and content of bioactive compounds dramatically influences their antimicrobial properties; consequently, all botanicals and their derived products and extracts should be properly characterized and standardized through analytical techniques, such as GC, HPLC–NMR, HPLC–MS, GC–MS, and CE–MS.

17.5.2 Extraction Methods Bioactive components, including essential oils, proteins, terpenoids, flavonoids, and phenols, among others, can be obtained from one or more plant parts: leaves, flowers, peels, barks, seeds, roots, fruits, rhizomes, and gums and oleoresin exudations. The composition of the essential oils from different parts of the plant may vary widely (Burt 2004). Vegetable matrices are very complex, and the type of structure in which the bioactive is located depends on the plant type and is plant family-specific. For instance, essential oils may occur in specialized secretary structures, modified parenchyma cells, resin canals, oil tubes, or gum canals (Dalla Nora and Borges, 2017). Thus, many techniques are usually performed before or during the extraction to allow a better recovery or quality of the bioactive compounds (Gallego et al., 2019). A variety of methods are commercially used or have been proposed for the extraction and isolation of plant natural compounds; extensive studies on the subject have been compiled in many reviews (Azmir et al., 2013; Thakur et al., 2017; Gallego et al., 2019; Zhang et al., 2018b; Selvamuthukumaran and Shi, 2017). Extraction is the first step to separate the desired natural compounds from the plant matrix and can be performed by using conventional and non-conventional methods. Classical extraction techniques (solvent extraction, maceration, and hydro-distillation) are the most widely utilized methods on a commercial basis and are mainly based on the extracting power of different solvents and the application of heat and/or mixing. Major disadvantages of these methods are long extraction times, requirement of large volumes of solvents, low extraction selectivity, and damage to thermo-labile compounds. Techniques that involve high temperatures may cause the loss of volatile compounds due to long extraction times, oxidation and degradation of unsaturated or esterified compounds by thermal or hydrolytic reactions, and yield losses. To overcome these limitations, new techniques that require lower organic solvent amounts and energy consumption and shorter extraction times, and exhibit higher selectivity have been developed; some of these non-conventional techniques are super-critical fluid extraction, pressurized liquid extraction, microwave-assisted extraction, ultrasound-assisted extraction, enzyme-assisted extraction, and pulsed electric field-assisted extraction; these techniques may be scalable and can be coupled to other extraction processes (Gallego et al., 2019). Many of these advanced extraction techniques comply with green chemistry principles (Chemat et al., 2012). Extracts are complex and usually contain a variety of natural components, and further separation could be required. Separation of extract components can be performed by methods based on the difference in molecular size (membrane filtration, gel filtration chromatography) or on the ionic strength (ion-exchange chromatography) as well as using other modern separation techniques such as molecular

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distillation, supercritical fluid chromatography, preparative gas chromatography, molecular imprinted technology, and multi-dimensional chromatographic separation (Zhang et al., 2018b). Extraction techniques may affect the physicochemical properties of essential oils and alter the natural proportion of the chemical constituents as in their original state. Process parameters, such as extraction solvent and temperature, can dramatically change the composition of the extracts. The extraction conditions may be modulated to change the selectivity according to the target compound. In addition, the coupling of processes has been employed to make the extraction more efficient (Gallego et al., 2019). Thus, there is a wide array of possibilities going from the extraction method or the combination of extraction methods to the selection of process variables to obtain selected bioactive compounds. Accordingly, the chemical composition of extracts can be affected in a different way and so their antimicrobial activities (Azis et al., 2018; Elyemni et al., 2019; Tao et al., 2014).

17.5.3 Interaction with Food Matrix Most of natural antimicrobials demonstrated excellent activities against microorganisms in model media and beverages but failed to be as effective when used in complex foods. The antimicrobial may require up to 1000 times the in vitro dose (nutrient broth, planktonic media) to be equally effective in vivo situations (complex foods), overcoming the low thresholds of these compounds. The antimicrobial efficacy was found to be a function of food ingredients and structure; active compounds tend to interact with food components such as proteins, fats, sugars, and salts, and/or be adsorbed onto the multiple interfaces of foods. Hence, only part of the total compound amount is available to perform the antimicrobial action. High fat concentration was found to have an adverse influence on the antimicrobial action of various essential oils in hotdogs, milk, and other foods (Singh et al., 2003; Cava et al., 2007; Canillac and Mourey, 2004). Regarding the effect of carbohydrates and starch, the reports are somehow contradictory, although the negative impact on antimicrobial effects would also occur at high concentrations (Devlieghere et al., 2004; Ofman et al., 2004; Gutiérrez et al., 2008). Tassou et al. (2000) and Min et al. (2010) reported a negative impact of high protein contents. Weiss et al. (2015) proposed a useful “structural” explanation on why preservatives lose activity in foods; in contrast with planktonic media where structure is homogeneous, foods are in generally highly structured and may simultaneously contain various phases with different physical properties and chemical compositions. Two types of physical interactions between antimicrobials and food ingredients may occur: electrostatic and hydrophobic ones. Antimicrobial compounds carry a certain charge depending on pH, temperature, and ionic strength and can electrostatically interact with proteins, forming soluble or insoluble complexes. Hence, antimicrobials, bound to the proteins, are no longer homogeneously distributed in the food and are not available for acting on the microorganisms. Hydrophobic interactions arise between permanent or induced dipoles and promote the association between polar molecules, that tend to form cluster-like structures, while non-polar molecules tend to form a separate phase where hydrophobic molecules associate with other hydrophobic ones. For example, phenolic compounds associated to proteins and non-polar molecules may partition into lipid phases, hindering a direct interaction with microorganisms and rendering antimicrobials less effective. The interaction of antimicrobials with food will result always in a loss of antimicrobial activity excepting when both the antimicrobials and the microorganisms are located in the same substructures. Hence, considering the difficulty in predicting the interaction between antimicrobials and components in complex foods, each food needs to be individually assessed (Weiss et al., 2015).

17.6 Increasing the Efficacy of Natural Antimicrobials from Plants Besides their high variability in composition and concentration of active substances, low water solubility, high volatility, limited stability against chemical or physical degradation, uncontrolled release, interaction with food matrix/components, and strong organoleptic properties limit the application of essential oils and other bioactive compounds in food systems. Some strategies employed/suggested to partially overcome these deficiencies are the application of essential oils in gaseous (vapor) phase, the use of the

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antimicrobials in combination with other factors (the so-called “hurdle technology”), and the employment of novel delivery systems.

17.6.1 Application of Essential Oils in Gaseous Phase Many bioactive compounds of essential oils are volatile, a property that makes them useful as possible antimicrobials for stored foods (Tzortzakis, 2010). Antimicrobial activity depends on the composition and volatility of the compounds. Due to the hydrophobic nature of essential oils, water is not a good solvent for these substances, and a higher effectivity at relatively lower concentrations than in liquid phase could be expected. Treatments in gaseous phase exhibit also antimicrobial activity over larger areas than those in liquid phase. As the essential oils are not directly added to foods, they may have a relative minor effect on sensory attributes. Various studies about the application of plant volatile compounds have been carried out in recent years. Tyagi and Malik (2011) compared the effectiveness of E. globulus essential oil against 14 food-spoilage bacteria and molds in liquid and vapor phases. The significant high antimicrobial activity detected in the vapor phase was attributed to the higher percentage of monoterpene hydrocarbons in the vapor than in the liquid phase (54.7 vs 44.5 %). Mexican oregano, thyme, and mustard essential oils in vapor phase were tested against various bacterial and fungal species; evaluated microorganisms exhibited different sensitivity to tested oils, the bacterial strains being the most resistant to thyme and oregano essential oils (Reyes-Jurado et al., 2019a). The growth of Botrytis cinerea and Rhizopus stolonifer conidia in artificially inoculated strawberries was significantly reduced by tee tree oil vapor at 0.9 g/L, and treated berries maintained a fresher quality than untreated strawberries during storage (20°C, 3 days); tee tree oil induced fruit defense responses as shown by the induction of H2O2 levels and several defense-related enzymes in the first period of incubation (Shao et al., 2013a,b). Allyl isothiocyanate (AITC, 0.81–1.41 μg/ml, gas phase) was evaluated for preserving mung bean sprouts and fresh-cut iceberg lettuce; although AITC indeed showed microbicidal or microstatic activity against the native flora of both fresh produce, color and respiratory activity were severely affected (Kramer et al., 2018). Exposure of radish sprouts inoculated with Listeria monocytogenes (6.3 log CFU/g) to oregano, thyme, or cinnamon bark essential oils gases (78–625 μL/L; 24 h, 30°C) resulted in a very small decrease in counts; even at the highest concentration, the microorganism was not completely inactivated. This lower antimicrobial activity of essential oils compared to the effects obtained in a laboratory medium was attributed to: (a) a reaction of the antimicrobial compounds of the essential oils with organic matter on the surface of the sprouts, and (b) structural protection by the produce; L. monocytogenes may be embedded in the stomata and crevices of the sprout and/or internalized into the open tissue of the sprout (Lee et al., 2018). Interesting, many studies reported a synergistic effect when two or more essential oils were applied in combination in gaseous phase. The use of different essential oils, which develop synergistic interactions at low concentrations, could minimize their sensory impact. Tunc et al. (2007) studied the effect of three aroma compounds (carvacrol, cinnamaldehyde, and allyl isothiocyanate), ethanol, and SO2, individually and in binary combinations, on the growth of Penicillium notatum, simulating an active packaging and the effect of a modified atmosphere enriched with antimicrobial compounds; tested aroma compounds were more effective than ethanol and SO2; six combinations were synergistic, enabling the reduction of the concentration of each antimicrobial and reducing their organoleptic impact. Combinations of clove and mustard essential oils demonstrated a synergistic interaction in vitro and in vivo (strawberry) to inhibit Botrytis cinerea, a ubiquitous plant pathogen of fruit; moreover, the antimicrobial effect was greater in vivo than in vitro probably due to the low pH of the berries (Aguilar-González et al., 2015). In other study, the antifungal activities of the essential oils of 97 plants against Penicillium corylophilum, a moderately xerophilic mold, were tested in a laboratory medium and on beef jerky, an intermediatemoisture food; the combination of cinnamon bark, citronella, and may chang essential oil vapors had synergistic activity in inhibiting the growth of P. corylophilum on beef jerky (Ji et al., 2019). The antimicrobial activity of volatile plant compounds makes them extremely attractive, in particular because of their low toxicity at low concentrations, renewability, and biodegradability, for application as gaseous treatments to control postharvest diseases in an eco-chemical approach (“biofumigation”).

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Studies about the effect of different plant essential oils on disease development caused by fungi during storage of some commercially important horticultural products (papaya, tomato, kiwifruit, apple, avocado, strawberry, blueberry, and cucumber) were reviewed by Mari et al. (2016). In general, higher concentrations of essential oils provided in vitro a lower survival of the fungus. Besides the structural modification of hyphal morphology by partitioning the lipid layer of the cell membrane and the change in cell membrane permeability, some essential oils would act as a signaling compound to stimulate plant defense mechanisms.

17.6.2 Combination with Other Antimicrobials and/or Preservation Factors The addition of plant bioactive compounds at concentrations needed to control the microorganisms may adversely affect the food flavor. Moreover, their antimicrobial activity is usually moderate when applied as the only preservation factor. The use of combinations of bioactive compounds with other antimicrobials or other preservation factors (“hurdle technology”) as a strategy to overcome these limitations has been a point of intensive research. Overall, multiple disturbances of microbial homeostasis have been used/ suggested in different arrangements: (a) using a plant antimicrobial with other stressors simultaneously to prevent the growth of spoilage and pathogenic microorganisms, and (b) using one or more stressors (one may be a plant antimicrobial) simultaneously or in sequence to inactivate/injure target microorganisms, and then, in sequential mode, a plant antimicrobial compound alone or with more stressors to prevent the survival/proliferation of remaining refractory or sub-lethally damaged cells (Alzamora et al., 2016). Hundreds of papers in this topic have been published in recent years (Burt, 2004; Berdejo et al., 2019; Calo et al., 2015; Coutinho de Oliveira et al., 2015). Although the effect of combinations strongly depends on the type and amount of microorganisms, the doses of the stressors, and the food matrix, some general points can be addressed to facilitate the selection of hurdles or stressors: • Many essential oils showed synergistic antimicrobial activity when applied simultaneously in a combined approach, or when used with traditional synthetic food preservatives, probably because of the effect on different cell targets. • Many combinations of essential oils with non-thermal technologies or mild heat resulted in synergistic lethal effects associated with modifications of microbial cell envelopes, which facilitated the access of the antimicrobial to the target and/or cell membrane disruption and loss of membrane potential. • Synergism or additivity was observed due to the microbicidal effect of the non-thermal or other physical stressors and the subsequent microstatic effect of the natural antimicrobial during storage.

17.6.3 Delivery Systems Over the last years, many studies have been carried out on the development and potential application of delivery systems of active components in the food, pharmaceutical, and cosmetic industries. Results have been compiled in several excellent review articles and book chapters (Fu et al., 2016; Prakash et al., 2018a,b; Weiss et al., 2009; Castro-Rosas et al., 2017; Donsi and Ferrari, 2016). Many potential benefits have been claimed for these systems as compared with the direct addition of antimicrobial compounds: enhanced long-term chemical and physical stability (impaired interaction with food ingredients, lower losses of volatile compounds), improved solubility in aqueous phase and thus better bioavailability of active compounds, decreased impact on food sensory properties, release control, dispersion uniformity, and rapid penetration of bioactive compounds into the target sites (Fu et al., 2016; Pavoni et al., 2019). Overall, these delivery systems are still relatively new technologies and only limited information is available. Most of the studies have been in culture medium models, and there are very few studies in food matrices; they did not consider the interactions between the delivery system and the food matrix, their sensory impact, the technological aspects for the incorporation of antimicrobial structures, and the toxicological aspects of formulated nanostructures (Fu et al., 2016; Brandelli and Taylor, 2015). Other points

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to be addressed are the economics of the nanoscale system, the consideration of food-grade ingredients for their formulation, and the physical stability of the delivery systems throughout the shelf life of the product (Weiss et al., 2009). However, interest in this subject is growing worldwide, and it is expected that within the next few years encapsulated antimicrobials will be available at the commercial level. Delivery systems can be classified as: (a) emulsion-based, (b) nanosized carrier-based, and (c) packaging films or coating-based (Fu et al., 2016).

17.6.3.1 Emulsions Colloidal delivery systems based on microemulsions, nanoemulsions, and Pickering emulsions are increasingly being investigated to encapsulate, protect, and deliver lipophilic bioactive components in the food and pharmaceutical industries (McClements, 2012). Conventional emulsions, thermodynamically unstable, consist of liquid-liquid dispersions of two immiscible compounds in which one is dispersed in the form of small droplets (0.1–20 μm) to another one, and an emulsifier that forms an interfacial layer that improves the stability of the system. Pickering emulsions are conventional emulsions stabilized by solid particles, which gives them greater resistance to destabilization mechanisms, while nanoemulsions are conventional emulsions with mean droplet size ranging from 20 to 200 nm. In contrast, microemulsions are thermodynamically stable, and consist of isotropic liquids formed by mixing oil, water, and surfactant (emulsifier) together; they are characterized by the small size of the disperse phase (d < 100 nm). Due to their exceptional solubility properties, microemulsions are able to deliver both lipophilic and hydrophilic compounds (Pavoni et al., 2019). The small size of the droplets (d < 200 nm) in micro- and nanoemulsions results in a number of potential benefits for certain applications: long-term stability, high optical clarity, and enhanced transport of active molecules through biological membranes. Only a limited number of studies addressed the use of micro- and nanoemulsions in real food systems, i.e. by direct mixing with a liquid food (milk and fruit juices), washing the food surface with the antimicrobial aqueous dispersions (fresh lettuce, spinach leaves, and pears), infusion in porous food matrices (pork sausages and zucchini), and coating with a biopolymeric layer containing the antimicrobial dispersion (apple pieces, broccoli florets, plums, green beans, rucula leaves, and sliced bread) (Donsi and Ferrari, 2016; Fu et al., 2016). Regarding Pickering emulsions, they have been used for the protection and release of various bioactive compounds, including flavonoids and thyme oil (Fu et al., 2016). The antimicrobial efficacy of nanoemulsions is strongly influenced by essential oil components, tested microbial strain, emulsion formulation, and food matrix. Emulsion formulation affects the transport of essential oils to the microbial cell membrane and their interaction with the molecular sites of the membrane. In some cases, encapsulation in nanoemulsions inhibited the antimicrobial activity (Donsi and Ferrari, 2016; Terjung et al., 2012; Nielsen et al., 2017).

17.6.3.2 Nanosized Carriers/Coatings Nanoparticles (particle size less than 1 μm) which could be used as coating material/carrier agents for the plant bioactive compounds include, among others, liposomes, polymer nanoparticles, and solid-liquid particles (Brandelli and Taylor, 2015; Fu et al., 2016; Prakash et al., 2018a,b; Weiss et al., 2009). Liposomes are nano-vesicles with an internal aqueous core rounded by a self-assembled coat of amphiphilic lipids and can be used for the encapsulation and delivery of hydrophilic, hydrophobic, and amphiphilic molecules. Polymer nanoparticles refer to polymer nanospheres and nanocapsules, where bioactive molecules may be adsorbed at the surface or encapsulated within the particle in the case of nanospheres, or confined to an aqueous core and surrounded by the polymeric shell around them in the case of nanocapsules. Solid-liquid particles have a relatively rigid core consisting of hydrophobic lipids that are solid at room temperatures, surrounded by a monolayer of phospholipids. Some examples of nanoparticle delivery systems are the incorporation of essential oils of Artemisia arborescens L. and Zataria multiflora in solid-liquid particles, Oreganum dictamnus L. in phosphatidylcholine liposomes, Atractylodes macrocephala L. in phosphatidylcholine and cholesterol liposomes,

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and lime in nanocapsules (Prakash et al., 2018a,b); in all these cases, enhanced antimicrobial activities compared to the free essential oil form were demonstrated.

17.6.3.3 Packaging Films/Coatings Antimicrobials derived from plants can be added directly to the food or incorporated into active packaging systems. The incorporation of antimicrobial agents in active packaging has been extensively investigated to control their diffusion and maintain them in adequate concentration on food surfaces. An active packaging (AP) system can be classified into two main types: (a) non-migratory AP, which contains antimicrobial agents that show their activity without true migration into foods, and (b) active-releasing packaging, which allows the controlled migration of antimicrobial agents in direct contact with the food surface (non-volatile migration), or the release of volatile antimicrobials which act in the headspace around the food surface and in the food itself after absorption (volatile migration) (Khaneghah et al., 2018a,b). Numerous methods have been investigated to develop antimicrobial packaging systems (Figure 17.2). Among them are: (a) the addition of volatile antimicrobial compounds in a sachet or pad in order to released them within the packaging headspace, (b) the incorporation of antimicrobial compounds (volatile and non-volatile) into the structure of package polymers, (c) the coating or absorption of the antimicrobial compounds onto the packaging, (d) the use of polymers (e.g. chitosan) with antimicrobial properties, and (e) the addition of antimicrobial compounds into edible coatings. The incorporation of antimicrobial agents into synthetic (e.g. petroleum-based polymers) and biobased polymers (e.g. bio-polyethylene, alginate, cellulose, and polylactic acid (PLA) among others) represent good strategies for increasing the shelf life of packed food products (Khaneghah et al., 2018b; Scaffaro et al., 2018b) The antimicrobial efficiency depends on the release rate of the antimicrobial

FIGURE 17.2  Some methods evaluated for the release of antimicrobials compounds in active packaging.

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agents from the packaging. The most critical factors for controlling the release rate and their transfer into the foodstuff are the method of incorporation, the nature of the polymeric matrix, the mechanism of release, food properties, temperature, and environmental relative humidity (Wu et al., 2018). Methods commonly utilized for the incorporation of antimicrobial agents into polymeric materials include thermomechanical processes, such as melt extrusion, or a casting procedure, which involves the dissolution of the polymer and the antimicrobial substance into an appropriate solvent, followed by the evaporation of the solvent. For thermal-sensitive antimicrobials such as essential oils and organic acids, coating has emerged as a useful and alternative incorporation system. Coating consists of the application of at least one layer of fluid containing the antimicrobial substance onto the surface of a polymeric matrix. Another technique utilized is the covalent immobilization to the polymeric material of antimicrobial agents containing reactive functional groups, such as antimicrobial enzymes, peptides, polyamines, organic acids, etc.; some advantages of this method are the reduction of the amount of the antimicrobial added to the polymer and the minimization of antimicrobial activity losses (Khaneghah et al., 2018a,b). There is an increasing tendency of exploring the application of bio-based packaging due to environmental concerns. In this context, polylactic acid-based antimicrobial materials received considerable attention due to the good mechanical properties, renewability, biodegradability, and biocompatibility of PLA (Llorens et al., 2015; Scaffaro et al., 2018a). Sacaffaro et al. (2018b) carried out an extensive review of the incorporation of essential oils and other antimicrobial compounds of natural origin to PLA by different techniques; they highlighted the need to conduct more in vivo studies to further assess the effectiveness of the PLA-based antimicrobial systems. Chitosan, a natural polysaccharide, has also potential applications in active food packaging systems due to its excellent film-forming ability and its intrinsic antimicrobial properties. Chitosan is soluble in diluted aqueous acidic solutions due to protonation of their amino groups; this cationic character confers its antimicrobial properties and its ability to carry and slow-release functional ingredients. Chitosan-based films have been evaluated in the form of pure chitosan films or combined with other biopolymers or synthetic polymers (Wang et al., 2018a). The inclusion of gallic acid into chitosan-based films significantly increased antioxidant activity, antimicrobial activity, and tensile strength of the film, and decreased water vapor and oxygen permeability (Wu et al., 2016a; Xie et al., 2014). Chitosan film enriched with cinnamon oil also showed good antimicrobial activity (Wang et al., 2011). Other tested oils, such as olive oil, rosemary essential oil, and oregano essential oil, also improved the mechanical and barrier properties, and the antioxidant activity of chitosan films (Jian et al., 2012; Pelissari et al., 2009). The addition of natural antimicrobial compounds into polymeric edible coatings produced with polysaccharides, proteins, or lipids obtained from renewable agricultural resources and/or food-processing wastes has also been investigated to control the diffusion of these compounds into different foodstuffs. Edible coatings produced by other biopolymers such as chitosan have also been extensively studied (Aloui and Khwaldia, 2016; Irkin and Esmer, 2015). These coatings consist of thin layers of biopolymers that are applied on food surfaces by different mechanical procedures, such as spraying, brushing, dipping, or by new techniques such as layer-by-layer assembly (Dhall, 2013; Poverenov et al., 2014; BilbaoSainz et al., 2018). Several studies have been reported on the use of biopolymer-based coatings carrying different plant-derived antimicrobial compounds to preserve the quality and increase the safety of many fresh and minimally processed fruit and meat products (Fernández- Pan et al., 2014; Aloui et al., 2014; Aloui and Khwaldia, 2016; Duran et al., 2016; Shakila et al., 2016; Valdés et al., 2017). However, for practical applications some shortcomings have been reported such as the weak adhesion of coating materials to the hydrophilic surface of the food, degradation of the antimicrobial agent, or its rapid desorption through coating materials (Aloui and Khwaldia, 2016). In recent years, with the advances in the field of nanotechnology, the incorporation of nanoemulsions containing antimicrobial compounds into edible coating solutions has been investigated as an effective strategy to improve their distribution and enhance their adhesion to the surface of solid foods (Kim et al., 2014; Salvia-Trujillo et al., 2015; Wu et al., 2016b; Acevedo- Fani et al., 2017). Another innovative and promising approach is the use of antimicrobial multilayer coatings which rely on incorporating antimicrobial compounds into multilayered/nanolaminate systems formed using the electrostatic deposition technique; these multilayer coatings are able to act as

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efficient antimicrobial delivery systems, due to their inner and barrier layers that could control the diffusion rate of natural antimicrobial compounds embedded in the matrix layer (Mantilla et al., 2013; Sipahi et al., 2013; Moreira et al., 2014; Aloui and Khwaldia, 2016).

17.7 Toxicity of Natural Antimicrobials and Its Evaluation and Regulatory Aspects Studies to determine the dose to elicit the antimicrobial effect are necessary for each food. Once the dose is selected, the impact on organoleptic characteristics must be assessed. It is also important to evaluate whether such dose will be toxic or not to the consumers. Variability in the composition of botanical products affects their safety assessment. Plant extracts contain more than one component, from which one or more will be the component(s) of interest (due to their benefit), but others might involve an adverse effect (Schilter et al., 2003). Unlike the internationally accepted approaches for the safety evaluation of food additives or food contaminants (WHO, 1987), in which in vitro or in vivo studies with animals lead to the determination of health-based guidance values of intake and consider large uncertainty factors, a similar approach in the case of botanical products may not be appropriate (Schilter et al., 2003). Tests in the case of botanical products would involve high amounts of food to be administered to the animals in order to obtain uncertainty factors that are comparable to those obtained through the administration of pure compounds (Schilter et al., 2003). A guidance for the safety assessment of botanical products used in food and food supplements, consistent with the existing food regulations at that time, was proposed by an expert group of the Natural Toxin Task Force of the European Branch of the International Life Sciences Institute (ILSI Europe); they considered the framework of risk assessment as a basis for the safety evaluation of these products (Schilter et al., 2003). The document highlights the importance of standardization of the products, considers data regarding history of use, use of the product for different applications, intended use, dietary exposure, dietary consequences, and refers to hazard identification and characterization, risk characterization, and post-launch monitoring, and includes a decision tree as a guidance tool for the determination of the information that needs to be considered for the safety evaluation of these products. Other publications complement the criteria for safety assessment, such as the decision tree proposed by Walker (2004) as well as examples of assessments by van den Berg et al. (2011) and Speijers et al. (2010). Plant extracts/essential oils intended to be utilized in foods are generally regulated as food additives, food flavorings, or classified as “generally recognized as safe” (GRAS) substances, after fulfilling the requirements for each case. Specific databases of safe compounds, permitted concentrations, and permitted food uses are provided by different regional authorities (Malhotra, Keshwani, and Kharkwal, 2015). In the European Union (EU), the procedure for authorization of a flavoring substance is common to the one established for food additives and enzymes (European Commission, 2008a, 2008b). Annex I of Regulation 1334/2008 (European Commission, 2008b) lists the flavoring substances that can be utilized in foods. It is suggested to refer to its amendments in EUR-Lex for updates (European Union, 2019), and the Food Flavorings Database is also available (DG SANTE, 2019; European Commission, 2019). In the case of active packaging, all components not available in the positive lists of the regulations and not recognized as a food additive must be below 10 µg per kg of food or food simulant (Chibane et al., 2018), according to the EU regulation for food-contact materials (European Commission, 2009, 2011). The United States Food and Drug Administration (FDA) refers to food additives and GRAS substances in the Code of Federal Regulations, Title 21 (U.S. Food and Drug Administration, 2018). There is also the “Substances Added to Food” inventory (U.S. Food and Drug Administration, 2019), which includes ingredients regulated by the FDA, such as food additives, flavoring substances, and GRAS substances, as well as substances formerly utilized (prohibited substances, delisted color additives, and no longer GRAS substances); the inventory is only a partial list of food ingredients, and inclusion of information from non-FDA entities does not indicate an FDA approval or evaluation of this use. Even though some plant extracts/essential oils/botanical products constituents have been approved for their use in foodstuffs and classified in one of the categories mentioned, some intoxications reported

Naturally Occurring Compounds – Plant Sources

563

with botanical products were due to contamination with other plant species or through misidentification of species (Schilter et al., 2003). Other cases of toxic effects were reported, such as allergic contact dermatitis when used frequently as in the case of aromatherapy (Carson and Riley, 2001; Bleasel, Tate, and Rademaker, 2002; Schaller and Korting, 1995; Trattner, David, and Lazarov, 2008). Cytotoxicity of Pimenta dioica and Rosmarinus officinalis essential oils was observed when evaluated ex vivo by Lorenzo-Leal et al. (2019) using the model of human-derived macrophage THP-1 cells. An in vivo experiment, however, showed a different result: a wound treatment of diabetic mice showed recovery when the animals were treated with Rosmarinus officinalis essential oil (Abu-Al-Basal, 2010). Wright et al. (2019) reported that Kakadu plum (Terminalia ferdinandiana) fruit and leaf extracts did not have any toxic effects against nauplii stage Artemia franciscana crustaceans but recommended that further toxicity studies in human cell lines be performed before the extracts are used commercially. Using a similar toxicity assessment using brine shrimp (Artemia salina) larvae, Kayode et al. (2018) found that Adansonia digitata leaf and stem-bark essential oils were non-toxic. Carvacrol showed a concentration-dependent toxic effect in vitro (Llana-Ruiz-Cabello et al., 2014), despite being registered as flavoring by the EU, FDA, and the Joint Food and Agriculture Organization/ World Health Organization (FAO/WHO); therefore, potential risks associated with the use of essential oils constituents should be continuously monitored (Donsì and Ferrari, 2016). In addition to the in vitro studies of the antimicrobial effect and the in vitro/in vivo toxicity evaluation, further research is needed regarding the interaction of the botanical products with the food matrix, the delivery systems of their active compounds, and their stability throughout shelf life. These data and the standardization of the composition of botanical products will be convenient for their safe use in foods at doses needed to produce the desired antimicrobial effect.

17.8 Application in Food and Sensory Analysis For many years, many of the published reports regarding the application of phenolic compounds (antioxidants and constituent of extracts and essential oils) as antimicrobials had been accomplished in model and laboratory systems while recently more studies have been carried out in real foods. The growing consumer demand for safe and natural products has led to the increase in publications focused on the study of plant extracts, essential oils, and their components, as antimicrobials in real food systems including different green leaves; fresh and processed tomatoes; a variety of whole and sliced fruits and vegetables; fruit juices and jams; nuts, seeds, and beans; flours and bread; fresh, cooked, and processed meats; cheeses, milk, and puddings; salsas and salad dressings. Table 17.3 displays some of the studies published between 2015 and 2019. The essential oils of spices and plants, as well as their major components, are more effective in microbiological media than when evaluated in real foods (Gutierrez, Barry-Ryan and Bourke, 2009). In most cases the inhibitory concentrations found in model systems increase significantly when evaluated with the same microorganisms in actual foods and, in consequence, few of the applications of phenolic antioxidants as antimicrobials have been successful (Kabara and Eklund, 1991). This reduction in the effectiveness observed in vivo represents an important limitation to the use of essential oils and phenolic antioxidants as antimicrobial agents in foods (Juven et al., 1994). Ribes et al. (2016) reported a difference between the effectiveness of oil-in-water emulsions of clove and cinnamon leaf essential oils against Aspergillus flavus, A. niger, and Penicillium expansum when tested in strawberry jams and in agar media; the time to observe the effectiveness of the essential oil was significantly longer in jams. Authors attributed this behavior to the difference in matrix structure and water contents and the possible antagonistic interactions with ingredients present in the jam. Yuan and Yuk (2018) found that 32 mg/ mL of Syzygium antisepticum plant extract was able to inhibit and retard staphylococcal growth at 10°C and ambient temperature, respectively, in cooked chicken; MIC values obtained in vitro for the extract against the same S. aureus strains were reported by the authors as 0.125 mg/mL, 256 times less than the MICs needed in the food. Interactions among phenolic groups and proteins, lipids, and aldehydes could explain, at least partially, the reduction of the antimicrobial effect of essential oils where major constituents are phenols.

Garlic crops

Watermelon juice Guavas

Fresh cabbage Shredded cabbages Kale leaves Fresh-cut cauliflower

Lettuce

Fresh-cut red chard

Romaine lettuce leaves Iceberg lettuce and chard

Mature green tomato Tomatoes

Cherry tomatoes

Fruits, Vegetables, and Roots Lamb’s lettuce

Food Evaluated

S. Typhimurium, S. aureus, and E. coli Total mesophilic bacteria, mold and yeast, and thermotolerant coliforms White rot (Sclerotium cepivorum Berk)

Pseudomonas fluorescens indole-3-acetic aciddependent type III secretion system E. coli O157:H7 Total mesophilic microorganisms E. coli and Pichia pastoris E. coli O157:H7 and L. monocytogenes L. monocytogenes, E. coli O157:H7, and total yeasts and molds

E. coli O157:H7 Salmonella enterica

Rotting rate of tomatoes Fusarium solani and Rhizopus stolonifer Total microbial count E. coli O157:H7 Mixed inoculum of E. coli, L. monocytogenes, and S. enterica L. monocytogenes and Salmonella Typhimurium

Mesophilic aerobic bacteria, yeasts, and lactic acid bacteria Aspergillus flavus, A. oryzae, A. niger, and Alternaia alternata Botrytis cinerea

Target Microorganisms

Combination of thymol-eugenol and carvacrol-eugenol rinse Thyme, oregano, and lemongrass essential oil in vapor phase Carvacrol nanoemulsion and acidic electrolyzed water Cinnamon leaf essential oil emulsions UV-C, gamma irradiation combined with oregano or lemongrass essential oil in formulation with citrus extract and lactic acid Trans-cinnamaldehyde nanoemulsions Edible chitosan-cassava starch coatings with Lippia gracilis (LGRA106 and LGRA107 genotypes) essential oil Essential oils from Tagetes minuta L., Tagetes filifolia L., Origanum vulgare L. Origanum x majoricum, and Laurus nobilis L. and their binary combinations with iprodione

Perillaldehyde, the main constituent of Perilla frutescens essential oil Quinoa protein-chitosan edible film containing thymol nanoemulsion Encapsulated allyl isothiocyanate Combination of thymol and salicylic acid Essential oil from bark and leaf of Adansonia digitata Ultrasound assisted with oregano essential oil Essential oils of Origanum vulgare L. and Rosmarinus officinalis L. alone and in combination Cinnamon bark oil emulsion washing and ultraviolet-C radiation Cold nitrogen plasma and clove oil alone and in combination Ultrasound treatment combined with oregano and thyme essential oils Caryophyllene-rich white and black pepper essential oils

Oregano and thyme essential oils

Tested Agent(s)

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3

(Continued)

Camiletti et al. (2016)

Jo et al. (2015) De Aquino et al. (2015)

Yuan et al. (2019) Hyun et al. (2015) Sow et al. (2017) Kang et al. (2019) Tawema et al. (2016)

Myszka et al. (2017)

Cui, Ma, and Lin (2016) Millan-Sango et al. (2016)

Park et al. (2018)

Wu et al. (2015) Kong et al. (2016) Kayode et al. (2018) Millan-Sango et al. (2015) Barbosa et al. (2016)

Robledo et al. (2018)

Tian et al. (2015)

Siroli et al. (2015)

References

564 Antimicrobials in Food

Strawberry jams

Peach (Prunus persica L. Batsch) slices Fresh-cut melons Strawberries

Grape berries (Vitis vinifera L. x V. labruscana Bailey) Mangaba fruit

Raw apple juice Reconstituted orange juice Table grapes

Fungal decay percentage and total yeast and mold counts Aspergillus flavus, A. niger, and Penicillium expansum A. flavus, P. expansum, Zygosaccharomyces rouxii, and Z. bailii Aspergillus niger

S. Typhimurium, mesophilic aerobes, yeasts and molds Total aerobic mesophilic bacteria, yeast and mold, and Bacillus cereus Mesophiles, Pseudomonas spp., Enterobacteriaceae, and yeasts and fungi Total microbial count and mold and yeast counts Colletotrichum acutatum

Total aerobic mesophilic and psychrophilic bacteria, Enterobacteriaceae, lactic acid bacteria, and yeast and molds Saccharomyces cerevisiae L. monocytogenes and Salmonella enterica serovar Typhimurium A. flavus and P. expansum Alicyclobacillus acidoterrestris Yeast-mold growth

Fresh-cut carrot slices

Orange and pomegranate juices Cantaloupe juice

Aspergillus caronarius E. coli O157:H7, S. enterica, and L. monocytogenes Aletrnaria alternata

Apples and pears slices Organic cantaloupes Dragon fruit (Hylocereus undatus)

Target Microorganisms

Aflatoxin B1 production by A. flavus Penicillium expansum

Licorice Golden Delicious apples

Food Evaluated

References

Frazão et al. (2017)

Myrcia oata Cambeseedes essential oils

Oil-in-water emulsions containing cinnamon bark oil, zinc gluconate and transferulic acid

Clove and cinnamon leaf essential oils in oil-in-water emulsions Cinnamon bark-xanthan gum emulsions

Citral-based edible coatings Carboxymethylcellulose coating with Lippia sidoides essential oil Alginate-limonene edible coatings

Posidonia oceanica and green tea extract

Oh et al. (2017)

Ribes et al. (2018)

Ribes et al. (2017)

Ribes et al. (2016)

Dhital et al. (2018)

Oliveira et al. (2019)

Piva et al. (2017)

(Continued)

Basak and Guha (2017) de Pascoli et al. (2018) Takama and Korel (2017)

Sánchez-Rubio et al. (2016) Sarkar et al. (2017)

Martínez-Hernández et al. (2017)

Kapetanakou et al. (2018) Zhang et al. (2016) Castro et al. (2017)

Li et al. (2016) Frankova et al. (2016)

Thermo-ultrasound and cinnamon leaf essential oil Starch octenyl succinate stabilized emulsions with nisin and/ or thymol Betel leaf (Piper betle L.) essential oil Piper peltatum and Piper marginatum Alginate preharvest spray and postharvest coating with and without vanillin Lemongrass oil droplet size in chitosan emulsion as coating

Litsea cubeba essential oil Clove, cinnamon, lemongrass, and oregano essential oils combined with warm air flow Sodium alginate-cinnamon essential oil coatings Organic thyme oil emulsion Cinnamon (Cinnamomun zeylanicum) and clove (Eugenia caryophyllus) essential oil Carvacrol-loaded chitosan-tripolyphosphate nanoparticles

Tested Agent(s)

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 565

Meat and Seafood Ground beef

Oats

Maize kernels

Corn kernels

Valencia rice grain

Loaf bread Bread Par-baked wheat and sourdough bread Cake

Wheat flour

Clover (Trifolium resupinatum) sprouts

Tomato paste Cereals, Grains, and Nuts Young coconut liquid endosperm Peanut, cashew, almond, walnut pistachio, and hazelnut Stored peanut seeds Almonds Tofu Fresh green beans (Phaseolus vulgaris L.)

Food Evaluated

Methicillin-resistant Staphylococcus aureus Listeria monocytogenes

Total aerobic bacteria, total coliforms, and molds and yeasts Mycotoxin production by Aspergillus parasiticus and Fusarum poae A. parasiticus growth and aflatoxin reduction Mold spoilage Aspergillus niger and Penicillium paneum Aerobic mesophilic bacteria, yeast and molds, and Salmonella Alternaria alternate, Bipolaris oryzae, Fusarium graminearum, F. equiseti, and F. verticillioides A. parasiticus, Fusarium tricinctum, F. verticilloides, A. alternata, and Gibberela zeae Fusarium graminearum growth and deoxynivalenol and zearalenone production Aspergillus flavus, A. parasiticus, and A. clavatus

A. flavus Salmonella Enteriridis PT30 and S. Tennessee K4643 S. aureus E. coli and S. Typhimurium

Yerba mate (Ilex paraguarensis) aqueous extracts Clave (Syzygium aromaticum) and cinnamon (Cinnamomus cassia) essential oils

Vapor phase essential oil of cinnamon leaves, thyme, clove, oregano, and ginger

Cananga odorata essential oil

Gaseous allyl isothiocyanate

Allyl, benzyl, and phenyl isothiocyanates Lavender and melissa waste Thyme essential oil Microencapsulated and free thyme (Thymus vulgaris) essential oil Clove (Syzygium aromaticum L. Merr. & Perry) essential oil

Gaseous allyl isothiocyanate

1500 ppm malic acid and 75 ppm nisin Isothiocyanates generated in oriental and yellow mustard flours (E)-2-hexenal Cinnamonum cassia oil Liposome-encapsulated clove oil Carvacrol incorporated in modified chitosan coating, gamma irradiation, and modified atmosphere packaging alone or in combinations Thyme (Thymus vulgaris) essential oil

E. coli O157:H7 Aflatoxins produced by Aspergillus parasiticus

Tested Agent(s) Carvacrol and potassium sorbate

Salmonella enterica serovar Typhimurium

Target Microorganisms

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3 (CONTINUED) References

(Continued)

Burris et al. (2015) Khaleque et al. (2016)

Božik et al. (2017)

Kalagatur et al. (2018)

Tracz et al. (2017)

Santamarina et al. (2016)

Saladino et al. (2017) Vasileva et al. (2018) Debonne et al. (2018) Gonçalves et al. (2017)

Nazareth et al. (2016)

Shirvani et al. (2016)

Ma et al. (2017) Tsai et al. (2017) Cui et al. (2015) Severino et al. (2015)

Garbriel and Estilo (2015) Hontanaya et al. (2015)

Batista et al. (2019)

566 Antimicrobials in Food

Fresh poultry meat

Raw chicken fillets

Low/sodium sliced vacuum/packed turkey breast ham

Doner kebabs Sheep burgers

Chinese sausages

Sausage

Portuguese sausages paínhos and alheiras Turkish fermented sausages (sucuk)

Dry fermented sausage-chouriço

Cooked sausage

Ground chicken Beef meatballs

Food Evaluated

Fungus community assessed by high-throughput sequencing Listeria monocytogenes Total viable counts, Pseudomonas spp., Enterobacteriaceae, and lactic acid bacteria Lactic acid bacteria, total psychrotrophic bacteria, Listeria innocua, thermo-tolerant coliforms, Salmonella sp., and Staphylococcus sp. Listeria monocytogenes, psychrotrophic and aerobic mesophilic bacteria, lactic acid bacteria, Pseudomonas spp., Enterobacteriaceae, and molds and yeasts Total mesophilic aerobic bacteria, psychotropic aerobic bacteria, and total coliforms

Salmonella spp. Staphylococcus aureus, Listeria monocytogenes, Salmonella Enteritidis, and Campylobacter jejuni Total aerobic mesophilic bacteria, yeasts and molds, E. coli, Clostridium spp., and total Enterobacteriaceae count Salmonella spp., L. monocytogenes, S. aureus, and lactic acid bacteria Total viable aerobic bacteria, E. coli, Staphylococcus spp., Salmonella spp., and Listeria monocytogenes Surface mold, total mesophilic bacteria, Micrococcaceae, Enterobacteriaceae, yeasts and molds, and lactic acid bacteria Salmonella Typhimurium, Listeria monocytogenes, Staphylococcus aureus, Shigella flexneri, and Escherichia coli Listeria innocua

Target Microorganisms

References

Chitosan/montmorillonite nanocomposites incorporated with rosemary essential oil

Sodium alginate, galbanum oleoresin gum, and Ziziphora persica essential oil-based edible coatings

Carvacrol and high-pressure processing

Grapefruit seed extract Oregano extract

Tosati et al. (2018)

Hydrogels made of turmeric residue and gelatin or cassava starch and gelatin with purified curcumin and UV-A light treatment Flos Sophorae (Sophora Japonica L. flower buds) or rutin

(Continued)

Lauriano de Souza et al. (2019)

Hamedi et al. (2017)

de Oliveira et al. (2015)

Haskaraca et al. (2019) Fernandes et al. (2016)

Tang et al. (2019)

Cruz-Gálvez et al. (2018)

Soncu et al. (2018)

Catarino et al. (2017)

García-Díez et al. (2016)

Šojić et al. (2015)

López-Romero et al. (2018) Pesavento et al. (2015)

Potato starch-based coating with Hibiscus sabdariffa acetone and methanol extracts

Bay, garlic, nutmeg, oregano, rosemary, and thyme essential oils Whey protein coating containing Origanum virens essential oil Chitosan coatings with thyme or rosemary essential oils

Gallic acid, eugenol, and heating Thymus vulgaris, Origanum vulgare, Cinnamomum zeylanicum, Rosmarinus officinalis, and Salvia officinalis Nutmeg (Myristica frangrans) essential oil

Tested Agent(s)

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 567

Rainbow trout (Oncorhynchus mykiss)

Minced trout fillets

Organic beef jerky

Ready to cook pork chops

Reconstructed chicken product

Raw and thermally processed chicken meat

Cooked chicken

Ostrich meat

Pork loins Black wildebeest biceps femoris muscles Lamb meat

Refrigerated beef steaks

Beef meat

Marinated chicken meat

Food Evaluated

L. monocytogenes, total mesophilic and psychrotrophic bacteria, Pseudomonas spp., P. fluorescens, Shewanella putrefaciens, lactic acid bacteria, and Enterobacteriaceae Total viable count, Enterobacteriaceae counts, lactic acid bacteria, H2S-producing bacteria Total viable count, lactic acid bacteria, and Pseudomonas spp.

