Handbook of Essential Oils: Science, Technology, and Applications, 3rd Edition [Third ed.] 2020014781, 9780815370963, 9781351246460

Table of contents :
Cover
Half Title
Title Page
Copyright Page
Contents
Editors
Contributors
Chapter 1: Introduction
Chapter 2: History and Sources of Essential Oil Research
2.1 Ancient Historical Background
2.2 First Systematic Investigations
2.3 Research during the Last Half Century
2.3.1 Essential Oil Preparation Techniques
2.3.1.1 Industrial Processes
2.3.1.2 Laboratory-Scale Techniques
2.3.1.3 Microsampling Techniques
2.3.2 Chromatographic Separation Techniques
2.3.2.1 Thin-Layer Chromatography
2.3.2.2 GC
2.3.2.3 Liquid Column Chromatography
2.3.2.4 Supercritical Fluid Chromatography
2.3.2.5 Countercurrent Chromatography
2.3.3 Hyphenated Techniques
2.3.3.1 Gas Chromatography-Mass Spectrometry
2.3.3.2 High-Resolution GC-FTIR Spectroscopy
2.3.3.3 GC-UV Spectroscopy
2.3.3.4 Gas Chromatography-Atomic Emission Spectroscopy
2.3.3.5 Gas Chromatography-Isotope Ratio Mass Spectrometry
2.3.3.6 High-Performance Liquid Chromatography-Gas Chromatography
2.3.3.7 HPLC-MS, HPLC-NMR Spectroscopy
2.3.3.8 Supercritical Fluid Extraction–Gas Chromatography
2.3.3.9 Supercritical Fluid Chromatography-Gas Chromatography
2.3.3.10 Couplings of SFC-MS and SFC-FTIR Spectroscopy
2.3.4 Identification of Multicomponent Samples without Previous Separation
2.3.4.1 UV Spectroscopy
2.3.4.2 IR Spectroscopy
2.3.4.3 Mass Spectrometry
2.3.4.4 13C-NMR Spectroscopy
References
Chapter 3: Sources of Essential Oils
3.1 “Essential Oil–Bearing Plants”: Attempt of a Definition
3.2 Phytochemical Variation
3.2.1 Chemotaxonomy
3.2.2 Inter- and Intraspecific Variation
3.2.2.1 Lamiaceae (Labiatae) and Verbenaceae
3.2.2.2 Asteraceae (Compositae)
3.3 Identification of Source Materials
3.4 Genetic and Protein Engineering
3.5 Resources of Essential Oils: Wild Collection or Cultivation of Plants
3.5.1 Wild Collection and Sustainability
3.5.2 Domestication and Systematic Cultivation
3.5.3 Factors Influencing the Production and Quality of Essential Oil-Bearing Plants
3.5.3.1 Genetic Variation and Plant Breeding
3.5.3.2 Plant Breeding and Intellectual Property Rights
3.5.3.3 Intraindividual Variation between Plant Parts and Depending on the Developmental Stage (Morpho- and Ontogenetic Variation)
3.5.3.4 Environmental Influences
3.5.3.5 Cultivation Measures, Contaminations, and Harvesting
3.6 International Standards for Wild Collection and Cultivation
3.6.1 GA(C)P: Guidelines for Good Agricultural (and Collection) Practice of Medicinal and Aromatic Plants
3.6.2 ISSC-MAP: The International Standard on Sustainable Wild Collection of Medicinal and Aromatic Plants
3.6.3 FairWild
3.7 Conclusion
References
Chapter 4: Natural Variability of Essential Oil Components
4.1 Manifestation of Variability
4.2 Variability at Different Taxonomic Levels
4.2.1 Species
4.2.2 Populations
4.3 Connections of Chemical Diversity with Other Plant Characteristics
4.3.1 Propagation and Genetics
4.3.2 Morphological Characteristics