Total viable counts, lactic acid bacteria, Enterobacteriaceae, and Pseudomonas spp. Penicillium corylophilum

Total viable counts Total viable counts, lactic acid bacteria, and coliform counts Total viable counts, lactic acid bacteria, and Pseudomonas spp. Total viable counts, lactic acid bacteria, Staphylococcus aureus, and Escherichia coli Staphylococcus aureus and methicillin-resistant S. aureus Aerobic and anaerobic mesophilic bacteria, fecal coliforms, lactic acid bacteria, Staphylococcus aureus, Salmonella spp., Escherchia coli, and Salmonella Enteritidis Mesophilic and psychrotrophic aerobic counts

Total viable count, lactic acid bacteria, Brochothrix thermosphacta, Pseudomonas spp., total coliforms, Escherichia coli, yeasts and molds Psychrotrophic bacteria, Brochothrix thermosphacta, Pseudomonas spp., and Enterobacteriaceae Total viable counts, Pseudomonas spp., lactic acid bacteria, Enterobacteriaceae, and yeasts and molds

Target Microorganisms

References

Volpe et al. (2015) Raeisi et al. (2015)

Carboxymethyl cellulose coatings incorporated with Zataria multiflora Boiss. essential oil and grape seed extract

(Continued)

Kakaei and Shahbazi (2016)

Ji et al. (2019)

Zhang et al. (2018a)

Serrano-León et al. (2018)

Stojanović-Radić et al. (2018)

Yuan et al. (2019)

Hedayati Rad et al. (2018)

Pabast et al. (2018)

Fang et al. (2018) Shange et al. (2019)

Langroodi et al. (2018)

Sirocchi et al. (2017)

Karam et al. (2019)

Carrageenan-based edible coatings with lemon essential oil

Cinnamon bark, citronella, and may chang essential oils and their combinations in vapor phase Chitosan-gelatin film incorporated with grape seed extract and Ziziphora clinopodioides essential oil

Chitosan films containing peanut skin and pink pepper residue Chitosan edible coatings with bamboo vinegar

Basil and/or rosemary essential oils

Chitosan coatings with free or nano-encapsulated Satureja khuzestanica essential oil Kefiran/waterborne polyurethane film incorporated with Zataria mltiflora and Rosmarinus officinalis essential oils Syzygium antisepticum extract

Chitosan edible coating with Zataria multiflora essential oil and hydroalcoholic extract of sumac in modified atmosphere packaging Gallic acid/chitosan coating Oregano essential oil

Rosemary essential oil and modified atmosphere packaging

Thymol and carvacrol combined with air storage and vacuum packaging

Tested Agent(s)

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3 (CONTINUED)

568 Antimicrobials in Food

Skim milk Pasteurized milk Vanilla cream pudding Seasonings Salsa (tomatoes, jalapeño peppers, onions, garlic, and distilled vinegar) Mayonnaise-based salad dressing

Milk

Deepwater pink shrimp (Parapenaeus longirostris Lucas 1846) Dairy Cheese Low-fat cut cheese

Salted sardines

Black sea bream (Acanthopagrus butcheri Munro) fillets Atlantic salmon (Salmo salar)

Grass carp (Ctenopharyngodon idellus) Sea bass (Dicentrarchus labrax)

Bream (Megalobrama ambycephala)

Semi-fried coated rainbow trout (Oncorhynchus mykiss) Abalone (Haliotis discus hannai Ino)

Food Evaluated

E. coli O157:H7, S. Typhimurium, and L. monocytogenes S. pombe and Z. bailii

Listeria monocytogenes Staphylococcus aureus, psychrophilic bacteria, and yeasts and molds Staphylococcus aureus, Bacillus licheniformis, and Enterococcus hirae Farrow and Collins Pseudomonas putida and Staphylococcus spp. Foodborne isolated Staphylococcus aureus Five strain mixture of Listeria monocytogenes

Mesophilic and psychrophilic bacteria and Pseudomonas spp. Total microbial counts, mesophilic rod lactic acid bacteria, Enterobacteriaceae, staphylococci and halophilic bacteria Total viable counts, psychrotrophic bacterial counts, and Enterobacteriaceae

Total viable count, psychrophilic bacterial counts, and Pseudomonas spp. Total viable count, Pseudomonas spp., H2Sproducing bacteria, Enterobacteriaceae, and lactic acid bacteria Total viable count, psychrophilic bacteria, lactic acid bacteria, Enterobacteriaceae, Pseudomonas spp., and H2S-producing bacteria Pseudomonas spp. and Aeromonas spp. Total colony counts, total aerobic counts, and psychrotrophic counts Shewanella spp.

Target Microorganisms

References

Kim and Kang (2017) Monu et al. (2016)

Carvacrol, eugenol, and trans-cinnamaldehyde

Bevilacqua et al. (2016) Shi et al. (2017) Lianou et al. (2018) Carvacrol and/or ohmic heating treatments

Odorless citrus extract Nisin and cinnamaldehyde Cinnamon extract and storage temperature

Dannenberg et al. (2016) Artiga-Artigas et al. (2017)

Pink pepper tree (Schinus tereinthifolius Raddi) essential oil Sodium alginate-mandarin fiber-oregano essential oil edible coatings Free and nanoemulsified thyme essential oil

Jemaa et al. (2017)

Alparslan et al. (2016)

Alfonzo et al. (2017)

Alves et al. (2018)

Wright et al. (2019)

Huang et al. (2017) He et al. (2019)

Nisar et al. (2019)

Hao et al. (2017)

Raeisi et al. (2016)

Edible gelatin coating enriched with orange (Citrus sinensis (L.) Osbeck) leaves essential oil

Chitosan films with grape seed extract and carvacrol microcapsules Lemon essential oil micro-emulsions

Kakadu plum (Terminalia ferdinandiana) extracts

Cinnamon bark oil Superchilling storage-ice glazing and clove essential oil

Pectin-based coatings enriched with clove essential oil

Shallot (Allium ascalonicum L.) fruit and ajwain (Trachyspermum ammi (L.) Sprague) seed extracts Sodium alginate coating with bamboo leaf extract or rosemary extract

Tested Agent(s)

Recent Studies on the Application of Plant Essential Oils and Extracts as Antimicrobials in Real Food Systems

TABLE 17.3 (CONTINUED)

Naturally Occurring Compounds – Plant Sources 569

570

Antimicrobials in Food

Juven et al. (1972) showed that oleuropein antimicrobial activity could be reduced by the addition of triptone and/or yeast extract to the culture medium. Tassou and Nychas (1994) demonstrated that inoculum size, oleuropein concentration, and pH influenced significantly S. aureus growth and lag time while proving that the efficiency of phenolic compound antimicrobial action was reduced in foods with relatively low protein content; the principal cause of antimicrobial activity loss could be probably the solubilization of these compounds in the lipidic phase of the medium, reducing its availability to act as an antimicrobial. Aureli et al. (1992), evaluating the antimicrobial capacity of thyme against L. monocytogenes growth in model systems and in a real food, found that essential oil antilisteric efficiency decreased when used in vivo (ground pork meat) in comparison with the behavior in laboratory media (solid and liquid). Spencer et al. (1988) reported that the interaction or complex formation between phenols and proteins depends partially on protein characteristics, pH, and on the phenolic group contained by the molecule. This interaction takes place by hydrogen bonds between phenolic groups and peptides and also by hydrophobic interactions. The interactions of aldehydes with proteins have been extensively studied, since protein addition to aldehyde solutions can decrease the effective concentration of these groups (Cha and Ho, 1988; Hansen and Heinis, 1991; Montgomery and Day, 1965). Citral (lemon flavor component) concentration was reduced almost 100% when 5% of casein or soy protein isolate was added to aqueous solutions; 68% initial vanillin concentration, measured by HPLC, was lost after 26 hours in drinks containing aspartame (Tateo et al., 1988). The presence of fats and/or proteins in food could also provide a protective layer for bacteria while the physical structure of foods may reduce the diffusion of active compounds to target microorganisms (Perricone et al., 2015). Carvalho et al. (2015) observed that the effect of thyme essential oil against S. aureus and L. monocytogenes was reduced in a semi-solid cheese model in comparison with a cheesebased broth; the effect was attributed in part to the limited diffusion of the EO in the solid media. Burris, Higginbotham, and Stewart (2015) observed that higher concentrations of yerba mate extracts were needed to inactivate methicillin-resistant S. aureus in ground beef with high fat content. The effectiveness of plant-derived antimicrobials in a certain food matrix is different for different bacterial strains; Bajpai et al. (2019) studied the effect of the biflavonoid ametoflavone, isolated from Nandina domestica, against S. aureus and E. coli inoculated in two different food models: minced chicken breast meat and apple juice after 10 days of storage at 4°C. It was found that MIC levels of ametoflavone (62.5 and 125 μg/mL for S. aureus and E. coli, respectively) were more effective against E. coli in apple juice but more effective against S. aureus in chicken. Stojanović-Radić et al. (2018) observed that basil essential oil has a higher effect against Salmonella when it is applied in fresh chicken meat, while its combination with rosemary essential oil or rosemary essential oil alone are more efficient in the drier, thermally processed meat. The considerably higher levels of essential oils or plant extracts needed in food products to reach the desired antimicrobial effect will most likely affect sensory characteristics of the product and potentially turn it unacceptable. Thus, it is important to consider which essential oil/food combinations will be of liking to the consumer (Van Haute et al., 2016). Only four (thyme, holybasil, cassia, and clove) out of nine tested essential oils were acceptable to consumers at the required concentration to have an antimicrobial effect in chicken sausages (Sharma et al., 2017). Frankova et al. (2016) studied the effect of oregano, clove, and cinnamon oils in vapor phase in combination with warm airflow as treatment for apples inoculated with P. expansum; the highest tested concentration of oregano essential oil (16 μL/L) was the most effective treatment after 21 days of storage. That same concentration of oregano essential oil was shown to negatively affect the organoleptic properties of treated apples, in contrast with treatments with other studied essential oils or lower concentrations of oregano which were satisfactorily evaluated; thus, it was concluded that the most effective concentration of oregano essential oil cannot be utilized against the growth of phytopathogenic fungi on apples (Frankova et al., 2016). In orange and pomegranate juices, concentrations of cinnamon leaf essential oil above 0.04 mg/mL were found to have negative changes in aroma and flavor which made them unacceptable for consumers; the accepted concentration was 0.02 mg/mL which, in combination with ultrasonic treatment (24 kHz; 105 μm; 33.31 W/mL 30 min) at 50°C, resulted in 2.52 and 2.81 log reductions of S. cerevisiae in orange and pomegranate juice, respectively (Sánchez-Rubio et al., 2016).

Naturally Occurring Compounds – Plant Sources

571

The use of natural antimicrobials in combination with other environmental stress factors not only can enhance their antimicrobial properties but also makes possible the development of products that consumers are demanding by reducing the amounts of synthetic or natural antimicrobials needed to assure microbial stability. The combined effect of carvacrol at acceptable sensory levels (200 ppm) with highpressure processing (600 MPa/180s/25°C) was able to significantly extend the shelf life of low-sodium sliced vacuum-packed turkey breast ham (de Oliveira et al., 2015). It has been demonstrated that vanillin (4-hydroxy-3-methoxy-benzaldehyde) inhibited microbial growth in laboratory media and fruit purées stored at 25–27°C (López-Malo et al., 2000). Promising results have been obtained by Cerrutti et al. (1997) and Cerrutti and Alzamora (1996) in strawberry (pH 3.4, aw 0.95) and apple purées (pH 3.5, aw ≅ 0.99) preserved by combined factors (hurdle technology); strawberry purée with 3000 ppm vanillin and reduced aw inhibited the native and inoculated flora (Saccharomyces cerevisiae, Zygosaccharomyces rouxii, Z. bailii, Schizosaccharomyces pombe, Pichia membranaefaciens, Botrytis sp., Byssochlamys fulva, Bacillus coagulans, and Lactobacillus delbrueckii) for at least 60 days of storage at 25°C, while in apple purée with 2000 ppm vanillin, a germicidal effect was observed on inoculated S. cerevisiae, Z. rouxii, and D. hansenii. Cerrutti and Alzamora (1996) also reported that 3000 ppm of vanillin in a banana purée with pH 3.4 and aw 0.98 was inhibitory for S. cerevisiae, Z. rouxii, and D. hansenii. Castañón et al. (1999) evaluated the effects of vanillin (1000 or 3000 ppm) or potassium sorbate (1000 ppm) addition on the microbial stability during storage at 15, 25, or 35°C of banana purée preserved by combined methods (aw 0.97, pH 3.4); native flora (standard plate, yeasts and molds) counts during storage at different temperatures in the control purées (without antimicrobial addition), and in those containing 1000 ppm vanillin, demonstrated that a reduced storage temperature (15°C) was not enough to detain or delay the spoilage of the purée; after six days of storage at 15°C, native yeasts and molds reached counts of 104 CFU/g (about 3 log cycles greater than the initial count), and these purées were at this time sensorially unacceptable (odor and textural changes accompanying the microbial spoilage). The addition of 1000 ppm of vanillin increased the lag phase up to ≅ 16 days at 15°C, and the time to detect the microbial spoilage was extended to around 21 days. In the presence of 3000 ppm of vanillin or 1000 ppm potassium sorbate no microbial growth (< 10 CFU/g) was detected after 6 days and up to 60 days of storage (15, 25, or 35°C). Results obtained by Cerrutti and Alzamora (1996), Cerrutti et al. (1997), and Castañón et al. (1999) demonstrated that the addition of vanillin in combination with a slight reduction of aw and pH may be a promising method for fruit purée preservation and confirmed the antimicrobial properties of vanillin; they found that the protein and fat contents of the fruit partially determined the vanillin concentration necessary to obtain a sound product. The relatively high protein (1.2%) and fat (0.3%) contents in banana, compared with those of other fruits, explain the necessity of a higher vanillin concentration to obtain the same antimicrobial effects as in the other studied fruits. Castañón et al. (1999) reported results of sensory evaluation of banana purées containing 3000 ppm of vanillin or 1000 ppm of potassium sorbate; mean scores corresponded to products with a good overall acceptability with scores around 6 (like slightly). Studied purées were significantly different (p < 0.05) in odor (the one with vanillin was preferred) and flavor (the one with potassium sorbate was better) whereas there was not significant difference (p < 0.05) in color and overall acceptability. The flavoring characteristics of vanillin are well-accepted and have demonstrated compatibility with many fruits in concentrations up to 3000 ppm (Cerrutti and Alzamora, 1996). The microbial stability of mango juice (pH 4.9) supplemented with extracts of ginger (main antimicrobial compounds: zingerone, gingerol, and shogaol) and nutmeg (main antimicrobial compounds: myristicin and sabinene) was investigated during 3 months at room temperature by Ejechi et al. (1998); the combination of heating (15 min at 55°C) and 4% v/v of each spice extract inhibited microbial growth (yeasts and non-spore-forming bacteria) and created products with acceptable taste. Surface disinfection of tomatoes using cinnamic aldehyde was studied by Smid et al. (1996); whole tomatoes were dipped for 30 min in a solution containing 13 mM cinnamic aldehyde and then stored at 18°C in sealed plastic bags. The combination of the treatment with the natural phenolic compound and packaging under modified atmosphere reduced spoilage-associated fungi and bacteria on the tomatoes’ surface, increasing their shelf life up to 11 days. Batista et al. (2019) observed that the synergistic interaction of carvacrol and potassium sorbate in tomato paste eliminated Salmonella spp. at the third day of storage with no effects on physicochemical properties and a sensory acceptance index of 67% from a nine-point hedonic scale. A less severe thermal process could

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be applied to young coconut liquid endosperm to preserve sensory attributes while inhibiting E. coli O157:H7 growth if combined with malic acid and nisin treatments (Gabriel and Estilo, 2015). Strategies to reduce the amount of essential oil needed to ensure the microbiological safety of food products include the encapsulation of essential oils into emulsions or their utilization as components of active packaging systems instead of using them as ingredients in the food products. This in turn diminishes the unacceptable organoleptic changes in food that accompany elevated essential oil concentrations (Hyldgaard, Mygind and Meyer, 2012). Gonçalves et al. (2017) confirmed that the encapsulation of thyme essential oil decreased its minimal inhibitory concentration (MIC) values when tested in vitro, reasoning that the protective micro-environment that the particle wall provides retards the volatilization of the bioactive compounds, thus rendering the system more efficient to inhibit microbial growth. This functional barrier generated between the core and wall material helps to maintain the biological function of the core by avoiding chemical and physical reactions (Bakry et al., 2016). Martínez-Hernandez, Amodio, and Colelli (2017) reported that, unlike sliced carrots treated with a carvacrol solution, treatments with carvacrol-loaded chitosan-tripolyphosphate nanoparticles at the same concentration exhibited the lowest aroma score related to carvacrol off-flavors; encapsulated carvacrol was also effective in controlling microbial growth throughout 13 days of storage. Lee et al. (2019) found that encapsulation of rosemary extract in chitosan and γ-poly glutamic acid nanoparticles was able to improve its dispersibility in barley tea and enhanced its antimicrobial activity against B. subtilis. Chang et al. (2017) used sachets containing polyvinyl alcohol-oregano essential oil microcapsules to be utilized in iceberg lettuce containers as active packaging to control Dickeya chrysanthemi, yeasts and molds, and mesophilic aerobic bacteria; it was observed that the sachets allowed effective antimicrobial activity of oregano essential oil under high humidity and temperature conditions. Other important aspects to consider from the use of essential oils and plant extracts as antimicrobials in food systems are the effects that food processing techniques such as cooking or pressure could have on the antimicrobial activity of the compounds (Nikmaram et al., 2018). In this sense, Wang et al. (2018b) developed β-cyclodextrin and porous starch microcapsules for the encapsulation of clove oil; their microcapsules could endure 30 min of boiling and still present inhibition of mold spores on meat products. Given that food-contact surfaces are an important source of food contamination when not properly cleaned (Rouger, Tresse and Zagorec, 2017), as well as biofilm tolerance to chemical biocides and concerns about the environmental toxicity of residues from classical synthetic sanitizers (Bridier et al. 2015), recent studies have focused on the use of compounds extracted from aromatic plants; Rodrigues et al. (2017) and Tapia-Rodriguez et al. (2017) applied oregano essential oil and carvacrol to prevent S. aureus biofilm formation as well as biofilm formation and quorum sensing of P. aeruginosa, respectively. Cui et al. (2016) used salvia oil nanoliposomes to control S. aureus biofilm formation on milk containers. Plant bioactive compounds with antimicrobial properties are also being studied for their application as feed supplements for farm animals as antibiotic substitutes intended to improve animal health (GuilGuerrero et al., 2016), over the concern that antibiotic use in livestock has contributed to the emergence and transmission to humans of antibiotic-resistant bacteria. Careful consideration has to be given since compounds like carvacrol, oregano, and thyme essential oils have a broad-spectrum antimicrobial activity and, while potent against pathogens, they can also have an effect on beneficial ruminant bacteria (Benchaar et al., 2008). Of interest are the studies reviewed by Hassoun and Ҫoban (2017) since the effect of the use of essential oils and their components as fish dietary additives extended the delay of microbial growth in fish meat post-mortem.

17.9 Final Remarks The use of spices, herbs, plants, essential oils, and related phenolic compounds is limited due to the high minimal inhibitory concentrations needed in actual foods with high protein and/or fat contents, which impart undesirable flavor and/or odor. These undesirable effects can be minimized if the natural compound is utilized in combination with other environmental stress factors in the frame of the “hurdle technology.” Table 17.4 presents some general guidelines to be followed for selecting plant-derived

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TABLE 17.4 Factors That Influence Plant-Derived Antimicrobial Selection and Effectiveness Antimicrobial Characteristics Possible synergistic, antagonist, or additive interaction effects with other antimicrobial factors (composition of food) Initial contamination level

Handling and distribution

Toxicological and Legal Aspects, Solubility, Sensory Impact, Cost Moisture content, fat and protein content, water activity and pH, presence of other inhibitors (acids, salts, smoke, antimicrobials), interactions with food matrix and other food additives Sanitary conditions of ingredients and raw materials, sanitary conditions of equipment, processing conditions, type of potential growing microorganisms Length of storage, temperature of storage, packaging

antimicrobials. In this way, considering the consumers’ interest in more “natural” foods, the potential for applications in minimally processed fruit and vegetables appears to be good (Alzamora et al., 1995, 2016). However, for a wider and more rational use of these natural compounds, some points should be addressed: • The extraction methodology utilized to obtain the essential oil or extract. • The response of key microorganisms to the multi-target preservation system in vitro and then evaluation of its efficacy in vivo. • The form in which the plant, herb, spice, essential oil, or extract will be incorporated in the foods without adversely affecting sensory, nutritional, and safety characteristics, and without increasing significantly the formulation, processing, or marketing costs of the minimally processed product to which they are added.

17.10 Acknowledgments Authors López-Malo, Paris, Lastra-Vargas, and Palou gratefully acknowledge financial support from the National Council for Science and Technology (CONACyT) of Mexico (Project CB-2016-01-283636) and Universidad de las Américas Puebla (UDLAP Projects 2409 and 3555). Authors Alzamora, Coronel, and Gómez thankfully acknowledge financial support from the National Council of Scientific and Technical Research (CONICET) of Argentina and Universidad de Buenos Aires.

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18 Naturally Occurring Compounds – Animal Sources Jarret Stopforth and Travis Kudron CONTENTS 18.1 Introduction................................................................................................................................. 596 18.2 Lactoperoxidase........................................................................................................................... 597 18.2.1 Molecular Properties.................................................................................................... 597 18.2.1.1 Occurrence and Biosynthesis....................................................................... 597 18.2.1.2 Chemistry and Structure.............................................................................. 597 18.2.1.3 Stability........................................................................................................ 598 18.2.2 Antimicrobial Activity.................................................................................................. 598 18.2.2.1 Mode of Action............................................................................................ 598 18.2.2.2 Specificity.................................................................................................... 599 18.2.3 Applications in Food..................................................................................................... 600 18.3 Transferrins................................................................................................................................. 601 18.3.1 Lactoferrin, Lactoferricin B, and Activated Lactoferrin.............................................. 601 18.3.1.1 Molecular Properties................................................................................... 602 18.3.1.2 Antimicrobial Activity................................................................................. 603 18.3.1.3 Applications in Foods.................................................................................. 608 18.3.1.4 Safety and Tolerance.....................................................................................610 18.3.2 Ovotransferrin...............................................................................................................611 18.4 Immunoglobulins.........................................................................................................................612 18.4.1 Lactoglobulins...............................................................................................................612 18.4.1.1 Molecular Properties....................................................................................613 18.4.1.2 Antimicrobial Activity..................................................................................614 18.4.1.3 Applications in Food.....................................................................................615 18.4.2 Ovoglobulins..................................................................................................................616 18.4.2.1 Molecular Properties....................................................................................616 18.4.2.2 Antimicrobial Activity..................................................................................617 18.4.2.3 Applications in Food.....................................................................................617 18.5 Avidin...........................................................................................................................................618 18.5.1 Molecular Properties.....................................................................................................618 18.5.1.1 Occurrence and Biosynthesis........................................................................618 18.5.1.2 Chemistry and Structure...............................................................................618 18.5.1.3 Stability.........................................................................................................618 18.5.2 Antimicrobial Activity...................................................................................................619 18.5.2.1 Mode of Action.............................................................................................619 18.5.2.2 Specificity.....................................................................................................619 18.5.3 Applications in Food......................................................................................................619 18.6 Lactolipids....................................................................................................................................619 18.6.1 Molecular Properties.................................................................................................... 620 18.6.1.1 Occurrence and Biosynthesis....................................................................... 620

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18.6.1.2 Chemistry and Structure.............................................................................. 620 18.6.1.3 Stability........................................................................................................ 620 18.6.2 Antimicrobial Activity...................................................................................................621 18.6.2.1 Mode of Action.............................................................................................621 18.6.2.2 Specificity.....................................................................................................621 18.6.3 Applications in Food......................................................................................................621 18.7 Defensins..................................................................................................................................... 622 18.7.1 Molecular Properties.................................................................................................... 622 18.7.1.1 Occurrence and Biosynthesis....................................................................... 622 18.7.1.2 Chemistry and Structure.............................................................................. 622 18.7.2 Antimicrobial Activity.................................................................................................. 623 18.7.3 Applications in Food..................................................................................................... 623 18.8 Chitosan....................................................................................................................................... 623 18.9 Other Antimicrobials of Animal Origin..................................................................................... 624 18.9.1 Pleurocidin.................................................................................................................... 624 18.9.2 Casocidin...................................................................................................................... 624 18.9.3 Lysozyme (Consult Chapter 14, “Lysozyme” for More Information).......................... 624 18.9.4 Lipids............................................................................................................................ 624 References............................................................................................................................................... 625

18.1 Introduction Natural antimicrobials include agents found in plants, microbes, insects, and animals (Naidu, 2000; Sofos et al., 1998). This chapter focuses on the natural antimicrobials that exist in animals where they evolved as host defense mechanisms and may exhibit antimicrobial activity in foods as natural ingredients or be extracted and used as additives in other foods. In general, animal-derived antimicrobials have been known for many years and, thus, there has been much interest in the isolation and purification of these compounds to be used as food antimicrobials or as adjuncts to other antimicrobials. The discussion in this chapter targets mainly those antimicrobials isolated from milk, especially of bovine origin, and from poultry eggs, since these are the major food products produced by animals for human consumption that possess antimicrobial activity devised by nature for protection and immune development in the neonate. The antimicrobials isolated from these products are generally broad-spectrum agents providing protection to the neonate against bacteria, fungi, parasites, and viruses, and in some cases are the only means of transferring protective factors to the unborn or newly born young (e.g., bovine colostrum is the only source of immune factors including antibodies from mother to newborn) (Floris et al., 2003). Maternal transfer of immune factors and antimicrobial substances provides the offspring with acquired immunity, leaving them relatively immunocompetent against diseases. Substances with antimicrobial properties from the mother, such as lactolipids in bovine milk, are not strictly immune factors and may possess both nutritional and protective qualities. Major challenges faced by the industry include difficulty in the production of these compounds in large enough quantities to be effective in a food and the incorporation of these compounds in foods while minimizing undesirable interactions, undesirable effects, or subsequent inactivation of their desirable effect. It should also be noted that the use of these compounds as antimicrobials in foods may not have the same effectiveness as if the compound were a natural ingredient in food. This is especially true in milk where the total antimicrobial activity is not necessarily a function of individual antimicrobials but probably the additive or synergistic effect of multiple antimicrobials (e.g., lactoferrin, lactoperoxidase, lactoglobulins). Despite the challenge of obtaining effective activity of antimicrobial compounds in a food matrix, there has been commercial application of various compounds isolated from animal products including the fortification of dairy products with milk antibodies, the fortification of dairy products including milk with substrates to facilitate lactoperoxidase production, and the use of monoacylglycerols from milk as biopreservatives in many foods. Furthermore, there are many proposed applications of antimicrobial compounds in foods or consumables including: (i) passive immunization with milk- and

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egg-derived antibodies; (ii) the addition of transferrins, lactoperoxidase, and immunoglobulins in oral healthcare products; and (iii) the use of lactolipids, immunoglobulins, and transferrins as ingredients in infant formulas. Products derived from animal sources, in particular eggs and milk, are usually characterized as generally recognized as safe (GRAS) compounds; however, these products do need to be evaluated for potential allergenic properties in sensitive individuals. The following sections provide a summary of the most important naturally occurring antimicrobial compounds found in animal tissues and in particular in bovine milk and poultry eggs.

18.2 Lactoperoxidase Lactoperoxidase (LP), a hemoprotein present in milk, tears, and saliva (Tenovuo and Pruitt, 1984), is the most abundant enzyme in bovine milk (Reiter, 1985a). The peroxidase activity associated with bovine milk was first demonstrated by Arnold in 1881 and the protein, termed lactoperoxidase, was isolated by Theorell and Akeson in 1943 (Naidu, 2000). The lactoperoxidase–thiocyanate–hydrogen peroxide interaction constitutes what is referred to as the LP system, wherein hydrogen peroxide serves as a substrate for LP in oxidizing thiocyanate (SCN–) and iodide ions, resulting in the generation of highly reactive oxidizing agents (Thomas, 1985; Naidu, 2003). The association of LP with the inhibition of microbial growth was first demonstrated by Wright and Trammer (1958), while characterization of the complete LP system including enzymes and substrates occurred later (Jago and Morrison, 1962; Reiter et al., 1963). Additionally, the LP system possesses hexokinase and glyceraldehyde-3-phosphate dehydrogenase activities, which may contribute to the antimicrobial action of the system (Carlsson et al., 1983). The LP system has the ability to inhibit bacteria, fungi, parasites, and viruses and is, thus, considered a broad-spectrum natural antimicrobial contributing to protecting the gut of weaning calves from enteric pathogens, protecting the mammary gland from disease, and indeed preserving milk (Bjorck et al., 1979; Reiter and Bramley, 1975; Reiter et al., 1980).

18.2.1 Molecular Properties 18.2.1.1 Occurrence and Biosynthesis LP is synthesized and secreted by ductal epithelial cells of the mammary gland and other exocrine glands (Parkos, 1997). The compound constitutes approximately 1% (10–30 µg/ml) of the whey proteins in the milk (Reiter, 1985a, b) and levels may be influenced by feeding practices, udder irritation, and estrogen levels (Janota-Bassalik et al., 1977; Kern et al., 1963; Kiermeier and Kuhlmann, 1972). The level of LP in bovine milk is about 20 times higher than that of human milk and changes constantly during the postpartum. Thiocyanate, which is required for the antimicrobial activity of the LP system, may be present in significant amounts in saliva, milk, and airway secretions, while hydrogen peroxide may be generated by microbial flora, usually bacteria, resident in sites such as the oral cavity, respiratory tract, or mammary gland or in situ by epithelial cells in these regions (Reiter and Perraudin, 1991). In bovine milk, the initial concentration of LP in colostrum is low, increasing to a peak at 4–5 days postpartum, after which it declines to a level considered relatively high and remains unchanged at that level during lactation (Kiermeier and Kuhlmann, 1972). In order to combat infections, the concentrations of LP and SCN– increase in milk from infected bovine udders as compared with normal, healthy udders (JanotaBassalik et al., 1977).

18.2.1.2 Chemistry and Structure It has been determined that bovine LP is comprised of a single peptide chain, with eight disulfide bonds contributing to the rigidity of the molecule (Sievers, 1980). The single polypeptide chain contains 612 amino acid residues with a molecular weight of about 80 kDa (Cals et al., 1991; Paul and Ohlson, 1985). The polypeptide chain contains 15 half-cystine residues and 4 or 5 potential N-glycosylation sites, and carbohydrate moieties account for almost 10% of the molecular weight (Cals et al., 1991). LP is a

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heme-containing enzyme that shares 50–70% amino acid sequence homology (particularly among the active-site related residues) with myelo-, thyro-, and eosinophil peroxidases (Dull et al., 1990), all of which have a protoporyphyrin IX catalytic center (Fenna et al., 1995; Sievers, 1979). The covalent linkages between heme and the protein are well-represented in the myeloperoxidase X-ray crystal structure (Fenna et al., 1995; Zeng and Fenna, 1992); however, little work has been done concerning the crystallographic data for LP. Although the details regarding the heme-binding site in LP have not been established, the heme structure has been studied in terms of its electron transfer mechanisms (Nakamura et al., 1986) as the heme moiety is essential for the development of the oxidation-reduction reaction associated with LP activity. The presence of an odd number of half-cystines supports the theory that a heme thiol is released from this enzyme by a reducing agent and suggests that the heme is bound via disulfide links to the peptide chain (Naidu, 2000), indicating that there are no free thiol groups present in the enzyme molecule (Ekstrand, 1994). The iron content of LP is 0.07% which corresponds to one iron atom per LP molecule as part of the heme group (Kussendrager and van Hooijdonk, 2000). The molecular conformation of LP is thought to be stabilized by the strong binding of a calcium ion (Kussendrager and van Hooijdonk, 2000). Although earlier research (Sievers, 1980) revealed that leucin is found on the N-terminus of the LP polypeptide chain, Watanabe et al. (2000) found that different preparations of natural LP may have different N-terminal amino acid residues. This heterogeneity may be a result of variation in terms of isolation methods (i.e., disk-electrophoresis and ion-exchange chromatography); however, the finding suggests that the change in N-terminus structure does not affect the conformation of LP. Other findings of this research using circular dichroism (CD) revealed that the secondary structure (α-helix) of LP is influenced by peptide bonds and that the protein is rich in β-sheet structure (Watanabe et al., 2000).

18.2.1.3 Stability It was reported that the LP system stored in airtight containers lost only 35% of the initial thiocyanate concentration during 18 months and that the system was strong enough to kill 106 CFU/ml of 4 test organisms. When the LP system was stored in the presence of air it lost thiocyanate activity after 7 d but after 516 days was still able to kill inocula of 106 CFU/ml Pseudomonas aeruginosa, Staphylococcus aureus, and Candida albicans and Escherichia coli within 2 h, 4 h, and 1 week, respectively (Bosch et al., 2000). It has been shown that during pasteurization, whole milk loses about 75% of its LP activity, while the purified LP was rendered unstable after 15 min of exposure (Wutrich et al., 1964). Research (Herandez et al., 1990) indicated that heat denaturation of LP in milk, whey, permeate, and buffer started at about 70°C (close to the temperature at which the native structure of the enzyme unfolds to follow first-order kinetics) and that the calcium ion concentration influenced the heat sensitivity of LP. The heat stability of LP is lower under acidic (pH 5.3) conditions and may be related to the release of calcium from the molecule (de Wit and van Hooijdonk, 1996). LP is deactivated during storage at pH 3 with partial denaturation at 2000 mg/l in bovine and human milk, respectively, as well as two major reservoirs of mammals, including a circulatory pool in the secondary (specific) granules of polymorphonuclear neutrophils at 15 to 30 μg/107 cells, depending on the age of the individual (Baggliolini et al., 1970; Baveye et al., 1999; Bennet and Kokocinski, 1978; Levay and Viljoen, 1995; Naidu, 2000), and a stationary pool on the mucosal surfaces (van Hooijdonk et al., 2000; Levy, 1996; Levay and Viljoen, 1995; Lonerdal and Lyer, 1995). The presence of LF in mucosal surfaces originates from epithelial cells or sub-mucosal glands that excrete LF by the influence of the parasympathetic nervous system (Naidu, 2000; Raphael et al., 1989; Testa, 2002). Examples of mucosal surfaces where LF can be found are lactating breast tissues, gastric tissues, and duodenal epithelial cells (Mason and Taylor, 1978). LF accounts for 11.5% of the total secretory proteins excreted by bronchial glands (Harbitz et al., 1984) and 25% of the total tear protein produced by lachrymal glands (Kijlstra et al., 1983). In addition, concentrations of 0.2 to 1.0 mg/ml are found in seminal plasma (Witchman et al., 1989) and have been found in the spermcoating antigen (Ashorn et al., 1986). The highest levels of LF (5–7g/l) are observed in colostrum and gradually decrease by almost sevenfold in mature milk during lactation (Hennart et al., 1991; Hirai et al., 1990; Levay and Viljoen, 1995; Lonnerdal and Iyer, 1995; Montagne et al., 1998; Playford et al., 2000). In plasma it is normally found at low levels (approximately 0.2–1.6 μg/ml) (Levay and Viljoen, 1995; Steijns and van Hooijdonk, 2000; van der Strate et al., 1999). Milk from mastitic cows contains higher levels of LF than that from healthy ones. In addition, the concentration of LF is higher when infection is caused by Staphylococcus aureus and streptococci species rather than when caused by coagulase-negative staphylococci or Corynebacterium bovis (Hagiwara et al., 2003; Kai et al., 2002), indicating the antimicrobial role of the compound and its potential to prevent disease. Increase of LF in plasma may occur during acute-phase host responses, such as infection, inflammation, or toxic shock (Klasing, 1984; Baynes et al., 1986; Naidu, 2000; Qadri et al., 2002). van der Strate et al. (1999) established a linear correlation between concentrations of LF in plasma and counts of neutrophils. Elevated levels of LF in plasma are a primary indication of an inflammatory state, such as septicemia and endotoxemia (Gutteberg et al., 1988) and are commonly followed by an increase of LF in serum, thus making LF levels a reliable diagnostic tool for pathological conditions (Qadri et al., 2002). Increases of LF both in plasma and serum also correlate with increases in C-reactive protein during meningococcal septicemia (Gutteberg et al., 1984), cystic fibrosis protein during cystic fibrosis (Barthe et al., 1989), cholecystokinin secretion during pancreatitis (Dite et al., 1989), or lysozyme during localized juvenile periodontitis (Friedman et al., 1983). The iron-chelating properties of LF are an additional reason for increased clinical levels of LF (e.g., during rheumatoid arthritis) (Blake et al., 1981; Bukhardt and Schwingel, 1986) as well as in various pathogenetic malignancies (especially carcinomas) when iron uptake by neoplastic cells is necessary (Barresi and Tuccari, 1987).

18.3.1.1.2 Physico-Chemistry LF is a single polypeptide chain with a molecular weight of 75 to 80 kDa and consists of approximately 690 amino acid residues (Baker et al., 2000; Metz-Boutigue et al., 1984; Powel and Ogden, 1990; Rey et al., 1990). The amino acid composition of LF includes 16–18 different amino acids (of which 8 are basic); the frequency and types of amino acids depend on the mammalian species (Naidu, 2000). The reported isoelectric point (pI value) for LF varies with the measuring method. According to electrophoretic, chromatofocusing, and Rotafors methods, the pI of LF is around 8.0 (Groves, 1960; Shimazaki et al., 1993; Szuchet-Drechin and Johnson, 1965), although it may range from 5.5 to 10.0 (Naidu, 2000). In general, LF is considered a basic protein (with an expected pI higher than 7.0), because its N-terminal region contains multiple arginine (arg) and lysine (lys) residues that make this region extremely basic (Vogel et al., 2002). The heat resistance (known to be stable at 90°C for 60 min) and heat-induced enthalpy of LF change depends on its iron-binding status and the pH of the substrate. The iron-saturated form of LF (i.e., holo-form [87 ± 3°C]) is more stable than the non-saturated form (i.e., apo-form [67 ± 3°C]) (Naidu, 2000; Paulsson et al., 1993; Reiter and Oram, 1967; Rossi et al., 2002).

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Neutral and basic pH enhances thermal denaturation of LF, whereas LF is more heat-stable at low pH, especially around 4.0 (Saito et al., 1994). The holo-form of LF is more resistant to proteolysis than the apo-form (Brock et al., 1976). Moreover, binding of Ca2+ via the carboxylate groups of the protein surface increases the chemical and thermal stability of bovine LF (Rossi et al., 2002). Specifically, the presence of Ca2+ causes an approximate 9°C increase in the denaturation temperature of both holo- and apo-forms of LF (Rossi et al., 2002).

18.3.1.1.3 Structure LF is characterized by an amphipathic structure with strong cationic properties on the N-terminus (Bellamy et al., 1992). The three-dimensional structure of human and bovine LF have been determined to a 2.8 Å resolution (Naidu, 2000; Thakurta et al., 2003), and a high degree of homology of this protein has been observed between species. LF is folded into similarly sized globular and homologous terminal N- and C-lobes, which are stabilized by intrachain disulfide bonds and linked by an α-helix that provides flexibility (Baker et al., 1987; Baveye et al., 1999; Naidu, 2000; Vorland, 1999). The symmetry of the lobes derives from gene duplication, and they are further divided into two similar domains of about 160 amino acids (Testa, 2002). Each lobe contains glycosylation sites (the number varies among mammalian species) in homologous position and has the capacity to bind one Fe3+ with high affinity (Kd = 1020 M–1) together with one CO32– or HCO3 – ion that is held electrostatically to an arginyl side group (Naidu, 2000). Each iron atom is coordinated to four protein ligands in accordance with the carbonate or bicarbonate ion. Certain areas within the molecular structure serve as potential cavities for binding not only of carbonate but also of larger anions. Specifically, the HCO3 – occupies a pocket between the iron and two positively charged groups and serves to neutralize this positive charge, thus facilitating binding of iron ions. Using X-ray structure analysis it has been concluded that the multi-domain nature and the flexibility of LF allow for binding of various metals with no significant changes in the total structure (Naidu, 2000). Apart from the sites for iron absorption, metal-binding sites for other cations, such as Cu(II), Cr(III), Mn(III), and Co(III), are available within the structure of LF similarly to serum transferrin (Ainscough et al., 1980; Hirose, 2000; Naidu, 2000). The anion-binding properties of LF are affected by the type of metal ion that is already bound. Anions that have been investigated for their potential to bind LF in the presence of Fe2+ or Cu2+ include carbonate, citrate, oxalate, and hybrid carbonate–oxalate complexes (Brodie et al., 1994). The role of the citrate/bicarbonate ratio is very crucial for the apo- to holo-form transition of LF (Nonnecke and Smith, 1984), that controls the iron-binding process and, hence, the biological activity of LF. LFcin is produced by peptic hydrolysis (also mentioned as cleavage degradation) of bovine or human LF (Bellamy et al., 1992; Tomita et al., 1994) with different proteases. Bovine LFcin (commonly termed “lactoferricin B” [LfcinB]), was found to consist of 25 amino acid corresponding to the sequence of residues 17 to 41 close to the N-terminal of the molecule (Pierce et al., 1991; Tomita et al., 1994; Wakabayashi et al., 1992). Human LFcin (commonly termed “lactoferricin H” [LfcinH]) was found to consist of 47 amino acid residues, corresponding to the sequence of residues 1 to 47 at the N-terminal of the human LF (Bellamy et al., 1992; Metz-Boutigue et al., 1984). Both LFcinB and H have an almost circular structure consisting of a loop of 18 amino acid residues that links two linear subfragments, and have molecular weights of 3126 and 5558, respectively (Bellamy et al., 1992; Metz-Boutigue et al., 1984; Pierce et al., 1991). The linkage of subfragments occurs through a disulfide bond between the two terminus cysteine residues of the loop (Bellamy et al., 1992).

18.3.1.2 Antimicrobial Activity 18.3.1.2.1 Mode of Action The diversity of substrates in which LF is present, its coexistence with other physiological substances, and the regulatory role of LF as acute-phase reactants explain its multifunctionality and its wide spectrum of antimicrobial activity (Naidu, 2000; van Hooijdonk et al., 2000). It is considered that the antimicrobial activity of LF is generally dependent on its protein conformation (structural characteristics and spatial orientation) and substrate conditions (Naidu and Arnold, 1997). In particular, two basic biochemical

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properties of LF contribute to its antimicrobial effect and involvement in the host defense: the extremely powerful iron-binding capability, and its strong interaction with other molecules and surfaces. The amphipathic structure and the net positive charge, especially on the strongly cationic N-terminus region of LF, are considered deterministic for its interaction with microbial membranes (Bellamy et al., 1992). Binding of LF occurs via specific membrane receptors that exist for example in the leukemic lines, the activated T-lymphocytes, the monocytes-macrophages, the brush-border membranes, the parenchymal liver cells, the breast epithelial cells, platelets, and neuronal cells (Testa, 2002). Specific molecules that interact with LF are glycosaminoglycans of epithelial milieu (e.g., heparin sulfate in mucins), collagens, fibronectins, and DNA of mammalian cells (Naidu, 2002; Nichols et al., 1990; Wu et al., 1995). As previously mentioned, LF is a broad-spectrum antimicrobial, and modes of action appear to be common for different types of microorganisms. Among the studies conducted to elucidate the mechanisms of antimicrobial activity, most have been performed with bacteria and viruses although work has been done with fungi and protozoa. The following section introduces potential mechanisms by which LF may inhibit microorganisms, specifically bacteria and viruses. LF may be bacteriostatic or bactericidal, and iron deprivation is the most likely underlying mechanism for the direct bacteriostatic effect of LF (Naidu, 2000) as demonstrated by in vitro studies. However, bacteria such as Escherichia coli, Moraxella catarrhalis, Neisseria spp., and Vibrio spp. have potential defensive mechanisms to counteract iron depletion, mainly by the formation of siderophores that mediate iron uptake. Alternatively non-siderophore-mediated iron uptake by bacteria may occur either through outer-membrane protein receptors (Neisseria meningitis) that recognize the complex of LF-iron and internalize the chelated iron cation (Mickelsen et al., 1982; Naidu, 2000; Schryvers and Morris, 1988; Tranter, 1994), or by the production of reductases as is the case with L. monocytogenes (Cowart and Foster, 1985). Moreover, the bacteriostatic activity of LF may be reduced by factors that are irrelevant to microbial resistance, such as inappropriate citrate/bicarbonate ratios, given that bicarbonate is essential for iron chelating, whereas citrate competes with LF for iron and, hence, makes the iron available for growth by the organisms (Reiter, 1978b). The bactericidal activity of LF is iron-independent and relates to direct binding of positively charged LF on negatively charged microbial outer membranes, which results in dispersion of lipopolysaccharides (or negatively charged fatty acids), increase in membrane permeability, and eventually cell death (Arnold et al., 1980; Baveye et al., 1999; Caccavo et al., 2002; Ellison et al., 1988; Naidu and Arnold, 1997; Pellegrini 2003; Rossi et al., 2002; Yamauchi et al., 1993). Ellison et al. (1990) suggested that Ca2+ and Mg2+ are the major cations that modulate the LF-induced damage in Gram-negative bacteria, and this conclusion was confirmed by Rossi et al. (2002). The binding of LF to microbial cell surfaces, however, is enhanced by bacterial receptors that exist in a variety of Gram-positive bacteria (e.g., lipoteichoic or teichoic acids, surface layer proteins, peptidoglycan components, heat shock proteins [Helicobacter pylori]) and Gram-negative bacteria (e.g., lipid A, whereas the pore-forming outer membrane proteins [porins] are a very common binding target of LF) (Amini et al., 1996; Baveye et al., 1999; Caccavo et al., 2002; Dhaenens et al., 1997; Erdei et al., 1994; Naidu, 2000; Naidu and Arnold, 1994; Naidu et al., 1993; Sallmann et al., 1999; Vorland et al., 1999c). Likewise, antibiotic potentiation has also been demonstrated in vitro as a result of increase in the permeability of outer membranes (Naidu and Arnold, 1994; Vorland et al., 1999c; Wakabayashi et al., 2002). An additional antimicrobial function that stems from the interaction of LF with the outer membrane of Gram-negative bacteria is the inhibition of microbial attachment to sub-epithelial matrix proteins, or alternatively the detachment of bacteria from mucosal surfaces (Kawasaki et al., 2000; Naidu and Bidlack, 1998). The blocking of cell-surface attachment factors, such as fimbriae and other adhesins, and inhibition of the colonization-factor antigens synthesis are associated with this antimicrobial mechanism (Naidu and Bidlack, 1998; Naidu, 2002). Moreover, LF binds on many tissue surfaces with higher affinity than cell-surface anchors of pathogens (Naidu, 2002). Bellamy et al. (1993) demonstrated an additional mechanism derived from membrane disruption (similar to that of polymyxin B) related to the inhibition of proline uptake. Optimal binding (dose-dependent) to membranes and biocidal effect was obtained at pH 6.0 and pH 7.5 for E. coli and Bacillus subtilis, respectively. The interaction of LF and LFcin with microbial membranes has also been found to account for their fungicidal activity (Bellamy et al., 1993; Wakabayashi et al., 1996).

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Lactoferrin has been studied in vivo and in vitro against E. coli O157:H7. The mechanisms used against E. coli O157:H7 and others are bacteriostatic, bactericidal, and anti-adhesion effects. By use of these three mechanisms lactoferrin has the potential to decrease colonization pressure on farms and in the long run potentially decrease E. coli O157:H7 illness in humans. There have been no reports of bacteria developing lactoferrin resistance (Rybarczyk et al., 2017). Another interesting aspect of bovine lactoferrin is that it has been shown to be safe for preterm neonates while holding promise to reduce major morbidities in preterm infants (Embleton et al., 2013). Many new techniques for food preservation are being researched and developed. Techniques such as HHP, irradiation, ohmic heating, cold plasma, and ultrasound are some of the newest in the last 10 years that have been worked on, yet not much information is available on the effect of a variety of compounds such as milk proteins and enzymes such as lactoferrin. As lactoferrin is susceptible to degradation by heat treatment some of these new techniques may be useful in preserving lactoferrins’ useful properties. Also lactoferrin may be able to enhance the safety and efficiency of the new technologies as well in addition to its existing biological function (Franco et al., 2018). More research is needed with these new technologies as food matrix interactions with lactoferrin can affect its properties. A more detailed investigation of the mode of action of both LF and its hydrolysates, oriented towards the effect of LF on liposomes, electrochemical potential (ΔΨ), and pH gradient of Gram-negative pathogens, has become available in the literature. Many studies aim to fully elucidate all the potential underlying mechanisms of antimicrobial activity, as well as to detect synergism phenomena between LF and other compounds. It was shown that 0.1 to 6.4 μM of iron-free LF increased permeability of the outer and inner membrane of E. coli, reduced by 50% or dissipated the electrical potential and the pH gradient, and also caused selective ion permeabilization on liposomes of E. coli (Aguilera et al., 1999, 2003). Different amino sequences of LFcinB were found to enter the cytoplasm of S. aureus and E. coli within 15 min of exposure, whereas lower quantities of LFcinB were also found in the cell wall (Haukland et al., 2001). Penetration and attachment to the cell wall was proven to be time- and concentration-dependent. Ulvatne et al. (2001) illustrated that 30 to 100 μg/ml of LFcinB, corresponding to concentrations below the minimum inhibitory concentrations (MIC) or minimum bactericidal concentrations (MBC), caused morphological changes of cells and depolarized cytoplasmic membrane and destabilized liposomes, causing leakage and fusion of liposome contents, whereas no cell lysis was observed. These indications emphasize that membrane destabilization is the major mechanism for the antimicrobial activity of LFcinB. Investigation of the synergism between LFcinB and antibiotics has provided useful indications about potential clinical applications as well as for elucidation of the mode of action of LFcinB and/or common antibiotics. The antimicrobial effect of antibiotics, which are usually excluded by the outer membrane of Gram-negative bacteria (e.g., erythromycin, vancomycin, and penicillin) may be enhanced by LFcin which destabilized bacterial membranes and facilitated their penetration (Diarra et al., 2002; Vorland et al., 1999b). Recent studies have further elucidated the mode by which LFcinB destabilizes bacterial membranes. Hossain et al. (2019) present the importance that rapid permeabilization of bacterial plasma membranes has in cessation of the cells based on cell length during interaction with LFcinB. The researchers further found that LFcinB-induced local rupture of lipid bilayers and rapid permeabilization of Gram-negative bacterial cell membranes is significantly affected by membrane potential. Other antibiotics, such as polymyxins which act on membranes, may compete with LFcin with a moderated antimicrobial effect. Similarly, the prevention of entry of ribosome-targeting aminoglycosides (gentamycin) into cells due to disturbance in the respiratory chain of the cytoplasmic membrane caused by LFcinB may also result in a reduction of the antimicrobial effect of antibiotics (Vorland et al., 1999a). For the above reasons, LFcinB acted synergistically with penicillin against S. aureus and with erythromycin against E. coli, whereas antagonism was evident between gentamicin and LFcinB against S. aureus (Vorland et al., 1999a). In another study, minocyclin and other compounds (e.g., acids, alcohols, and acylglycerols) enhanced the antimicrobial activity of LFcinB against antibiotic-resistant strains of S. aureus (Wakabayashi et al., 2002). The effect of sequence of application of LFcin and antibiotics was studied in vitro by Haukland and Vorland (2001), who suggested that LFcinB had no overwhelming effect when applied at the post-antibiotic stage against E. coli and S. aureus. Microbial resistance to LF is a potential response, and a possible resistance mechanism for E. coli and S. aureus involves the

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production of proteases that decompose LFcin (Ulvatne et al., 2002). Additionally, the presence of Mg2+ ions in the growth environment of cells prior to or during exposure to LFcin may also offer protection against subsequent exposure to LFcin by altering the properties of the outer membrane (Masschalk et al., 2003). With respect to antiviral activity of LF, according to Naidu and Bidlack (1998) and van der Strate et al. (2001), the potential mechanisms for antiviral activity are the following: (i) prevention of viral infection by direct binding of LF to virus particles (e.g., envelope proteins); (ii) interference with virus docking into cells by binding of LF either to sulfate proteoglycans (HSPGs), or to viral receptors of the host cells that are used by the virus for intracellular entry; (iii) inhibition of virus proliferation; (iv) intracellular activity of LF likely associated with interference with antigen synthesis of virus (e.g., during infection by rotavirus); and (v) other indirect mechanisms, such as cell cytopathy, regulatory function of LF on myelopoiesis, and cytopathy during viral infection (e.g., friend virus complex, inhibition of viral hemagglutination, like human influenza virus). Factors such as the stage of infection (i.e., duration of infection before addition of LF) (Vorland, 1999), other synergistically acting compounds, such as zidovudine (Viani et al., 1999), and the saturation of LF with different metals (e.g., Zn2+, Mn2+, Fe3+) are crucial for the antiviral outcome of LF (van der Strate et al., 2001). Finally, LF is important in immunoregulation and specifically in the first line of the host defense system, which is an interaction of neutrophils, lymphocytes, macrophages, and their secretory products. Synergism with antibodies, or the activation of a complex series of reactions, is associated with the contribution of LF to the overall protective immune response after infection or inflammation (Levay and Viljoen, 1995; Lonnerdal and Iyer, 1995; Sanchez et al., 1992; Machnicki et al., 1993; Ward et al., 2002; Zagulski et al., 1989). Specifically, when contamination with a pathogen occurs, polymorphonuclear neutrophils capture the invader (phagocytosis) and specific granules release low iron-saturated LF (6–8%) into the blood, which in turn creates an hypoferremic environment (depletion of iron), thus preventing the pathogen from acquiring sufficient iron for growth (Miyauchi et al., 1998; Sanchez et al., 1992; Ward et al., 2002). The affinity of LF with lipopolysaccharides (LPS) is also beneficial, since binding of LF to lipid A, the toxin moiety of LPS of Gram-negative pathogens, prevents the toxic shock induced by interaction of this lipid with monocytes/macrophages (Baveye et al., 1999; Caccavo et al., 2002). A review of mechanisms related to the modulation and amplification of the inflammatory process by LF that was presented by Baveye et al. (1999) listed the following mechanisms: (i) enhancement of phagocytosis by preventing the deactivation of complement factor C3; (ii) inhibition of extracellularly formed hydroxy-radicals by sequestration of free iron; (iii) enhancement of polymorphonucleocytes (PMNs) recruitment to inflammatory sites; (iv) regulation of the proliferation and differentiation of immune cells (suppression of myelopoiesis) by suppressing the production of interleukins IL-1 and IL-6 by monocytes; (v) inhibition of platelet aggregation; and (vi) promotion of the recruitment and activation of immune cells in inflammatory sites by blocking the release of other cytokines (e.g., the tumor necrosis factor alpha-TNF-α) (Crouch et al., 1992; Machnicki et al., 1993; Mattsby-Baltzer et al., 1996; Shinoda et al., 1996; van Hooijdonk et al., 2000; Vorland, 1999; Wagstrom et al., 2000 ; Zagulski et al., 1989). A number of studies reviewed by Ward et al. (2002) illustrated that inhibition of TNF-α by LF may protect human and mice from allergen-induced skin inflammation. Moreover, it is shown that LFcin and some of its acylated derivatives inhibit the lipid peroxidation in liposomes (Wakabayashi et al., 1999a). In one of the latest reviews on LF, Wakabayashi et al. (2003) used terms such as immunosuppression and immunostimulation to describe the above immunoregulatory activities of LF and LFcin or its derivatives with shorter amino acid sequences. Immunostimulation, and specifically the stimulation of phagocytic and cytotoxic properties of macrophages, was also termed the opsonic effect (Naidu, 2000). A list of major binding targets of these peptides, including LPS, heparin, DNA, glycosaminoglycan, etc., and some more specific mechanisms, such as the induction of apoptosis in monocytic and mycloid leukemic cells, or the activation of kinase CK2, commonly representative of different LFcin residues (Maekawa et al., 2002; Roy et al., 2002; Yang et al., 2002). ALF as an immobilized derivative of LF possesses identical mechanisms for antimicrobial activity, summarized into three major categories: (i) blocking of microbial adhesion; (ii) bacterial detachment; and (iii) microbial growth inhibition (Naidu, 2002). The interaction of ALF with the outer membranes of bacteria, interference with adhesin/fimbrial synthesis or colonization factors, and competition with

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bacteria in adhesion on tissue-matrix components are the properties responsible for the first two of the above mechanisms. Iron deprivation with further consequences on ATP synthesis and cellular multiplication accounts for the growth inhibitory mechanism, and the interaction of ALF with nucleic acid is likely associated with its antiviral activity.

18.3.1.2.2 Specificity LF is known to possess a wide antimicrobial spectrum including Gram-positive and Gram-negative bacteria, such as B. subtilis, E. coli O157:H7, E. coli O111, Helicobacter pylori, Klebsiella pneumoniae, L. monocytogenes, Micrococcus flavus, P. aeruginosa, S. aureus, Proteus mirabilis, and S. Typhimurium (Bellamy et al., 1993; Naidu, 2000; Viejo-Diaz et al., 2003; Wakabayashi et al., 2003), yeast of the genus Candida spp. (Kuipers et al., 2002; Samaranayake et al., 1997; Ueta et al., 2001; Wakabayashi et al., 1996, 1998), fungi, such as Rhodotorula rubra (Andersson et al., 2000), Penicillium spp. (Liceaga-Gesualdo et al., 2001), Trichophyton spp. (Wakabayashi et al., 2000), RNA and DNA, enveloped or nonenveloped viruses (Naidu, 2000; van der Strate, 2001; Vorland, 1999), and protozoa, including Toxoplasma gondii (Isamida et al., 1998), Giardia lamblia (Turchany et al., 1995), and Tritrichomonas foetus (Grab et al., 2001). For determination of the MIC and MBC of bovine and human LF as well as their hydrolysates (i.e., LFcinB and LFcinH) the standard microdilution method of Vorland et al. (1999b) in 1% peptone water has been used by the majority of researchers. In general, hydrolysates of LF (i.e., LFcins) and especially LFcinB are more inhibitory than the original LF (Naidu, 2000). It is important to note that LFcin of human origin is more active than LFcin of bovine, murine, and caprine origin (Vorland et al., 1998). According to Bellamy et al. (1992), who performed a screening on MIC of bovine and human LF as well as the corresponding LFcins against E. coli O111, the MICs of human and bovine LF were 2000 µg/ml and 3000 µg/ml (37µM), respectively; whereas the MICs of LFcinH was 100 µg/ml (ca. ten-fold lower than that of LF), and that of LFcinB was 6 µg/ml (almost 1000-fold lower than LF). In the same review, the MIC of LFcinB was found lower than the MIC of bovine LF against a variety of other microorganisms, including K. pneumoniae, L. monocytogenes, P. aeruginosa, and S. aureus. In a more recent study, Kimura et al. (2000) reported that the MIC of Korean goat LF against E. coli O111 was 5000 µg/ml (i.e., higher than that of bovine LF but lower than the MIC of sheep and horse LFs) (Lee et al., 1997). The results of this study also agree with the observed differences between the antimicrobial activity of bovine LF and its hydrolysate, since the MIC of Korean goat LFcin was only 100 µg/ml (i.e., similar to that of LFcinH and 50-fold lower than that of Korean goat LF). Ulvatne et al. (2001) demonstrated that P. mirabilis was more resistant to LFcinB than E. coli. In a study of Recio and Visser (2000) the antibacterial effects of the apo- and holo-forms of bovine, ovine, and caprine LF were comparatively evaluated in a medium containing 1% peptone and 1% glucose, against E. coli and M. flavus. Of the apo-lactoferrins, caprine LFcin had the highest effectiveness against both microorganisms, followed by bovine and then ovine LF. Holo-forms of LF were almost ineffective. Griffiths et al. (2003) comparatively investigated the effect of the apo-form and the holo-form bovine and human LF against E. coli and S. Typhimurium in coculture with probiotic bacteria, such as bifidobacteria. A conclusion of high importance for the protective role of intestinal flora was that the apoform of LF retarded the growth of enteric pathogens without affecting the growth of probiotic bacteria. This conclusion is beneficial for the potential application of LF in foods compared to antibiotics, given the deleterious effect of the latter to the intestinal flora of humans. The antimicrobial effect of several LFcinB analogs (i.e., peptides with shorter amino acid sequences) has also been considered by many research groups with varied results related to their effectiveness compared to the original LFcinB (Chapple et al., 1998; Chen et al., 2003; Hoek et al., 1997; Kang et al., 1996; Odell et al., 1996; Rekdal et al., 1999; Strøm et al., 2002a, b; Ueta et al., 2001; Wimley and White, 2000). Schibli et al. (1999) reported that the six-residue center of LFcinB is responsible for the antimicrobial activity of the whole molecule. Usually, the examined analogues are 11- or 15-residue LFcin derivatives synthesized by the incorporation of certain amino acids on residues 6 and 8 (for addition of tryptophan), 5 and 9 (for addition of arginine), or other residues (e.g., lysine) of LFcin (Chen et al., 2003; Haug and Svendsen, 2001; Kang et al., 1996; Strøm et al., 2000, 2002a, b). The antimicrobial activity of these analogues depends on the type of incorporated amino acids, the terminus of LFcin where addition of

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amino acids is made (N-terminal analogues show higher variability on antimicrobial effectiveness than C-terminus analogues), as well as the charge and the lipophilicity of the resulting analogue ( Strøm et al., 2000, 2002a). Tryptophane derivatives seem to be the most effective analogues, probably due to the high membrane affinity of this amino acid (Haug and Svendsen, 2001; Strøm et al., 2002b; Wimley and White, 2000). Wakabayashi et al. (1999b) demonstrated that N-acylated and D-enantiomer derivatives of LFcin possess increased antibacterial and antifungal activity as indicated by the higher MICs of LFcinB. The MICs of these derivatives ranged from 3 to 12 μg/ml against bacteria, such as E. coli, P. aeruginosa, S. aureus, and the fungus Trichophyton mentagrophytes, whereas significantly higher MICs, ranging from 25 to 100 μg/ml were observed for C. albicans. Vorland et al. (1999a) made a comparative test on the MIC of LFcin and its peptides synthesized by L- and D-amino acids, on spheroplasts of E. coli, P. mirabilis, and protoplasts of S. aureus as well as on the above microorganisms with intact cell walls. The D-enantiomer of LFcin was more active than the L-enantiomer. LFcin (MIC 30 μg/ml and MBC 80 μg/ml) and its D-enantiomer (MIC and MBC equal to 30 μg/ml) had the same effect on S. aureus, but the D-enantiomer with MIC and MBC equal to 5 μg/ml was far more effective than LFcin against E. coli. None of the tested peptides was effective against P. mirabilis. Spheroplasts and protoplasts were more sensitive than their counterparts, suggesting that the cell wall has a protective role against LFcin. Lower temperatures (22 and 28°C) sensitized bacteria to all tested peptides in comparison with 37°C, and similar results for the effect of the D-amino acid counterpart of LFcin were reported by Ulvatne and Vorland (2001). The antifungal properties of LF and LFcins have also been investigated. Wakabayashi et al. (1996) reviewed the antifungal spectrum and fungicidal mechanism of LFcinB. Candida species, due to their clinical significance as the causative agents of many types of candidosis, are the most extensively used organisms in studies for the investigation of the antifungal effect of LF and LFcins. Indeed, the antifungal effect of LF, LFcin, and LFcin residues alone (Xu et al., 1999) or in combination with other compounds (e.g., azole antifungal agents) (Wakabayashi et al., 1996, 1998), or in the form of sodium alginate tablets (Kuipers et al., 2002), against C. albicans (Ueta et al., 2001), and other species (e.g., C. glabrata or C. krusei) has been well-established (Samaranayake et al., 1997). An inhibitory effect has also been reported for other fungi (Wakabayashi et al., 2003), including Rhodotorula rubra (Andersson et al., 2000), spores of Penicillium sp. (Liceaga et al., 2001), and Trichophyton spp. (Wakabayashi et al., 2000). Additionally, LF and its hydrolysates have been shown in vivo (in mice) to inhibit protozoa, such as Toxoplasma gondii (Isamida et al., 1998), Giardia lamblia (Turchany et al., 1995), and tumor metastasis (Yang et al., 2002; Yoo et al., 1997). Some protozoa, however, such as Tritrichomonas foetus are reported to sequester the iron necessary for survival from the host (cattle) LF (Grab et al., 2001). Moreover, LF has antiviral activity against a wide range of human and animal RNA- and DNA-viruses, enveloped or not (Naidu, 2000; van der Strate, 2001; Vorland, 1999), such as hepatitis C virus (Ikeda et al., 1998, 2000), cytomegalovirus (Andersen et al., 2001), rotavirus (Superti et al., 1997), friend virus (Vorland, 1999), poliovirus (Marchetti et al., 1999), respiratory syncytial virus (Grover et al., 1997), human immunodeficiency virus (Swart et al., 1996; 1998; Vorland, 1999), human influenza virus (Kawasaki et al., 1993), spleen focus forming virus (Hangoc et al., 1987), feline immunodeficiency virus (Sato et al., 1996), and herpes simplex virus (Hammer et al., 2000; Hasegawa et al., 1994). In contrast, LFcin does not always demonstrate antiviral activity (Ikeda et al., 2000).

18.3.1.3 Applications in Foods LF is available in ready-to-use form such as liquid or spray-dried powder. In the past two decades, its metal-chelating property has been the primary claim for the application of LF in several infant food formulas in South-East Asian markets (Satue-Gracia et al., 2000). Nandi et al. (2002) successfully induced expression of LF in transgenic rice grains at levels up to 0.5% by linking a synthetic LF gene to rice glutelin 1 promoter for future application in infant formulas. The pharmaceutical applications of LF and its hydrolysates LFcinB and LFcinH are also well-established (Clare et al., 2000, 2003; Yamamoto et al., 2003), as well as their potential benefits in curing infected fish and seafood (Gallardo-Cigarroa et al., 2000; Kakuta, 2000; Koshio et al., 2000). Oral administration of bovine LF is reported to enhance

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the survival of rainbow trout, red sea bream, and goldfish upon infection with Aeromonas salmonicida, Cryptocaryon irritans, and Ichthyopthirius multifiliis, respectively (Kakuta, 2000). However, the application of these compounds in foods is still limited so far due to the requirement of high doses to obtain preservative effect. For instance, the susceptibility of LF to pH, elevated levels of calcium or phosphates, excess of cations (especially iron), and improper citrate/bicarbonate ratios are factors with the potential to decrease the activity of LF (Naidu, 2002). The antimicrobial activity may also be reversed by trypsin, ferrous sulfate, magnesium sulfate, and hematin (Reiter and Oram, 1967). In addition, phenomena related to the risk of denaturation, structural alterations, or even charge-induced aggregation during the isolation of LF pose further limitations in the application of lactoferrin in foods. Therefore, research has been focused on the discovery of alternatives to overcome the limitations in application of LF. Potential alternatives include: (i) activation of LF under conditions that protect its structure and minimize the negative impact of milieu conditions, thus resulting in activated lactoferrin; and, (ii) LF digestion derivatives (i.e., LFs and their analogues with smaller amino acid residues) in an effort to evaluate the potential of compounds, which are released naturally as a result of proteolysis of LF and LFcin, respectively. Investigation of the antimicrobial properties of LF and LFcins in foods is still in its infancy, with a few published studies available. A common conclusion drawn from these studies is that the activity of LF is reduced due to the excess of metal cation that saturates the peptide. Thus, the presence of other compounds with chelating properties, such as EDTA, or the dilution of cations concentrations are necessary to enhance LF activity (Branen and Davidson, 2000; Chantaysakorn and Richter, 2000; Masschalck et al., 2003). Alternatively, the combination of LFcin with other emerging technologies, such as high hydrostatic pressure, has been proven promising, especially when prevention of saturation of LFcin with ions is not feasible (Masschalck et al., 2001, 2003). LFcinB was primarily tested in ground beef at concentrations of 50 or 100 μg/ml, where it was found to cause a maximum of 2 log10 CFU/g reduction at 4 or 10°C (Venkitanarayanan et al., 1999). The positive contribution of EDTA on the antimicrobial activity of LFcin was demonstrated by Branen and Davidson (2000) against E. coli and L. monocytogenes in TSB and in a medium containing 1% peptone, 0.05% yeast extract, and 1% glucose. LFcinB at concentrations of 1600 μg/ml was unable to inhibit the growth of E. coli at 37°C, whereas the addition of 100 or 400 μg/ml EDTA, depending on the strain of E. coli, totally prevented growth. Similar results were obtained against L. monocytogenes; however, lower concentrations of LFcin were necessary for inhibition due to the higher sensitivity of L. monocytogenes compared to E. coli. However, there seems to be a dispute on whether EDTA enhances LF activity in foods. Murdock and Matthews (2002) found no antimicrobial effect by the combination of the two compounds in UHT milk acidified to pH 4.0 and stored at 37°C, even at higher concentrations of LFcin and EDTA (i.e., 4000 μg/ml and 10 mg/ml, respectively). With respect to the increase of LFcin activity as a result of reduced cation concentration, Chantaysakorn and Richter (2000) found that 5000 and 10,000 μg/ml of LFcin had no effect on E. coli in carrot juice, but significantly retarded growth when added in a filtrate of carrot juice through 500 or 10,000 Da molecular weight rejection membranes and totally inhibited growth in dialysate of the filtrate. Considering the decrease in cation concentration in the filtrate and further reduced cation concentration in the dialysate of carrot juice, there is a likely negative correlation between antimicrobial activity and presence of metal cations. Emerging preservation technologies, such as high hydrostatic pressure, when combined with LF have been proven promising in the effort to overcome the limitations in its application (Masschalck et al., 2001, 2003). High pressure (155 to 400 mPa) was reported to enhance the bactericidal activity of bovine LF and LFcinB against E. coli, Salmonella Enteritidis, S. Typhimurium, Shigella sonnei, Shigella flexneri, P. fluorescens, and S. aureus in potassium phosphate buffer at 20°C (Masschalck et al., 2001). High pressure up to 300 mPa as a single treatment caused 1 to 2 log10 reductions, whereas when applied in combination with antimicrobial peptides approximately 2 additional log reductions were sustained for all microorganisms. LFcinB caused consistently higher reductions than bovine LF, apart from the combination with 400 mPa, where bovine LF indicated higher reductions than LFcinB. The same group found that a combination of 100 or 270 mPa for 15 min with 20 μg/ml of LFcin in phosphate buffer increased reduction S. Typhimurium and P. aeruginosa by 1 to 2 log and 3 to 5 log (depending on strain) in comparison with the single application of high pressure and LFcinB, respectively. Recently Federico et al.

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(2015) and Baruzzi et al. (2015) demonstrated the significant antimicrobial effect of LFcinB to reduce pseudomonads on ready-to-eat vegetables. ALF is a novel LF derivative that has gained increasing interest as an alternative of LF, due to the limitations in application of the latter in foods. However, since ALF technology was established (Naidu, 2001), there is still limited information on the efficacy of this compound in foods. In a review by Naidu (2002), preliminary research data related to the detachment efficacy and bacteriostatic effect of ALF against E. coli in broth, beefsteaks, and fresh beef are shown. ALF and LF at a concentration of 1% were comparatively evaluated for their efficacy to detach collagen-bound E. coli and delay its growth. The ALF had a 2.7 log higher detachment efficacy than LF and caused 17.4 h more stasis on E. coli than LF according to impedimetric data in tryptic soy broth. Accordingly, MIC of ALF and LF was 62 μg/ml and >1000 μg/ml, respectively. Similar results demonstrating the better bacteriostatic performance of ALF compared to LF were obtained against 4 log contamination of E. coli on beef steaks. ALF was also tested in combination with current meat decontamination intervention strategies (cold/hot water and/or organic acids) for its efficacy to detach E. coli from fresh beef. Specifically, a set of sequential decontamination treatments (i.e., cold water for 10 s, hot water at 82°C for 30 s, and 2% lactic acid for 10 s) was applied alone or followed by spraying of 1% solution of ALF for 10 sec to detach an approximate E. coli population of 7 logs. The regular sanitizing assembly alone resulted in 72.2% detachment of E. coli per gram of beef tissue, whereas additional spraying with ALF increased the detachment efficacy to almost 100%/g of beef tissue. ALF was deposited on the meat surface as a fine mist created by electrostatic or high-pressure liquid spray-nozzles in a flow adjusted by digital controllers. ALF has also demonstrated activity against other pathogenic microorganisms, such as L. monocytogenes, Salmonella spp., and some meat spoilage organisms including Pseudomonas spp. and Klebsiella spp. Sensory evaluation of strip loins treated with ALF, stored under vacuum below 3.3°C and periodically exposed to retail display within a period of 35 days, showed that ALF might extend retail display life by 1.7–2.5 days. This conclusion was based on evaluation of lean and fat color, percentage discoloration, and overall appearance of strip loin samples treated with ALF compared to nontreated samples. Furthermore, Ransom et al. (2003) comparatively evaluated the effect of single or sequential dipping into solutions of ALF, LF, and lactic acid (2%) on the survival and growth of E. coli O157:H7, L. monocytogenes, and S. Typhimurium, inoculated pre- or post-dipping, on vacuum-packaged bologna at 10°C for 33 days, as well as on beef plates, beef carcasses adipose, and beef lean tissues stored aerobically at 12°C for 2 to 29 days. It was shown that dipping beef bologna slices into solutions of ALF after inoculation enhanced the inactivation of E. coli O157:H7, and inhibited the growth of L. monocytogenes compared to untreated slices after 33 days of vacuum-packaged storage at 10°C. Treatment with ALF before inoculation delayed the growth of S. Typhimurium, whereas post-inoculation treatment prevented growth during storage; however, dipping into LF did not affect the growth of S. Typhimurium and L. monocytogenes compared with samples dipped in water. Of the treatments evaluated, lactic acid was the most effective treatment in inhibiting or reducing pathogen populations on all products, followed by ALF applied either pre- or post-inoculation, exclusively on bologna, while LF did not appear to affect survival, or growth of the bacterial populations. The same study also showed that dipping into ALF followed by dipping into 2% lactic acid was the most effective treatment for decontaminating inoculated beef carcass adipose tissue.

18.3.1.4 Safety and Tolerance Bovine LF as a naturally occurring compound in milk has been consumed for years at levels ranging from 50 to 75 mg/day by humans of all ages (children, teens, and adults). The adverse effects of LF on iron absorption, modulation of microflora, and prevention of infection have been investigated with human studies, whereas animal studies involved acute toxicity (Nishimura, 1991), oral toxicity (Nishimura, 1997, 2000), and Ames assay (Kawai and Tanaka, 1997). No adverse effects on iron absorption, modulation of microflora, and prevention of infection are reported for daily intake of LF ranging from 0.3 to 1.0 g/kg/day for 11 days to 5 months for infants and 1.7 mg/kg/day to 60 mg/kg/day for a single-dose study for a total of 8 weeks for adults. Likewise, no adverse effects were evident in animal studies.

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The regulatory frames of LF and ALF are based on their natural occurrence in milk and consequently in beef tissue, since both of them are consumed by humans. Initially, LF was characterized as GRAS by the U.S. Food and Drug Administration (USFDA) (GRAS proposal), and DMV International (De Melkindustrie Veghel International) suggested that its use is considered safe in sports and functional foods at a level of 100 mg/product serving, estimating that the daily uptake of LF from that use is 1.0 g per person per day (GRN 000077). In a previous GRAS notice (GRN 000067) National Beef suggested the use of bovine milk lactoferrin as a GRAS compound in uncooked beef (carcasses, subprimals, and finished cuts of beef) as a component of an antimicrobial spray at concentrations up to 2%. The estimated daily uptake of LF from this use was 4.1 mg per person per day. However, allergenic and immunological data related to both GRAS notices emphasized the necessity of stating the source of LF (e.g., cow milk) in the ingredient statement of the products that contain this compound in order to prevent consumption of these products by people who are allergic to milk or its constituents. In a GRAS Notice (GRN 000130), ALF Ventures (the company that commercially produces and markets ALF) suggested that milk-derived LF can be considered GRAS as a component of an antimicrobial spray for dressed beef, which will subsequently be rinsed to reduce the residues of milk-derived LF, even for people who are allergic to milk, and, hence, disclaiming its presence in ingredients statements is no longer necessary. Milk-derived ALF was also considered GRAS (21 CFR.170.36[f]) and permitted at levels of 65.2 mg/kg of beef (Naidu et al., 2003) and, accordingly, in October of 2001, the USDA approved the use of ALF on fresh beef. However, the regulatory status of ALF and LF in other countries is not clear yet (Naidu, 2002). The European Union has issued a directive (83/417/EEC) for the use of proteins derived from milk, but LF is not currently included. Therefore, permitted use of LF in foods may only result from the consideration of LF as a milk protein (provision 79/112/EEC). Asian countries, such as Japan, South Korea, and Taiwan, have listed LF among other natural compounds in the List of Existing Food Additives; however, its use (in Taiwan) is limited to some specific nutritional foods under the condition “only for supplementing foods with an insufficient nutritional content and may be used in appropriate amounts according to the actual requirements” (Naidu, 2002).

18.3.2 Ovotransferrin Ovotransferrin (OTF, also called conalbumin) is an iron-binding monomeric glycoprotein that constitutes at least 10–12% of the total egg white solids (Beuchat and Golden, 1989; Parkinson, 1966). There are many similarities between OTF and LF and thus the following section focuses only on those characteristics unique to OTF. The isolation and purification of OTF may be accomplished using solvent fractionation and chromatographic methods (ion-exchange chromatography or metal-affinity chromatography). Structural variability in terms of molecular size, amino acid composition, and visible/ultraviolet (UV) absorption spectra of OTF is evident among different birds and/or strains (Clark et al., 1963; Itoh et al., 1979; Lush, 1961; Osuga and Feeney, 1968). Many similarities exist between all transferrin family proteins, including OTF (i.e., a total number of 680 to 700 amino acid residues subdivided in two halves, one C-lobe and one N-lobe with 35 to 40% internal homology) (Testa, 2002). Moreover, the reported homology between OTF and human LF and serum transferrin is 49% and 51%, respectively. OTF is specified by the same gene as serotransferrin, but differs from the serotransferrin of the same species only in its glycan part (Testa, 2002). Like LF, OTF reversibly binds two Fe3+ ions per molecule concomitantly with two bicarbonate anions. Its high affinity for iron (~1030 M–1) renders the latter unavailable for bacteria and, hence, inhibits their growth (Tranter and Board, 1984; Valenti et al., 1983a). Therefore, the stoichiometric balance of iron in the environment is essential to maintain the effectiveness of OTF (Conner, 1993; Valenti et al., 1983b). It is well-known that saturation of OTF with iron reduces its effectiveness against many Gram-negative bacteria in hen egg albumen; however, OTF remains effective against Gram-positive bacteria, including lysozyme-resistant strains, at 30 or 39.5°C regardless of the presence or absence of iron (Tranter and Board, 1984). The antimicrobial activity of OTF is highly dependent on milieu conditions and the target organism. Alkaline pH and elevated temperature close to the physiological temperature of birds (~40°C) enhance the antimicrobial activity of OTF (Tranter and Board, 1984).

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OTF is mainly considered to have bacteriostatic activity, although there is evidence of a biocidal effect independent of iron depletion, against a wide range of bacteria, such as E. coli, Klebsiella spp., Proteus spp., Pseudomonas spp., and S. aureus (Valenti et al., 1982, 1983a, 1984). It has been reported to extend the lag phase and reduce the growth rate of many Gram-positive and Gram-negative microorganisms, with the latter being less sensitive. Of the Gram-positive bacteria, Bacillus sp. and micrococci are the most sensitive groups (Board, 1969). An inhibitory effect against some yeasts species, such as Candida sp., has also been observed (Valenti et al., 1983a, 1985). Despite the negative impact of iron-saturation on the antimicrobial activity of OTF, the complex of OTF with other metal cations is reported to increase its antimicrobial effectiveness. Among the complexes of OTF with metal cations tested in vitro, the OTF-Zn2+ complex indicated the highest antimicrobial effectiveness (Valenti et al., 1983b). OTF is heat-sensitive and 80% of the activity is lost with heating to 70–79°C for 3 min (Banwart, 1979), or 60°C for 5 min (Tranter, 1994). However, the presence of ions, such as phosphate or citrate at pH values above 6.0 (Nakamura and Omori, 1979), or saturation with iron in the presence of an anion, increases the resistance of OTF to heat and proteolytic action and disulfide reduction by thiols (Banwart, 1979; Williams et al., 1985).

18.4 Immunoglobulins Immunoglobulins (Ig), or antibodies, are a complex, heterogeneous mixture of glycoproteins that are produced by plasma cells (lymphocytes or immunocytes) and are present on the B-cell membrane or secreted by plasma cells (Goldsby et al. 2000). Ig are the effectors of the immune system responsible for binding specific antigen molecules which are foreign to a host system to enable an immune response and clearance of the foreign substances and any associated harmful effects. Ig are able to recognize, bind, and occasionally neutralize bacteria, viruses, polysaccharides, nucleotides, peptides, and proteins (Bostwick et al., 2000). Due to the diversity and great multitude of foreign molecules or antigens (Ag) in nature, there is a need for a large number of Ig to recognize and counter all the different antigens that may exist. The great diversity of Ig molecules originates from subtle structural differences in their antigen combining sites, or variable regions, accounting for unique antigen binding specificities (Goldsby et al., 2000). The structural differences in the regions other than the antigen combining sites, constant regions, are related to the different effector functions mediated by antibodies, like complement activation or binding to one or more of the Ig receptors expressed on monocytes and granulocytes (Goldsby et al., 2000; Wilson and Stanfield, 1994). The main functions of Ig are thus, (i) Ag binding – Ig binding specifically to one or more closely related antigens via the antigenic determinant (Stanfield and Wilson, 1995); and (ii) effector functions – the Ig has no direct biological effect on the Ag and secondary “effector functions” are sequestered to aid in the clearance of the Ag and are usually preceded by Ag binding. Effector functions may include (i) complement fixation – a group of serum proteins that participate in an enzymatic cascade that generates the cytolytic membrane-attack-complex and results in lysis of cells through the release of biologically active molecules; and (ii) binding to various cell types such as phagocytic cells, lymphocytes, platelets, mast cells, and basophils that have receptors that bind Ig and activate the cells to perform a clearance function (Stanfield and Wilson, 1995). Those functions associated with lacto- and ovoglobulins mainly complement activation and Ig-augmented activity including agglutination, opsonization, adherence-blocking, and neutralization to aid in the clearance of microorganisms.

18.4.1 Lactoglobulins Colostrum, as a source for transporting immune factors including Ig from mother to newborn, has been known for more than 100 years. While colostrum remains an important source, in humans, rabbits, and rodents, passive immunity is conferred mainly through the transfer of immune factors via the uterus. In contrast, for livestock (pigs, horses, sheep, and cattle), colostrum serves as the major source of immunity (Bostwick et al., 2000; Butler, 1994; Hilpert et al., 1987; Reddy et al., 1988; Stephan et al., 1990; Yolken

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et al., 1985). The use of commercially produced and purified Ig as antimicrobials for the prevention and treatment of microbial diseases in humans was proposed by Weiner et al. (1999), and use in food systems for preservation has been suggested (Korhonen et al., 1998; Pellegrini, 2003; Zommara et al., 2002). The limiting factors for the use of Ig from animal products are the cost of purification and the need for hyperimmunization to achieve broader specificity (Bostwick et al., 2000). Ig may be derived from a number of commercial sources, including colostrum and milk from different livestock, eggs, and cell culture; however, the most practical sources are bovine milk and colostrum followed by avian eggs (Bostwick et al., 2000).

18.4.1.1 Molecular Properties 18.4.1.1.1 Occurrence Ig may be characterized according to their existence in the host, either as serum fractions or as secretory Ig, and since this chapter discusses antimicrobials in food originating from animal sources the focus will be on secretory Ig from lacteal secretions (milk and colostrum) and avian eggs. There are five general classes of Ig, namely IgG, IgA, IgM, IgE, and IgD, and another specific class, IgY, which is exclusively found in eggs (Bostwick et al., 2000). The Ig found in bovine and colostral whey include: (i) IgA; (ii) IgG with two major subclasses IgG1 (comprising approximately 75% of IgG in milk), IgG2, and IgG fragments; (iii) IgM; (iv) J-chain or component; and (v) the free secretory component (Butler, 1994). The nomenclature of Ig is based on their immunological cross-reaction with reference proteins of human origin and was proposed by the World Health Organization (Butler, 1983; Butler et al., 1971). The protein content of bovine colostrum and milk is higher than that from humans, with Ig levels in bovine colostrum higher than that of humans and, conversely, higher Ig levels in human milk than found in bovine milk (Butler, 1994; Hanson et al., 1993; Kulkarni and Pimpale, 1989). IgG accounts for the majority of the Ig fraction in colostrum and milk (ca. 75–80%) followed by IgM and IgA at similar levels (Butler, 1994; Hanson et al., 1993; Kulkarni and Pimpale, 1989). 18.4.1.1.1.1 Structure  All Ig molecules are symmetrical glycoproteins of approximately 180 kD that are monomers or polymers of four polypeptide chains comprising two identical non-glycosylated light chains (L) (each approximately 20 kD) and two identical glycosylated (with mannoses and N-acetylglucosamine) heavy chains (H) (each approximately 50–70 kD) linked together with disulfide bonds (Butler, 1974; Lascelles, 1979; Vasilov and Ploegh, 1982). The H and L chains of each molecule have a constant (C) and a variable (V) region. The V regions are composed of approximately 100 amino acids near the N-terminal while the C region makes up the remainder of the molecule toward the C-terminal (Butler, 1974; Edelman, 1973; Lascelles, 1979). The variable N-terminal regions of both the H and L chains are considered to be important in the specificity of the Ig to which it can bind, whereas the complement fixation, membrane transport, species-specific, and class-specific Ag determinants are related to the C region of the H chains (Butler, 1974; Lascelles, 1979). The primary difference between the Ig molecules found in bovine colostrum and milk is in the structure of the heavy chains and especially in the V regions (Bostwick et al., 2000).