4.4 Morphogenetic and Ontogenetic Manifestation of the Chemical Variability
4.5 Origin of Essential Oil Variability
4.6 Chemotaxonomic Aspects
4.7 Considerations for Proper Assessment of Natural Variability
References
Chapter 5: Production of Essential Oils
5.1 Introduction
5.1.1 General Remarks
5.1.2 Definition and History
5.1.3 Production
5.1.4 Climate
5.1.5 Soil Quality and Soil Preparation
5.1.6 Water Stress and Drought
5.1.7 Insect Stress and Microorganisms
5.1.8 Location of Oil Cells
5.1.9 Types of Biomass Used
5.1.10 Timing of the Harvest
5.1.11 Agricultural Crop Establishment
5.1.12 Propagation from Seed and Clones
5.1.13 Commercial Essential Oil Extraction Methods
5.1.14 Expression
5.1.15 Steam Distillation
5.1.16 Concluding Remarks
Acknowledgments
References
Chapter 6: Chemistry of Essential Oils
6.1 Introduction
6.2 Basic Biosynthetic Pathways
6.3 Polyketides and Lipids
6.4 Shikimic Acid Derivatives
6.5 Terpenoids
6.5.1 Hemiterpenoids
6.5.2 Monoterpenoids
6.5.3 Sesquiterpenoids
6.6 Synthesis of Essential Oil Components
References
Chapter 7: Analysis of Essential Oils
7.1 Introduction
7.2 Classical Analytical Techniques
7.3 Modern Analytical Techniques
7.3.1 The Use of Gas Chromatography and Gas Chromatography-Mass Spectrometry (GC-MS), and Linear Retention Indices in Essential Oil Analysis
7.3.2 Fast Gas Chromatography for Essential Oil Analysis
7.3.3 Gas Chromatography-Olfactometry for the Assessment of Odor-Active Components of Essential Oils
7.3.4 Gas Chromatographic Enantiomeric Characterization of Essential Oils
7.3.5 Multidimensional Gas Chromatography (MDGC)
7.3.5.1 Multidimensional Gas Chromatographic Techniques (MDGC)
7.3.5.2 Multidimensional Preparative Gas Chromatography
7.3.5.3 Multidimensional Gas Chromatography Coupled to Isotope Ratio Mass Spectrometry (MDGC-IRMS)
7.3.6 Comprehensive Two-Dimensional Gas Chromatography and Multidimensional Liquid-Gas Chromatography
7.3.6.1 Analysis of Essential Oils through Comprehensive Two-Dimensional Gas Chromatography (GC × GC)
7.3.6.2 On-Line and Off-Line Coupled Liquid Chromatography-Gas Chromatography
7.3.7 Liquid Chromatography, Liquid Chromatography Hyphenated to Mass Spectrometry, and Multidimensional Liquid Chromatographic Technique in the Analysis of Essential Oils
7.4 General Considerations on Essential Oil Analysis
References
Chapter 8: Use of Linear Retention Indices in GC-MS Libraries for Essential Oil Analysis
8.1 Introduction
8.2 Retention Index Theories
8.3 Linear Retention Indices Present in the Databases
8.4 Accuracy of Retention Indices
8.5 Retention Index Compilations
8.6 Research Software
8.7 Application of Retention Indices in GC-FID and GC-MS Essential Oil Analyses
8.8 Gas Chromatographic Enantiomer Characterization Supported by Retention Indices
8.9 Retention Indices Applied to Multidimensional Gas Chromatographic Analysis
8.10 Concluding Remarks
References
Chapter 9: Safety Evaluation of Essential Oils: Constituent-Based Approach Utilized for Flavor Ingredients—An Update