18.4.1.1.2 Biosynthesis The genes coding for the H and L chains for the production of an Ig molecule are located on different chromosomes and there are separate genes coding for the V and C regions (Bostwick et al., 2000). The genes for the V and C regions on the same chromosome have discrete coding segments or exons separated by noncoding segments or introns, and the mRNA produced from these genes undergoes a series of splices to remove the introns which moves the V and C regions closer together and results in a functional H or L chain (Tonegawa et al., 1976). After synthesis the H and L chains are glycosylated mainly with mannose oligosaccharide (some linked to N-acetylgalactosamine) at asparagines primarily, as well as serine and threonine (Kemp et al., 1983). The assembly of the H and L chains into a complete molecule occurs when cysteine molecules form disulfide bonds, through either the formation and crosslinking of two H chains or the combination of an H and L chain (Buxbaum et al., 1971; Parkhouse, 1971). The Ig

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molecules are synthesized in a class of lymphocytes known as beta cells (B-cells) which may differentiate into plasma cells when activated and secrete Ig (Bostwick et al., 2000; Goldsby et al., 2000). The process of generating mucosal Ig that is found in bovine colostrum and milk begins with the uptake of Ag by M-cells of the Peyer’s patches in the intestine or lamina propria of the bronchi (Goldsby et al., 2000). The Ag is presented to B-cells in lymphoid tissue where it is bound preferentially by membrane-bound IgD on the B-cells, activating the B-cells and causing them to secrete IgM, which later switches to IgG (Goldsby et al., 2000). The primed B-cells may then proliferate and differentiate into plasma cells and be directed via lactation hormones to the mammary gland where they produce mucosal Ig in colostrum and milk (Goldsby et al., 2000; Swain et al., 1999).

18.4.1.1.3 Stability The stability of Ig molecules during processing is affected by thermal treatment (Dominguez et al., 1997; Li-Chan et al., 1995; Lindstrom et al., 1994), and although Ig is heat-sensitive, a substantial proportion of the Ig activity will survive pasteurization (Chen and Chang, 1998). The use of high-temperature/shorttime (HTST) pasteurization techniques results in only 10–30% loss of Ig activity, while ultrahigh temperature (UHT) processing and evaporation destroy almost all the specific immune activity (Kummer et al., 1992; Li-Chan et al., 1995). Rapid inactivation of Ig starts at temperatures exceeding 65°C, and it has been demonstrated that at 81°C most of the antiviral activity is lost (Mainer et al., 1999). Anema (2000) indicated that thermal denaturation of Ig is retarded with increasing levels of milk solid concentration. The specific antimicrobial activity of Ig appears to not be affected by storage temperature, retaining its activity up to 12 months storage at 4, 20, and 37°C (Husu et al., 1993).

18.4.1.2 Antimicrobial Activity 18.4.1.2.1 Mode of Action Bovine colostrum and milk may contain many naturally occurring antimicrobial substances including the antibody-complement system and complementary antigen-antibody binding activity (Pakkanen and Aalto, 1997; Regester et al., 1997; Reiter, 1985a). The antibody-complement system is considered to be one of the major antimicrobial activities in colostrum (Mueller et al., 1983). The antibodies or Ig absorbed from colostrum after birth in combination with the complement system have a crucial role in conferring passive immunity to a newborn calf (Butler, 1986; Staak, 1992). The Ig-mediated antimicrobial activity of complement is well-established in colostrum where all the complement components are present, while in milk the activity is not always demonstrated due to the fact that not all components are present (Eckblad et al., 1981; Husu et al., 1993; Korhonen et al., 1995; Reiter and Brock, 1975). The complement system consists of over 20 different proteins involved in an enzymatic cascade that may be activated by Ag–Ig interactions (classical pathway), by certain carbohydrates (lectin pathway), or by surfaces not protected by natural inhibitors (alternative pathway) (Korhonen et al., 2000). Complement may kill microorganisms, clear immune complexes, or induce and enhance antibody responses through lytic functions of the membrane-attack complex generated by the cascading enzymes (Fearon, 1998). The activation of the classical pathway involves binding of the first component of complement to Ag–Ig interactions or directly to microbes, while in the absence of Ig, complement activation occurs through the lectin pathway with lectins bound to pathogen surfaces or through the alternative pathway, involving the presence of bacterial cell membrane components such as lipopolysaccharides (Holmskov and Jensenius, 1996; Turner, 1996). Although Gram-positive and some Gram-negative bacteria are resistant to complement, most Gram-negative bacteria are sensitive to the lytic actions of complement and both may become inactivated due to opsonization and subsequent opsonophagocytosis (Rautemaa and Meri, 1999). Aside from complement activation, the major Ig-augmented antimicrobial activity occurs through the highly specific, reversible, noncovalent Ag–Ig interactions responsible for immobilizing target Ag as well as the sequestration of other components of the immune system to remove the Ag (Bostwick et al., 2000). The mechanisms that may be used by Ig to target an Ag include agglutination, opsonization, adherence-blockade, toxin and virus neutralization, or stasis and cidal activity. Agglutination is

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brought about by the flexibility in Ig molecules, allowing cells with common surface Ag to be crosslinked to one another, and usually involves flagella and pili and, to a lesser extent, the cell membrane. Opsonization or covering of target Ag by Ig molecules may serve to signal effector immune cells to assist in targeting, enhancing phagocytic activity, and even destroying foreign bodies. Ig may bind to specific cell surface components on microorganisms and form an adherence-blockade, preventing the microbe from binding to its host cell surface receptor. Although Ig are rarely cidal, they do tend to display stasis more often by binding to microorganisms and causing alterations or disturbances within the cells by impairing growth, multiplication, and other cellular processes until other immune effectors may aid in killing or removal of the Ag. Additionally, Ig may serve to neutralize toxins and viruses by binding to various portions of the cells or molecules and thereby preventing receptor-mediated internalization of the Ag molecules.

18.4.1.2.2 Specificity Ig molecules are one of the most effective antimicrobial agents in their ability to target a wide spectrum of pathogenic agents including bacteria, viruses, fungi, protozoa, toxins, and other proteinaceous or polysaccharide molecules due to the polyclonal nature of their composition. The two major effects on pathogenic agents involve blockage of attachment and invasiveness or immobilization followed by the destruction or removal of Ag regardless of its nature and origin.

18.4.1.3 Applications in Food Commercial whey or colostral Ig has been used for many years as a feed supplement for farm animals, mainly newborns, to combat contagious diseases, and have been proven useful particularly against diarrheal diseases (Korhonen et al., 2000). Commercial products containing milk Ig have been developed and marketed for human use in the prevention and/or treatment of microbial disease (Ebina, 1992; Reddy et al., 1988). The main sources of Ig for commercial production are bovine milk and colostrum due to their availability and safety as compared with serum-derived analogues (Korhonen et al., 2000). Hyperimmune milk or whey derived from the immunization of cows with specific antibodies has been produced and found to contain higher levels of Ig than that of non-immunized cows (Casswall et al., 1998; Cordle et al., 1991; Facon et al., 1995; Greenberg and Cello, 1996; Kelly et al., 1996; Tacket et al., 1992) and has indeed been marketed in Asian countries for many years (Casswall et al., 1998, 2000; Hilpert et al., 1987). The extent of the increase in specific activity of Ig appears to be dependent on the specific organism and agent used for vaccination (Li-Chan et al., 1995; Facon et al., 1995; Tomita et al., 1995). Li-Chan et al. (1995) found increased ELISA activity for only one of five bacterial Ags in immunized cows in comparison to the levels of activity in milk from commercial dairies. Tomita et al. (1995) found that although cows vaccinated with a lipopolysaccharide-protein conjugate derived from E. coli J5 enhanced serum antibody titer to the organism, it did not enhance whey IgG titers. Several studies have suggested that feeding hyperimmune milk increases Ig against the immunizing bacteria and may reduce disease (Boedeker et al., 1987; Chernokhvostova et al., 1990; Murosaki et al., 1991). In early studies, Stephan et al. (1990) found that Ig preparation from the first colostrum of multiple cows displayed antimicrobial activity against a number of microorganisms and when tested in situ in a human host conferred Ig against Cryptosporidium (Shield et al., 1993). Ormrod and Miller (1991) reported that Ig from the milk of dairy cows immunized with a multivariate bacterial vaccine had anti-inflammatory activity in the rat hind-paw edema assay. Kobayashi et al. (1991) and Ishida et al. (1992) showed that hyperimmune milk conferred a higher survival rate for mice given lethal doses of irradiation as compared to control mice not receiving the milk. Although bovine colostrum and milk appear to be the ideal sources of Ig for human use, they originate from a foreign species and, as such, are limited to application only against oral and gastrointestinal pathogens or for topical applications in humans (Korhonen et al., 2000). The immunoglobulin β-lactoglobulin in combination with hypericin was effective at reducing S. aureus by up to 5 logs. The complex of hypericin with β-lactoglobulin, a protein carrier that has full compatibility with dairy processes, indicates that it may be used as a disinfectant for food manufacturing and handling (Rodríguez-Amigo et al., 2015).

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18.4.2 Ovoglobulins Much like mammalian species that produce Ig in serum and lactations, domestic avian species such as chickens, turkeys, and ducks produce Ig in serum and in their eggs (Rose et al., 1974). The Ig in the avian serum is transferred to the yolks of eggs to provide the offspring with acquired immunity to avian diseases and other Ag, leaving the newly hatched chick relatively immunocompetent (Rose and Orlans, 1981). Although avian sera contains three main Ig (IgG, IgM, and IgA), the Ig found in egg yolk is referred to as IgY (Leslie and Clem, 1969) due to the different structure and immunological properties as compared with mammalian IgG (Akerstrom et al., 1985; Higgins, 1975; Jensenius et al., 1981; Kobayashi and Hirai, 1980). In comparison with mammalian IgG, IgY is much larger (Kobayashi and Hirai, 1980), more acidic, less rigid, and does not fix complement (Higgins, 1975).

18.4.2.1 Molecular Properties 18.4.2.1.1 Occurrence and Biosynthesis Avian blood contains at least three distinguishable kinds of Ig, including IgG, IgM, and IgA, of which IgG comprises approximately 75% of the total Ig (Leslie and Martin, 1973). IgG (known as IgY in egg yolk) is transferred from the maternal serum to the egg yolk and subsequently to the circulation of the chick through the endoderm of the yolk sac, while IgM and IgA are secreted into the ripening egg follicle and incorporated into the egg via the oviduct (Locken and Roth, 1983; Patterson et al., 1962). Subsequently, the transfer of Ig to the embryonic gut occurs when amniotic fluid is swallowed and thereafter may provide passive immunity to the newly hatched chick (Losch et al., 1986). Hens lay on average of 240 eggs per annum, and the associated production of IgY totals 24 g, which is relatively high in comparison with certain mammals such as rabbits, mice, and goats (Hatta et al., 1997; Leslie and Clem, 1969). In a study comparing two species of chickens, it was determined that although the ratio of yolk weight to egg white was similar in both types of chickens and the total content of IgY in the yolk was relatively constant during 18 weeks, the level was influenced by hen species, egg weight, and egg production per day (Li et al., 1998).

18.4.2.1.2 Chemistry and Structure IgY, much like the Ig, found in serum consists of two heavy (H) chains and two light (L) chains and has a molecular weight of approximately 180 kDa (Parvari et al., 1988). Unlike the H chains of mammalian IgG which have three constant (C) regions and a hinge region, IgY has four C regions and no hinge (Parvari et al., 1988). The content of β-sheet structure in the constant region of IgY is lower than that of mammalian IgG, and the flexibility of the region corresponding to the hinge region in IgG was less than that of IgG (Ohta et al., 1991; Shimizu et al., 1992). A lack of disulfide linkage in the IgY L-chain, lower flexibility in the hinge region, and other structural properties (molecular size, intramolecular bonding, and domain conformation) may all influence the lower molecular stability of IgY as compared with IgG (Shimizu et al., 1992).

18.4.2.1.3 Stability The stability of IgY was compared to that of mammalian IgG when exposed to adverse pH and temperature environments (Hatta et al., 1993; Otani et al., 1991; Shimizu et al., 1988, 1992, 1993). In doing so, it was determined that above 70°C, IgY was more sensitive to the effects of heat than IgG, and the maximum temperature of denaturation for IgY was 73.9°C as compared with 77°C for IgG (Hatta et al., 1993). Shimizu et al. (1988, 1992, 1993) found that heating for 15 min at 65°C or higher, especially at temperatures above 75°C, reduced the activity of IgY. Otani et al. (1991) reported that at pH 2 and 3, IgY activity was more sensitive to the acidic conditions than IgG. Similarly, it was found that the activity of IgY was decreased at pH values below 3.5 and almost lost at values of 3.0, while at the opposite extreme, alkaline conditions up to pH 11 did not affect activity whereas incubation at pH 12 and above resulted in a significant decrease (Shimizu et al., 1988, 1992, 1993). It is well-established that although IgY is sensitive to pepsin digestion, it is relatively resistant to digestion by trypsin or chymotrypsin and that overall

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it is more sensitive to digestion by all three enzymes in comparison to IgG (Hatta et al., 1993; Otani et al., 1991; Shimizu et al., 1993).

18.4.2.2 Antimicrobial Activity 18.4.2.2.1 Mode of Action The essential mechanism for Ig to clear foreign material including pathogenic microbes is related to the specific binding to complementary Ag. The Ag–Ig interaction is often referred to as the “lock-andkey” mechanism for the specific noncovalent interactions including hydrogen bonds, hydrophobic bonds, coulombic interactions, and van der Waals forces (Sim et al., 2000). The target Ag may be composed of amino acids, nucleic acids, carbohydrates, or lipids, and it is the strength of association and complementarity which determines the overall binding strength (Sim et al., 2000). IgY has mostly been documented for its antiviral and antibacterial activity achieved by the exclusion of these pathogenic agents from infection of host cells. In order for infection to develop in a host, the etiologic agent needs to form an association with the host cells either by contact or by subsequent internalization. Viruses express surface receptor molecules specific for the host cell membrane to initiate and assist in internalization of the infective material and, thus, the potential for preventing this is the ability of Ig to specifically bind to the viral receptor and block infection, a process termed viral neutralization (Goldsby et al., 2000). Bacterial pathogens are similar to viruses in that most bacteria need to form an adherence with the host cell before the infection process is initiated and the adhesion to the host cell usually triggers invasive mechanisms of the bacteria or toxin production, which may be internalized to cause infection. It was demonstrated that anti-fimbriae Ig prevented the attachment of bacteria to piglet intestinal epithelia by blocking the mucosal receptor, interfering with binding to mucins, or neutralizing the colonization factor (Imberechts et al., 1997; Jin et al., 1998; Wanke et al., 1990).

18.4.2.2.2 Specificity IgY has been shown to be specific against infectious pathogens of bacterial or viral origin. Specific IgY activity against pathogenic agents has been elucidated from in vitro studies in hens immunized with pathogens and through passive immunization of animals with specific IgY (Sim et al., 2000). Antibacterial effects involving the use of specific IgY in vivo or in vitro have included use of E. coli, Edwardsiella tarda, P. aeruginosa, Salmonella spp., Streptococcus mutans, and S. aureus (Akita et al., 1998; Hamada et al., 1991; Hatta et al., 1994, 1997; Ikemori et al., 1992; Jin et al., 1998; Ozpinar et al., 1996; O’Farrelly et al., 1992; Wiedemann et al., 1991; Yokoyama et al., 1992; Yoshiko et al., 1996). Antiviral effects involving the use of specific IgY in vivo or in vitro have included the use of rotavirus, coronavirus, and IBD-virus (Ebina, 1996; Eterradossi et al., 1997; Ikemori et al., 1997; Kuroki et al., 1993, 1994, 1997).

18.4.2.3 Applications in Food IgY may find application as a microstatic agent in preventing the growth of pathogenic bacteria using specific polyclonal Ig to effectively reduce or neutralize bacterial proliferation in food products and especially meat, thereby reducing the food safety risk associated with these products (Sim et al., 2000). It was suggested that IgY could be used as an ingredient for foods or even mouthwashes to prevent the colonization of invading microorganisms (Hatta et al., 1997). Furthermore, IgY may be used as a food adjuvant to control bacterial growth and prevent attachment of microorganisms to the intestinal epithelium (Sim et al., 2000). Akita and Nakai (1993) proposed the use of IgY in infants under 6 months as the reduced acidity of infant gastric acid would allow the molecule to maintain its protective effect against peptic digestion. However, numerous studies have documented the role of colostrum and milk in conferring protection to newborns (Jason et al., 1984), and others examined the potential of nonmaternal Ig for passive immunization of the GI tract (Hatta et al., 1997; Yokoyama et al., 1992). Passive immunization may be the most valuable application of Ig and involves the introduction of pathogen-specific IgY into a host to combat infectious diseases (Hatta et al., 1997). However, for passive immunization to be implemented,

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large amounts of Ig would need to be administered to an individual, and this may be achieved through commercial-scale production from eggs laid by hens (Hatta et al., 1997).

18.5 Avidin Avidin is a 66-kDa positively charged glycoprotein isolated from various avian egg whites and egg jelly of invertebrates (Green 1964; Gyorgy et al., 1942). Avidin received much attention due to its anti-nutritive effects (Green, 1975); however, subsequent interest has revolved around its ability to bind up to four biotin molecules and form stable complexes (Green, 1964, 1975). The interaction between avidin and biotin is the strongest known protein–ligand binding found in nature (Boas, 1927). The discovery of an antibacterial molecule, streptavidin, produced by Streptomyces spp. (a 60-kDa avidin analogue) yielded a similar primary structure compared to avidin and indeed confirmed early suspicions that avidin possesses antimicrobial properties (Bayer and Wilchek, 1990).

18.5.1 Molecular Properties 18.5.1.1 Occurrence and Biosynthesis Avidin is a minor protein in avian albumen accounting for only 0.05% of the egg white (Mine, 2000). Avidin production occurs in the goblet cells of the epithelium of the oviduct exclusively in laying hens, indicating that the production is regulated by functioning of the ovaries and specifically the hormone progesterone, and several steroids (Elo and Korpela, 1984; Mine, 2000).

18.5.1.2 Chemistry and Structure Avidin is a basic glycoprotein consisting of four identical subunits each with an approximate molecular weight of 16 kDa (approximately 66 kDa for the molecule), and has a pI of approximately 10 (Green, 1975; Woolley and Longsworth, 1942). Each subunit consists of 128 amino acid residues and a single intramolecular disulfide bond between cystine residues at positions 4 and 83 (DeLange and Huang, 1971). Each subunit is organized in an eight-stranded antiparallel orthogonal β-barrel with extended loop regions providing the biotin-binding pocket (Livnah et al., 1993; Pugliese et al., 1994). Avidin binds one biotin molecule to each of the four comprising subunits, specifically in the highly complementary polar, protein-core pocket, and in the absence of biotin the site may be partially occupied by a water molecule (Gitlin et al., 1988; Pugliese et al., 1994). The structure of avidin reveals the existence of several surface-exposed lysine and arginine residues, which may contribute to the basic nature of the molecule (Hendrickson et al., 1989). Avidin possesses a carbohydrate moiety (about 10% of the total molecular weight) that is comprised of a single oligosaccharide chain with four or five mannose units and three N-acetylglucosamine residues linked to Asp-17 of each polypeptide subunit (Kett et al., 2003).

18.5.1.3 Stability Avidin may be resistant to treatment with iodine applied at neutral pH, acetylation of the amino groups, and esterification of the carboxyl groups; however, inactivation of the molecule may result from oxidation with H2O2 in the presence of Fe2+ or treatment with formaldehyde in the presence of alanine or hydroxylamine at 50°C (Fraenkel-Conrat and Fraenkel-Conrat, 1952). Although avidin is relatively stable over a wide range of pH and temperature in regards to its biotin-binding ability, all four tryptophans associated with the molecule are rapidly oxidized at pH 4.0 and biotin-binding activity is lost when at least two are destroyed (Green, 1963). Avidin may also lose stability at low ionic strength, with 0.1M HCl, 0.1 M sodium-dodecyl-sulfate, and 6 M guanidine HCl resulting in dissociation of avidin subunits and loss of biotin-binding ability (Green, 1963). In addition, it has been shown that avidin may be denatured at temperatures exceeding 70°C; however, the avidin–biotin complex offers stabilization to the molecule up to 100°C (Gitlin et al., 1988; Pugliese et al., 1994).

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18.5.2 Antimicrobial Activity 18.5.2.1 Mode of Action Although the antimicrobial activity of avidin has not been established, it has been suggested that the compound is involved in antimicrobial responses based on the production of streptavidin by Streptomyces in the derivation of an antibiotic system and supported by initiation of avidin production at the site of tissue injury in chickens (Elo and Korpela, 1984). It is proposed (Elo et al., 1978; Korpela, 1984) that avidin production and its secretion by macrophages is induced during inflammation and cellular damage and as such may constitute a host-defense factor for bacterial and viral infection. Studies (Miller and Tauig, 1964) indicating increased amounts of avidin in chicken tissues after intraperitoneal and intravenous administration of E. coli support the view that avidin is directed toward combating microbial infection. Korpela (1984) revealed that bacterial binding of avidin was independent of the saturation of its biotinbinding site and in E. coli the receptor was the porin protein of the outer membrane. Porins extend through the outer membrane of Gram-negative bacteria and are indeed among the most abundant proteins of the cell, constituting an integral part of the structure and function, and since no porins or similar analogues exist in Gram-positive bacteria it is assumed that porin is the only avidin-binding component of the cell envelope (Lugtenberg and van Alphen, 1983). It has been hypothesized (Campbell et al., 1972; Eakin et al., 1941; Herts, 1946) that due to the high affinity avidin has for biotin, it may function as an antimicrobial by rendering it unavailable to biotin-requiring microbes, and indeed it has been shown to inhibit the in vitro growth of biotin-requiring yeasts and bacteria (Green, 1975). Furthermore, binding of extracellular/intracellular biotin may even render it unavailable to the animal cell, thereby resulting in decreased activity of biotin enzymes and altered cellular metabolism and growth (Messmer and Young, 1977; Miller and Tauig, 1964; Wood and Barden, 1977).

18.5.2.2 Specificity Although there is very little documentation of the inhibitory effects of avidin on pathogenic bacteria, Korpela (1984) reported on the in vitro binding of avidin to various Gram-negative and -positive bacteria. The Gram-negative bacteria found to bind avidin were E. coli, K. pneumoniae, P. aeruginosa, and Serratia marcescens, while the Gram-positive bacteria that bound avidin were S. aureus and S. epidermis.

18.5.3 Applications in Food Since little is known about the antimicrobial activity of avidin, there are no current applications of this glycoprotein in food as an antimicrobial. However, avidin is used in the avidin-biotin system as a diagnostic tool in immunoassays, and there is potential for the application of avidin as an antimicrobial compound in foods (Mine, 2000).

18.6 Lactolipids Lipids, especially fatty acids, isolated from milk and other dairy products possess both nutritional and protective qualities (Kabara, 1978). Detail on the antimicrobial activity of lipids and their esters is covered in Chapter 11 (Medium-Chain Fatty Acids (>C8) and Monoesters). Lipids were identified to possess antimicrobial activity in the mid nineteenth century when it was evidenced that fatty acids generated during the ripening of cheese prevented neurotoxin production by C. botulinum (Grecz et al., 1959). In neonates, the antimicrobial activity conferred by milk lipids is due to the release of antimicrobial fatty acids and monoglycerides from milk triglycerides present in fat globules, which constitute 98% of milk fat (Hamosh, 1991, 1995; Jensen, 1995). Other lipids of animal origin that have provided evidence of antimicrobial activity include epidermis-derived skin lipids shown to inactivate S. aureus (Bibel et al., 1989; Miller et al., 1988), free fatty acids at mucosal surfaces shown to inactivate pneumococci (Coonrod, 1987), and

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porcine intestinal lipids shown to inactivate Clostridium perfringens (Fuller and Moore, 1967). Lipids may serve to inhibit the establishment, multiplication, and proliferation of pathogenic microorganisms in a host, and their effects may be enhanced by interaction with other antimicrobial factors present in milk (e.g., lactoferrin, lactoperoxidase, lactoglobulins) (Mandel and Ellison, 1985; Watanabe et al., 1984).

18.6.1 Molecular Properties 18.6.1.1 Occurrence and Biosynthesis Fatty acid biosynthesis occurs after a meal when the body is energy rich (Drackley, 2000). The events after consumption involve the generation of adenosine triphosphate (ATP) by glycolysis, generation of NADPH (the reduced state of nicotinamide adenine dinucleotide phosphate) by the pentose phosphate pathway, and the storage of glucose as glycogen. Any excess glucose is converted to fatty acids and stored as triacylglycerols (Drackley, 2000). The majority of fatty acid biosynthesis occurs in the cytosol of the liver, although there is limited biosynthesis in other tissues, including adipose tissue, the brain, and mammary glands (Drackley, 2000; Jensen, 1996). The antimicrobial activity in milk is contained mainly in long-chain unsaturated fatty acids and the medium-chain saturated fatty acids (Lampe and Isaacs, 2000). The presence of fatty acids in milk originates from the diet, the mobilization of stored fatty acids, and synthesis in the mammary glands (Jensen, 1996). The diet of lactating animals affects the proportion of fatty acids present in the milk, as a high-energy meal will increase production of medium-chain fatty acids and thereby promote the antimicrobial properties of the milk (Francois et al., 1998; Spear et al., 1992).

18.6.1.2 Chemistry and Structure Fatty acid molecules are best defined by a description of the length of the carbon chain (number of carbon molecules) and the number of double bonds present and their exact positioning (Drackley, 2000). Common fatty acids are straight-chain compounds and usually possess an even number of carbon atoms (Drackley, 2000). Fatty acid chain-lengths are classed into three categories based on their length, namely short-chain fatty acids with a chain length between 2 and 4, medium-chain fatty acids with between 6 and 10 molecules, and long-chain fatty acids commonly with 12 to 24 carbon molecules but potentially up to 80 (Garrett and Grisham, 1999). The simplest fatty acids have no unsaturated linkages and cannot be modified by hydrogenation or halogenation; these are referred to as saturated fatty acids (Garrett and Grisham, 1999). The presence of double bonds between carbon molecules defines the chains as unsaturated and specifically monounsaturated if only one double bond is present and polyunsaturated if two or more double bonds are present (Garrett and Grisham, 1999). Alternatively, fatty acids in animals may possess branched chains or contain a variety of other functional groups including acetylenic bonds, epoxy-, hydroxy-, or keto-groups, and even ring structures (Garrett and Grisham, 1999). Monoglycerides or monoacylglycerols are fatty acid monoesters of glycerol and are found in very low amounts in cell extracts as an intermediate product in the degradation of triacylglycerides or diacylglycerides during lipolysis (Garrett and Grisham, 1999). Monoglycerides are the most polar components of simple lipids since they possess only one hydrocarbon chain and two alcohol groups and, thus, need careful manipulation to prevent their loss in hydrophilic solutions and on chromatographic columns (Ledoux et al., 2000; Precht et al., 2001). Furthermore, monoglycerides possess detergent properties and as such they easily form micelles in water solutions (Garrett and Grisham, 1999).

18.6.1.3 Stability The presence of proteins, especially albumin, may reduce the antimicrobial activity of fatty acids through specific and nonspecific binding (Shibasaki and Kato, 1978). The antimicrobial activity of specifically unsaturated fatty acids may also be reduced by the presence of other surface-active agents like cholesterol (Ammon, 1985; Kabara, 1978). Since the antimicrobial activity of short-chain fatty acids is due to the undissociated form rather than the anionic form, their activity is highly pH-dependent, with activity

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decreasing as the degree of dissociation increases (Kabara, 1978; Lundblad and Seng, 1991). The effect pH has on antimicrobial activity is observed mainly with short- and medium-chain fatty acids where the minimum inhibitory concentration of short-chain fatty acids increases while that of medium-chain acids decreases with corresponding increases in pH (Kabara, 1978). The activity of long-chain unsaturated fatty acids as well as monoglyceride esters and ethers of short-chain fatty acids appear to be unaffected by pH alterations (Kabara, 1978).

18.6.2 Antimicrobial Activity 18.6.2.1 Mode of Action The antimicrobial products produced by the hydrolysis of milk triglycerides are fatty acids and monoglycerides (Thormar et al., 1987). Milk lipids are mainly antiviral, and it has been shown that short- and long-chain saturated fatty acids have minimal antiviral activity, while medium-chain saturated and longchain unsaturated fatty acids are strongly antiviral (Thormar et al., 1987). Monoglycerides of fatty acids produced by hydrolysis of milk lipids are also effective at inactivating enveloped viruses at five to ten times lower concentrations than their corresponding fatty acid (Thormar et al., 1987). In addition, the antiviral activity of fatty acids and monoglycerides is additive (Isaacs and Thormar, 1990). Isaacs and Thormar (1990) demonstrated that the virus envelope is the target for lipid-dependent viral inactivation. Furthermore, research (Noseda et al., 1989; Verdonck and van Heugten, 1997) indicates that antimicrobial lipids destabilize the membrane of cells. The antimicrobial activity of lipids through membrane destabilization has also been demonstrated against fungi and bacteria (Kabara, 1978; Shibasaki and Kato, 1978). The mechanisms by which lipids target microorganisms may be through disruption of the bacterial cell wall or membrane or envelope in viruses, blockage of receptor–ligand interactions, inhibition of intracellular replication, or by inhibition of an intracellular target (Lampe and Isaacs, 2000).

18.6.2.2 Specificity The antimicrobial activity of milk lipids has been demonstrated on a variety of enveloped viruses including herpes simplex virus, influenza virus, respiratory syncytial virus, measles virus, vesicular stomatitis virus, visna virus, mouse mammary tumor virus, dengue virus types 1–4, cytomegalovirus, Semliki forest virus, Japanese B encephalitis virus, and human immunodeficiency virus (Lampe and Isaacs, 2000). Milk lipids have also been shown to inactivate Gram-positive bacteria including B. cereus, B. subtilis, C. botulinum, Corynebacterium spp., L. monocytogenes, Micrococcus spp., Pneumococcus spp., S. aureus, S. epidermis, Streptococcus spp., and Gram-negative bacteria including C. trachomatis, E. coli, N. gonorrhoeae, P. aeruginosa, and S. Enteriditis (Hernell et al., 1986; Isaacs et al., 1990; Kato and Shibasaki, 1975; Lampe et al., 1998; Qu et al., 1996; Reiner et al., 1986; Rohrer et al., 1986; Rabe et al., 1997; Wang and Johnson, 1992, 1997; Wang et al., 1993). In addition, milk lipids display antimicrobial activity against fungi including Aspergillus niger and Trichoderma viride, yeasts, including Alternaria spp., C. albicans, C. utilis, Cladosporium spp., Kluveromyces marxianus, and S. cerevisiae, and the protozoal pathogen G. lamblia (Isaacs et al., 1990; Kato and Shibasaki, 1975; Marshall and Bullerman, 1986; Sofos et al., 1998).

18.6.3 Applications in Food The antimicrobial activity of lipids has been used in food preservation for decades (Grecz et al., 1959; Shibasaki and Kato, 1978). The majority of the lipids isolated from animal products such as milk are considered as GRAS chemicals and can as such find application as antimicrobials in food products (Lampe and Isaacs, 2000). Monoacylglycerols have increased the shelf-life of various foods including soy sauce, miso, sausage, Worcestershire sauce, sponge cake, and noodles (Shibasaki, 1982; Sofos et al., 1998). In addition to these foods, the lauric acid ester of monoacylglycerol has shown antimicrobial potential in seafood salad, Camembert cheese, and various flesh foods including deboned chicken meat, minced fish, refrigerated beef roasts, and turkey frankfurter slurries (Baker et al., 1985; Hall and Mauer,

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1986; Unda et al., 1991; Wang and Johnson, 1997). Monoacylglycerols may also be combined with other antimicrobials (e.g., nisin) to increase their effectiveness in foods (Sofos et al., 1998). Considering that monoacylglycerols lower the heat resistance of certain bacteria and fungi, they may find application in decreasing the heat-treatment requirements of some foods (Kimsey et al., 1981; Sofos et al., 1998). A proposed application of antimicrobial lipids has been the use in infant formulas to provide protection after hydrolysis of the triglycerides into antimicrobial fatty acids and monoglycerides in the gastrointestinal tract following consumption (Isaacs et al., 1995).

18.7 Defensins Defensins are a group of antimicrobial peptides folded in a characteristic β-sheet and a framework of six disulfide-linked cysteines (Ganz, 2003; Selsted et al., 1985). Defensins are widely distributed in nature and indeed in mammalian epithelial cells and leukocytes, often at high concentrations, and have a broad spectrum of antimicrobial activity (Ganz, 2003; Selsted et al., 1985; Tiwari et al., 2009). Some of the key defensins conferring antimicrobial activity are protamine and magainin (Humblot et al., 2009). Protamine and magainin have been shown to be effective against bacteria and fungi (Humblot et al., 2009).

18.7.1 Molecular Properties 18.7.1.1 Occurrence and Biosynthesis Defensin peptides have been found in all mammals examined, in chickens, and in turkeys, and are abundant in cells and tissues active in host defense against microorganisms (Brockus et al., 1998; Harwig et al., 1994; Zhao et al., 2001). The highest (>10 mg/ml) concentration of defensins is usually in the granules or storage organelles of leukocytes (Ganz, 2003). Upon ingestion of microorganisms by leukocytes into phagocytic vacuoles, the vacuoles fuse with the granules and the granule material is delivered onto the target organism (Ganz, 2003). Another site containing high (>10 mg/ml) concentrations of defensins are the Paneth cells, specialized cells of the small intestine, that contain secretory granules which are released into intestinal pits or crypts (Ganz, 2003; Harder et al., 1997). Other cells containing defensins at lower concentrations (10–100 μg/ml) include barrier and secretory epithelial cells, which may produce defensins constitutively or only upon infection, and to a lesser extent cells of the immune system such as monocytes, macrophages, and lymphocytes (Agerberth et al., 2000; Harder et al., 1997; Ryan et al., 1998).

18.7.1.2 Chemistry and Structure There are two main defensin subfamilies, α- and β-defensins, that differ in peptide segment length between the six cysteine residues and the cysteine pairs connected by disulfide bonds (Hill et al., 1991; Hoover et al., 2000; Sawai et al., 2001; Zhang et al., 1992; Zimmermann et al., 1995). Defensins exist as a triple-strand β-sheet that have the distinctive fold brought about by the disulfide bonds between cysteine residues (Hoover et al., 2000; Sawai et al., 2001). The amino acid sequence and composition of defensins are highly variable, however, the cysteine framework is conserved in each defensin subfamily (Ganz, 2003). Most α- and β-defensins possess clusters of positively charged amino acids, although their distribution in the molecule is variable (Ganz, 2003). Subcellular storage organelles such as leukocytes and Paneth cells are rich in negatively charged glycosaminoglycans (Parmley et al., 1986). The majority of α- and β-defensins from leukocytes and Paneth cells contain arginine as the main cationic amino acid, while β-defensins secreted from epithelial cells contain similar amounts of arginine and lysine (Fromm et al., 1995; Kostoulas et al., 1997). The general synthesis of α-defensins occurs through the encoding of tripartite pre-propeptide sequences to produce an amino acid precursor consisting of 90–100 amino acids with an amino (N)-terminal signal sequence (approximately 19 amino acids), an anionic pro-piece (approximately 45 amino acids), and a carboxy (C)-terminal mature cationic defensin

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(approximately 30 amino acids) (Ganz, 2003). The synthesis of β-defensin precursors is similar, consisting of a signal sequence at the N-terminus, a short or absent pro-piece, and the mature defensin peptide at the C-terminus; however, it lacks the anionic pro-piece which is prominent in the α-defensin precursor (Ganz, 2003).

18.7.2 Antimicrobial Activity Defensins are mainly antibacterial and antimycotic, especially at low ionic strength conditions, low concentrations of divalent cations, plasma proteins, and other interfering substances (Ganz, 2003; Lehrer and Ganz, 2002; Selsted et al., 1993). Under these optimal conditions, defensins may be active at very low (1–10 μg/ml) concentrations (Ganz, 2003). Defensins have also been shown to be effective against some enveloped viruses (Daher et al., 1986). The main mechanism responsible for antimicrobial activity is attributed to the permeabilization of target membranes, and subsequent cell leakage, inhibition of RNA, DNA, and protein synthesis, and decreased cellular viability (Ganz, 2003).

18.7.3 Applications in Food Although defensins are antimicrobial peptides that may be isolated and purified from animals, there is, other than the antimicrobial activities as a natural ingredient in raw food products, no known application of the peptides as an additive in foods.

18.8 Chitosan There is a significant amount of literature referencing the use of animal-derived peptides with antimicrobial properties; however there also are a number of polysaccharides and lipids from animals that also show antimicrobial effects. Chitosan [(1-4)-2-amino-2-deoxy-β-D-glucan] comprises a series of polymers with differing ratios of glucosamine and N-acetylglucosamine. Chitosan is a marine polysaccharide derived from alkaline-hydrolysis of the chitin in the exoskeleton of crustaceans and arthropods. It is also a component of fungal cell walls. Chitosan is polycationic and has been shown to have antifungal (Ben-Shalom et al., 2003) and antibacterial (Fernandes et al., 2008) properties. Chitosan inhibits the growth of foodborne bacteria and fungi including lactic acid bacteria, Bacillus cereus, Staphylococcus aureus, Listeria monocytogenes, Salmonella Typhimurium, Escherichia coli, Shigella dysenteriae, Byssochlamys spp., and Zygosaccharomyces bailii (Yang et al., 2005; Chung et al., 2011; Davidson et al., 2013). Minimum inhibitory concentrations for bacteria and yeasts vary widely depending on the molecular weight of the polymer, degree of acetylation, pH, temperature, and interfering compounds (e.g., proteins and lipids). The antimicrobial mode of action of chitosan is not completely understood, although the polycationic nature of the polymer, the spacing between moieties, and the hydrophilicity have all been identified as having significant impact on the overall activity of the molecule and as contributors to the mechanism of microbial inhibition (Kong et al. 2010). Even though the mode of action is not completely understood, the two suggested mechanisms are (i) the binding of cationic chitosan to sialic acid in phospholipids, consequently restraining the movement of microbiological substances (Chatterjee et al., 2005), and (ii) penetration of oligomeric chitosan into the cells of microorganisms and prevention of the growth of cells by preventing the transformation of DNA into RNA (Sashiwa and Aiba, 2004). The use of chitosan alone in food safety is somewhat limited due to its low solubility at neutral and alkaline pH and interference of the food matrix with the ionic interactions required to inhibit the bacterial cell (Hu and Gänzle, 2019). However it has been extensively studied for its use as a biodegradable film having antimicrobial properties (Sundaram et al., 2016, Torlak and Sert, 2013. The chitosan matrix can be formed into films, gels, fibers, beads, and nanoparticles (Dutta et al., 2012). Edible films cast from chitosans also have been infused with other antimicrobials to enhance the inhibition of microorganisms and improve applicability. For example, incorporating lysozyme in a chitosan film resulted in significant inhibition of Escherichia coli O157:H7 and Listeria monocytogenes, and lowered numbers of both

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pathogens in broth. Chitosan films with added essential oil of oregano reduced viable L. monocytogenes and E. coli O157:H7 on inoculated bologna samples stored for 5 d at 10°C (Zivanovic et al. 2005). Park et al. (2005) found that chitosan-based coatings infused with potassium sorbate applied to strawberries produced significant reductions in Rhizopus and Cladosporium spp. as compared with controls. A chitosan coating with 0.1% lauric arginate, 0.1% EDTA, and 1% cinnamon oil was effective for inactivating >3 log CFU/cm2 Escherichia coli O157:H7 and Listeria monocytogenes immediately after coating and reduced Salmonella enterica to below the detection limit on cantaloupes during a 14-day storage (Ma et al., 2016). This coating also reduced total molds and yeasts to the detection limit. Chitosan with nisin incorporated has been shown to have antimicrobial activity against several Gram-positive pathogenic bacteria (see Chapter 8, “Nisin”). Chitosan can potentiate the efficacy of other limiting technologies such as high pressure and heat, thereby creating multiple hurdles (Kumar et al., 2009). Food quality can also be improved by chitosan outside of antimicrobial activity in the retardation of lipid oxidation, melanosis, and plant metabolism even if antimicrobial activity is limited (Hu and Gänzle, 2018).

18.9 Other Antimicrobials of Animal Origin 18.9.1 Pleurocidin As discussed in this chapter, there are a number of antimicrobial peptides of animal origin that can act quickly to destroy the cellular lipid bilayer membranes even of fast-growing microorganisms such as Gram-negative and Gram-positive bacteria (Tiwari et al., 2009) while also exhibiting antifungal and antiviral activities (Aires et al., 2009). Indeed pleurocidin has been shown to effectively combat a number of bacteria and fungi including E. coli O157:H7, L. monocytogenes, P. expansum, S. cerevisiae, and V. parahemolyticus (Burrowes et al., 2004). Pleurocidin is found in the skin mucus membrane of the winter flounder (Pleuronectes americanus).

18.9.2 Casocidin Much like the antimicrobial peptides (i.e., lactoperoxidase, lactoferricin B) mentioned earlier in this chapter, casocidin is another antimicrobial peptide found in milk. Casocidin is produced by the hydrolysis of S2-casein via the enzyme chymosin found in rennet. Antibacterial activity has been demonstrated against Staphylococcus spp., Sarcina spp., B. subtilis, Diplococcus pneumoniae, and Streptococcus pyogenes (Szwajkowska et al., 2011). Peptides such as casein A and casein B have been shown to inhibit Cronobacter sakazakii which has possible market application in milk-based formulations for neonates (Norberg et al., 2011). Furthermore, peptides from S2-casein, S1-casein, and k-casein have demonstrated antibacterial effects against E. coli and B. subtilis (Elbarbary et al., 2012).

18.9.3 Lysozyme (Consult Chapter 14, “Lysozyme” for More Information) As discussed previously, lactoferrin, lactoferricin B, lactoglobulins, and lysozyme together have demonstrated synergistic effects against bacteria and fungi. Lysozyme, however, when used alone has demonstrated strong bacteriolytic effects against food-spoilage organisms (Potter et al., 2005). Lysozyme targets primarily Gram-positive bacteria as the cell wall consists of peptidoglycans (Tiwari et al., 2009), while Gram-negative bacteria have demonstrated some resistance due to their outer membrane that acts as a physical barrier. The use of surfactants and chelating agents such as EDTA has proven to assist lysozyme in antibacterial effects targeting Gram-negative bacteria due to their action disruption of the outer membrane (Branen and Davidson, 2004).

18.9.4 Lipids There is very little work done on elucidating the antimicrobial properties of animal-derived lipids. Research has shown that lipids present in milk have mild antibacterial and antifungal effects (Shin et

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al., 2007). The combination of purified monoglycerides and fatty acids, especially those found in milk, has demonstrated almost twice the antibacterial effect than when applied alone (Isaacs, 2001). Specific animal lipids that have shown promise as antibacterial agents are eicosapentaenoic and docosahexaenoic acid (Shin et al., 2007).

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Yolken, R.H., Losonsky, G.A., Vonderfect, S., Leister, F., and Wee, S.-B. 1985. Antibody to human rotavirus in cow’s milk. N. Eng. J. Med. 312:605–610. Yoo, Y.-C., Watanabe, S., Watanabe, R., Hata, K., Shimazaki, K.-I., and Azuma, I. 1997. Bovine lactoferrin and lactoferricin, a peptide derived from bovine lactoferrin, inhibit tumor metastasis in mice. Jpn. J. Cancer Res. 88:184–190. Yoshiko, S.K., Shibata, K., Yun, S.S., Yukiko, H.K., Yamaguchi, K., and Kumagai, S. 1996. Immune functions of immunoglobulin Y isolated from egg yolk of hens immunized with various inceftious bacteria. Biosci. Biotech. Biochem. 60:886–888. Zagulski, T., Lipinski, P., Zagulska, A., Broniek, S., and Jarzabek, Z. 1989. Lactoferrin can protect mice against a lethal dose of Escherichia coli in experimental infection in vivo. Brit. J. Exp. Pathol. 70:697–704. Zeng, J., and Fenna, R.E. 1992. X-Ray crystal structure of canine myeloperoxidase at 3 Å resolution. J. Mol. Biol. 226:185–207. Zhang, X.L., Selsted, M.E., and Pardi, A. 1992. NMR studies of defensin antimicrobial peptides. 1. Resonance assignment and secondary structure determination of rabbit NP-2 and human HNP-1. Biochem. 31:11348–11356. Zhao, C., Nguyen, T., Liu, L., Sacco, R.E., Brogden, K.A., and Lehrer, R.I. 2001. Gallinacin-3, an inducible epithelial beta-defensin in the chicken. Infect. Immun. 69:2684–2691. Zimmermann, G.R., Legault, P., Selsted, M.E., and Pardi, A. 1995. Solution structure of bovine neutrophil beta-defensin-12: The peptide fold of the beta-defensins is identical to that of the classical defensins. Biochem. 34:13663–13671. Zivanovic, S., Chi, S., Draughon, F.A. 2005. Antimicrobial activity of chitosan films enriched with essential oils. J. Food Sci. 70:M45–51. Zommara, M.A., Toubo, H., and Imaizumi, K. 2002. Supplementing bovine milk immunoglobulin G prevents rats fed on a vitamin E-deficient diet from developing peroxidation stress. Ann. Nutr. Metab. 46:97–102.

19 Use of Antimicrobials as Processing Aids in Food Processing Emefa A. Monu and Jairus R. D. David CONTENTS 19.1 Importance.................................................................................................................................. 647 19.2 Regulatory Perspectives.............................................................................................................. 648 19.3 Types of Processing Aids............................................................................................................ 650 19.4 Processing Aids in Meat and Poultry Products........................................................................... 650 19.4.1 Foodborne Illness and Microorganisms of Concern in Meat and Poultry................... 650 19.4.2 Animal Post-Evisceration Carcass Decontamination...................................................651 19.4.2.1 Carcass Washes............................................................................................ 652 19.4.2.2 Processing Aids in Scald and Chilling Water............................................. 654 19.4.2.3 Post-Chill Processing Aids Added for Antimicrobial Activity................... 654 19.4.3 Addition of Processing Aids During Further Processing of Meat and Poultry Products........................................................................................................................ 655 19.4.3.1 Non-RTE Meat and Poultry Products.......................................................... 655 19.4.3.2 RTE Meat and Poultry Products.................................................................. 655 19.4.4 The Role of Acidifiers in Microbial Reduction of Meat and Poultry Products............ 656 19.4.5 Summary...................................................................................................................... 657 19.5 Decontamination of Fresh and Fresh-Cut Produce..................................................................... 657 19.5.1 Fresh Produce – Benefits and Risks............................................................................. 657 19.5.2 Fresh Produce – Foodborne Illness and Microorganisms of Concern......................... 657 19.5.3 Washing and Sanitizing Treatments for Fruits and Vegetables.................................... 658 19.5.3.1 Chlorine....................................................................................................... 659 19.5.3.2 Peroxyacetic Acid (PAA)............................................................................. 659 19.5.4 Summary...................................................................................................................... 660 References............................................................................................................................................... 660

19.1 Importance Food-processing aids are a large category of ingredients that are approved for use in food processing operations to facilitate the batching and production of products, such as plating agents, antifoaming and anticaking agents, acidifiers, and antimicrobials – e.g., lactic acid and chlorine, etc. Approved antimicrobial processing aids play a very important role in ensuring food microbiological quality and safety in the upstream production and pre-processing of food. Food safety is built on a series of processing techniques and applied compounds that each decrease the microbial load of the targeted food. There may not be one that completely eliminates resident microorganisms (thermal processing, irradiation, and certain “non-thermal” processes are the only techniques which can truly accomplish this), but together they act to reduce the microbial load at the front end so that the safety and quality are not dependent solely on the final kill step applied by the processor. Several processes are validated for a 5D or 6D reduction (5–6 log reduction) of the target microorganisms, so if the microbial load of the incoming product is higher than 647

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that, there will still be quality and safety issues in the final food product. Most of the treated end products are intermediate or starting products for manufacturing a range of other products that may or may not be ready to eat (RTE); additional preparation and cooking may be needed at the point of consumption. Many antimicrobial interventions function more effectively when applied to lower microbial loads. As they are often applied during the initial and intermediate portions of processing and subsequent steps may remove or dilute the antimicrobials, it is thus important that processing aids are fast-acting. It is also important that an antimicrobial has low toxicity, even in concentrated form, as this can affect the employees who work with the substance, and may also require that extensive pretreatment is done to any waste water that contains the compound before discharge into the environment, as directed by the US Environmental Protection Agency (EPA). Therefore, there are several characteristics processors look for in an antimicrobial processing aid:

1) Fast-acting 2) Degrades over time so that residual concentrations are not present in the final product 3) Effective at low concentrations so that there will not be significant amounts in the final product 4) Does not pose a health hazard to employees 5) Low toxicity to consumers 6) Cost-effective

For each category of food product, there are common types of antimicrobials used. For example, for beef carcass washes, lactic acid is most commonly used, whereas for poultry products chlorine is traditionally used. Peracetic acid has rapidly become popular for application at various stages of primary processing including sprays, rinses, and chill water. Other chapters of this book have discussed compounds and microorganisms that are added to foods or food-contact surfaces for the sole purpose of their antimicrobial functionality. This chapter will discuss processing aids used as antimicrobials in food processing operations. The role of antimicrobial processing aids will be illustrated using two diverse unit operations with specific objectives: animal postevisceration decontamination, and decontamination of fresh produce and fresh-cut produce. In both of these unit operations processing aids are intended and used for the reduction of the microbial bioburden rather than total elimination. This level of microbial reduction may be adequate for spoilage control and assurance of shelf-life quality, but may or may not be adequate for the control of foodborne human pathogens and assuring food safety. Processing aids are a small and flexible hurdle and to be effective should always be used in conjunction with all other recommended best practices of Good Agricultural Practices (GAPs), Current Good Manufacturing Practices (cGMPs), and Hazard Analysis Critical Control Point (HACCP) throughout the supply chain; animal husbandry, slaughtering, and handling and storage; and agronomy, harvest, and post-harvest handling and storage.

19.2 Regulatory Perspectives The definition and context of application of processing aids is defined in 21 CFR 101.100(a)(3) (Food and Drug Administration, 2018). The three contextual categories of processing aids are:



(a) substances that are added during the processing of a food but are removed in some manner from the food before it is packaged in its finished form; (b) substances that are added to a food during processing, are converted into constituents normally present in the food, and do not significantly increase the amount of the constituents naturally found in the food; or (c) substances that are added to a food for their technical or functional effect in the processing but are present in the finished food at insignificant levels and do not have any technical or functional effect in that food. (Food and Drug Administration, 2018)

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The definition illustrates that it is not only the inherent nature of the substance, but its contextual contribution to the final food product that determines whether it is an additive or a processing aid. For example, if sulfur dioxide is used in the apple peeling and slicing stage of the production of an apple pie to reduce the browning of the apples, it would be a processing aid rather than an additive. It is not present in significant amounts in the final product (most of it will be driven off during baking), has no sensory or functional effect in the final product, and was only used as an aid to the manufacturing of the product (Saltmarsh and Insall, 2013). If, however, that same sulfur dioxide was added as an antimicrobial agent to wine, it would be considered an additive rather than a processing aid. Processing aids, by definition, do not need to be labeled along with other ingredients on the package and for the United States Department of Agriculture (USDA) label review and approval process. The use of any antimicrobial ingredient that does not comply with the three contexts described above as per 21 CFR 101.100(a)(3) should be considered as a direct or indirect food additive and should be declared explicitly in the ingredient listing (see Chapter 1, “Food Antimicrobials—An Introduction,” Table 1.3). Although the inclusion of processing aids on product labels is not legally required in most countries, many companies still disclose the inclusion of these substances on their product label voluntarily. This practice is due to the needs and concerns of customers and consumers of the affected company. For example, maltodextrin is a processing aid commonly used in plating liquid antimicrobials to convert them into powder form when spray drying, and does not need to be labeled in the ingredient listing. However, many companies have chosen to list maltodextrin as an ingredient for increased transparency and to build trust with consumers. Due to the way they are defined, processing aids do not need to be included on food labels, which is why they often get overlooked when discussing food antimicrobials. The above definition was determined by the Food and Drug Administration (FDA), which regulates most of the U.S. food supply. The U.S. Department of Agriculture (USDA), which regulates meat and eggs, does not have its own definition listed in its regulations; for the purpose of determining whether a substance is a processing aid, the FDA definition is used. The United States is not the only country to look at processing aids in this manner. Mexico defines a processing aid as a substance or material, excluding instruments, utensils and additives, that is not consumed as a food ingredient by itself and is used intentionally in the production of raw materials, foods or their ingredients to achieve a technological function during the treatment or processing and that can lead to the unintentional presence of residues or derivatives in the final product

(Magnuson et al., 2013). The Codex Alimentarius Commission, a joint international food standards program between the Food and Agriculture Organization and the World Health Organization, defines processing aids as any substance or material, not including apparatus or utensils, and not consumed as a food ingredient by itself, intentionally used in the processing of raw materials, foods or its ingredients, to fulfil a certain technological purpose during treatment or processing and which may result in the non-intentional but unavoidable presence of residues or derivatives in the final product.

(Codex Secretariat, 2018). The European Union (European Commission, 2008) and Australia (Food Standards Australia New Zealand, 2008) also use similar definitions. However, some countries such as Canada have no regulatory definition of the term processing aid, although Health Canada does have a working definition of the term that is similar to that used by Codex and does not require them to be included on the ingredient label (Canadian Food Inspection Agency, 2018). Countries also differ in how they classify specific antimicrobials. For example, in Canada ozone gas can be applied as a processing aid to poultry meat carcasses (up to 10 ppm) and poultry parts (up to 200 ppm) (Health Canada Bureau of Chemical Safety Food Directorate, 2018). However, in the U.S., such application is considered a secondary direct food additive by the USDA (US Department of Agriculture Food Safety Inspection Service, 2019).

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19.3 Types of Processing Aids Processing aid substances include cleaners and sanitizers, water-treatment chemicals, catalysts, exchange resins, desiccating agents, filtering agents, washing and peeling agents, emulsifying agents, and head space/packaging gases. They are often applied in meat and poultry processing, fruit and vegetable processing, water treatment, and included in a variety of further processed foods and beverages. Seafood products are unique in that they serve as both a source and an application of processing aids. Enzymes harvested from the wastes of fish processing (such as protease inhibitors) can then be used in seafood and other types of products. Although cleaners and sanitizers have antimicrobial properties, they will not be discussed here in detail as they are not applied to the food. Various processing aids have purposes in multiple commodities, for example, acidified sodium chlorite is used in poultry, red meat, seafood, and fresh produce. Examples of some processing aids, their approved usage levels in the United States, and applications are shown in Table 19.1.

19.4 Processing Aids in Meat and Poultry Products 19.4.1 Foodborne Illness and Microorganisms of Concern in Meat and Poultry Meat is a perishable product. There are several bacteria that can cause spoilage of meat products and reduce the quality even at refrigeration temperatures. These bacteria may be naturally present on the carcass, or may be introduced through contact with knives, equipment, and surfaces in the processing environment. Improper employee handling techniques may also increase the number of bacteria on meat products. There are many bacteria that are commonly found on fresh meats and poultry. Those that cause spoilage include slime-inducing Pseudomonas, Lactobacillus, Enterococcus, and Brochothrix; Shewanella, which can cause greening; and Clostridium, which can cause bone taints (Nychas et al., 2008; Sohaib et al., 2016). Although fresh meat is usually not RTE and proper cooking times and temperatures will eliminate most pathogenic bacteria, if these microorganisms are present in fresh meat they may still pose a threat to consumer health due to improper cooking temperatures, improper handling in the home leading to cross contamination, and improper storage temperatures (Mataragas et al., 2008; Ravishankar et al., 2010). There are several microbial hazards associated with raw meat and poultry. These include Salmonella, Campylobacter, Clostridium perfringens, and Shiga toxin-producing Escherichia coli (Borch and Arinder, 2002; Mead, 2004). Prevalence rates for these organisms vary by country and product. For example, an Ethiopian study from two beef-processing plants found E. coli O157:H7 in 0.54% of skin and internal carcass swabs and in 0.8% of samples from retail stores (Abdissa et al., 2017). Similarly, analysis of carcasses at a vertically integrated cattle operation in Mexico showed E. coli O157:H7 in 0.4% of carcasses post-processing (Narvaez-Bravo et al., 2013). In the EU, from 2007 to 2009, fresh meat was found to have a prevalence of 0.3–2.3% and 0.1–0.7% of STEC and O157 E. coli, respectively (Takkinen et al., 2011). In the U.S., prevalence rates of STEC in post-processing beef carcasses have been shown as 10% (Arthur et al., 2002). In the U.S., the USDA has set performance standards (Food Safety and Inspection Service, 2016) for the presence of viable Salmonella and Campylobacter in comminuted (ground) and whole carcasses of chicken and turkey as well as chicken parts (Table 19.2) and has established E. coli O157:H7 and six other strains of Shiga toxin-producing E.coli (STEC) as adulterants in ground beef, with zero tolerance for any presence of these E.coli in these types of products (Food Safety and Inspection Service, 2011). In combination with including antimicrobials in the finished product and utilizing processing techniques such as thermal processing, irradiation, acidification, and drying to destroy these pathogens, it is important to control both spoilage and pathogenic microorganisms through the primary processing chain. In order to keep pathogens to acceptable levels in the final product, it is necessary to employ several techniques. To ensure this, the USDA Food Safety and Inspection Service (FSIS) established requirements for meat and poultry facilities to implement a Hazard Analysis and Critical Control Point plan. This plan involves the development of a “system of preventive controls designed to improve the safety of their products” (Food Safety and Inspection Service, 1996). Processing aids play a vital role

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Use of Antimicrobials as Processing Aids in Food Processing TABLE 19.1

Some Select Examples of Approved Antimicrobial Applications of Processing Aids in Various Foods in the United States Substance

Intended Use

Aqueous solution of citric and hydrochloric acids adjusted to a pH of 1.0 to 2.0

Poultry carcasses, parts, trim, and organs

Aqueous solution of peroxyacetic acid, hydrogen peroxide, acetic acid, and 1-hydroxyethylidene-1, 1-diphosphonic acid (HEDP) Blend of lactic acid, polysorbate 80, and xanthan gum

Poultry scald tanks

Bacteriophage preparation (Salmonella-targeted)

Ready-to-eat (RTE) poultry products prior to slicing and on raw poultry, including carcasses and parts applied as a spray Meat or poultry product (including ground, formed, or whole muscle meat) that will be heat-treated and processed to be NRTE or RTE

Combination of natural source of nitrite and natural source of ascorbate

Lauramide arginine ethyl ester (LAE) dissolved in ethanol or water Colicin protein preparation

Acidified sodium chlorite Potassium acid tartrate Sodium pentachlorophenate

Processing of beef heads and carcasses

Fresh cuts of beef and pork

Nine recombinant proteins intended for use singly or in combination as an antimicrobial spray on meat products Spray or dip for fresh produce pH lowering in wine and other beverages Preservative for ammonium alginate, a processing aid in the manufacture of polyvinyl chloride emulsion polymers for use as articles or components of articles that contact food

Amount An aqueous solution of citric and hydrochloric acids adjusted to a pH of 1.0 to 2.0 applied as a spray or dip with a minimum contact time of 2 to 5 seconds, pH measured prior to application Level of peroxyacetic acid will not exceed 220 ppm, hydrogen peroxide will not exceed 110 ppm, and 1-hydroxyethylidene-1, 1-diphosphonic acid (HEDP) will not exceed 13 ppm Not to exceed 5% lactic acid solution, not to exceed 0.07% polysorbate 80, and not to exceed 0.05% xanthan gum; applied as a spray; exposure time 5–30 seconds, pressure 20–60 psi, temperature 18–55°C Bacteriophage preparation (Salmonella-targeted) applied as a spray at 106 to 107 plaque-forming units (pfu) per gram of food product For use as a component in the product formulation at (1) a rate of a minimum 75 ppm of nitrite from natural sources and minimum 500 ppm of ascorbate from natural sources or (2) a rate of a minimum 100 ppm of nitrite from natural sources and minimum 250 ppm of ascorbate from natural sources by weight of the finished food product Applied and dried to the inside of packaging at a concentration not to exceed 105 ppm LAE by weight of finished food product Colicin protein preparation applied as a spray at a rate of 1–10 mg/kg

500–1200 ppm at pH 2.3–2.9 Not specified At temperatures not exceeding room temperature, ≤0.5 % w/w of ammonium alginate solids

Source: USDA (U.S. Department of Agriculture Food Safety Inspection Service, 2019), and 21 CFR (Food and Drug Administration, 2018a, 2018c)

in these systems and, as seen in Table 19.3, can be applied anywhere from the fresh carcass to the final in-package product.

19.4.2 Animal Post-Evisceration Carcass Decontamination Meat and poultry products go through several stages; first-processing (also known as primary processing) involves the conversion of the live animals to carcasses and parts such as roasts, wings, organ meats,

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Antimicrobials in Food TABLE 19.2 Performance Standards for Raw Poultry in Processing Plants in the United States Maximum % Positive (Performance Standard) Product Broiler chicken carcass Turkey carcass Comminuted chicken Comminuted turkey Chicken parts

Salmonella

Campylobacter

9.8 (5 out of 51) 7.1 (4 out of 56) 25 (13 out of 52) 13.5 (7 out of 52) 15.4 (8 out of 52)

15.7 (8 out of 51) 5.4 (3 out of 56) 1.9 (1 out of 52) 1.9 (1 out of 52) 7.7 (4 out of 52)

Source: FSIS (Food Safety and Inspection Service, 2016).

etc. Further processing converts the carcasses and parts into other products such as frozen fried chicken, burgers, deli meats, canned soups, etc. (Fletcher, 2004). Processing aids are one strategy applied during meat and poultry processing to achieve pathogen reduction. After evisceration and before further processing, they can be applied as carcass washes, or be added to water used in chilling or scalding of the carcass.

19.4.2.1 Carcass Washes One application of processing aids in meat and poultry first-processing facilities is as antimicrobials in the carcass washes applied post-slaughter. Water is used to remove fecal matter, hair/feathers and microorganisms, and in carcass chilling and is therefore a prime vehicle for antimicrobial compounds. In the U.S. poultry industry, it has been estimated that 26 L of water is used per bird (Northcutt and Jones, 2004), which is a significant amount considering that many plants produce over 1 million birds per week. Beef carcasses are sprayed, which is always applied from the top of the carcass to the bottom. Other red meat carcasses such as goat, pork, veal, and lamb are washed in a similar manner, but application times are shorter due to the smaller sizes of the carcasses. The beef carcass is first washed with warm or hot water (usually for 2 minutes) followed by a 5-minute drip. This drip time is necessary to allow the excess film of water to be removed from the carcass surface, which would impede the interaction of the antimicrobial applied in the following step with any bacteria on the carcass surface. The antimicrobial rinse is applied for about a minute and can be made up of different compounds. Organic acids are usually used as antimicrobial agents during carcass washing. The most common antimicrobial rinse for beef carcasses is a 2% lactic acid solution, which has been shown to result in a 3.5, 4.7, and 5.0 log reduction of E. coli O157:H7, Salmonella Typhimurium, and Campylobacter spp., respectively, on beef carcasses (Flowers, 2006). Acetic acid is also commonly used at 2%, and commercial mixes such as Fresh Bloom™ that include several organic acids are also available. However, there are several mixtures of antimicrobials that are approved as safe and suitable ingredients by the Food Safety Inspection Service (FSIS) for use as sprays or dips on meat carcasses (Table 19.3). This list is a living document that is constantly being updated as companies or researchers apply for approval. One commercially available example is Amplon, which contains sulfuric acid and sodium sulfate and is approved for use as a processing aid as a spray, dip, or wash for poultry parts and carcasses and has exhibited a 0.8–1.2 log reduction of Salmonella on inoculated chicken wings (Kim et al., 2017; Scott et al., 2015). Poultry carcass washing takes place at several points of first-processing. The outside of the bird is rinsed after de-feathering, and there is often an inside wash as well where nozzles are inserted into the abdominal cavity. Another wash can be applied just prior to the chilling of the carcass (Barbut, 2016). Inside-outside bird washers (IOBW) and dip tanks are then used both before and after the chilling step to create multiple hurdles (Singh et al., 2015). As in red meat production, organic acids can be used in poultry carcass washes. One study showed that wash sprays containing >5% lactic acid significantly reduced Salmonella counts on chicken thighs with or without skin (Ramirez-Hernandez et al., 2017). Similarly, a dip treatment of acidified lactic acid (ALA), containing 0.5% lactic acid, reduced S. Heidelberg and Campylobacter jejuni in ground chicken frames (Moore et al., 2017).

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Use of Antimicrobials as Processing Aids in Food Processing TABLE 19.3

Some Select Examples of Processing Aids Deemed Safe and Suitable for Use in Meat and Poultry Products by the USDA Substance Aqueous potassium hydroxide solution Aqueous solution of citric and hydrochloric acids adjusted to a pH of 0.5 to 2.0 Aqueous mixture of peroxyacetic acid (PAA), hydrogen peroxide (HP), acetic acid, sulfuric acid (optional), and 1-hydroxyethylidene-1,1diphosphonic acid (HEDP), catalyzed with sulfuric acid

Blend of citric acid, phosphoric acid, and hydrochloric acid

Aqueous solution of acidic calcium sulfate and lactic acid Acetic acid Aqueous solution of citric acid and hydrochloric acid Aqueous solution of citric acid and hydrochloric acid

Aqueous mixture PAA, HP, acetic acid, and HEDP

Lauramide arginine ethyl ester (LAE) dissolved in either ethanol or water

Blend of citric acid and sorbic acid in a 2:1 ratio

Intended Use Hide-on carcass wash in spray cabinet Meat carcasses, parts, trim, and organs (1) In process water used for washing, rinsing, or cooling whole or cut meat including carcasses, parts, trim, and organs; (2) in process water or ice for washing, rinsing, storing, or cooling of processed and pre-formed meat products Poultry carcasses

Continuous spray or dip on raw poultry carcasses, parts, giblets, and ground poultry Chicken livers Processed and comminuted red meat products in an enclosed mixing, grinding, and/or blending system. Permeable and impermeable casings of meat and poultry products applied as a spray, dip, or immersion to casings prior to opening, removal, or slicing operations (1) For use in: brines, sauces, and marinades applied either on the surface or injected into processed or unprocessed, cooked or uncooked, whole or cut poultry or parts and pieces; (2) surface sauces and marinades applied on processed and preformed meat and poultry products Packaged fresh cuts of beef and pork

Reduction of the microbial load of purge trapped inside soaker pads in packages of raw whole muscle cuts of meat and poultry

Amount Wash solution used at a final concentration of 0.01–0.40 % w/w Adjusted to a pH of 1.0 to 2.0 applied as a spray or dip for a contact time of 2 to 5 seconds (1) PAA not to exceed 1800 ppm, HP not to exceed 600 ppm, and HEDP not to exceed 22.5 ppm; (2) Not to exceed 495 ppm PAA, 165 ppm HP, and 14 ppm HEDP

Citric acid 1.87%, phosphoric acid 1.72%, and hydrochloric acid 0.8% applied as a spray with a minimum contact time of 1 to 2 seconds and allowed to drip from the carcasses for 30 seconds Acidic calcium sulfate sufficient for purpose; lactic acid not to exceed 5.0 % and 55°C Immersion dip at a concentration of up to 5% and not to exceed 2 minutes Solution adjusted to a pH of 0.5–2.0

Solution adjusted to a pH 1 µm) (Jesorka & Orwar, 2008). Solid lipid nanoparticles (SLNs) are another platform that can be useful for the distribution of antimicrobials. These delivery systems are generally in the range of 50 to 1000 nm, formed by solid lipids at room temperature and a surfactant for emulsification. The materials used in the formulations are solid lipids such as fatty acids (palmitic, decanoic, behenic acids), triglycerides (trilaurine, tripalmitin, trimiristine), sterols (cholesterol), monoglycerides (glyceryl monostearate), and waxes (cetyl palmitate). Various types of surfactants are used for emulsification, including lecithin, phosphatidylcholine, cholate, glycolate, and poloxamer 188 (Zhang, Pornpattananangkul, Hu, & Huang, 2010). Nanostructured lipid carriers (NLCs) have been introduced as the next generation of the SLNs to overcome the potential limitations of SLNs. They are distinct from SLNs by the composition of the solid matrix, as the lipid phase in NLCs contains both solid and liquid lipids at ambient temperature. In fact, NLCs are modified cohorts of SLNs that present a mixture of solid and liquid phase (oil) resulting in an unstructured matrix, which improves the stability and loading capacity (Ganesan & Narayanasamy, 2017; Naseri, Valizadeh, & Zakeri-Milani, 2015). The general types of SLNs and NLCs are summarized in Figure 20.2. Solid lipid nanoparticles have several unique characteristics that make them attractive as antimicrobial delivery systems. The excipients of SLNs are occlusive, meaning that a thin film is rapidly formed with the application on a surface, reducing the evaporation of water and retaining moisture from the surface. This property promotes an enhancement of the active substance penetration into the substrate because of the components of SLNs (Ghasemiyeh & Mohammadi-Samani, 2018). The production of SLNs is relatively simple and allows reproducible production on a large scale, without the need for organic solvents.

20.4.1 Preparation Methods Most preparation methods of lipid-based delivery systems result in heterogeneous mixtures of lipid vesicles, including MLVs and LUVs. In general, more homogeneous mixtures can be obtained by using heat or the application of shear forces such as sonication or extrusion. The challenge is to achieve the formation of particles with adequate size, low dispersion, and good encapsulation efficiency (Patil & Jadhav, 2014).

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FIGURE 20.2  Types of solid-lipid nanoparticles (SLNs) and nanostructured lipid carriers (NLCs). Reproduced from Ganesan and Narayanasamy (2017), with permission from Elsevier.

FIGURE 20.3  Liposome formation by the reversed-phase method. (1) Phosphatidylcholine is dissolved in an organic solvent. (2) A portion of aqueous solution containing chitosan is added. (3) An emulsion of reverse micelles containing chitosan is obtained under sonication. (4) The organic solvent is evaporated. (5) The organogel is obtained. (6) Vesicles containing chitosan are formed after water addition and shaking. Reproduced from Mertins, Lionzo, Micheletto, Pohlmann, and Silveira (2009), with permission from Elsevier.

The most used methods for antimicrobial encapsulation into liposomes are the film hydration method and the reverse micelle method. The film hydration method starts with a solution of the phospholipid in an organic solvent and after solvent evaporation, a thin lipid film is formed in the wall of the flask. An aqueous solution containing the antimicrobial substance is added under agitation, which often results in MLVs. Ultrasound, heat, or extrusion is then applied to obtain LUVs and then SUVs (Malheiros, Daroit, & Brandelli, 2010a). On the other hand, the reverse micelle method includes the phospholipid dissolution in the organic solvent and the addition of the aqueous phase with the active substance (Figure 20.3). A W/O emulsion

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FIGURE 20.4  Production of nanometric liposomes using microfluidic approaches. (A) Schematic representation of microfluidic device for the production of liposomes of controlled size by hydrodynamic focusing. (B) Schematic representation of the microfluidic drop system for microfluidic fabrication of water-in-oil-in-water double emulsion drops with ultrathin shells.

is formed with the phospholipid at the interface. Next, the mixture is ultrasonicated and the reverse micelles are formed, where water droplets are coated by the phospholipid. After evaporation, many micelles collapse and a high-viscosity organogel is formed. In the final stage, the liposomes are formed by adding water and stirring (Mertins, Sebben, Pohlmann, & da Silveira, 2005). More recently, the hydrodynamic focusing approach in microfluidic devices has been developed and applied for the production of liposomes (Patil & Jadhav, 2014). The formation of the liposome is carried out by the introduction of a channel containing the phospholipids dispersed in alcohol, generally ethanol or isopropanol, through a central channel. Then, the constriction of that channel occurs by two perpendicular water lines (Figure 20.4A). The hydrodynamic focus of the central channel causes controlled diffusion of the alcohol to the aqueous phase and vice versa. This results in a change of solubility and selforganization of the phospholipids into bilayers and finally into SUVs (Jahn, Vreeland, DeVoe, Locascio, & Gaitan, 2007; Zizzari et al., 2017). The production of liposomes by microfluidic drop systems has also been described (Figure 20.4B). Microfluidic devices are used to generate W/O/W (water-in-oil-in-water) double emulsions of sub-micron size. Phospholipids are solubilized in partially water-soluble organic solvents to accelerate the solvent evaporation process. With the evaporation of the intermediate phase, giant liposomes are formed (Trantidou, Friddin, Salehi-Reyhani, Ces, & Elani, 2018). Typical methods of SLN preparation include high-pressure homogenization, atomization, mixing with high shear force, and ultrasound (Naseri et al., 2015). Spray drying (atomization) is an economical alternative to freeze-drying that is used to convert an aqueous SLN dispersion into a powdered product. This method causes particle aggregation due to the high temperature, shear forces, and partial melting of the particle during the process. Lipids with melting point(s) higher than 70°C are recommended for spray drying. The greatest advantage of high-speed stirring and sonication in SLN production is related to the easy access to the required equipment, which is common in many laboratories (Ganesan & Narayanasamy, 2017). The problem of these methods is the generation of broader particle size distributions, ranging into

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the micrometer range. Such variability results in physical instability, including particle aggregation upon storage. Potential metal contamination due to sonication is also a significant problem in ultrasound-based methods. To achieve stable formulations, high-speed stirring and ultrasound should be used in combination and performed at high temperatures (Zhang et al., 2010).

20.4.2 Applications Liposomes have been employed for a long time as a drug delivery system, efficiently carrying antimicrobial substances (Brandelli, Pinilla, & Lopes, 2017c). The liposome encapsulation of antimicrobials intended for food use, such as nisin, lysozyme, curcumin, and essential oils, among others, has been described as well (Lopes & Brandelli, 2018). Nisin and other food-grade antimicrobials showing a broad antimicrobial activity may have impaired effectiveness in food systems due to undesirable interaction with some food components or enzyme inactivation (Malheiros et al., 2010a). Liposomes encapsulating natural antimicrobials have been shown to be active against food pathogens. For example, nisin has been successfully incorporated into liposomal systems prepared from different phospholipids, soy lecithin, and marine lecithin (Imran et al., 2015; Taylor, Gaysinsky, Davidson, Bruce, & Weiss, 2007). In addition, the bacteriocin pediocin PA-1/AcH, belonging to the Class II bacteriocins known as “antilisterial” peptides, was incorporated into phosphatidylcholine liposomes. The nanostructured pediocin showed antimicrobial activity against diverse Listeria spp. (de Mello et al., 2013). Some examples of liposomes as antimicrobial carriers in food systems were previously described. Phosphatidylcholine/phosphatidylglycerol liposomes encapsulating nisin and EDTA were tested in liquid medium, resulting in a significant extension of the lag phases of L. monocytogenes and E. coli O157:H7. The bacteriostatic inhibition of these pathogens for at least 48 h was reported (Taylor, Bruce, Weiss, & Davidson, 2008). In another study, the lag phase of L. monocytogenes Scott A incubated at 5 or 20°C in fluid milk was significantly increased with the addition of encapsulated nisin (Schmidt, Holub, Sturino, & Taylor, 2009). Similar inhibition of L. monocytogenes was observed in whole or skimmed fluid milk at 7 and 30°C after the addition of liposome-encapsulated nisin (Malheiros, Daroit, da Silveira, & Brandelli, 2010b), garlic extract (Pinilla, Noreña, & Brandelli, 2017), or nisin/lysozyme mixtures (Lopes, Barreto Pinilla, & Brandelli, 2019). Phosphatidylcholine liposomes containing nisin were also able to control L. monocytogenes in cheese (Malheiros, Sant’Anna, Barbosa, Brandelli, & Franco, 2012). Solid lipid nanoparticles (SLNs) are ideal for topical applications and commonly used in the administration of drugs that are extremely insoluble in water, to combat superficial fungal infections (Gupta & Vyas, 2012). In addition to topical applications, SLNs are also used as capsules for oral administration, for example, to combat infections of the gastrointestinal tract by Pseudomonas spp. This utility can be extended to deliver poorly water-soluble antimicrobials, such as essential oils, in food systems. SLNs are interesting systems for the delivery of hydrophobic bioactive compounds that can be prepared using food-grade lipids. In this regard, nisin-loaded SLNs were produced with Inwitor 900 (glyceryl monostearate) by three cycles of high-pressure homogenization and using poloxamer 188 and sodium deoxycholate as surfactant and co-surfactant, respectively (Prombutara, Kulwatthanasal, Supaka, Sramala, & Chareonpornwattana, 2012). These nanostructures promoted the controlled release of nisin, inhibiting L. monocytogenes for up to 20 days during in vitro release studies.

20.5 Polymer-Based Delivery Systems Biocompatible and biodegradable polymers have been widely used for the controlled release of antimicrobial compounds. Such delivery systems can be developed in the form of nanoparticles or composite films and coatings. A variety of biodegradable polymers has been used to prepare delivery systems, including synthetic polymers such as poly(lactic acid) (PLA), poly(lactic-co-glycolic acid) (PLGA), poly(ε-caprolactone) (PCL), and natural polymers including proteins such as gelatin, casein, and whey proteins, or polysaccharides such as cellulose, starch, alginate, and chitosan (Kumari, Yadav, & Yadav, 2010).

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Polymeric nanoparticles have some distinctive characteristics. They have structural stability and well-defined properties (such as size and zeta potential), which are achieved by precise control over the polymer size, surfactant, and solvent used during their synthesis. In addition, the surface of polymeric nanoparticles generally contains functional groups that can be modified with the addition of a drug or a ligand. This modification allows the development of targeting strategies by the attachment of specific affinity ligands such as antibodies and lectins at the particle’s surface. Dendrimers are nanoparticles defined as highly ordered globular branched macromolecules (Boas & Heegaard, 2004). The structure of a dendrimer consists of three regions: the nucleus (or focus), the dendritic structures (highly branched), and the external surface that contains functional groups. The first synthetic methods for branched amines were described in 1978, then the patents of highly branched poly-L-lysine dendrimers appeared, and in 1984 the synthesis of the first family of polyamidoamine (PAMAM) appeared (Abbasi et al., 2014). Dendrimers have several unique properties that produce nanoparticles for the diffusion of antimicrobials. The highly branched dendrimers provide an enormous surface to size ratio and, therefore, allow a high reactivity with microorganisms in vivo. The drug can be loaded in the hydrophobic structure or by electrostatic interaction with a multivalent hydrophilic surface. In addition, the dendrimer can be antimicrobial by itself, for example, when quaternary ammonium compounds are used as terminal functional groups, which are known for their capacity to disrupt bacterial membranes. PAMAM dendrimers are the most studied for the supply of antimicrobials, and some studies show that the antifungal and antibacterial activities of some existing chemotherapeutics can be improved by dendrimer encapsulation (Kalomiraki, Thermos, & Chaniotakis, 2015). A polymeric film or coating can be defined as a thin, continuous layer of polymeric material formed or placed on food or a food component. Besides acting as protective barriers, films can be used as carriers of antimicrobial compounds, thus enhancing the shelf-life of the food product and potentially promoting health benefits (Brandelli et al., 2017a). Biopolymer-based films and coatings usually exhibit lower moisture barriers when compared to petroleum-based films due to their hydrophilicity. To improve moisture barrier characteristics, hydrophobic compounds such as essential oils can be included in the formulations prepared from hydrophilic materials, not only increasing the resistance to moisture, but also providing them with antimicrobial properties. Antimicrobial films and coatings made from polysaccharides have been demonstrated to improve the physicochemical and microbiological quality of fresh-cut fruits and vegetables (Cé, Noreña, & Brandelli, 2012; Ramos, Fernandes, Silva, Pintado, & Malcata, 2012). Nanofiber mats have also been developed as antimicrobial delivery systems. Nanofibers are produced from polymers specifically treated to form filaments with a diameter in the nanometer range. Different materials can be used to produce nanofibers, including both synthetic and natural polymers. Nanofibers have great potential for the delivery of antimicrobials because they have a very high surface to volume ratio (Wei, 2012). Antimicrobial loading onto the nanofiber can occur by encapsulation during the electrospinning process or by simple physical adsorption onto the nanofiber surface, generally given by hydrogen and van der Waals bonds. Another possibility is the adsorption of nanoparticles loaded with the antimicrobial on the surface of the fibers. A layer-by-layer assembly method allows a few nanometers’ coverage of nanofiber surface by the deposition of polyelectrolytes. Charged polysaccharides such as heparin or chitosan can be used (Brandelli, 2012).

20.5.1 Preparation Methods Polymeric nanoparticles can be prepared using mainly two methods that consist of self-assembly and emulsification. Self-assembled nanoparticles formed by copolymers are organized into hydrophobic and hydrophilic segments. In general, the hydrophobic segment forms the core containing the drug and the hydrophilic segment provides protection for the core against degradation. The general preparation method is by solvent displacement. In that process, the encapsulating polymer and the drug payload are dissolved in a water-miscible solvent, such as acetonitrile or acetone, and then added to an aqueous suspension. As the solvent evaporates, the copolymers undergo a nanoprecipitation that traps the active substance (Vauthier & Bouchemal, 2009). Another technique of polymeric nanoparticles preparation is emulsification-polymerization, which forms nano-capsules. In this method, the monomer is dissolved in the polymerization medium in the

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presence of a surfactant, polymerization initiators are added, and the drug can be adsorbed to the nanocapsules during the polymerization or covalently conjugated after displacement of the solvent. As the solvent evaporates, the copolymers undergo a nanoprecipitation that traps the active substance (Zhang et al., 2010). Other methods for preparation of polymeric nanoparticles are interfacial polymerization, salting-out, coacervation, solvent diffusion, and self-assembling macromolecules (Vauthier & Bouchemal, 2009; Yadav, Kumari, & Yadav, 2011). Dendrimers can be chemically synthesized by two different methods, either divergent or convergent (Boas & Heegaard, 2004). In the divergent approach, the dendritic structure is synthesized from the core as the initial point and built up generation by generation. The alternative convergent method starts from the surface and terminates at the core, where the dendrimer segments are coupled together. In the convergent approach, only a small number of reactive sites are functionalized in each step, giving a smaller number of possible side-reactions per step (Figure 20.5). Polymer-based films containing antimicrobials for food packaging applications can be produced by the casting method or by thermal processing. The production of thin polymer films by casting methods has been employed for a long time. This technique involves the distribution of a film-forming solution onto a uniform surface, followed by the solvent evaporation, and finally the formed film is peeled off the support surface. Casting methods have been widely employed to produce biopolymer-based antimicrobial packaging films, mostly using starch, chitosan, carboxymethyl cellulose, and gelatin as polymer matrices (Siemann, 2005). The production of packaging films by thermal processing, such as extrusion, melt blending, and injection molding, has been extensively described and can also be applied to incorporate antimicrobials into thermoplastic polymers. The thermal stability of the antimicrobial compound and compatibility with the polymer matrix are critical aspects to be considered in thermal processing. Since melting temperatures must be applied, these thermal processes use often use temperatures above 140°C, and therefore conventional petroleum-based thermoplastics, such as low-density polyethylene (LDPE), polypropylene (PP), and poly-lactic acid (PLA), are employed (Gavara, 2015). Both casting and thermal processes have been used to incorporate natural antimicrobials such as the bacteriocin nisin into packaging films (Imran et al., 2012; Zehetmeyer et al., 2016). Several methods are described for the production of ultra-fine fibers, but the electrospinning method is the most effective and straightforward for producing nanofibers in large quantities. In this method, one electrode is located in the solution containing the polymer and the other is linked to a collector, which is generally a metal tube or plate. The charge forms a Taylor cone at the tip of the needle and with the acceleration of the polymer solution the solvent evaporates and forms the nanofiber (Wei, 2012). A

FIGURE 20.5  Schematic representation of dendrimer synthesis pathways. Top: divergent strategy. Bottom: convergent strategy. Reproduced from Boas and Heegaard (2004), with permission from Royal Society of Chemistry.

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schematic representation of the electrospinning apparatus is shown in Figure 20.6. Coaxial electrospinning is used to produce nanofibers with a core-shell structure. Nanofibers produced by coaxial electrospinning that have been previously demonstrated include polyethylene oxide (shell) and polydodecyl thiophene (core). Nanofibers formed by nano-emulsification are prepared from an aqueous phase containing the hydrophilic polymer mixed into an organic phase containing the hydrophobic polymer that makes the coating. This nano-emulsion is then subjected to electrospinning to produce the nanofibers (Xu et al., 2006). Poly-ethylene glycol-poly lactic acid (PEG-PLA) copolymer has been used previously in the fabrication of nanofibers via the nano-emulsion method (Heunis & Dicks, 2010). Another method is the co-electrospinning, where the active substances are present in the polymer suspension. An example is the conjugation of the antimicrobial peptide (Ser-Glu-Glu)3 with polyethylene oxide. During the electrospinning, the peptide is polarized with orientation to the nanofiber surface (Yoo, Kim, & Park, 2009). The quality and characteristics of the nanofiber depend on several factors, such as manufacture temperature, viscosity, surface tension of the suspension, and the intensity of the electric field. Thicker fibers are obtained by increasing the polymer concentration, for example, 4% polyvinylpyrrolidone (PVP) produces fibers of 20 nm and 8% PVP produces 50 nm nanofibers, while 10% PVP results in 300 nm. Also, the morphology of the nanofibers can change according to the concentration of the polymer. For instance, PLA nanofibers prepared from 1 or 3% suspensions can result in a smooth morphology or a bead-on-string shape, respectively (Heunis & Dicks, 2010). Nanofibers are amenable to surface modification methods and consequently broadening their applications, including targeted drug delivery, also including delivery to food-contaminating microorganisms. Several methods to modify synthetic polymers are used to improve the diffusion capacity of drugs from nanofibers, such as wet chemical methods (Brandelli, Lopes, & Boelter, 2017b). For example, the NaOH treatment of PLA nanofibers increases the ability to bind calcium (for bone-regeneration purposes). The reaction of diamines with polyester nanofibers can increase drug-loading capacity through the adhesion of the charged surface. Plasma treatment is also a technique that has been used. For example, treatment with oxygen or air plasma can produce carboxyl and amino groups on their surfaces. The treatment of PCL nanofibers with argon or air plasma has been used to increase the number of carboxyl groups and shown to increase cell adhesion and proliferation (Yoo et al., 2009). The introduction of functional groups on the surface of nanofibers can also be done by graft polymerization (copolymer) (Yoo et al., 2009). In general, UV or plasma treatment is used to generate free radicals for polymerization.