9.1 Introduction
9.2 Constituent-Based Evaluation of Essential Oils
9.3 Scope of Essential Oils: Used as Flavor Ingredients
9.3.1 Plant Sources
9.3.2 Processing of Essential Oils for Flavor Functions
9.3.3 Chemical Composition and Congeneric Groups
9.3.4 Chemical Assay Requirements and Chemical Description of Essential Oil
9.3.4.1 Intake of the Essential Oil
9.3.4.2 Analytical Limits on Constituent Identification
9.3.4.3 Intake of Congeneric Groups
9.4 Safety Considerations for Essential Oils, Constituents, and Congeneric Groups
9.4.1 Essential Oils
9.4.1.1 Safety of Essential Oils: Relationship to Food
9.4.2 Safety of Constituents and Congeneric Groups in Essential Oils
9.5 Guide and Example for the Safety Evaluation of Essential Oils
9.5.1 Introduction
9.5.2 Elements of the Guide for the Safety Evaluation of the Essential Oil
9.5.2.1 Introduction
9.5.2.2 Prioritization of Essential Oil According to Presence in Food
9.5.2.3 Organization of Chemical Data: Congeneric Groups and Classes of Toxicity
9.5.3 Summary
References
Chapter 10: Metabolism of Terpenoids in Animal Models and Humans
10.1 Introduction
10.2 Metabolism of Monoterpenes
10.2.1 Borneol
10.2.2 Camphene
10.2.3 Camphor
10.2.4 3-Carene
10.2.5 Carvacrol
10.2.6 Carvone
10.2.7 1,4-Cineole
10.2.8 1,8-Cineole
10.2.9 Citral
10.2.10 Citronellal
10.2.11 p-Cymene
10.2.12 Fenchone
10.2.13 Geraniol
10.2.14 Limonene
10.2.15 Linalool
10.2.16 Linalyl Acetate
10.2.17 Menthofuran
10.2.18 Menthol
10.2.19 Menthone
10.2.20 Myrcene
10.2.21 a- and ß-Pinene
10.2.22 Pulegone
10.2.23 a-Terpineol
10.2.24 Terpinen-4-ol
10.2.25 a- and ß-Thujone
10.2.26 Thymol
10.3 Metabolism of Sesquiterpenes
10.3.1 Caryophyllene
10.3.2 Farnesol
10.3.3 Longifolene
10.3.4 Patchoulol
References
Chapter 11: Central Nervous System Effects of Essential Oil Compounds
11.1 Overview
11.2 Introduction
11.2.1 Translatability and Reproducibility
11.3 Review Methodology
11.3.1 Literature Review
11.3.2 Identification of EOs and/or their Compounds with Psychopharmacologic Action
11.4 Compounds from EOs with Psychopharmacology Potential
11.4.1 Alcohols
11.4.2 Phenols and Aromatic Methyl Ethers
11.4.3 Hydrocarbons
11.4.4 Carbonyl Compounds
11.4.5 Monoterpene Epoxides and Furanes
11.4.6 Nitrogenated Compounds
11.5 Compounds from EOs with Anxiolytic Properties
11.5.1 Assessing Anxiolytic Properties
11.5.1.1 Methods
11.5.1.2 Mechanisms of Action
11.5.2 Anxiolytic Properties of EO Compounds
11.6 Hypnotic Properties of Compounds from EOs
11.6.1 Assessing Hypnotic Properties
11.6.1.1 Methods
11.6.1.2 Mechanisms of Action
11.6.2 Hypnotic Properties of EO Compounds
11.7 Compounds Reported to Possess Anticonvulsant Effects
11.7.1 Assessing Antiepileptic Properties
11.7.1.1 Methods
11.7.1.2 Mechanisms of Action
11.7.2 Anticonvulsant Properties of EO Compounds
11.8 Antidepressant Properties
11.8.1 Assessing Antidepressant Properties
11.8.1.1 Methods
11.8.1.2 Mechanisms of Action
11.8.2 Antidepressant Properties of OE Compounds
11.9 Neuroprotector Properties of Compounds from EOs
11.9.1 Assessing Neuroprotective Properties
11.9.1.1 Methods
11.9.1.2 Mechanisms of Action
11.9.2 Neuroprotective Properties of EO Compounds
11.10 Concluding Remarks and Perspectives
References
Chapter 12: Effects of Essential Oils on Human Cognition
12.1 Introduction
12.2 Activation and Arousal: Definition and Neuroanatomical Considerations
12.3 Influence of Essential Oils and Fragrances on Brain Potentials Indicative of Arousal
12.3.1 Spontaneous EEG Activity
12.3.2 Contingent Negative Variation
12.4 Effects of Essential Oils and Fragrances on Selected Basic and Higher Cognitive Functions
12.4.1 Alertness and Attention
12.4.2 Learning and Memory
12.4.3 Other Cognitive Tasks
12.5 Conclusions
References
Chapter 13: Aromatherapy: An Overview and Global Perspectives
13.1 Introduction
13.2 The Image of Aromatherapy: Treat or Treatment?
13.3 Defining Aromatherapy: Is There Consensus?
13.4 Aroma Therapy or Aroma Care?
13.5 Aromatherapy: Scientific or Energetic?