FIGURE 20.6  Schematic representation of an electrospinning device showing its component and the resulting nanofiber mat (representative SEM image) collected after the process. Reproduced from Brandelli, Lopes, and Boelter (2017b), with permission from Elsevier.

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20.5.2 Applications Polymeric nanoparticles encapsulating natural antimicrobials as delivery systems have been widely studied. The great interest in polymeric nanoparticles is associated with their advantages not only regarding their capacity to encapsulate a vast selection of hydrophobic and hydrophilic molecules, but also due to their ability to release them in a controlled or triggered fashion. Polymeric particles for delivery systems consist of a shell formed by the polymer chain, surrounding the entrapped core containing the bioactive compound, enhancing the stability of the latter (Borel & Sabliov, 2014). Furthermore, there is a variety of synthetic and natural polymeric materials available that are considered biodegradable, biocompatible, and non-toxic, which have been used to encapsulate different natural antimicrobials, including bacteriocins, enzymes, essential oils, and phenolic compounds. These delivery systems have been shown to improve antimicrobial activity of the entrapped compound against both Gram-positive and Gramnegative pathogens including L. monocytogenes, E. coli, and Salmonella enterica (Hill & Gomes, 2014; Hill, Taylor, & Gomes, 2013b; Ozdemir et al., 2018; Pereira et al., 2018; Pola et al., 2019; Silva, Hill, Figueiredo, & Gomes, 2014). Consequently, there is a plethora of polymeric material and antimicrobial compound combinations available to develop antimicrobial delivery systems for specific applications. Therefore, polymeric nanoparticle-based delivery systems development is expected to continue expanding in the future. Polymer-based antimicrobial films and coatings have been extensively studied in the last decades, and several examples of applications have been described in the literature. These include the utilization of either conventional or natural polymers, and effective films have been produced by the incorporation of antimicrobials such as essential oils, phenolic compounds, bacteriocins, and lysozyme among others (Azeredo, 2013; Brandelli et al., 2017b; Brasil, Gomes, Puerta-Gomez, Castell-Perez, & Moreira, 2012; Mantilla, Castell-Perez, Gomes, & Moreira, 2013; Martiñon, Moreira, Castell-Perez, & Gomes, 2014; Rhim, Park, & Ha, 2013; Sipahi, Castell-Perez, Moreira, Gomes, & Castillo, 2013; Vasile, 2018). Packaging materials having controlled-release properties for multiple active compounds, showing both antimicrobial and antioxidant properties, have been developed. Active films prepared with a blend of zein with fatty acids showed lower initial release rates for model bioactive compounds, namely (+)-catechin and lysozyme, than the zein-derived control films. Both catechin and lysozyme release rates were affected by modification of the fatty acid chain length, while the number of double bonds in the fatty acid influenced only the catechin release rate. The analysis of film morphology indicated that the controlledrelease properties of the blend films was mostly related to the microspheres formed within their matrix and encapsulation of bioactive compounds (Arcan & Yemenicioğlu, 2014). Cinnamaldehyde and cinnamon bark extract have been recognized by their strong antimicrobial activity against diverse foodborne pathogens, such as S. aureus, E. coli, Bacillus cereus, and P. aeruginosa (Gomes, Moreira, & Castell-Perez, 2011; Hill, Gomes, & Taylor, 2013a; Hill et al., 2013b; Loquercio, Castell-Perez, Gomes, & Moreira, 2015; Siddiqua, Anusha, Ashwini, & Negi, 2015). Different concentrations of this natural antimicrobial compound were incorporated in sorbitan monooleate/chitosan films. The obtained films ranged from 145 to 345 nm in thickness, and these structured chitosan/cinnamaldehyde materials exhibited the ability to control bacterial colonization on a variety of surfaces, including food process equipment surfaces. NMR analysis suggested that cinnamaldehyde was mostly physically incorporated into the hydrophilic chitosan films. Spin-coating can be used to incorporate and deliver high loads of cinnamaldehyde from ultrathin chitosan films (Rieger, Eagan, & Schiffman, 2015). These natural bioactive films show potential for use as bioactive materials in food packaging.

20.6 Inorganic Particles-Based Delivery Systems Inorganic particles have been extensively investigated over the past few decades. Several metallic and metal oxide nanoparticles are recognized by their antimicrobial activities, but growing evidence indicates the potential of inorganic nanoparticles as effective carriers for multiple types of antimicrobial substances. The potential of silver nanoparticles as an important antimicrobial agent has been demonstrated by several studies, which showed their usefulness to combat different infectious diseases and can

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be used as novel nanomedicine (Brandelli, 2012; Rai, Yadav, & Gade, 2009). In addition, other metallic nanoparticles, including noble metal nanoparticles and metal oxide nanoparticles, have been also investigated for their antimicrobial potential and effectiveness as antimicrobial drug carriers (Rai, Ingle, Gupta, & Brandelli, 2015). Inorganic clay nanoparticles are layered aluminum phyllosilicates containing neutral or negatively charged layers. Nanoclays are often employed in the manufacture of nanocomposite materials, functioning as reinforcement materials in polymeric films and improving their functional properties. Some active molecules with specific dimensions can be entrapped into montmorillonite nanosheets or halloysite nanotubes (Unalan, Cerri, Marcuzzo, Cozzolino, & Farris, 2014). Aquasomes are described as a class of drug carriers based on three-layer organized nanostructures (Umashankar, Sachdeva, & Gulati, 2010). A ceramic core has the surface non-covalently modified by a carbohydrate, which is exposed to the adsorption of a hydrophilic drug. Disaccharides, such as trehalose and cellobiose, have been used for their protective properties against desiccation and to help maintain the structural integrity of the drug (Banerjee & Sen, 2018).

20.6.1 Preparation Methods The preparation of inorganic nanoparticles is completed by either chemical, physical, or biological synthesis. Biological synthesis has gained space because it is associated with green synthesis in which plant extracts or microorganisms are used and, consequently, less toxic processes with minimum release of hazardous materials into the environment. The chemical reduction of metal ions is the most common and easy method for the preparation of the metallic nanoparticles. The chemical transformation of the metal ions into metal nanoparticles can be achieved using wet chemical synthesis, a photochemical process, by employing liquid crystal, polymer templates, or solution-based methodologies (Chouhan, 2018; Lee & Jun, 2019). Usually, the chemical synthesis process of metal nanoparticles in solution employs the following three main components: (a) metal precursors (often inorganic salts like AgNO3 AgCl, AgClO4), (b) reducing agents (e.g., NaBH4, N,N-dimethylformamide, N2H4, formaldehyde), and (c) stabilizing/capping agents (e.g., citrate, 2-mercapto ethanol, polyols). The different reducing power of different reductants may have a significant influence in determining the final shape of nanoparticles (Chouhan, 2018). The synthesis of metallic nanoparticles by green methods is carried out with a salt that has the metal and the plant extract or microorganism (Singh, Kim, Zhang, & Yang, 2016). In general, these methods generate heterogeneous nanoparticles with low yield. The optimization of parameters is done by the control of time, mixing rate, temperature, pH, and aeration. After optimization, stable and homogeneous nanoparticles with good performance are often obtained. By manipulating the parameters of the process, it is possible to control of shape and morphology of the nanoparticles (Figure 20.7). For example, pear extract results in triangular and hexagonal AuNPs at alkaline pH, but these do not form under acidic pH conditions (Ghodake, Deshpande, Lee, & Jin, 2010). There are numerous synthesis methods reported that allow precise control of nanoparticle size (Singh et al., 2016), specially for the synthesis of Ag and AuNPs. For example, a fine control of the size of the nanoparticles was achieved through the control of pH, temperature, and concentration of Ag nitrate, citrate, and borohydride (Figure 20.8). The synthesis was carried out in two stages, the first reduction at 60°C and the second one at 90°C. With the adjustment of synthesis parameters homogeneous AgNPs ranging from 5 to 100 nm were achieved (Agnihotri, Mukherji, & Mukherji, 2014).

20.6.2 Applications The potential applications of metal nanoparticles have been described in many areas, including the biomedical, pharmaceutical, agriculture, environmental, and chemical fields. They can be used as drug carriers, imaging devices, diagnostics, cosmetics, optical devices, and sensors. Silver nanoparticles are known for their antimicrobial activity and have been used in applications to eliminate microorganisms in air filters, water, and medicine. The Ag+ ions are effective at millimolar concentrations while the Ag nanoparticles in the nanomolar range, showing that they are much more effective antimicrobials (Rai et al., 2009). In one antimicrobial application study, a combination of Ag nanoparticles and lysozyme

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FIGURE 20.7  Different approaches available for the biological synthesis of metal nanoparticles.

was produced as a colloidal suspension to cover medical instrumentation. These nanocomposite materials showed antimicrobial activity against various bacteria, reducing the viability of S. aureus, Bacillus anthracis, Klebsiella pneumoniae, and Acinetobacter baylyi by >1.5 log10-cycles in up to 3 h (Eby,

FIGURE 20.8  Schematic representation of size-controlled silver nanoparticles synthesized by employing the coreduction approach. Reproduced from Agnihotri et al. (2014), under the terms of the Creative Commons Attribution 3.0 Unported License (http​:/​/cr​​eativ​​ecomm​​ons​.o​​rg​/ li​​cence​​​s​/ by/​​3​.0/)​.

Luckarift, & Johnson, 2009). The modification of silver and other metallic nanoparticles with biopolymers may facilitate their use for different applications such as catalysis, optics, drug delivery, and antimicrobial activity. In particular, improved solubility and stability can be useful for antimicrobial delivery systems (Rai et al., 2015). Gold nanoparticles (NPs) are recognized for their antimicrobial activity and as a drug carrier and diagnostic tools. Additionally, they are characterized by a high electron density and strong optical absorption (Daniel & Astruc, 2004). The development of gold NPs decorated with biomolecules has been conducted by surface modification. Peptides with cellular internalization signaling such as TAT (transactivator of transcription, rich in Lys and Arg) or NLS (nuclear localization sequence) have been adsorbed with success onto gold NPs. The interaction of biomolecules occurs by chemosorption, electrostatic interactions,

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and hydrophobic interactions. Gold NPs have a very high affinity for thiol groups, with covalent bondtype energy of approximately 200 kJ/mol (Sperling & Parak, 2010). Magnetic and metal oxide nanoparticles are also relevant as delivery systems and sensing tools. These nanoparticles have been used for targeted drug delivery and diagnostic methods. Magnetic NPs can be visualized by MRI, conducted by a magnetic field, and heated by a magnetic field for drug release (Arruebo, Fernández-Pacheco, Ibarra, & Santamaría, 2007). Inorganic nanoparticles based on clay minerals or silica not only have been extensively employed as nanofillers in composite packaging, but also these materials could be used as carriers for antimicrobial substances. Montomorillonite and halloysite nanoclays have been successfully evaluated as carriers for the bacteriocins nisin and pediocin (Meira, Zehetmeyer, Werner, & Brandelli, 2017). In addition, nisin physically adsorbed or covalently attached to silica nanoparticles showed antimicrobial activity against S. aureus (Behzadi et al., 2018).

20.7 Important Properties for Antimicrobial Delivery Systems and Methods of Characterization 20.7.1 Antimicrobial Loading and Encapsulation Efficiency The entrapment efficiency provides an indication of the amount of antimicrobial drug that is successfully entrapped/adsorbed into the nanoparticles. The loading capacity helps one to find the effective drug content of nanoparticles after their separation from the medium. The encapsulation efficiency and the actual loading capacity expressed in percentage are calculated according to Equations 20.1 and 20.2, respectively.

Encapsulation Efficiency ( % ) =



Loading capacity ( % ) =

Drug added - Free ( untrapped ) drug Drug added

(20.1)

Entrapped drug weight ´1100 (20.2) Nanoparticles weight

For example, if the encapsulation efficiency is 40%, it means that 40% of the drug included in the formulation is entrapped into the nanoparticles. If the loading capacity is 40%, it means that 40% of the nanoparticles weight is composed of the drug, i.e., each 1 mg nanoparticles contains 0.4 mg drug. Consequently, the selection of an encapsulation method and the encapsulant materials are usually driven to achieve a high encapsulation efficiency in order to minimize antimicrobial drug and encapsulant materials wastage, while loading capacity can still be low as it is dependent on the molecular weight of the encapsulant materials (Tao, Hill, Peng, & Gomes, 2014). The encapsulation efficiency of the delivery systems depends upon different factors associated with the process parameters of the preparation method (e.g., stirring rate, ultrasound energy, temperature, rate of solvent removal, drying process type) and the formulation, such as concentration of the components, solubility of polymer in solvent, drug–polymer interaction, and miscibility of components, among others (Hill et al., 2013b; Loquercio et al., 2015; Oliveira, Angonese, Ferreira, & Gomes, 2017a; Oliveira, Mezzomo, Gomes, & Ferreira, 2017b; Ozdemir et al., 2018; Pereira et al., 2015; Pereira et al., 2018; Pola et al., 2019). These parameters have been mostly studied for polymeric nanoparticles (Vauthier & Bouchemal, 2009). A low encapsulation efficiency has been related with slow solidification of particles, which can be associated with a high solubility of the polymer in the organic solvent, low solubility of the organic solvent in water, low concentration of the polymer, high dispersed phase to continuous phase (DP/CP) ratio, and/or slow solvent removal rate. By contrast, a high encapsulation efficiency is often associated with fast solidification of particles, related with low solubility of the polymer in the organic solvent, high solubility of the organic solvent in water, high concentration of the polymer, low DP/CP ratio, and fast solvent removal rate (Jyothi et al., 2010).

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20.7.2 Size, Zeta Potential, and Morphology The particle size is a key property of any delivery system since it is related to the surface to volume ratio and may influence the release kinetics of the payload release of the active compound. Several methods have been used for the determination of particle size in nanotechnology, including light scattering techniques and microscopy methods (Fairhurst & Weiner, 2018). Particle size determination by dynamic light scattering (DLS) has been widely used. This method is based on measuring light scattering intensity fluctuations, which are time-related and occur around a mean intensity value, being caused by the particles that are moving in the suspension under the influence of Brownian motion. The intensity fluctuation recorded is temporally correlated with its own delayed value, resulting in a decaying intensity autocorrelation function from which the translational diffusion coefficient can be determined (Stetefeld, McKenna, & Patel, 2016). With the optical properties of the particles, equivalent volume- and number-weighted distributions, a hydrodynamic equivalent spherical diameter is obtained. As DLS is a widely applied method, commercial DLS instruments are relatively accessible at laboratories. This apparatus is equipped with a red light-emitting He–Ne laser, and the scattered light is collected by a photodiode detector located at a backward scattering angle of 173° (Ruf, 1993). In particle-tracking analysis (PTA), also known as nanoparticle-tracking analysis (NTA), individual nanoparticles that are moving under Brownian motion in suspension are tracked. The sample is irradiated by a laser and a video of the light scattered by the particles is recorded by a light-sensitive camera through a magnifying objective. From this video, the Brownian motion of the particles is reconstructed and the translational diffusion constant, and lastly, the size is inferred from the particle paths. In order to distinguish the individual particles, the sample must be diluted to ensure the mean particle distance is larger than the diffraction limit of the microscope. Thus, the measurements are performed at a much higher dilution (around 1500×) than other methods (Filipe, Hawe, & Jiskoot, 2010; Gross, Sayle, Karow, Bakowsky, & Garidel, 2016). The centrifugal liquid sedimentation (CLS) method measures the sedimentation time of nanoparticles under increased gravitational forces and uses the sedimentation time to estimate the supposed Stokes particle diameter. Before the measurements, a transparent rotating disc is partially filled with a sequence of liquids of decreasing density, creating a density gradient. Then, a few microliters of nanoparticle suspension are injected into the center of the rotating disc. If the nanoparticles have a higher density than the density gradient, they will sediment moving radially to the outer edge of the disc. The sedimentation time is monitored by a detector near the outer edge, which records the loss of light intensity of a laser beam passing through the disc (Langevin et al., 2018). Small-angle X-ray scattering (SAXS) is a method that estimates the angular distribution of an X-ray beam scattered off the suspended particles in the forward direction under small angles. The scattering contrast is caused by electron density differences in the sample. The scattering data is given by the scattering intensity, l(q), as a function of the momentum transfer by the Equation 20.3:

q ( q) =

4p sin q (20.3) l

where λ is the wavelength of the X-ray beam and 𝜃 is half of the scattering angle. In the case of monodispersed particle suspensions, the scattering curve I(q) shows noticeable fluctuations, which depend on the particle diameter and can be evaluated by fitting the scattered intensity with a mathematic model. Consequently, this analysis can provide the size distribution of nanoparticle suspensions (Sakurai, 2017). Information on the aggregation and physical state of the nanoparticles is relevant to define the most appropriate method for particle size determination. If the particles are suspended in a stable medium and have not formed aggregates, either DLS or transmission electron microscopy (TEM) would provide good single particle size information. If the nanoparticles exist as a dry powder and formed aggregates, then a reasonable single particle size measurement may be obtained either by TEM or by calculating an average particle diameter using a measured Brunauer–Emmett–Teller (BET) surface area (Fairhurst & Weiner, 2018). It is important to mention that the eventual difference between calculated BET surface area value

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and TEM results is due to a small amount of surface area being uncounted due to the particle aggregates. In the case of measuring aggregate particle size, a combination of microscopy and light scattering methods, either dynamic or static light scattering, would be appropriate. Since a significant amount of energy is required to disperse the aggregate nanoparticle materials, single particle size is not obtainable using a DLS technique (Ponce, Mejía-Rosales, & José-Yacamán, 2012). Surface charge of the nanoparticles plays a key role in the preparation of a stable particle suspension as well as on the selection of substrates and surfaces for particle distribution (Guerrini, Alvarez-Puebla, & Pazos-Perez, 2018). Surface electrical properties of the particle and the substrate can be expressed by their zeta potential values. Thus, the measurement of zeta potential has become critical for delivery system research. However, the interpretation of nanoparticle electrophoretic mobility, especially that of ligand-coated particles, can be an intricate task. Despite the inherent complexity of the data, key concepts from colloidal science can help to refine valuable information from electrophoretic analysis. Zeta potential can be influenced by the ionic strength and pH value of the bulk solution, and by the functional groups associated with the particle surface (Clogston & Patri, 2011). Besides the characterization of particle size, microscopy techniques may provide useful information about particle structure and morphology. Microscopic methods, both TEM and scanning electron microscopy (SEM), are possibly the most popular for nanoparticle imaging analysis. The similarities in operation and data output may suggest that these techniques are often treated as interchangeable, but in fact the applications to which they are best suited differ considerably, as do their respective magnification ranges and image resolution capabilities. Electron microscopy works by bombarding a sample with a stream of electrons and monitoring the resulting transmission (TEM) or scattering (SEM) effects (Egerton, 2005; Yao & Wang, 2005). These electrons are detected and converted into magnified images. This information is often processed by an image analysis software, generating particle size data for individual particles, size distributions for the entire dispersion, and several shape and morphological parameters (Woehrle, Hutchison, Özkar, & Finke, 2006). The main difference in data output between SEM and TEM is the way in which the images are resolved. While SEM produces three-dimensional images of particles, TEM produces two-dimensional images at higher magnifications that require further interpretation. Despite that images rendered by TEM are two-dimensional, this technique is capable of delivering much higher resolution (approximately three orders of magnitude higher). Surface and shape analysis can be better assessed by SEM, particularly in applications such as quality control of colloidal nanoparticles, or for evaluating surfaces and structural features of nano-sized powdered materials. As TEM can provide a higher resolution, it may be better suited for particle size analysis in the nano- to sub-nanoscale region (Yao & Wang, 2005). Nanoparticle analysis can also be performed using atomic force microscopy (AFM), which is one of the most popular scanning probe microscopy methods. The operating principle of AFM is based on a functionally sharp tip (probe) placed at the end of a flexible cantilever beam that is brought into physical contact with the sample surface. The cantilever deflects in proportion to the force of interaction. A piezoelectric transducer allows positioning and scanning via the probe in three dimensions over the sample with very precise movements; a response system detects the interaction of the probe with the sample. Scanning across the surface, the probe follows the bumps and grooves formed by the atoms on the surface. A topography of the surface can be generated by monitoring the deflections of the flexible cantilever beam (Henry, 2005). The size of isolated nanoparticles of spherical shape deposited on a perfectly flat substrate can be determined from the AFM image by measuring the nanoparticle image height. This number is not biased by probe-sample convolution effects and can give precise nanoparticle size results. However, particle size analysis is not so straightforward if the nanoparticles are deposited on rough or curved surfaces, and therefore it may be difficult to implement particle analysis in AFM imageprocessing software. Another effect strongly influencing the AFM analysis of nanoparticles is particle agglomeration and self-ordering on the substrate. In addition, the interaction of nanoparticles with surfaces and the AFM probe can be evaluated through probe tip functionalization (Ong & Sokolov, 2007). Chemically functionalized tips gain selectivity and sensitivity to perform interactions at the molecular level. Besides, the magnitude of these interactions can be analyzed to allow the detailed mapping of surface chemistry at nanoscale resolution (Fritz, 2008; Garcı ́a & Pérez, 2002).

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20.7.3 Controlled-/Triggered-Release Properties Controlled release of antimicrobial substances can be achieved by their incorporation either in dissolved or dispersed form of the above-discussed delivery systems. The systems can be customized according to the desired final release profile and/or targeted organism(s) to deliver the antimicrobial at a predictable rate, in pulsatile manner, in the form of extended, triggered, and target-responsive forms. The investigation of novel antimicrobial formulations and delivery systems must achieve the essential understanding of the drug release mechanism by applying transport- and time-dependent kinetic equations. Several products based on controlled release are available, mostly for clinical applications, but limited explanation of their releasing mechanisms is provided. Mechanisms of controlled drug release can be roughly classified based on the release of active agents from delivery systems, namely: diffusion, degradation, swelling followed by diffusion, and active efflux (Gomes et al., 2011; Hill et al., 2013a; Oliveira et al., 2017a). However, ad libitum antimicrobial-releasing systems often exhibit a burst release followed by a long slow release with sub-lethal concentrations, which could lead to antimicrobial resistance in bacterial strains. To circumvent this, delivery systems that respond to an external stimulus such as pH, temperature, magnetic field, light, and electric pulses have been designed. For example, stimuli-responsive polymers could provide methods to trigger antimicrobial release when foods are stored at a temperature that promotes microbial growth, or spoilage occurs, providing a shift in pH that raises the likelihood of pathogen growth. Antimicrobials would be released when they are needed the most, rather than being metabolized or degraded before they are in contact with pathogenic bacteria. Dual-stimuli responsive delivery systems have been demonstrated that combine pH- and temperature-responsive polymers via crosslinking, for hydrophobic antimicrobial compounds delivery, improving their antimicrobial effects (Hill et al., 2013a). In this study, for the same pH and temperature conditions, the chitosan-co-PNIPAAM nanoparticles were significantly more potent bacterial inhibitors against both pathogens (S. enterica and L. monocytogenes) and also exhibited a faster cinnamon bark extract release over time as well as slightly higher entrapment efficiency (Hill et al., 2013a). A recent study developed and optimized pH-responsive nanoparticles based on PLGA-chitosan for triggered release of antimicrobials (Pola et al., 2019). Optimized nanoparticles characterization indicated a satisfactory TCIN encapsulation (33.20 ± 0.85%), spherical shape, pH-responsive controlled release, with faster release in the presence of CHIT at low pH, and enhanced antimicrobial activity against both pathogens (Pola et al., 2019) (Figure 20.9). One attractive type of such delivery systems responds to the presence of adhering bacteria to release antimicrobial content. Recently, a series of such “self-defensive”, bacterially triggered coatings has been developed, responding to bacterial presence only when and where needed (Cado et al., 2013; Komnatnyy, Chiang, Tolker-Nielsen, Givskov, & Nielsen, 2014; Pavlukhina et al., 2014; Zhuk et al., 2014). Both enzymes (Cado et al., 2013; Francesko et al., 2016), and acids (Andersen & von Meyenburg, 1980; Smith, 1991) excreted by bacteria have been used as triggers for antimicrobial release to combat adhering bacteria. In all cases, the pH-responsive “activation” of the delivery systems was clearly demonstrated when the coating was exposed to differing strains of S. aureus or E. coli, both known to acidify the medium in which they grow as a result of secretion of lactic and acetic acid, respectively (Andersen & von Meyenburg, 1980; Smith, 1991). Albright et al. (2017) recently demonstrated pH- and otherwise bacterially triggered polymer delivery systems that were directly linked to bacterial presence and pH activation through localized acidification. In this study, localized interfacial acidification induced by adhering bacteria was demonstrated and correlated with pH-triggered release of gentamicin and polymyxin B, by making use of antibiotic-loaded poly(metacrylic acid) (PMAA) hydrogel coatings (Albright et al., 2017). All of these mechanisms employ the physical transformation of constituents of the system when they are placed into a biological environment. Although chemically driven drug delivery systems are feasible, they involve chemical modifications with active agents and carrier vehicles that need regulatory approval. In addition, adequate toxicology and safety evaluation including clinical trials are required before approval for final application. Therefore, simple delivery systems with approved active compounds and excipients are often utilized in the preparation of the controlled antimicrobial delivery systems used for food and biomedical applications.

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FIGURE 20.9  Schematic diagram of the PLGA/trans-cinnamaldehyde (TCIN)/chitosan (CHIT) nanoparticles development with pH-responsive behavior for the triggered release of antimicrobials. Reproduced from Pola et al. (2019), with permission from Elsevier.

The widespread development of controlled-release systems intended for numerous applications makes it quite complicated to classify and describe each system with its respective mechanism. Release kinetics for the majority of the antimicrobial delivery systems currently investigated can be explained by diffusion or empirical models (Benita, 1996). These systems can be classified as: (a) monolithic devices where the antimicrobial agent is dispersed in a polymer matrix and its release is controlled by diffusion along the matrix, and (b) reservoir systems (membrane-controlled devices), where the antimicrobial agent is contained in a core that is surrounded by a thin polymer membrane. Release to the surrounding environment occurs by diffusion through the rate-controlling membrane. Monolithic devices and reservoir systems have recently been employed and delivery systems based on this scheme have proven their efficacy in model foods (Huynh & Lee, 2015). There are also chemically controlled release approaches that include matrix erosion, combined erosion with diffusion, desorption of adsorbed drug, and drug covalently attached to a polymer. Moreover, externally controlled devices are also available that include magnetic microspheres, antibody recognition, and electrically powered microelectromechanical systems (MEMs) that are controlled via wireless communication to release the drug (Benita, 1996; Nisar, Afzulpurkar, Mahaisavariya, & Tuantranont, 2008; Razzacki, Thwar, Yang, Ugaz, & Burns, 2004; Tsai & Sue, 2007). Chemically and externally controlled-release approaches have been widely studied and applied in the biomedical field, while less explored in food safety applications, which could be associated with the inherent cost of these approaches.

20.7.4 Antimicrobial Delivery System Efficacy Evaluation The initial in vitro confirmation of antimicrobial activity for an antimicrobial delivery system is often carried out by agar diffusion or broth dilution methods based on visualization of inhibition zones of microbial growth and optical density changes over time during incubation, respectively (Brandelli & Taylor, 2015). The main advantages of agar diffusion assays over other methods are the low cost, simplicity, facilitating the testing of large numbers of microorganisms and antimicrobial agents, and the easy interpretation of results (Balouiri, Sadiki, & Ibnsouda, 2016). The use of standard methods is recommended for a quantitative measurement of the inhibitory activity of antimicrobial agents. Many approved guidelines for dilution antimicrobial susceptibility testing of microorganisms are available, and the most

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recognized standards are supplied by the Clinical and Laboratory Standards Institute (CLSI, 2018) and the European Committee on Antimicrobial Susceptibility Testing (EUCAST, 2003). Dilution methods are most appropriate for the determination of MIC values, since they offer the possibility to estimate the concentration of the tested antimicrobial agent in the agar (agar dilution) or broth medium (macro- or micro-dilution). The MIC is defined as the lowest concentration of an antimicrobial agent that completely inhibits growth of the microorganism in tubes or microdilution wells as detected by the absence of visible turbidity. The determination of minimum bactericidal concentration (MBC) or minimum fungicidal concentration (MFC) is the most common method for the estimation of bactericidal or fungicidal activity. The MBC is defined as the lowest concentration of antimicrobial agent needed to kill 99.9% of the final inoculum after incubation for 24 h under standardized conditions (Balouiri et al., 2016). Time-kill testing is most suitable for determining the bactericidal effects of an antimicrobial agent. This test is a robust methodology for obtaining information about the dynamic interaction between the antimicrobial substance and the targeted microbial organism(s). Time-kill testing provides useful information about time- or concentration-dependent antimicrobial effects (Tsuji, Yang, Forrest, Kelchlin, & Smith, 2008). This assay is more laborious than MBC testing and is generally performed in research settings when a new antimicrobial is being evaluated, to confirm unusual MBC results, or as an effort to explain treatment failure when bactericidal action is mandatory (Brandelli & Taylor, 2015). Methods based on colorimetric or fluorescent probes, associated with active microbial metabolism, are also used to test antimicrobial susceptibility. The tetrazolium/formazan (2,3,5-triphenyltetrazolium chloride, TTC) couple is a particular redox system acting as proton acceptor or oxidant. This redox system has been widely employed in different sectors of the biological sciences. In the presence of microorganisms, TTC is reduced to red formazan, which is directly proportional to the amount of viable active cells. Thus, the TTC method is considered as a comparatively fast method for evaluating the antibacterial activity of antimicrobial agents. In TTC assay, less than 12 h was required to reach susceptibility results and fewer bacterial counts are required (Moussa, Tayel, Al-Hassan, & Farouk, 2013). Similar tetrazolium salts such as MTT and XTT are widely used to estimate cell viability and proliferation in toxicity assays of nanoparticles. However, this method can be influenced by some conditions such as acidic pH and nanomaterials. Some nanoparticles can interfere in the results by inducing the formation of reactive oxygen species, which can lead to inaccurate evaluation (Wang, Yu, & Wickliffe, 2011). Adenosine triphosphate (ATP) is a key energy-storing molecule in all living cells, and it is present in relatively constant amounts in a cell. Consequently, ATP quantification has been used to estimate the microbial population in a sample, and the ATP bioluminescence assay was developed to measure the ATP produced by bacteria or fungi. This assay is based on the luciferase-catalyzed reaction of luciferin and ATP, thus quantifying existing ATP, indicating the presence of metabolically active cells (Ivancic et al., 2008). In the presence of ATP, luciferase converts D-luciferin to oxyluciferin that generates light. The amount of the emitted light is measured by a luminometer and expressed as relative light unit (RLU), which can be converted into RLU per mole of ATP. Thus, a linear relationship between cell viability and measured luminescence can be obtained.

20.7.5 Comparison among Delivery Systems – Advantages/Disadvantages A comparison among different delivery systems is important in order to provide information and assist the selection of antimicrobial delivery systems, and consequently for these systems to fully penetrate the food safety market and be used in industry (Hill et al., 2017). Table 20.1 shows a summary of advantages and disadvantages based on the important performance characteristics for delivery systems aforementioned, namely: encapsulation efficiency, loading capacity, size and size distribution, and controlled-/triggered-release ability, and challenges such as storage stability. Some important differences among delivery systems that are not summarized in Table 20.1 are cost, ease of application, i.e., embedded in packaging or as a food additive, and toxicity (these parameters vary too considerably to generalize in a table format). Each of these is critical for the adoption of a delivery system by the food industry.

• • • • •

• • • • • • • • •

Solid lipid nanoparticles

Polymer-based Dendrimers

Liposomes

• • • • • • • • • • • • • • • •

Nano-emulsion

Delivery System

High level of control over size (usually ranging from 1.1 nm to 9 nm) Hydrophilic and hydrophobic drugs Controllable surface functionality High surface/size ratio Intracellular delivery capability High reactivity Can be modified to acquire targeted drug delivery Charged end groups – polyvalence Intrinsic antimicrobial activity

Compatible with drugs that are extremely insoluble in water Hydrophilic and lipophilic drugs Ideal for topical and oral applications Biocompatible Organic-solvent free

Versatile: hydrophilic, hydrophobic, amphiphilic molecules Controlled release Visually transparent 20 to 500 nm diameter Superior stability compared to emulsions High surface area – improved delivery Solubility of hydrophobic antimicrobial compounds Reduced toxicity of encapsulated molecule Transdermal, ocular, and parenteral administration Ease of production process Versatile: hydrophilic, hydrophobic, amphiphilic molecules Can be tailored to site-specific delivery Controlled release Low toxicity Reduce toxicity of encapsulated materials Biocompatibility

Advantages

• • • • • •

• • • • • • • • • • • •

• • • • • • • •

Disadvantages

(Continued)

Potential toxicity Cytotoxicity Hemolysis Promote cell lysis Limited efficiency with large molecules Functionalization is required to reduced toxicity and increase entrapment efficiency

Polydispersed size distribution High-cost production Low loading capacity Fast release rates Low storage stability Susceptible to oxidation Limited solubility Limited loading capacity Low entrapment efficiency Premature release Instability during storage Potential toxicity for parenteral application

Stabilizers and surfactants are needed Low kinetic/thermodynamic stability Susceptibility to hydrolysis Use of organic solvents Organic solvents residues Toxicity issues Weak physical stability Low resistance to changes in temperature, pH, or ionic strength

Summary of Advantages and Disadvantages Based on Key Performance Characteristics of Delivery Systems*

TABLE 20.1

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• • • • • • • • • • • • •

• • • • • • • • •

Use of both synthetic and natural polymers High surface area High porosity Morphology can be controlled by the concentration of polymer Amendable to surface modification methods High mechanical strength Cost-effective Allows application of intrinsic antimicrobial polymers Great capacity of encapsulation of vast diversity of hydrophobic and hydrophilic molecules High entrapment efficiency Controlled-release and triggered-response capability Long-lasting release Cost-effective Particle configuration increases the stability of the bioactive compound Use of intrinsic antimicrobial polymers Several production methods Intrinsic antimicrobial activity Tunable to controlled release and triggered response Control of nanoparticle size (usually between 5 and 100 nm) High surface area Easily functionalized Stable (AuNP)

Advantages

• • • • • • •

• • • • • • •

Potential toxicity Limited colloidal stability (AgNP) Require photoactivation (TiO2NP) Limited drug loading Require purification steps Use of strong chemical reagents Non-biodegradable

Use of organic solvents Burst release Polydispersed size distribution Aggregation/flocculation problems Susceptible to pH and temperature changes Require purification steps Batch-to-batch variation

• Depends on temperature, viscosity, surface tension, intensity of the electric field • Small particles can be released during degradation • Release of toxic byproducts during degradation • Limited application

Disadvantages

* Compiled from the following references: Abbasi et al., 2014; Borel & Sabliov, 2014; Claire et al., 2015; Kesharwani, Jain, & Jain, 2014; Kriegel, Kit, McClements, & Weiss, 2010; Kumari, Yadav, & Yadav, 2010; Liu & Fréchet, 1999; Lovelyn & Attama, 2011; Malheiros, Daroit, & Brandelli, 2010a; Miller, Wang, Benicewicz, & Decho, 2015; Mozafari, Johnson, Hatziantoniou, & Demetzos, 2008; Otoni, Pontes, Medeiros, & Soares, 2014; Rai, Ingle, Gupta, & Brandelli, 2015; Sherje, Jadhav, Dravyakar, & Kadam, 2018; Simões et al., 2017; Svenson, 2009; Weber, Zimmer, & Pardeike, 2014; Wei, 2012; Zhang, Pornpattananangkul, Hu, & Huang, 2010.

Inorganic particles

Nanoparticles

Nanofibers

Delivery System

Summary of Advantages and Disadvantages Based on Key Performance Characteristics of Delivery Systems*

TABLE 20.1 (CONTINUED)

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20.8 Conclusions and Future Trends Delivery systems such as nano-emulsions, nanoliposomes, solid-liquid polymeric and inorganic nanoparticles, and nanofibers have been used to enhance the antimicrobial efficacy and utilization of natural antimicrobials in foods. Their ability to control-/trigger-release the antimicrobial compound and to reduce the interactions of the antimicrobial payload with food components in the food matrix provides great promise for food safety applications and their use in the replacement of synthetic preservatives. While there are many materials that can be used for the preparation of delivery systems in food matrices, the choice of an appropriate encapsulant and preparation method is driven by cost, availability of materials, and performance characteristics (including encapsulation efficiency, loading capacity, size and size distribution, controlled-/triggered-release ability, and storage stability). Future research should focus on developing delivery systems that are based on all-natural components (i.e., encapsulant and antimicrobial compounds) and also on exploring different preparation methods to optimize the antimicrobial activity of active compounds over the entire food product shelf-life. In addition, the discovery of new natural antimicrobial compounds and their combination that could be applied in foods and also understanding of their interaction with the food matrix would be valuable. Furthermore, the development of triggered mechanisms of antimicrobial release in food systems would enable enhancement of antimicrobial efficacy. This approach enables antimicrobial release-on-demand based on the presence of targeted microorganism(s) in a highly localized way, consequently reducing unwanted interaction between the antimicrobial and the food matrix, which dilutes antimicrobial concentration, and which may subsequently induce antimicrobial adaptation by the microbe. Finally, the development of effective antimicrobial delivery systems in combination with non-thermal and thermal food processing such as high pressure, plasma treatment, microwave, irradiation, high-temperature/short-time pasteurization, among others, to promote a synergistic response for microbial safety protection would benefit the food industry. There are many different needs in the food chain associated with food safety and food security. The scientific community continues to contribute and develop a large library of antimicrobial delivery systems that have the potential to be used to ensure food safety. These delivery systems must be beneficial to consumers and the food industry, while complying with regulatory agencies’ requirements in order to be adopted. It is time to apply these different delivery system technologies to the benefit of society.

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21 Hurdle Technology – or Is It? Multifactorial Food Preservation for the Twenty-First Century J. David Legan and Jairus R. D. David CONTENTS 21.1 Introduction ................................................................................................................................ 695 21.2 Benefits of the Hurdle Concept................................................................................................... 696 21.3 Drawbacks of the Hurdle Concept.............................................................................................. 697 21.4 Alternative “Multifactorial” Metaphors...................................................................................... 697 21.4.1 The Swiss Cheese Model.............................................................................................. 697 21.4.2 The Pole Vault: Hurdles Upended................................................................................ 699 21.5 Risks............................................................................................................................................ 699 21.6 Microbial Modeling to Quantify Factors and Control Risk.........................................................701 21.6.1 Pasteurized Processed Cheese Spread and Control of Clostridium botulinum............701 21.6.2 Ready-to-Eat Meat Products and Listeria Control....................................................... 703 21.7 Multifactorial Preservation in Action......................................................................................... 704 21.7.1 Market Examples of Multifactorial Products............................................................... 706 21.7.1.1 Co-Extruded Cereal Bar with Fruit Core................................................... 706 21.7.1.2 Optimization of “Non-C. botulinum” Thermophilic Cook with Use of Nisin in Canned Foods................................................................................ 707 21.7.1.3 Bottled Water.............................................................................................. 708 21.7.1.4 Refrigerated Liquid Egg Containing Vegetable Blend............................... 708 21.7.1.5 High-Pressure Processed Products............................................................. 709 21.8 Future Development of Novel Opportunities...............................................................................710 21.9 Concluding Remarks....................................................................................................................711 21.10 Acknowledgments........................................................................................................................711 21.11 Disclaimer....................................................................................................................................712 References................................................................................................................................................712

21.1 Introduction For 30 years or more, consumers have demanded foods that are less severely processed, free from “chemical preservatives” (represented in Europe as absence of “E” numbers), formulated with “natural” ingredients to be more natural and healthier, and still convenient. While it is not always possible to deliver on those expectations, even attempting to do so requires detailed knowledge of food and ingredient technology and a sound understanding of food preservation. Food preservation is conceptually very simple: we must prevent microbes from growing to the point where they make the food unsafe or unpalatable. In principle, we can do this by killing any microbes and preventing them from re-contaminating the food or by reducing their growth rate to the point that they are not able to achieve harmful concentrations during the time for which we want the food to be palatable. At the same time, we must prevent chemistry and physics from reducing the food quality to an unacceptable

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extent. We have many more tools available for controlling the microbiology than we do for controlling the chemistry and physics, and in this chapter, we will explore, with examples, the benefits of combining multiple tools. But first, let us briefly review how food science arrived at the idea of “hurdle technology.” Traditional methods of food preservation, including drying, salting, smoking, fermenting to create acid or alcohol, etc., presumably “evolved” through an extended process of empirical observation and modification before the existence of microorganisms was even known. “Modern” methods of food preservation began with Nicolas Appert’s invention of the extended preservation of food by heating it in a sealed glass jar (Garcia and Adrian, 2009) and developed into the technology that today we recognize as canning. Later, Louis Pasteur’s discovery that wine could be prevented from spoilage by mild heating (ValleryRadot, 1919). This provided an example of a milder preservation process that would come to be recognized by the name “pasteurization.” The advent of mechanical refrigeration brought us reliable chilling and freezing, and the power of chemistry also brought synthetic organic acid preservatives at much lower cost than the same compounds produced naturally. As these technologies advanced through scientific research, the emphasis was naturally focused on each single innovation. Only a few of these technologies, such as thermal-sterilization, thermal-pasteurization, and irradiation (and more recently high-pressure process (HPP)-pasteurization), act by inactivating microorganisms. The other processes such as chilling, freezing, lowering aw, lowering pH, modified-atmosphere packaging, and the addition of antimicrobial preservatives act via inhibiting microorganisms to varying degrees. And as a final part of the preservation continuum, package integrity and hermetic seal restrict the access of microorganisms into the product. It is now well-known that use of these inhibitory factors in combination even at sub-lethal levels, and in lieu of extreme use of any single treatment, leads to improvements in desirable safety, quality, and nutritional attributes. This knowledge is critical to creating “premium” versions of existing products and developing new products, while, simultaneously, optimizing for microbial stability and food safety. This realization that multiple factors contribute to the preservation of a single food, and the association of that realization with the description of each individual factor as a “hurdle” can be attributed to Leistner and Rödel (1976) at a symposium on Intermediate Moisture Foods (IMF) in Weybridge, UK. These authors stated that The inhibition of micro-organisms in IMF does not solely depend on the aw, but the pH, Eh, t-value, F-value, preservative and the competitive microflora might be of importance too. How many of these “hurdles” and at what level they are needed for a sufficient inhibition of microorganisms during the desired storage period … depends not only on the types but also much on the number of organisms present.

The hurdle concept has been reviewed many times, from multiple angles (e.g. Khan et al., 2017; Leistner, 2000; Leistner and Gorris, 1995; Rahman, 2015; Singh and Shalini, 2016) as well as in earlier editions of this book, and the reader can find extensive information on multifactorial preservation systems in the research literature. Over time, the literature has more and more come to describe preservation based on multiple factors as “hurdle technology” (Leistner and Gorris, 1995; Chawla and Chander, 2004; Gupta et al., 2012; GarcíaGarcía et al., 2015; Moreira et al., 2017). As a pair of industry practitioners, the authors would argue that each “hurdle” may be the embodiment of a different technology and that the true power of the hurdle concept is in its call to understand all of the elements contributing to the preservation of a particular food, their relative contributions, and any interactions between them. This understanding can help us to develop preservation systems with the minimum impact on food quality for a particular amount of “preserving power.” The products developed using this principle may often be considered “minimally-processed products,” and hitherto have been considered “hurdle technology products.” In this review we have chosen to explicitly identify them as products of multifactorial preservation to recognize that they depend on a collection of technologies, and to make the connection with modern design principles, such as the use of multifactorial models, more intuitive.

21.2 Benefits of the Hurdle Concept The first benefit of the hurdle concept is simply that it requires us to understand all of the different factors contributing to the preservation of a particular product. When we do that, we realize that multiple

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FIGURE 21.1  “Traditional” illustration of the hurdles concept. The figure could be extended with additional hurdles or other hurdles could be substituted for those illustrated. (Modified from Scott, 1989.)

factors are involved in the preservation of most foods, even when we think primarily of a single factor as responsible for preservation. For example, with commercially sterile low-acid canned foods processed by heat we think of the thermal death profile as being the preserving factor, but preventing recontamination, and the climate of the area of distribution are also relevant. For example, for tropical distribution the intensity of the thermal process must be increased to eliminate thermophilic spores that might have been present in the raw materials. The second benefit is that once we understand the factors, we can determine the relative contribution of some or all of them, and design a preservation system optimized to provide the preserving power that we need with the lowest overall impact on the quality of the food. In principle, more barriers, each at “milder” intensity, can give the required safety and/or preservation with better perceived quality, particularly if there are any truly synergistic interactions between any of the factors.

21.3 Drawbacks of the Hurdle Concept The hurdle concept is not so much defined as, usually, illustrated in a “boiled eggs” diagram (Figure 21.1). This diagram illustrates that different factors (hurdles) may provide different amounts of “preserving power” shown by the height and/or width of the hurdle. It also indicates that the effort involved in overcoming the “preserving power” of one hurdle weakens the microbe so that it has less capacity to overcome the next hurdle. Eventually the microbe reaches a hurdle that it has been too weakened to overcome, and at that point preservation has been achieved. Many people, the authors included, do not find this model particularly intuitive or illustrative, possibly because we are familiar with the notion of hurdle racing, where athletes compete to see who can be the fastest to traverse a course obstructed by a number of hurdles. All hurdles are the same size and separated by a constant distance. The race winner is the one that traverses the course the fastest, and it is rare to see a runner fail to complete the course. Thus, there is immediate confusion when comparing our mental concept of a hurdle race (Figure 21.2) with the classical “hurdle technology” illustration (Figure 21.1). It may also in large part be because Figure 21.1 is unable to convey the possible interaction between a “gentler” inactivation process and multiple inhibitory factors, and combinations which may act additively and/or synergistically.

21.4 Alternative “Multifactorial” Metaphors 21.4.1 The Swiss Cheese Model In other areas of risk management, different models are used to illustrate the concept of adding multiple barriers to improve security. The Swiss cheese model of risk management and accident causation (Reason, 1990; Reason et al., 2001) (Figure 21.3) is used in many areas, for example health care (Perneger, 2005), aviation (Shorrock et al., 2006), and railroad safety (Underwood and

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FIGURE 21.2  Racing hurdles showing uniform height and separation: The question is not whether an athlete will clear them, but how fast?

FIGURE 21.3  The Swiss cheese model of accident causation. Each preservation factor, represented as a slice of Swiss cheese, is imperfect but product stability is secure provided that each barrier has a different failure mode. If there is a single failure mode for each barrier (the holes align) then the product will spoil or become unsafe. (Adapted from Reason et al., 2001.)

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Waterson, 2014). In this food-centered metaphor each preservation factor would be a slice of Swiss cheese. The presence of holes would indicate that the barrier does not control all risks. For example, one slice might represent a pasteurization step. A hole would indicate that the barrier is not complete against spore-formers. The next slice might indicate a reduced pH that restricts the growth of dangerous spore-formers but permits the growth of Alicyclobacillus, providing a spoilage risk. The next slice might represent purchasing specifications to ensure sourcing only of raw materials free from Alicyclobacillus spores, and so on. By gradually reducing the likelihood that multiple holes line up as additional barriers are added, the concept illustrates the benefits of multiple barriers, even if each may be incomplete by itself. Failure of the “final” barrier would clearly cause system failure, but note that failure of an earlier barrier may open a different path through the slices leading to failure but by a different mode.

21.4.2 The Pole Vault: Hurdles Upended The Swiss cheese model has considerable appeal, not least because a food is itself central to the metaphor. However, as we write, the field of food science has invested over 40 years (and how many tens of thousands of lectures?) in the hurdle concept. In an attempt to make the established metaphor more intuitive, we conceive it more as a high-jump or pole vault (Figure 21.4). Each new preservation factor or barrier (B1, B2, B3, etc.), causes “the bar to be raised.” The factors may contribute unequal additional heights but eventually enough factors accumulate such that the barrier cannot be overcome. During the competition, as the bar height is raised, the number of athletes able to clear the bar is progressively reduced. Think of this as progressively reducing the number of available failure modes. As the “preservation pressure” of additional factors increases, so too the number and types of organisms able to overcome the barrier and grow become smaller. At the extremes of preservation factors just before all growth is prevented there is usually only a single species able to overcome the pressure – the Olympic gold medalists of preservation resistance – e.g. as aw reduces, the last mold able to grow is Xeromyces bisporus, the most xerotolerant mold known. There are multiple potential microbial responses to consider once the bar is raised high enough to prevent the growth of any microbe (Figure 21.4). That organism may be injured to the point where it dies such that we do not need to consider it again, or it may be simply injured, able to persist in the food for either a short time, or a long time, until conditions permit its recovery, leading to failure of the preservation system. In presenting the Legan and David “Pole Vault” Model we aim to modify the established metaphor in a way that makes it more intuitive and more flexible; perhaps merely addressing our own mental challenges but, we hope, providing a service to food scientists and food microbiologists in general. However, we would like to keep the focus on the individual factors or barriers, since this is the level at which the system can be modeled and modified. We also want to emphasize that successful preservation increasingly relies on explicitly multifactorial preservation systems.

21.5 Risks In any preservation project, we must understand whether we are primarily concerned with mitigating safety risks, spoilage risks, or both. A thorough understanding of the intrinsic factors such as aw, pH, etc., helps us to determine the nature of the risk that we are managing, and detailed knowledge of the relationships between them allows us to understand how secure our preserving effect is in the event of typical variability in the magnitude of the individual factors. Developments in statistical modeling techniques and the ready availability of modeling software have allowed rapid progress in quantifying the impact of different factors on microbial growth within a food.

FIGURE 21.4  The Legan and David Pole Vault Model of preservation systems. Barriers (B1, B2, B3, and B4 alongside the poles) may be unequal in effects. Initially with only one factor, the bar can be easily cleared (A). As factors are added, it becomes progressively harder to overcome them (B, C), building to a point where the combined barrier (bar height) is too great to be overcome (D), and the ability of microorganisms to grow is prevented (E). Following failure to grow in the presence of multiple preservation factors, initially the microbes are injured (F). They might quickly recover (G), though this would suggest that the multifactorial preservation system is not yet robust enough. Alternatively, they might quickly die (I) or enter an extended period of quiescence and/or recovery (H1) from which they ultimately recover (H2), limiting product shelf life to an acceptable degree, or die (H3).

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21.6 Microbial Modeling to Quantify Factors and Control Risk 21.6.1 Pasteurized Processed Cheese Spread and Control of Clostridium botulinum Tanaka et al. (1986) explored factors affecting toxin production in pasteurized processed cheese spreads. In a series of experiments, they determined conditions permitting toxin formation by Clostridium botulinum over 42 weeks’ incubation at 30°C. They used a quadratic logistic regression model (not presented in their paper) to determine the boundary between toxin formation and non-toxic conditions and presented seven figures showing the boundary at different moisture contents. We present those boundaries compiled into a single contour plot in Figure 21.5 which compiles four factors: concentrations of NaCl and Na2HPO4, pH, and moisture content, all with three further factors fixed: time to end point, incubation temperature, and a pasteurization heat process. This work informed the safe formulation of pasteurized processed cheese spreads with multifactorial preservation for C. botulinum control and product safety assurance for many years. However, consumer tastes began to change, and lower fat formulations were demanded, and that created a need to review the basis of product safety. Many formulations of reduced-fat pasteurized processed cheese products were studied by Glass and Johnson (2004), including even more factors for the control of toxin formation by C. botulinum. Beginning with pasteurized processed cheese formulated using full-fat Cheddar, 30% reduced-fat Cheddar, or skim milk cheese and standardized to 59% moisture, pH 5.75, with 2.8% or 3.2% total salts, and 15% to 19% fat, they observed that toxin formation was delayed by 2 days in product formulated with skim milk cheese with 3.1% total salts. Next these authors compared the effect of fat levels and additional ingredients. They found that at similar fat levels (15.1–19.1%) the delay in toxin formation in skim milk pasteurized processed cheese was not statistically significant. However, when fat levels were < 1% in

FIGURE 21.5  Model from Tanaka et al. (1986) redrawn as a contour plot. Contours represent the combinations of % NaCl + Na2HPO4 and pH that lead to toxin production in 42 weeks at 30. °C at different moisture contents.

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skim milk pasteurized processed cheese, 13% in reduced-fat cheese, and 24% fat in full-fat cheese, toxin formation in the skim milk pasteurized processed cheese was delayed by 2 weeks relative to the other 2 formulations. Sodium lactate added at 1.5% significantly delayed toxin formation at all fat levels, though the delay increased as fat content decreased. Enzyme-modified cheese used as a flavor enhancer (1.5% addition) and monolaurin (0.05%) both delayed toxin formation in skim milk pasteurized processed cheese but not in the other formulations. So, this study identified three new formulation hurdles, adding to those previously noted by Tanaka et al. (1986), and identified some complex relationships between them, most notably that added lactate and monolaurin were more effective at low fat concentrations. Consumers again began to demand product changes, this time looking for lower sodium products in response to publicity about the relationship between sodium consumption and blood pressure and new US dietary guidelines recommending a maximum intake of 2300 mg of sodium per day (U.S. Department of Health and Human Services and U.S. Department of Agriculture, 2015). That created a new need to review the basis of product safety, this time with reduced-sodium formulations. A new model was created under the leadership of Glass et al. (2017). This determined probability and time to toxin formation over 56 weeks in a central composite design of 80 treatments, exploring the influence of seven factors: pH, moisture, Na2HPO4, NaCl, sorbate, fat, and fraction of replacement of sodium by potassium. The effects of pH, sorbic acid, moisture, Na2HPO4, and NaCl on toxin formation were all highly significant (P < 0.0001). Fat content and replacement of sodium by potassium were also significant, but less so (P = 0.0168 and P = 0.0526, respectively). Highly significant (P < 0.0001) pairwise interactions of sorbate with pH, and sorbate with moisture were identified along with a highly significant quadratic term for (Na2HPO4)2. Several other interactions were also identified, albeit with lower statistical significance (P = 0.0041–0.0526). These interactions demonstrate that, in this case, the overall effect of the several inhibitory factors was more than simply additive. Reviewing the output from all of the models (Table 21.1) we see that there are 10 factors whose effect on toxin formation is quantified (11 if we count NaCl + Na2PO4 as 1 factor, following Tanaka et al., 1986). In addition, incubation temperature was fixed, but would be expected to play a role, and end-point time to toxin was either fixed, or measured as a response variable, rather than being considered a factor contributing to safety. Moreover, the whole preservation system also relies on thermal pasteurization and packaging to protect the product from recontamination. Thus, we have a minimum of 13 identified factors contributing to the safety of these products (10 from Table 21.1, plus pasteurization, packaging, and storage temperature), and all for a “simple cheese spread.”

TABLE 21.1 Quantified Factors Controlling Toxin Formation by Clostridium botulinum in Pasteurized Processed Cheese Spreads Identified through Modeling and Designed Experimentation Tanaka et al. (1986) pH Moisture Na2HPO4 + NaCl

Glass et al. (2017)

Glass and Johnson (2004)

pH Moisture Na2HPO4 NaCl Fat

Fat Sodium lactate Monolaurin Enzyme modified cheese (EMC)

Fraction of replacement of Na by K Potassium sorbate

These factors were determined using a fixed incubation temperature and defined or measured end times, and all rely on the product having been thermally pasteurized and protected from recontamination by packaging.

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21.6.2 Ready-to-Eat Meat Products and Listeria Control Another class of products where modeling has been very influential in establishing safe formulation is ready-to-eat (RTE) cooked meats. Following the emergence of Listeria monocytogenes as a significant pathogen in the 1980s, and several large and costly recalls in the late 1990s, Listeria became the single largest cause of class I recalls (those with a reasonable probability that the hazard will cause adverse health consequences or death; Gorton and Stasiewicz, 2017). The meat-processing industry was determined to find improved ways to control Listeria and invested heavily in plant conditions and sanitation to reduce the risk of Listeria contamination, plus modified preservation systems to inhibit the growth of any Listeria that did manage to contaminate the product. Seman et al. (2002) used a central composite response surface design to investigate the effects of added sodium chloride (0.8 to 3.6%), sodium diacetate (0 to 0.2%), potassium lactate syrup (60% [wt/wt]; 0.25 to 9.25%), and finished-product moisture (45.5 to 83.5%) on the growth rate of L. monocytogenes in cured ready-to-eat (RTE) meat products stored at 4°C. Increasing concentrations of sodium diacetate (P < 0.11) and potassium lactate (P < 0.001) both gave significant reductions in the growth rates of L. monocytogenes. Increased finished-product moisture (P < 0.11) significantly increased growth rates, and sodium chloride had no significant effect. Thus three significant factors controlling the growth of Listeria were identified. Working with the same experimental data, Legan et al. (2004) first examined all growth curves and determined the time needed for 1 log of growth to occur, following the recommendations of the expert food microbiology panel of the Institute of Food Technologists (2003). This “time to growth” was used as the response variable in a boundary modeling approach with the same four continuous variables: added sodium chloride, sodium diacetate, potassium lactate syrup, and finished-product moisture. In addition, data generated (but not reported) by Seman et al. (2002) with the design repeated for ready-to-eat uncured meat products were included, giving a fifth, categorical, variable for cure status. Products were stored at 4°C. The results were modeled using a generalized regression approach. All five main effects were statistically significant: added sodium chloride, potassium lactate, and sodium diacetate increased time to growth. Increased finished-product moisture decreased time to growth, and “curing” increased time to growth. In addition, six two-factor interactions, and two quadratic terms were also significant, with all interactions excluding moisture further increasing time to growth, but those including moisture reducing it. The overall effect on the growth and no-growth space is shown in Figure 21.6 for cured products, and Figure 21.7 for uncured products. Excluded from both of these models was a temperature variable, but we know from long experience (and other models, e.g. Buchanan and Phillips, 1990; Buchanan et al., 1989) that increasing temperature increases the growth rate of L. monocytogenes. Products were vacuum-sealed to minimize exposure to oxygen and protected from recontamination by a laminated flexible package. Also excluded from the models was the lethality treatment applied to the meat during cooking, which limits the initial level of Listeria nominally to zero unless there is some failure of control between cooking and packaging. Hence, we can consider that there are nine factors at play here, of which eight protect the product from the growth of Listeria. Continuing the quest for improved and lower-cost Listeria control factors, Seman et al. (2008) built on the work of Glass et al. (2006, 2007), the insights on diacetate effectiveness from Legan et al. (2004), and their own unpublished observations, and designed a modeling study to investigate the combined effect of sodium benzoate (0.08 to 0.25%) and sodium diacetate (0.05 to 0.15%), with salt (0.8 to 2%) and finished-product moisture content (55 to 75%) on the growth of L. monocytogenes in RTE cured meat using time to 1 log of growth as the endpoint. Again, products were cooked (but inoculated for experimental purposes), vacuum-sealed in a laminated flexible package, and stored at 4°C. All main effects were significant except product moisture, which was significant when included in the two- and three-way interactions (P < 0.05). Sodium benzoate was more effective (lengthening time to growth) when used with increasing concentrations of sodium diacetate and salt and decreasing finished-product moisture. The reader is invited to read the ingredients list the next time they buy a package of RTE meat. If any of the ingredients listed above are seen, they will know that one of these models was influential in ensuring the safety of the product in their hands. In the event that they find a claim such as “contains no added nitrates or nitrites except those naturally occurring in added vegetable ingredient” they will know that

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FIGURE 21.6  Compilation of contour plots showing predicted weeks to growth against salt, moisture, sodium diacetate, and potassium lactate syrup concentrations for cured model products. Contours are plotted from 18 weeks in the lower left position rising at 6-week intervals to 48 weeks in the upper-right position in each individual plot. Shaded plots indicate salt and moisture concentrations from the model design treatments. Points within the individual graphs show associated diacetate and lactate syrup concentration where 1 log or more of growth was observed in the modeling experiments (◯), where less than 1 log of growth was observed in the modeling experiments (△), where 1 log or more of growth was observed in the validation data (×), and where less than 1 log of growth was observed in the validation data (◵). In these plots, the positions of validation data points are approximate, since the salt concentration, moisture content, or both were rounded to allow the points to show within this matrix. Where possible, rounding was in the direction of a reasonable decrease in growth potential so that points were more likely to show in the no-growth space, whether or not growth was observed. Two circled points show where more than 1 log but 2 logs or less of growth was seen in the modeling data. (Republished with permission of International Association for Food Protection, from Legan et al., 2004; permission conveyed through Copyright Clearance Center, Inc.)

the product was “cured” by a process involving conversion of the naturally high nitrate content of celery or another vegetable using a microbial culture and substituting the output for the sodium nitrite used in a “traditional” cure. This process is described in Husgen et al. (2008). Accompanying such a claim, they are likely to notice “cultured sugar” or “cultured corn sugar” and possibly vinegar in the ingredients list. This culture process produces lactic acid (neutralized to lactate), and if the vinegar is included in the fermentation it is simultaneously neutralized to diacetate, which means that the models above, or one of their successors are still helping to ensure safety.

21.7 Multifactorial Preservation in Action Severely processed products have a large margin of safety. However, when using multifactorial preservation, the applied process must be defined and rigorously controlled with full regulatory compliance and scientific understanding of the factors controlling product safety and stability. Globally this

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FIGURE 21.7  Compilation of contour plots showing predicted weeks to growth against salt, moisture, sodium diacetate, and potassium lactate syrup concentrations for uncured model products. Contours are plotted from 18 weeks in the lower left position rising at 6-week intervals to 48 weeks in the upper right position in each individual plot. Shaded plots indicate salt and moisture concentrations from the model design treatments. Points within the individual graphs show associated diacetate and lactate syrup concentration where 1 log or more of growth was observed in the modeling experiments (◯), where less than 1 log of growth was observed in the modeling experiments (△), where 1 log or more of growth was observed in the validation data (×), and where less than 1 log of growth was observed in the validation data (◵). In these plots, the positions of validation data points are approximate, since the salt concentration, moisture content, or both were rounded to allow the points to show within this matrix. Where possible, rounding was in the direction of a reasonable decrease in growth potential so that points were more likely to show in the no-growth space, whether or not growth was observed. (Republished with permission of International Association for Food Protection, from Legan et al., 2004; permission conveyed through Copyright Clearance Center, Inc.)

would involve compliance with local food law and the principles of risk management built into the Hazard Analysis and Critical Control Point (HACCP) system (Codex Alimentarius Commission, 1997). Within the US the additional bio-terrorism and deliberate adulteration/food fraud provisions included in the Hazard Analysis and Risk Based Preventive Controls (HARPC) provisions of the Food Safety Modernization Act (2011; FSMA) apply. Paradoxically, almost all products in the marketplace already rely on multiple factors to provide for their safety and preservation. Most products are: • Processed thermally or non-thermally as needed to inactivate microorganisms. • Formulated to lower aw and/or pH, and include antimicrobial preservatives to inhibit microbial growth. • Packaged to protect from recontamination, and sometimes with modified atmosphere to inhibit microbial growth. • Distributed with controlled temperature when this is needed to inhibit microbial growth.

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Hence it is rare to find a product at retail that is only processed by a single severe process. For example, even the “most processed” products, shelf-stable canned foods [low acid canned foods (LACF)] are thermally processed to “commercial sterility” (to inactivate microorganisms including mesophilic spores) in a double-seamed hermetically sealed container (to protect from post-process recontamination). Acidified canned foods receive a gentler thermal process (to inactivate vegetative microorganisms), along with lowered pH (to inhibit spore germination), and hermetic seal (to protect from recontamination). While designing and developing multifactorial products, co-optimization of quality and safety attributes is based on the design and precise delivery of:

1. The inactivation process 2. Inhibitory factors 3. Post-process protection including packaging and environmental controls 4. Additional factors listed in the HACCP program including critical control points (CCPs) and/ or control points listed in production directives and GMPs

It is important to emphasize that the factors or barriers apply to specific product/processing/packaging situations. When product design relies on modeling, any extrapolation beyond the model design space should be validated with microbial challenge studies and life cycle stability assessment.

21.7.1 Market Examples of Multifactorial Products Discussed below are examples of minimally processed products commercially available in the market place to demonstrate the use of multifactorial preservation to deliver products that are simultaneously optimized for food safety, stability, and quality. Outlined for each product are the design elements described above. Note that important information about commercial products might not appear in the scientific literature, but rather be disclosed through patents or patent applications, and there might be several applications or granted patents pertaining to a single product covering different aspects of manufacture, preservation, etc.

21.7.1.1 Co-Extruded Cereal Bar with Fruit Core This is an intermediate-moisture food (IMF) with a finished product aw of 0.65–0.67, intended to be stored and distributed at ambient temperature. The product consists of two co-extruded physical compartments with gradients of pH and aw. The outside is a cereal-based dough with near-neutral pH and low aw after baking. This surrounds a fruit core that has low pH and high aw. After extrusion, the assembly is baked, cooled, and packaged. The expected shelf life of compliant mold-free products is 9–12 months at ambient. Defective products have a reduced shelf life of about 1–2 months and fail through the development of visible mold growth on the product surface.

21.7.1.1.1 Inactivation Process Baking of coextruded raw product, to provide the required product texture and flavor characteristics, simultaneously inactivates non-spore forming vegetative pathogens and molds.

21.7.1.1.2 Inhibition Parameters The control of Brix and total soluble solids (TSS) of the formulated dough and the fruit core, along with control of the baking and cooling conditions, leads to the final product aw. The use of antimycotic preservative in both dough and fruit core boosts mold control, and the antimicrobial activity of the preservative is optimized through control of pH.

21.7.1.1.3 Production Directives • HACCP: cooling to 49.1–54.4°C (120–130°F); not to exceed 57.2°C (135°F) at exit prior to packaging

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• Finished product release criteria: aw of 0.65 to 0.67 and not to exceed 0.70 • Regulatory compliance: environmental monitoring of cooling zones and the air handling system, and proper package integrity and heat seal

21.7.1.1.4 Shelf-Life Failure Significant reduction in finished product ambient shelf life from 9–12 months to 1–2 months occurs due to the growth of xerophilic molds such as Aspergillus species, Wallemia sebi, and some Penicillium spp. on the product surface. Note that many food practitioners would recognize the Aspergillus colony appearance on low-aw product under their Eurotium teleomorph names. These names are no longer taxonomically valid (Houbraken et al., 2014: Hubka et al., 2013). This spoilage occurs because recontamination with aerial mold spores on exposed product can occur during post-bake cooling. If these re-contaminated products are packaged warm, condensation of moisture on the inside of the packaging material may be re-absorbed at the product surface, raising the aw to a level favorable for the surface growth of spoilage xerophilic molds.

21.7.1.2 Optimization of “Non-C. botulinum” Thermophilic Cook with Use of Nisin in Canned Foods The D value at 121°C for spores of Clostridium botulinum is 0.1 to 0.3 minutes. In the canning industry, a “12D process” is used to eliminate C. botulinum spores which is approximately 3 minutes at 121°C (12 × 0.3) and is commonly known as a “Bot Cook.” However, sometimes foods thermally processed to inactivate spore-forming pathogens exhibit high spoilage rates as a result of the growth of non-pathogenic thermophilic spore-forming bacteria. This most often occurs when canned foods are stored at high temperatures (38–52°C; 100–125°F) in tropical or desert areas where thermophilic aerobes and anaerobes can grow. In the food industry, this is commonly known as economic spoilage because there is no health hazard. The D value at 121°C for the causative spore-formers, Geobacillus stearothermophilus (Flat Sour) and C. thermosaccharolyticum, is about 4 minutes, and thus a 5D process for these extremely heat-resistant spores would result in an F0 of 20 minutes (David et al., 2013) or more than 6 times longer than the “Bot Cook.” If such a thermal process were utilized to inactivate these thermophilic spore-formers, a product of poor quality would result due to the many heat-induced physical and chemical changes. For the above reasons, nisin has been extensively used as a natural preservative in low-acid canned foods (LACF). By using nisin it is possible to control thermophilic spoilage when the cans are stored under warm conditions for prolonged periods, or to allow a reduction in thermal processing with an F0 of 5 minutes without the risk of thermophilic spoilage occurring. This reduction in total thermal process leads to improved product quality because of reduced thermal damage to the texture, taste, and appearance of certain vegetables, an increased throughput in the canning operation, and reduced energy consumption in the operation of retorts. High-acid foods such as canned tomatoes and tomato juice with pH below 4.5 can be processed with a gentler thermal process, as C. botulinum is not the target microorganism. However, spoilage can occur due to acid-tolerant, Gram-positive spore-formers such as Clostridium pasteurianum and Bacillus macerans, and nisin has been used to control the growth of such bacteria in these acidic foods (Abee and Delves-Broughton, 2003).

21.7.1.2.1 Inactivation Process • Thermal sterilization optimized for reduced “thermophilic cook,” but fully adequate for “12D Bot cook” • Adequate cooling of cans below 32–35°C (90–95°F) before palletizing and stacking to prevent “stack burn” in the warehouse, and potential for spore germination

21.7.1.2.2 Inhibition Process • Nisin is used to reduce the severity of the “thermophilic cook” and to conserve product quality.

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• Package integrity via double seam and hermetic seal. • Reduced pH control and use of nisin for processing tomatoes and tomato juice.

21.7.1.2.3 Production Directive and Ingredient Specifications • Incoming ingredients specification for spores – mesophilic, thermophilic, and aerobic and anaerobic • HACCP and regulatory compliance

21.7.1.3 Bottled Water In the United States, bottled water is regulated by the U.S. Food and Drug Administration (FDA), which defines bottled water as “water that is intended for human consumption and that is sealed in bottles or other containers with no added ingredients except that it may optionally contain safe and suitable antimicrobial agents.” As an option and within limitations, fluoride may also be added (Robin, and Feng, 2015).

21.7.1.3.1 Inactivation and Inhibition Process The FDA allows disinfection of all water types, typically by ozonation and/or ultraviolet light.

21.7.1.3.2 QA, GMP, and Production Directives • Water prior to ozonation must meet all microbiological, physical, chemical, and radiological quality standards for bottled water. • Post-ozonation, the residual ozone level must not exceed 0.4 mg/liter of bottled water. • Weekly coliform analysis of bottled water and source water; not more than 1 coliform per 100 mL. • If coliform is detected in source water or bottled water, the manufacturer must determine whether any of the coliform bacteria are E. coli. Bottled water containing E. coli is considered adulterated. Source water containing E. coli must be treated for the eradication of E. coli. • Microbiological testing of containers and closures should be done once every 3 months analyzing for heterotrophic plate count (HPC) and coliforms using 4 different containers and closures. All of the samples must be free of coliforms, and not more than one of the four samples may have HPC in excess of one colony per ml capacity of the container or one colony per square centimeter of surface area of closure.

21.7.1.4 Refrigerated Liquid Egg Containing Vegetable Blend Liquid whole egg pools the contents of multiple individual eggs, raising the risk of Salmonella contamination of the bulk liquid egg. For this reason, conditions for the pasteurization of liquid egg are defined in the regulations (e.g. 9 CFR 590.570; Egg Products Regulations, 1993). These specify both the temperature and time that must be met, but also allow for processes of equivalent lethality. Refrigerated liquid eggs containing vegetable blend are made by adding mixed vegetables into previously pasteurized liquid egg purchased from an upstream supplier. The blend must then be reprocessed so as to maintain safety and deliver a 90-day refrigerated shelf life while minimizing damage to the functional properties of the egg proteins. One method of preparing such a product was disclosed by Rapp (1989).

21.7.1.4.1 Inactivation Process The post-blending thermal process is optimized for protecting egg proteins while destroying Salmonella and spoilage microbes. Incoming liquid egg, previously pasteurized by the ingredient vendor, is shipped refrigerated in stainless totes. The apparently redundant “double pasteurization” is for compliance and for ensuring food safety integrity through the supply chain.

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21.7.1.4.2 Inhibition Parameters • Liquid whole egg has a slightly alkaline pH, typically around 7.2–7.9. • The natural antimicrobial nisin is added to control the germination of spores of Bacillus cereus and the growth of vegetative cells of Micrococcus, both tolerant of the mild thermal process intended for Salmonella destruction. • Final product is filled in a cleanroom using a filler equipped and configured for extended shelflife (ESL) operations. Packaging web and cap/pour fitment are sanitized prior to filling. • Finished product storage and handling refrigerated below 3.3°C (38°F).

21.7.1.4.3 QA, Ingredient Specifications, and Production Directives • Incoming pasteurized refrigerated eggs – temperature history and certificate of analysis (COA) • COA and microbiological specification for vegetable slurry, especially psychrotrophic and mesophilic spores and vegetative cells • Order of addition of nisin: nisin added while batching vegetable slurry stream prior to comingling with liquid eggs followed by final pasteurization • Full compliance with USDA, FSMA, HACCP, and GMPs

21.7.1.5 High-Pressure Processed Products High-pressure pasteurization (HPP) became commercially viable for food processing in the late 1990s, though the early experiments showing a lethal effect on vegetative bacterial cells were done in the late 1800s (Hite 1899). The major advantage of HPP is its ability to reduce or eliminate vegetative cells while maintaining a fresher taste and appearance when applied to suitable substrates. Additionally, HPP, like its thermal predecessor, does not kill spores, so additional physical or chemical factors are needed to protect from spore germination and outgrowth. Typically, these factors include refrigeration and ingredients applied during product formulation. Many different types of foods treated by HPP are available in grocery stores around the world, for example: • High-pressure pasteurized jams were introduced into the Japanese market in 1990 (Balasubramaniam et al., 2008). • One of the earliest US commercial HPP products was extended shelf-life guacamole: a preparation of avocado, introduced to the market in 1997 (Balasubramaniam et al., 2008). • “Natural” RTE meats are a significant market segment in the US (Balasubramaniam et al., 2008).

21.7.1.5.1 Inactivation Process In the context of multi-factorial preservation, HPP is unique in that it offers a non-thermal pasteurization in which the lethal principle (pressure) is transmitted instantaneously to all parts of a food product. For solid foods, this treatment is applied once the product has been sealed inside a package with at least some flexible component and the package is submerged in water or some other pressure-transmitting fluid. Lethality is a function of applied pressure (commercially commonly 600 MPa) and hold time (typically around 3 min). The lethality of HPP to L. monocytogenes in RTE meats was demonstrated by Youart et al. (2010), showing that 600 MPa for 3 min is more than enough to achieve the 2 log post-lethality treatment qualifying for reduced sampling under Alternatives 1 or 2 of the Listeria rule (U.S. Department of Agriculture Food Safety and Inspection Service, 2006).

21.7.1.5.2 Inhibition Parameters • HPP Jams: pH and aw are important factors preventing the outgrowth of any surviving spores. • HPP guacamole: low pH and refrigeration prevent spore germination.

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• Natural RTE meats: usually have “natural” variants of the lactates, diacetates, nitrites, etc., addressed in Section 21.6.2, above. These ingredients are produced by the fermentation of e.g. corn-derived sugars or bio-conversion of nitrates naturally present in some vegetable ingredients, particularly celery. • In all cases the sealed, flexible package protects from recontamination after processing until the package is opened. At that point maintenance of refrigeration and limits on time to consumption are needed.

21.7.1.5.3 Ingredient Specifications and Production Directives • • • •

Purchasing specifications including raw material microbial limits. Formulation design and control to achieve critical values of aw, pH, lactate, nitrite, etc. Critical limits on pressure and process time. HACCP plan including control of celery allergen when RTE meats receive HPP treatment in the same equipment as other products, e.g. at a co-manufacturing facility. • Temperature control.

21.8 Future Development of Novel Opportunities As consumers continue to demand ever more “natural” and “fresh-like” food products, industry innovators will do everything possible to satisfy that demand. As we have just seen, multifactorial preservation systems provide opportunities for effective preservation with maximum benefit to total product quality. However, these systems can be very complex. That means that in future we can expect modeling to play a more prominent role in optimizing the effectiveness of the factors that we already know about. The existence of public modeling portals such as the USDA Pathogen Modeling Program (http​​s:/​/p​​mp​.er ​​rc​ .ar​​s​.usd​​a​.gov​​/ PMPO​​​nline​​.aspx​), and databases with both models and validation data such as ComBase (www​.combase​.cc​/index​.php​/en/) are very helpful for guiding the design of more detailed modeling studies. As the number of variables increases, modeling becomes more challenging and expensive, and when we consider that some of the preservation systems discussed earlier have 10 to 13 identifiable factors, we can see that we may be pushing the limits of affordability for the food industry. Since the early 2000s, “Big Data” has been having large effects on consumer research and product design in a wide range of industries (Blackburn et al., 2017), and we might expect that it will be influential in a complex space like multifactorial food preservation. Of course, many factors besides microbial growth can affect food quality, but to this point quality is not typically considered by microbiological models. However, models of changes in food quality have now been developing for some time (Boekel, 2008). Combining microbial models with quality models will give us a greater ability to make an early assessment of the shelf life of newly developed products. One example of including multiple microbiological and food characteristics in a food safety model was presented by Halder et al. (2011). These authors describe an integrated software package that combines a physics-based process model with kinetic models of chemical and microbiological processes and a database with the composition and properties of many different foods. Then we encounter the reality that there have been few, if any, new preservation ingredients for foods developed in the past 50 years because of the costs involved in the research, and more recently because the US self-GRAS process has come in for some criticism. Also, concerns have been raised about the solvent extraction processes that can be involved in separating an antimicrobial from a plant source. A potentially interesting approach originating in the field of drug discovery arises from the recognition that microbial resistance to antimicrobial compounds is often connected with the ability of the cell to either exclude them, or pump them out. This active exclusion is driven by a number of multi-drug resistance (MDR) efflux pumps (Marquez, 2005). Inhibiting these efflux pumps can potentiate the activity of natural antimicrobials by two or more orders of magnitude (Tegos et al., 2002). Some MDR pump inhibitors are known (Garvey and Piddock, 2008; Stermitz et al., 2000), and in the pharmaceutical world this has

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led to identified approaches for overcoming antimicrobial resistance (Pagès and Amaral, 2009; Tegos et al., 2011). These approaches could be loosely described as looking for synergies of effect between unrelated materials. In the world of food science there are many published papers describing synergies between antimicrobials. Most are small studies (Ye et al., 2013; Guttierez et al., 2009; Nazer et al., 2005), and very few investigate the taste or flavor impact of the natural antimicrobials used. To change the trajectory of food science research into natural antimicrobials (particularly those derived from plant materials that might provide for “consumer-friendly” labeling), and enable the next generation of multifactorial preservation, we could adopt the high-throughput screening approaches (De La Fuente et al., 2006; Blondelle and Lohner, 2010) successfully used in the pharmaceutical industry for drug discovery, but must be aware that the path from novel compound, whether natural or synthetic, to market is long, complex, and expensive, encompassing as it does technical, regulatory, and consumer considerations. Another area to be very aware of is the dramatic progress in gene-sequencing technologies. According to the National Human Genome Research Institute (2019) the cost of sequencing a human genome approximately tracked Moore’s law in electronics (Moore 1965) which predicted a doubling in circuit complexity per year at the same minimum component cost. After 2008, when a switch from “Sanger sequencing” to “next-generation sequencing” took place, the cost per human genome decreased by three orders of magnitude more than Moore’s law’s prediction. The practical consequence of this cost reduction is that sequencing has become accessible for many activities that would have been unaffordable in 2008, and analysis of changes in the microbiome of a food product over time is a realistic way to track the progression of the spoilage microflora. Benson et al. (2014) showed how changes in the microbiome of chilled, fresh pork sausage over time were altered by incorporation of a lactate-diacetate preservative. Both with and without lactate-diacetate the ultimately dominant bacterial population was dominated by Lactobacillus graminis, but the population switch to this organism was much faster in the presence of lactate-diacetate and the peak population was lower. This correlated with a reduction in sensory changes over time. This improved ability to measure changes in the microbiome of foods as reviewed by Cao et al. (2017) and to understand the microbiome of food production environments (Bokulich and Mills, 2013) should lead to significantly deeper understanding of the mechanisms of food spoilage. Combining this understanding with advances in methods for modifying microbial population changes could give us the ultimate in multifactorial preservation systems.

21.9 Concluding Remarks The food industry’s understanding of multifactorial preservation systems has come a long way over the past 40 years. This progress has partly enabled, and partly been in response to, consumers’ demands for fresher, less processed, more natural products that retain all of the convenience and taste that they have come to expect. As this is a long-running trend, it seems unlikely to reverse any time soon, if ever. Product design using clean-label ingredients combined with multifactorial preservation is the best way that we know of to continue satisfying these consumer demands. We expect this field will continue to develop over the next 40 years and to incorporate at least some of the novel opportunities emerging from universities, research institutes, research consortia, and transferring from other industries. The best chance for success in developing “big ideas” is for all stakeholders in food safety and preservation to work collaboratively in the discovery and feasibility stages and reserve competition for applications and consumer product development.

21.10 Acknowledgments The authors wish to thank Heather Legan of BP America Inc. for discussions on risk management and the “Swiss cheese” model. Morgan Fields and Blake David were instrumental in developing the art work and drawing for “Pole Vaulting and Injured Patient Status” shown in Figure 21.4. Their patience and diligence in translating the authors’ concept through several iterations are acknowledged with gratitude.

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21.11 Disclaimer References to commercial products and trade names are made with the understanding that no discrimination and/or no endorsement by the authors or the organizations that they are involved with are implied.

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Pagès, J-M., and Amaral, L. 2009. Mechanisms of drug efflux and strategies to combat them: Challenging the efflux pump of Gram-negative bacteria. Biochemica et Biphysica Acta 1794:826–833. Perneger, T.V. 2005. The Swiss cheese model of safety incidents: Are there holes in the metaphor? BMC Health Serv. Res. 5:71–78. Rahman, M.S. 2015. Hurdle technology in food preservation. In Minimally Processed Foods Technologies for Safety, Quality and Convenience. Edited by Siddiqui, M.W. and Rahman, M.S. 17–33. Springer International Publishing, Switzerland. Rapp, H. 1989. Process of treating vegetables for use in a vegetable omelet mix. US Patent 4,910,036. Reason, J. 1990. The contribution of latent human failures to the breakdown of complex systems. Philos. Trans. R. Soc. Lond. B, Biol. Sci. 327:475–484. Reason, J.T., Carthey, J., and de Laval, M.R. 2001. Diagnosing “vulnerable system syndrome”: An essential prerequisite to effective risk management. Qual. Health Care, 10 (Suppl II): ii21–ii25. Robin, L.P., and Feng, P. 2015. Bottled water. In Compendium of Methods for the Microbiological Examination of Foods. Fifth Edition. Edited by Salfinger, Y., and Tortorello, M.L. 791–796. APHA Press, Washington, DC. Scott, V.N. 1989. Interaction of Factors to Control Microbial Spoilage of Refrigerated Foods. J. Food Prot. 52:431–435. Seman, D.L., Borger, A.C., Meyer, J.D., Hall, P.A., and Milkowski, A.L. 2002. Modeling the growth of Listeria monocytogenes in cured ready-to-eat processed meat products by manipulation of sodium chloride, sodium diacetate, potassium lactate and product moisture content. J. Food Prot. 65:651–658. Seman, D.L., Quickert, S.C., Borger, A.C., and Meyer, J.D. 2008. Inhibition of Listeria monocytogenes growth in cured ready-to-eat meat products by use of sodium benzoate and sodium diacetate. J. Food Prot. 71:1386–1392. Shorrock, S.T., Young, M., and Faulkner, J. 2006. Seeking and finding organizational accident causes: Comments on the Swiss cheese model. Aviation Safety Letter Issue 2. Transport Canada, Ottawa, ON. Singh, S., and Shalini, R. 2016. Effect of Hurdle Technology in food preservation: A review. Critical Reviews Food Sci. Nutrition, 56:641–649. Doi: 0.1080/10408398.2012.761594 Stermitz, F.R., Lorenz, P., Tawara, J.N., Zenewicz, L.A., and Lewis, K. 2000. Synergy in a medicinal plant: antimicrobial action of berberine potentiated by 5′-methoxyhydnocarpin, a multidrug pump inhibitor. Proc. Natl. Acad. Sci. USA 97:1433–1437. Tanaka, N., Traisman, E., Planinga, P., Finn, L., Flom, W., Meske, L., and Guiggisberg, J. 1986. Evaluation of factors involved in antibotulinal properties of pasteurized process cheese spreads. J. Food Prot. 49:526–531. Tegos, G., Stermitz, F.R., Lomovskaya, O., and Lewis, K. 2002. Multidrug pump inhibitors uncover remarkable ability of plant antimicrobials. Antimicrob. Agents and Chemother. 46:3133–3141. Tegos, G.P., Haynes, M., Stouse, J.J., Khan, M.Md.T., Bologa, C.G., Oprea, T.I., and Sklar, L.A. 2011. Microbial efflux pump inhibition: tactics and strategies. Curr. Pharma. Design 17:1291–1302. Underwood, P., and Waterson, P. 2014. Systems thinking, the Swiss Cheese Model and accident analysis: A comparative systemic analysis of the Grayrigg train derailment using the ATSB, AcciMap and STAMP models. Accid. Anal. Prev. 68:75–94. U.S. Department of Agriculture Food Safety and Inspection Service. 2006. Compliance guidelines to control Listeria monocytogenes in post-lethality exposed ready-to-eat meat and poultry products. Available at: http:​/​/www​​.fsis​​.usda​​.gov/​​oppde​​/rdad​​/ FRPu​​bs​/97​​- 013F​​/ LM​_R​​ule​_C​​ompli​​ance_​​Guide​​lines​​​_ May_​​ 2006.​​pdf. Accessed 02 September 2019. U.S. Department of Health and Human Services and U.S. Department of Agriculture. 2015. 2015–2020 Dietary guidelines for Americans. 8th edition. Available at: https​:/​/ he​​alth.​​gov​/d​​ietar​​yguid​​eline​​s​/201​​5​ /gui​​delin​​es​/ch​​apter​​-1​/ ke​​y​-re​c​​ommen​​datio​​ns/. Accessed 21 August 2019. Vallery-Radot, R. 1919. The Life of Pasteur. Translated by Devonshire, R.L. 113. London: Constable & Company. See https​:/​/ar​​chive​​.org/​​strea​​m ​/in.​​ernet​​.dli.​​2015.​​17390​​7​/201​​5​.173​​907​.T​​he​-Li​​fe​-Of​​-Past​​eur​​# p​​ age​/n​​135​/m​​ode​/2​​up. Accessed 26 September 2019. Ye, H., Shen, S., Xu, J., Lin, S., Yuan, Y., and Jones, G.S. 2013. Synergistic interactions of cinnamaldehyde in combination with carvacrol against food-borne bacteria. Food Control 34:619–623. Youart, A.M., Huang, Y., Stewart, C.M., Kalinowski, R.M., and Legan, J.D. 2010. Modeling time to inactivation of Listeria monocytogenes in response to high pressure, sodium chloride, and sodium lactate. J. Food Prot. 73:1793–1802.

22 Applications of Antimicrobials to Foods – A Food Industry Perspective Jairus R. D. David and Peter Taormina CONTENTS 22.1 Introduction..................................................................................................................................715 22.2 Research and Development Perspectives.....................................................................................716 22.2.1 Combination Studies......................................................................................................718 22.2.2 Standardization of Efficacy Determination...................................................................718 22.3 Considerations for Commercial Application of Antimicrobials in Food.....................................719 22.3.1 Phase 1 – Discovery or Proof of Concept..................................................................... 720 22.3.2 Phase 2 – Technology Development............................................................................. 720 22.3.2.1 Efficacy...................................................................................................... 720 22.3.2.2 Sensory Impact...........................................................................................721 22.3.2.3 Cost-in-Use.................................................................................................721 22.3.2.4 Capital Expenditure................................................................................... 722 22.3.2.5 Minimum Order Quantity (MOQ)............................................................ 722 22.3.2.6 Patent Landscape....................................................................................... 722 22.3.2.7 Regulatory Assessment............................................................................. 722 22.3.2.8 Challenge Studies...................................................................................... 723 22.3.2.9 Sensory and Shelf Life Study.................................................................... 723 22.3.2.10 The Production Process............................................................................. 723 22.3.2.11 Occupational Safety and Health Act (OSHA)........................................... 723 22.3.3 Phase 3 – Technology Transfer (Scale-up and Commercialization)............................ 724 22.3.3.1 Food Product Matrix................................................................................. 724 22.3.3.2 Beverages................................................................................................... 724 22.3.3.3 Bakery Products........................................................................................ 726 22.3.3.4 Fresh and Minimally Processed Fruits and Vegetables............................ 726 22.3.3.5 Dairy Products.......................................................................................... 727 22.3.3.6 Meat and Poultry Products........................................................................ 727 22.3.3.7 Sauces and Condiments............................................................................. 730 22.4 Conclusion....................................................................................................................................731 22.5 Acknowledgments........................................................................................................................731 Disclaimer................................................................................................................................................731 References................................................................................................................................................731

22.1 Introduction The objective of this chapter is to outline the important aspects of the application of antimicrobials to foods, including selection of the antimicrobial, determination of target microorganisms in a specific food matrix, efficacy testing against target microorganisms in vitro and in food matrices, and issues that must be addressed in the commercial application of the antimicrobial and product launch. Expectations of the 715

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performance of antimicrobials must be realistic and should include other considerations, such as effect on sensory and quality attributes of the food, cost and cost-in-use of the antimicrobial, and regulatory and labeling considerations, in addition to efficacy against target microorganisms in the food matrix. The “idea-to-launch” business framework and governance is recommended for pairing of a potential antimicrobial with a complex food matrix, along with clearly defined objectives, inputs, outputs, and technical success and business decision criteria. At the outset, it is vitally important to remember that the use of antimicrobials in food is meant to add to the robustness of existing safety and quality programs and production directives, and not to correct or mask poor practices (Davidson, et al., 2005). In fact, existing antimicrobials are not efficacious enough to overcome marginal or poor microbiological quality of a food. Thus, the effective use of antimicrobials in food begins with the existence of sound prerequisite programs, such as sanitation and Good Manufacturing Practices (GMPs), and appropriate food safety controls and process controls. It may be argued that prerequisite programs, and in fact any measures used to enhance the safety and quality of foods, can be viewed as hurdles and be included in a hurdle concept plan. While this is not the classical view when describing hurdle technology (Leistner, 2000), it may be practical to think in this way when setting up food protection programs in the manufacturing environment. Thus, any protective measure, including programs such as sanitation, GMPs, production directives, and even HACCP or food safety preventive controls, can be included in the hurdle concept. Similarly, antimicrobial systems do not preclude the need for downstream measures for food protection such as adequate temperature control during distribution, storage, display, and appropriate handling and preparation and use at the point of consumption.

22.2 Research and Development Perspectives Many companies have an idea of what microorganism(s) should be the target for antimicrobial control but not what type(s) of antimicrobial compound(s) may be useful against the target microorganism(s). Thus, the first step in selecting an antimicrobial is to determine its efficacy. Although many studies on the activity of antimicrobials have been published, it may be necessary to establish efficacy de novo. Because there are no standard methods for determining efficacy, researchers have generally used methods used by clinical microbiologists, such as agar diffusion assays, microbroth dilution assays, and the “time-kill” curves (Davidson and Parish, 1989). The appropriate methods for determining the activity of antimicrobials for food use are described in detail in Chapter 2 (“Methods for Activity Assay and Evaluation of Results”). Because many antimicrobials are hydrophobic or at least bear some degree of hydrophobicity, the commonly used agar diffusion assay that relies on consistent and rapid diffusion of compounds in the polar agar gel, may yield inaccurate results. Dilution assays are more appropriate for testing food antimicrobials. Reports in the literature on the efficacy of compounds tested with the agar diffusion method might be considered suspect unless these compounds have been highly purified and standardized, such as with nisin (Ripoche et al., 2006). In the evaluation of antimicrobials for potential use in foods, the suggested steps include in vitro testing in media to determine endpoints and dynamic inhibition, followed by application to foods and challenge studies. The endpoint assays, generally broth or agar dilution assays, involve adding the compound to a microbiological medium, adding the test microorganisms, and incubating for a specific time under conditions conducive to the microbes’ growth. This type of assay generates a “minimum inhibitory concentration (MIC),” or the concentration that prevents growth of the microorganism, as measured by a lack of turbidity (in broth) or colony formation on agar. A “minimum lethal concentration (MLC)” may be determined in the broth dilution assay by transferring media from tubes or wells where no growth occurred to fresh media. If no growth occurs in the fresh media, the assumption is that the microorganism was inactivated and thus that the concentration results in a 99.9% (3 log) reduction in microbial numbers. Obviously, both the MIC and MLC depend highly upon environmental growth conditions (e.g., pH, temperature) and the initial number of microorganisms. Following an endpoint assay to determine appropriate concentrations, one can determine the influence of the compound on dynamic growth by incubating the target microorganism in a microbiological broth

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medium and taking repeated samples over time to determine the changes in numbers of survivors. The plot of survivors over time is sometimes referred to a “time-kill” curve, a term used in clinical microbiology. From this type of assay, it can be determined what type of inhibition the test compound causes over time. The type may manifest itself in a number of different ways (Figure 22.1). Compared with the control, concentrations of an antimicrobial that are at the MIC may prevent an increase in the final cell number, delay and prolong lag phase, suppress growth for a time and then allow recovery, or cause lethality indicated by a reduction in cell numbers. A success criterion for further evaluation of an antimicrobial in such a test would likely be (a) an increase in lag phase, or (b) slower/delayed growth rate in exponential or logarithmic phase, or (c) lower final numbers of microorganisms in the stationary phase, or (d) some type of inactivation with no regrowth (David et al., 2013). One point to remember in these types of assays is that an antimicrobial neutralizer should be used in the medium/diluent being used for enumeration of survivors so as to avoid obtaining any false positive results. Before investing in elaborate and expensive challenge studies in actual food matrices, it is customary to assess the efficacy of promising antimicrobials in simple food systems. These studies may be done in culture tubes, using commercially sterile shelf-stable apple juice or ultra-high temperature (UHT)-sterilized shelf-stable 2% fat milk (Figure 22.2, Phase-2 Technical Success Criteria). These simple food models can be used to evaluate the effect of the food, including pH and binding of the antimicrobial by fat or protein. Obviously, antimicrobials that are bound or inactivated during thermal processing are not available to act against target microorganisms. Generally, one can expect the effect of juice to be similar to that of microbiological media because of the lack of protein and fat. In milk, there generally will be a dramatic drop in the activity of because of the high pH and binding by protein and fat. The purpose of these tests is to get an idea of what concentrations might be efficacious in the food product of interest. The next logical progression is to evaluate the antimicrobial in the actual food matrix of interest for testing the expected/desired shelf life and simulating production, processing, and packaging conditions present at the manufacturing plant. In some cases, commercially available antimicrobial products would have already undergone such testing in model systems against certain microorganisms of concern, and usage ranges may already be provided by the developer/vendor of the

FIGURE 22.1  Diagrammatic representation of the impact of antimicrobials use on microorganism growth in a model media system (David et al., 2013): (a) control without antimicrobial; (b) extended lag phase; (c) reduced growth rate and final level; (d) static [inhibition]; (e) cidal [inactivation]; (f) initial cidal effect followed by regrowth; Ni: initial number of target microorganism; Nf: final number of target microorganism.

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antimicrobial product. A product developer or microbiologist may decide if such data are suitable for the purpose of skipping the juice/milk model step, thereby saving time. However, often, when there are specific microorganisms of concern, such as a specific type of spoilage microorganisms, this may not be possible.

22.2.1 Combination Studies Antimicrobials when used alone are often generally not effective enough or may have a negative effect on food properties at concentrations needed to be effective when used alone. Thus, it is often desirable to use them in combinations of other antimicrobials or with other functional ingredients such as acidulants and emulsifiers. When combinations of antimicrobials are elevated, three outcomes are possible. A combination may be “additive,” i.e., the effect of the combined treatments is equivalent to the sum of the effects of the treatments acting independently. The two components can be “antagonistic” toward one another, actually resulting in a reduced efficacy of the combined treatments compared with their use independently. This might result, for example, from a chemical reaction between components to form a new, non-inhibitory, compound. The most desirable outcome is termed “synergistic,” in which the activity of the combination is enhanced compared with the sum of individual treatments. Measuring synergism in vitro is most easily done with a microtiter “checkerboard” assay and by calculating a fractional inhibitory concentration (FIC), defined as the concentration of each antimicrobial in combination (see Chapter 2, “Methods for Activity Assay and Evaluation of Results”) which produces inhibition of growth expressed as a fraction of the concentration that inhibits growth when the antimicrobial is used alone (Branen and Davidson, 2004; Davidson and Parish, 1989), or a fractional lethal concentration (FLC), defined as the concentration of each antimicrobial in the combination that produces lethality, expressed as a fraction of the concentration that is lethal when the antimicrobial is used alone (Techathuvanan et al., 2014). In foods, it is more difficult to determine synergistic activity, although a modified checkerboard assay is possible. Antimicrobials may be used with physical preservation processes to enhance the effectiveness of the process or as a safeguard for post-process contamination. Corbo et al. (2009) described potential process interactions as: (a) partial inactivation of the microorganism by the preservation process, followed by continued inhibition or inactivation by the antimicrobial during storage; (b) enhancement of the process inactivation of the microorganism by the antimicrobial or vice versa; or (c) totally independent effects. Such physical processes might include heat (pasteurization) or non-thermal processes, such as high hydrostatic pressure processing (HPP), high-pressure homogenization, or pulsed electric fields. Several studies have demonstrated that antimicrobials enhance the effectiveness of physical preservation processes in inactivating target microorganisms (Black et al., 2005; Calderon-Miranda et al., 1999; Liang et al., 2002; Mosqueda-Meglar et al., 2012; Nakimbugwe et al., 2006; Nguyen and Mittal, 2007; Pathanibul et al., 2009).

22.2.2 Standardization of Efficacy Determination One of the major needs in the arena of food antimicrobials, both natural and traditional, is the adoption of standard methods for the determination of efficacy. While medically important antimicrobials (i.e., antibiotics) and sanitizers have regulatory guidelines on efficacy evaluation, no such guidelines exist for food antimicrobials. In fact, there are no governmental standards concerning the efficacy of most commercially available antimicrobial food preservatives used as antimicrobials, with the exception of nisin and lysozyme. Thus, many commercial antimicrobial food preservatives, such as sorbate or benzoate, have not been evaluated for their intended purpose, i.e., inhibition or inactivation of microorganism. While this may not be a large problem if one is attempting to extend shelf life, it certainly is important if the compound is being used to control pathogenic microorganisms. Recommendations for the use of standard methods were called for over 20 years ago (Davidson and Parish, 1989) but to date, there has been no regulatory adoption. There are however regulatory requirements in the U.S. concerning the identification of preventive controls as part of a food safety plan, and if food antimicrobials are deemed

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as process preventive controls against pathogens for instance, a scientific validation of that control would be required. This could be a published study with similar parameters to the food in question or a challenge study in the actual food system.

22.3 Considerations for Commercial Application of Antimicrobials in Food Attempts at pairing a specific food matrix in need of a secondary barrier for food protection with a potential antimicrobial is very rarely a linear or straightforward exercise. In the food industry, several competing factors need to be reviewed and co-optimized to meet predetermined technical success criteria and business decision criteria, as illustrated in Figure 22.2. Key factors that must be considered include: (a) efficacy against target microorganisms in the food matrix during processing; (b) business case and justification; (c) cost-in-use; (d) sensory effects; (e) storage; (f) end use by consumers; (g) regulatory and labeling considerations; and (h) sustainable supply (David, 2015a). To achieve the goal of successful application of an antimicrobial, certain “technical success criteria” must be established up front for managing business expectations, cost structure, implementation at the manufacturing plant, and product launch. Figure 22.2 is a modified Stage® gate business process based on the “idea-to-launch” framework for product innovation and reducing time-to-market (Cooper and Edgett, 2012). The proposed framework is for the systematic pairing of a potential antimicrobial system with a food matrix, with clearly defined objectives, inputs, outputs, and success criteria for each of the three phases: PHASE-1 – Discovery (Proof of Concept), PHASE-2 – Technology Development, PHASE-3 –Technology Transfer (Scale-up and Commercialization).

FIGURE 22.2  A modified Stage Gate® business process based on the “idea-to-launch” framework, governance, and decision criteria for pairing a food product with a potential antimicrobial system for requisite food protection at an acceptable product cost structure (David et al., 2013; David, 2015b).

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22.3.1 Phase 1 – Discovery or Proof of Concept This phase consists of high-throughput screening of promising antimicrobials against target microorganisms via appropriate assays to determine MIC and MLC. Antimicrobials differ in their ability to inhibit or inactivate vegetative cells and spores of Gram-positive bacteria, Gram-negative bacteria, and yeasts and molds. As previously stated, the first step in choosing an antimicrobial is to correctly identify and characterize target spoilage and/or pathogenic microorganisms from food. In addition, one should have a good understanding of factory microbial ecology, including vectors, incoming bio-burden load in ingredients, and data trends from the environmental monitoring program. No single antimicrobial can control all types of bacteria, yeasts, and molds in all food matrices. Lower dose concentrations for MIC and MLC are indicative of higher efficacy. Also, an order of magnitude of reduction in microbial numbers relative to initial inoculum level at time zero can be approximated. Thus, for example, successful candidate antimicrobials causing a 4- to 5-log reduction would be moved to the next phase of technology development. It is customary to review those antimicrobials with a score of 1- to 3-log reduction for other good traits as well, such as polarity, pKa, sensory effects, GRAS status, etc. Even though most antimicrobials come with vendor-generated technical information and MIC and log reduction values, it is prudent for the user to re-check the MIC and MLC under desired environmental conditions of pH and temperature and against microorganisms isolated from product recall or spoiled product or factoryspecific environmental microbiome (Benson et al., 2014; David, 2015a). Vendor MIC or MLC data may look promising, but may not have undergone peer-review. Scientists evaluating food antimicrobials data should investigate the methods used in the development of such data, such as whether or not stressed cells were resuscitated using appropriate methods (conservative approach) or samples were plated only on selective and differential media. A quick test for antimicrobial impact on the odor and taste of the target product is essential. Usually, three levels of antimicrobials (MIC, below MIC, above MIC) are mixed with finished product to assess the concentration of the subject antimicrobial. Because the finished product is the basis for this quick test, it does not account for the impact of processing conditions on final product sensory characteristics or efficacy of the antimicrobial. Combination systems with other antimicrobials or other intrinsic or extrinsic hurdles may also help lower the use and dosage of individual antimicrobials for minimizing negative sensory impact and optimizing cost-in-use. In-company food technologists may also prepare multiple pilot batches to assess differing concentrations of antimicrobial on the resulting finished product’s sensory characteristics. Such evaluations can range from the highly analytical to less so, and such evaluation of antimicrobial impacts on finished product characteristics is not necessarily limited to only the largest food manufacturing companies.

22.3.2 Phase 2 – Technology Development This phase is where the bulk of the investment (resources and cross-functional teams), testing, and assessment work are staged and completed to facilitate making the “go/no-go” technical recommendations and business decision. Presented below are several work streams in a linear manner, but not rankprioritized. Almost all of the below-described work streams must be reviewed simultaneously to evaluate early failure, not to proceed or to further resource to expedite results.

22.3.2.1 Efficacy Interestingly, in the product development and product commercialization process, efficacy and validation are often the last consideration. Efficacy is often assumed, based upon prior experience, textbooks, or “taking an ingredient company’s word for it.” As such, technical personnel can be left scrambling to meet deadlines that well surpass the time needed to perform proper research and validation. Nonetheless, the demonstration of efficacy in product is essential to having a viable, wholesome, safe product in the marketplace that will not cause illness or spoil prematurely. For product renovation, efficacy studies should include both chemical antimicrobials currently used in formulation and potential replacement clean-label ingredients to assess the equivalency of efficacy in food matrix. As a rule of thumb, the

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efficacy of natural not-purified natural clean-label ingredients will be slightly lower than traditional purified chemical ingredients.

22.3.2.2 Sensory Impact Antimicrobials may leave a residual odor or taste that can potentially impact finished product sensory and quality attributes. Organic acids acetic and lactic acids are widely used in many different food systems and are used topically on some food but impart a noticeable flavor. Plant-based extracts from citrus or spice must match the flavor profile of the food product. Oregano, thyme, or rosemary extracts are sometimes used, but only in savory-type foods that mesh with those seasonings. Fermentates are byproducts of lactic acid bacterial fermentation that contain mixtures of organic acids and bacteriocins and are sold as dry blends on a maltodextrin carrier or in a syrup form. The pungency and acidity of fermentates may preclude the usage level and desired efficacy, especially if propionic acid is prevalent in the blend. These relatively obvious impacts should enable product developers to rule out certain antimicrobials as options early in the process. It is important to do a formal cutting of food product with antimicrobial concentrations equal to and less than MIC values established from Phase 1 work, to quickly evaluate any negative sensory impact. A more formal sensory cutting such as a consumer liking test (CLT) is required during Phase-3 prototyping and scale-up work streams (Figure 22.2). It may be important to develop a “tool box” of antimicrobials to appropriately pair with different food matrices, e.g., savory, dessert (sweet), and neutral. For example, sweet essential oils may be more appropriate for use in puddings and dessert pies, whereas essential oils from coriander, lemongrass, or mustards may be more compatible with savory sauces, meat and vegetable entrees, and pot pies. Combinations of antimicrobials need to be developed that may be more efficacious (additive or synergistic) and cause less impact on the sensory properties of the food. Taste and total quality in the final saleable product are not negotiable; antimicrobial candidates that perform well with respect to pathogen or spoilage microbe inhibition but which yield unpalatable flavor/odor outcomes will not be useful preservatives.

22.3.2.3 Cost-in-Use Cost-in-use is one of the business critical and sensitive constraints that should be evaluated early on for the determination of increased cost per unit of finished packaged product. The cost-in-use analysis should also include the cost of inert carriers (e.g., maltodextrin, vegetable oil, glycerol, NaCl, etc.), enablers (i.e., emulsifiers, surfactants, and wetting agents), and delivery systems for carrying the antimicrobial to target microorganisms in the food matrix. It is important to understand and develop specifications for incoming microbial loads (including bacterial spores) for the above ingredients. Often, natural and clean-label antimicrobials are more expensive than traditional chemical preservatives, and cost can be higher by a factor of ten or more. The vendor-provided cost per pound list price for Phase-1 successful antimicrobials helps one to assess whether the product in question can absorb upcharge per case of finished product, and thus to make a reasonable business case with marketing and other key stakeholders. The rule of thumb is that the cost of antimicrobials should be less than or equal to $0.015 to $0.025 per pound of finished packaged product. Higher costs of $0.03 to $0.07 per pound of premium-type or innovation product can be justified for antimicrobials meeting or exceeding many or all of the success criteria shown in Figure 22.2 (see Chapter 1, “Food Antimicrobials—An Introduction,” Table 1.7). It is important to recognize that the structure-function of the active agent is identically the same in both traditional chemical antimicrobials and clean-label natural formats (e.g., nitrite in sodium nitrite and nitrite in cultured celery juice powder; propionate in calcium propionate and cultured wheat starch). Also, traditional chemical antimicrobial preservatives have regulatory approval and a long history of use in commercial products (sorbates, benzoates, nitrites, propionates, etc.). However, some consumers prefer and demand premium foods that are not severely processed and made with clean-label natural ingredients including antimicrobial preservatives, despite the potential for increased cost of such products at retail over other foods made without such ingredients. As a result, manufacturers, retailers, and food service operators are responding to the clean-label movement with new and improved products (David, 2015b; see Chapter 1, “Food Antimicrobials—An Introduction,” Tables 1.4 and 1.5).

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In contrast to foods made with natural and/or clean-label antimicrobials, for many products, the goal is simply to have a fit-for-purpose product that appeals to cost-conscious customers, and highvolume retailers. In such cases, these consumers are not choosing based upon claims such as “no preservatives,” but instead are more concerned with price and budget management. For the processor, such products can be produced in high volume and with a relatively long shelf life with the use of traditional preservatives, such as a sodium benzoate and potassium sorbate blend (sometimes with sodium hexametaphosphate (SHMP) and/or EDTA (ethylenediaminetetraacetic acid) in sauces, condiments, and beverages, or such as a sodium lactate and sodium diacetate blend in ready-to-eat meat products).

22.3.2.4 Capital Expenditure Capital expenditure may also need to be factored in, such as a need for the installation of special mixing and dosing units at batching or at filler operations. For example, special mixing and dosing units are needed for the application of natamycin and dimethyl decarbonate (DMDC). Precise delivery of the approved antimicrobial into food is critical for compliance with rules on the regulatory agency-approved dose at the point of application. The regulatory agency-approved maximum limit is based on the concentration of antimicrobial at the point of application and not on residual activity in downstream finished product during or at the end of shelf life.

22.3.2.5 Minimum Order Quantity (MOQ) For antimicrobials imported from other countries, it is important for a procurement department to negotiate minimum order quantity (MOQ) with ingredient suppliers. The standard threshold is about 10,000-pound per shipment in 50 to 100-pound sub-container units. Often MOQ may greatly exceed annual usage of ingredient needed for producing and stocking finished product inventory, impacting working capital. Scheduling, stock keeping unit (SKU) volumes, ingredient inventory, ingredient shelf life, and market price will determine and justify accepting large volume MOQ from vendors.

22.3.2.6 Patent Landscape Patent landscape and intellectual property (IP) review are in order at this stage to ensure that there is room for creating unique and compelling consumer solutions and innovation, and to create substantial competitive advantage via IP and patents. It is important to ensure that the application of an antimicrobial or combination of antimicrobials in target categories of products is not protected by either technology or ingredient patents, or by ingredient-lock out or by ingredient-use patents. For example, there may be restrictions on the use of a lactate-diacetate system in certain types of fresh meat products or restriction on the use of nisin plus natamycin for the control of spoilage due to lactic acid bacteria in salad dressings. Due diligence is the key so as not to infringe on any domestic or international published patents or patent applications. It is possible to use ingredients protected by patents by way of paying royalties directly to the inventors or entity, or indirectly paid to the ingredient supplier as a part of the ingredient price.

22.3.2.7 Regulatory Assessment Regulatory assessment is also done at this stage to ensure that there are no “red flags” with regard to use, human safety, and toxicology of both parent antimicrobial and breakdown chemical compounds in buffers or food models. In addition, one needs to be cognizant of constraints due to regulatory limits set for most additives and GRAS antimicrobials (FDA, 2012; USDA-FSIS, 2012). At this phase, an assessment should be made of USDA and FDA boundaries for the labeling of antimicrobials in any subject food. Similar assessment should be made for antimicrobial use in foods manufactured in the U.S. that are destined for export to other countries.

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22.3.2.8 Challenge Studies Challenge studies are done in a food matrix with the antimicrobials at the pre-process, in-process, or post-process stages at storage and abuse conditions, and in packaging over 1× and 1.5× the normal shelf life of the product. The National Advisory Committee on Microbiological Criteria for Foods (NACMCF) (Anon., 2009) has developed guidelines for conducting challenge studies with regard to pathogen inhibition and inactivation that should be adapted when developing and executing challenge studies. For some meat products, it is important to conduct separate independent microbial challenge studies targeting specific pathogens – Listeria monocytogenes, and spores of toxigenic clostridia (Clostridium botulinum, Clostridium perfringens). Each challenge test will have specific success criteria, method of inoculation (surface versus in-product, or both), incubation and sampling protocols, total test duration, and cost. In addition, it is customary to test uninoculated treatment samples for indicator microorganisms to monitor the growth of natural microflora which may undermine product shelf life and affect sensory and quality attributes (Golden et al., 2017). For refrigerated foods, it is customary to test for both psychrotrophic spores and psychrotrophic vegetative cells (Benson et al., 2014; David, 2015a; Bozkurt et al., 2016).

22.3.2.9 Sensory and Shelf Life Study For proper change management, scale-up, and directional next steps, sensory and shelf life studies are done with a formulation containing preservative made on fully scalable pilot plant-scale equipment and/ or small batches of product made in the production plant. A consumer liking test (CLT) sensory testing of baseline control and treatment samples is done using a nine-point hedonic scale with preference scores and overall liking scores, and overall preference at 90% confidence interval (CI). The decision to move forward is based on parity for sensory and quality attributes – color, flavor, texture, mouthfeel, and other product-specific traits. Also, formulations may need to be adjusted for salt and sodium levels while using antimicrobials diluted with salt or sea salt (nisin, celery juice powder, etc.) for optimizing taste, label claims, and compliance. There are some additional considerations when using calcium propionate in products baked with fruit puree. For example, there can be syneresis of fruit core in coextruded cereal bar during baking due to interaction between calcium in calcium propionate and sodium in alginate in the fruit base (David, 2015b). Also the use of propionate at higher levels for mold-free extended shelf life may contribute to bitterness in baked goods, necessitating the use of other ingredients such as bitterness maskers. Bitterness maskers can be labeled as a natural flavor, but with significant upcharge per case of product.

22.3.2.10 The Production Process There may be several aspects to the production process to consider when selecting antimicrobials. Physical processes like in-container pasteurization can reduce or eliminate the need for antimicrobials. Heat treatments can render antimicrobials less (or, in some cases, more) efficacious. The configuration and surface area of the food can influence the selection of antimicrobials: smooth-surface products are more conducive to topical antimicrobial treatments. The degree to which the surface area of food components may be exposed to antimicrobial ingredients by being chopped, ground, blended, comminuted, or emulsified into foods is a critical factor in selection. The physical aspects of the packaging format can preclude the effective use of certain antimicrobials. For example, many fresh meats are now sprayed or dipped with processing aids like lauric arginate or octanoic/citric acid combinations that can cause a reduction in Salmonella or E. coli O157:H7. The efficacy of these antimicrobials depends not only on concentration and volume applied but also upon the application method (e.g., dip versus spray), contact time, and, for finished products, the capillary action necessary to distribute the antimicrobial across the product surface.

22.3.2.11 Occupational Safety and Health Act (OSHA) The human safety of operators in the warehouse and batching stations is important while handling powder, liquid, or frozen formats of antimicrobial preservatives. The requirements for handling and batching

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provided by vendors in the safety document sheet (SDS) should be duly followed with appropriate use of personal protective equipment (PPE) and proper ventilation. Also, disposal of out-of-date ingredients should be done per vendor recommendations. Additional precautions should be taken by operators while handling certain antimicrobials that may be considered allergenic or contain allergenic inert ingredients/ stabilizers. For example, antimicrobials derived from celery or mustard are considered allergenic in Canada and EU countries. Other routine precautions of line isolation, allergen wash-down procedures, and labeling and SKU palletizing should be followed through per company production directives.

22.3.3 Phase 3 – Technology Transfer (Scale-up and Commercialization) A determination of efficacy against target microorganisms in the food matrix of choice is a prerequisite for the further investment of resources. A desired success criterion needs to be formulated for these efficacy tests, as shown in Figure 22.2. Additionally, FSMA mandates scientific validation occur prior to actual commercial processing for process preventive controls, or within 90 days of startup, so this would be the opportune time for in-plant validations in commercial plants and/or pilot plants.

22.3.3.1 Food Product Matrix The efficacy of antimicrobials in foods is most easily determined and interpreted in carbohydrate-based beverages, followed by bakery products, fruits, vegetables and produce, dairy products, and meat, poultry, and seafood products (Davidson et al., 2013). Pilot-scale production tests or small batches in fullscale processing plants can be as simple as adding the preserving ingredient and collecting samples for evaluation. However, it is more likely that adding antimicrobials will require a great deal of forethought and planning as it influences other attributes of the food system that require a modification of the process. In either case, it is at this stage that the incorporation of the antimicrobial into or onto the food must result in adequate coverage and/or distribution and interaction with target microorganisms. A key objective of scaling and commercializing antimicrobial technologies into full-scale food production is confirming that the characteristics hold true, that the product impact is as expected at the time of processing and during shelf life, and that the naturally occurring microorganisms in the food system and processing environment are also affected similarly to laboratory strains.

22.3.3.2 Beverages Carbohydrate-based beverages such as fruit juices and soft drinks present the fewest challenges associated with incorporating antimicrobials. They are not dissimilar to microbiological media; these systems have relatively low pH and very low to non-existent protein and lipid contents, and they provide a homogenous liquid system for the dispersion of antimicrobials. The only hurdle for application in this type of food would be undesirable interactions of the antimicrobial with simple and complex carbohydrates and minerals (e.g., the chelator sodium hexametaphophate works well in unfortified fruit beverages, but not in calcium-fortified versions). Protection against the microbiological spoilage of beverages can be achieved using antimicrobials and/or processing techniques such as hot-filling, tunnel pasteurization, ultra-high temperature treatment (UHT), or pasteurization followed by aseptic packaging, and/or pasteurization followed by chilling the beverage. Generally, beverages with a pH < 4.6 can be preserved with antimicrobials, heat-processed, and filled into packages such that the product is not re-contaminated. For example, process techniques such as cold-filling with antimicrobials or pasteurization followed by cold-filling may be used to preserve this type of beverage. In a similar manner, this same beverage may be processed using non-preserved techniques such as hot-filling, tunnel pasteurization, pasteurization followed by aseptic filling, or even requiring the beverage to be chilled (i.e., under refrigeration following the pasteurization step). Beverages having a pH > 4.6 must be processed such that spores are destroyed using ultra-high temperatures followed by aseptic filling into packages or retorting sealed packages of product. One of the most common preservation systems for acidic, shelf-stable carbonated and non-carbonated soft drinks relies on weak acid preservatives (e.g., benzoic and/or sorbic acid). Benzoic and sorbic acids

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(and salts thereof) effectively inhibit yeast, bacteria, and molds with only a few exceptions. Weak acids in beverages exist in equilibrium between their dissociated and undissociated forms which are dependent upon the dissociation constant of the acid (pKa) and the beverage pH (Table 22.1). The pKa for benzoic acid is 4.19 and the pKa of sorbic acid is 4.76. A beverage pH below the pKa of the particular acid shifts the equilibrium towards the undissociated form (i.e., benzoic acid versus benzoate). The undissociated form is more efficacious against microorganisms; therefore, weak acid preservatives are most effective in the low pH range. The preservation properties of weak acids may be enhanced by the addition of chelating compounds to the beverage. For example, common chelating compounds added to beverages include calcium disodium ethylenediaminetetraacetic acid (EDTA) or one or more of the polyphosphates, such as sodium hexametaphosphate (SHMP). In high-nutrient non-carbonated products, such as those beverages containing juice, vitamins, and/or minerals, the weak acids are more likely to exert inhibition if used in conjunction with preservative enhancers. Weak acid preservation systems, however, have limitations. Genetic adaptation and subsequent tolerance by microorganisms may be one of the biggest concerns. Certain yeasts, such as Zygosaccharomyces bailli, Zygosaccharomyces bisporus, Candida krusei, and Saccharomyces cerevisiae, have specific genes that enable them to resist the weak acid preservatives and grow, despite their presence and regardless of the co-presence of EDTA or SHMP (Piper, 2001; Pitt and Hocking, 2009). Some bacteria, such as Gluconobacter spp., are also thought to be preservative-resistant. The levels of weak acids necessary to overcome this resistance have been shown to be far beyond regulatory limits on use levels. Spoilage of preserved teas, juice-containing beverages, and carbonated beverages is commonly due to preservativeresistant yeasts. Weak acids are also known to impart a sensory impact in the form of throat or mouth burn when used at levels needed to prevent spoilage. Although there are certain shelf-stable beverages where this may be acceptable, often this sensory perception is considered negative. In addition, non-government organizations and also some international government agencies have raised concerns regarding the use of weak acid preservatives in beverages and foods. Finally, some consumers take note of ingredients and certain niche markets prefer preservative-free products. Natural antimicrobials may be alternatives to traditional ways of preserving beverages. Antimicrobials that could be labeled as natural could also eliminate hot-fill requirements for unpreserved shelf-stable preservative-free beverages. Thus, it would be desirable to provide a natural antimicrobial and/or an antimicrobial system that inhibits the growth of microorganisms. Various antimicrobials are capable of maintaining shelf-stable carbonated sodas either by killing yeast and bacteria at the point of production or by the inhibition of yeast and bacteria growth in the product at ambient temperature. Challenge testing is recommended to validate microbial control and shelf-stability. Sodium benzoate added at about 300 ppm alone is sometimes sufficient for the preservation of carbonated sodas with pH 3.5 or less. At lower pH such as 2.5, lower sodium benzoate concentration such as 175 ppm may be adequate. As stated above however, certain yeasts and some bacteria can be resistant to weak acid preservatives. Typically, this does not happen in carbonated sodas, but it is possible. In such cases, the addition of potassium sorbate and SHMP or potassium sorbate and EDTA can help enhance the antimicrobial properties of benzoate and suppress even preservative-resistant yeasts and bacteria. These factors would influence choice of use and usage level. TABLE 22.1 Recommended pH Range for Some Commonly Used Antimicrobials Antimicrobial Agent Benzoic acid Sorbic acid Propionic acid Acetic acid Parabens Sulfites Nitrites

Recommended pH Range 2.5–4.0 3.0–6.5 2.5–5.0 3.0–5.0 3.0–9.0 2.5–5.0 5.0–5.5

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Cinnamic Acid (ppm)

3.0 3.5 4.0

40 50 50

Lauric Arginate (ppm) 2.5 2.5 10

(Adapted from Dai et al., 2010.)

Natural preservation may be a viable option. Cinnamic acid is a natural preservative that has been researched as a preservative against yeast and bacteria in acidic beverages such as teas and juice-containing beverages. Efficacy varies by pH; lower concentrations are needed to preserve against yeast at lower pH. Typically, 60 ppm would be the minimum needed to inhibit yeasts. Similarly, lauric arginate ester (LAE) is a C-12 fatty acid-based molecule that can be used in conjunction with cinnamic acid to preserve beverages. Efficacious combinations of LAE with cinnamic acid at different pH levels are given in Table 22.2. In some applications, the use of dimethyl dicarbonate (DMDC) (see Chapter 13, “Dimethyl Dicarbonate and Diethyl Dicarbonate”) can be an effective alternative to treating packaged beverages at the point of filling and sealing because it is slightly soluble in water, extremely reactive, and will quickly undergo hydrolysis reactions with organic matter (e.g., microbial cells). The remaining byproducts are alcohol and carbon dioxide; thus, there are typically no labeling requirements when used in accordance with current regulatory agency-approved levels, and it is considered a processing aid. Hours after use, DMDC is not detectable in the product. The challenge is to destroy target microorganisms quickly before the hydrolysis is completed. DMDC was first approved for addition to wines as a yeast inhibitor at a level not to exceed 200 ppm. DMDC can be used in carbonated, dilute beverages containing juice, fruit flavor, or both, in which the juice content does not exceed 50%, at a limit of 250 ppm. In order to ensure efficacy, DMDC is supplied to each individual bottle after filling and immediately prior to capping. Therefore, dosing equipment is needed in order to ensure each bottle receives the proper dose and that caps are applied immediately so that product cannot be recontaminated after DMDC has dissipated. This can be a significant up-front expense. Molds like Penicillium can become problematic if the hygienic conditions of the plant and equipment are not maintained. Most molds will be inhibited by sodium benzoate and/or potassium sorbate, but certain Penicillium spp. can become tolerant to sorbic acid if allowed to persist in the environment or equipment. Once these resistant molds reach the product in a package, they will potentially overcome the preservative and produce a colony or a mass within the package.

22.3.3.3 Bakery Products Bakery products may require antimicrobials to inhibit the growth of spoilage organisms and increase shelf life. These products have relatively low protein content, and surface application would inhibit spoilage fungi. Negative aspects of bakery products as to antimicrobial application are a neutral pH and non-homogeneity of the food system. Calcium propionate is commonly used to prevent mold growth on shelf-stable bakery products. If bakery products contain fillings or sauces, the utilization of weak acid preservatives can prevent microbial spoilage for several days at ambient temperature. However, scientific support for bacterial pathogen control would also be needed for ambient-stored products with a pH and aw combination that falls within the potentially hazardous foods ranges. Some breads intended for convenience store (c-store) distribution or distribution to warm climates may include calcium propionate, sorbic acid, and natamycin to protect against spoilage by molds.

22.3.3.4 Fresh and Minimally Processed Fruits and Vegetables The application of antimicrobials to minimally processed fruits should be similar to application to fruit juices. Fruits generally have low lipid and protein content, which decreases the opportunity for interactions. They generally also have lower pH and lower aw in their edible flesh, rind, and/or skin than

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vegetables, which may stress the target microorganisms. The only negatives to application on fruits are potential non-homogeneity as to composition and topography, as well as contamination sites of fruits. Those with higher pH (e.g., cantaloupe) may show decreased efficacy of antimicrobials, since low pH would not be working in concert with the antimicrobials to inhibit target microorganisms. Cut fruits intended for several hours or days of shelf life, and used at food service establishments, are preserved by use of a solution of citric acid, calcium chloride, sodium benzoate, and potassium sorbate to prevent not only microbial spoilage but also enzymatic browning through polyphenoloxidase (PPO) denaturation. Vegetables and fresh produce generally have low lipid and protein content but have high pH and aw and are highly non-homogenous as to composition and site of contamination. Produce can be treated with antimicrobials to reduce pathogen counts. Cut produce is treated with antimicrobials such as chlorine or peroxyacetic acid (PAA) via flume water or by spray application (see Chapter 19, “Use of Antimicrobials as Processing Aids in Food Processing”). Chorine that has been stabilized by a phosphoric acid and propylene glycol mixture has been shown to enhance the efficacy against Salmonella and E. coli O157:H7 in fresh-cut lettuce processing (Nou et al., 2011).

22.3.3.5 Dairy Products Dairy products are very difficult systems in which to achieve activity of antimicrobials because these products have high pH, high aw, and high levels of protein, fat, and divalent cations as interfering compounds, all of which can interact to reduce the activity of antimicrobials. As a result, antimicrobials have been applied to dairy products with varying success. Nisin has efficacy against C. botulinum and other clostridia in cheese for spoilage prevention during ripening. Nisin may also be used in refrigerated dairy products like sour cream. Natamycin is commonly used as a mold-inhibitor on sliced and shredded cheese.

22.3.3.6 Meat and Poultry Products Meat and poultry products may be the commodities with the highest number of inherent properties that will limit effective application of antimicrobials. These include a non-homogenous substrate surface (particularly for meat animal carcasses but also for derived further processed food products), neutral pH, and high concentrations of lipids and proteins. In addition to commodity effects, products formulated with gums, phosphates, titanium dioxide, or other additives may tend to bind or inactivate antimicrobials, leading to diminished efficacy and much higher cost-in-use. Developing appropriate antimicrobial delivery systems (see Chapter 20, “Delivery Systems”), such as encapsulation (e.g., emulsions, nanoparticles) can reduce their interactions with food components and increase interaction with the target microorganisms and allow for controlled release of the antimicrobial during the shelf life of the food (Davidson et al., 2013).

22.3.3.6.1 Fresh Meats and Poultry Products Fresh meats and poultry products could be formulated with various ingredients in order to sustain fresh, desirable organoleptic properties, and restrict microbial growth. Many raw meats and poultry items are enhanced for moisture, tenderness, and yield with solutions containing sodium phosphates and sodium chloride. These are not preservatives per se, but they do impart a change (e.g., buffering) to the muscle, and the process of introducing these solutions into the muscle can result in the translocation of bacteria from exterior surfaces to internal meat. Whole-muscle products that are distributed and displayed refrigerated may be moisture-enhanced by needled injection or vacuum tumbling to incorporate preserving ingredients and antioxidants as well as salt, phosphates, and, in some products, flavorings. Sodium lactate has been used for this purpose in moisture-enhanced pork (Banks et al., 1998; Brewer et al., 1995). Antioxidants like rosemary extract can be incorporated into the brine solution to delay the oxidation of lipids. Topical treatments to fresh meat and poultry show promise in reducing bacterial pathogens and may be selected to reduce risk. The application of 200 to 400 ppm LAE to chicken filets caused a 0.7 to 1.0 log reduction of Salmonella (Sharma et al. 2013a), but no significant reduction of Salmonella occurred in ground chicken treated with 200 ppm LAE (Sharma et al., 2013b). Other treatments with octanoic acid, levulinic acid, and caprylic acid, either individually or in combination with other antimicrobials, have similar outcomes against pathogens in meat and poultry.

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22.3.3.6.2 Raw Ground Products Raw, ground products could be combined with antimicrobials like acidified sodium chlorite, as a means to cause a reduction of pathogens like E. coli O157:H7 (Visvalingam and Holley, 2018). Some novel uses of plant-derived antimicrobials like blueberry extract (Das et al., 2017) and combinations of lemon juice and essential oils derived from herbs (Chung et al., 2018) have had moderate success for pathogen reduction in fresh meats at the levels below sensory impact.

22.3.3.6.3 Cooked, Ready-to-Eat Meats Cooked, ready-to-eat (RTE) meats are examples of substrates in which antimicrobials have inhibitory activity against multiple pathogens and spoilage microorganisms simultaneously. It has been well-established that the organic salts such as potassium and sodium lactate and sodium diacetate are inhibitory to L. monocytogenes in RTE meat (Mbandi and Shelef, 2001). There are three main mechanisms of action of organic salts against the pathogen: (a) lowering of the aw of the meat (Chen and Shelef, 1992); (b) organic salts in the undissociated acid form pass across the bacterial cell membrane and disrupt pH balance; and (c) feedback inhibition occurs when lactate and acetate force a shift of L. monocytogenes toward fermentative production of acetoin and away from aerobic respiration and the production of lactate and acetate (Stasiewicz et al., 2011). Further, it has been shown that diacetate, lactate, and product moisture content significantly impacted the growth rate of the pathogen, whereas the influence of sodium chloride was not statistically significant (Legan et al., 2004; Seman et al., 2002). Studies in ground pork showed that pH, aw, and organic salt were significant factors in the growth of L. monocytogenes (Vandeven, 2004; Zuliani et al., 2007). The MICs of sodium lactate and sodium chloride against five strains of Staphylococcus aureus were researched (Houtsma et al., 1993). The MICs for sodium lactate ranged from 268 to 625 mM (or about 2.5 to 5.8%), while the MIC for NaCl was 1873 mM (about 10%). Other researchers showed that 5% sodium lactate in a broth solution extended the lag phase, diminished the overall growth rate, and reduced the stationary phase ultimate cell concentration of S. aureus compared to broth at equal pH and aw with 2.6% NaCl (de Wit and Rombouts, 1990). Further, S. aureus was inhibited on ham formulated with 1.8% sodium lactate when held at 1 or 6°C (Jofré et al., 2008). Various organic salts are antimicrobial to Clostridium perfringens in meat and poultry. Calcium lactate at 1.0% or potassium lactate and sodium lactate at ≥2.0% controlled C. perfringens germination and outgrowth to