13.6 An Aromatic Evolution: Different Aromatherapy Styles
13.6.1 Subtle Aromatherapy
13.6.2 Self-Care Aromatherapy
13.6.3 Holistic Aromatherapy
13.6.4 Cosmetic Aromatherapy
13.6.5 Clinical Aromatherapy
13.6.6 Medical Aromatherapy
13.7 Different Cultures, Different Aromatherapy Styles
13.8 Evaluating Efficacy: Does it Work?
13.9 Aromatherapy: An Evolving Therapy
13.9.1 Personalized Aroma Inhaler Devices
13.10 Aromatherapy: A Safe Healing Modality?
13.10.1 Safety to the Therapist
13.10.2 Safety to the Patient/Consumer
13.11 Conclusion: Moving Toward Greater Integration
References
Chapter 14: Essential Oils in Cancer Therapy
14.1 Introduction
14.2 Cancer Cell Lines
14.3 Tests to Assess Cytotoxic Activity
14.3.1 MTT Assay
14.3.2 TUNEL Assay
14.3.3 Flow Cytometry
14.4 EOs against Diverse Human Cancer Cell Lines
14.5 EOs against Breast Cancer
14.6 EOs against Prostate Cancer
14.7 EOs against Liver Cancer
14.8 EOs against Lung Cancer
14.9 EOs against Melanoma
14.10 EOs against Colorectal Cancer
14.11 EOs against Ehrlich Carcinoma
14.12 EOs against Glioblastoma
14.13 Conclusion
References
Chapter 15: Antimicrobial Activity of Selected Essential Oils and Aroma Compounds against Airborne Microbes
15.1 Introduction
15.2 Methods
15.2.1 General
15.2.2 Studies in Small Examination Rooms
15.2.2.1 Examined Essential Oils
15.2.2.2 Examined Aroma Compounds
15.2.3 Studies in Large Examination Rooms
15.2.3.1 Examined Essential Oils
15.2.3.2 Examined Aroma Compounds
15.3 Results
15.3.1 Studies in Small Examination Rooms
15.3.1.1 Results for Essential Oils
15.3.1.2 Results for Aroma Compounds
15.3.2 Studies in Large Examination Rooms
15.3.2.1 Results for Essential Oils
15.3.2.2 Results for Aroma Compounds
15.3.3 Identified Airborne Microbes
15.4 Discussion
15.5 Conclusion
Acknowledgement and Dedication
References
Chapter 16: Quorum Sensing and Essential Oils
16.1 Introduction
16.1.1 Cell-to-Cell Communication
16.1.2 Quorum Sensing: Definition
16.2 QS In Bacteria
16.2.1 Gram-Negative Bacteria
16.2.1.1 The LUXRI -System
16.2.1.2 Vibrio harveyi
16.2.1.3 Vibrio parahaemolyticus
16.2.1.4 Pseudomonas aeruginosa
16.2.1.5 Chromobacterium violaceum
16.2.2 Gram-Positive Bacteria
16.2.2.1 Staphylococcus aureus
16.3 QS In Fungi
16.3.1 EOs as Biofilm Inhibitors in C. albicans.?
16.4 QS In Saccharomyces cerevisiae
16.5 QS In Viruses
16.6 QS and Biofilm Formation
16.7 QS Inhibitors (QSIs)
16.7.1 Practical Applicability of QS Inhibitors
16.8 EOs as QS Inhibitors
16.8.1 Principle of QSI Activity Testing of EOs
16.8.2 Materials and Methods
16.8.2.1 Sensor Strains
16.8.2.2 QS-Inhibition Detection Assays
16.8.2.3 Biofilm Inhibition
16.8.2.4 Other Assays
16.8.3 Research on EOs as QSI
16.8.3.1 Clove, Cinnamon, Peppermint, and Lavender Oil
16.8.3.2 Rose, Lavender, Geranium and Rosemary Oil
16.8.3.3 Oils of Piper Species
16.8.3.4 EOs from Columbian Plants
16.8.3.5 Tea Tree and Rosemary Oils
16.8.3.6 Clary Sage, Juniper, Lemon, and Marjoram Oils
16.8.3.7 Clove, Rose, Chamomile, and Pine Turpentine Oils
16.8.3.8 Clove Oil
16.8.3.9 Oregano and Carvacrol
16.8.3.10 Lemongrass and Cinnamon Oils
16.8.3.11 EOs from South American Species
16.8.3.12 Lavender Oil
16.8.3.13 Lippia alba Oils
16.8.3.14 Ferula and Dorema Oils
16.8.3.15 Hyptis dilatata Oil
16.8.3.16 Peppermint Oil and Menthol
16.8.3.17 Cinnamon Oil
16.8.3.18 Coriander and (S)-(+)-Linalool
16.8.3.19 Cinnamon Oil
16.8.3.20 Spice Oil Nano-Emulsions
16.8.3.21 Eucalyptus Oils
16.8.3.22 Mandarin Oil
16.8.3.23 Thyme Oil, Thymol and Carvacrol
16.8.3.24 Murraya koenigii CO2-Extract—A Preclinical Infectious Model
16.8.3.25 Lavender, Rosemary, and Eucalyptus Oils
16.8.3.26 Green Cardamom EO
16.8.3.27 Corsican Mentha suaveolens ssp. insularis Oil and Others
16.8.3.28 South African Oils Plus a Gas Chromatography-Based Metabolomics Approach
16.8.3.29 Thymus daenensis and Satureja hortensis Oils
16.8.3.30 Carum copticum EO
16.8.3.31 Cinnamomum tamala oil Combined with DNase
16.9 Overview of the Results
16.9.1 Positively Tested Oils
16.9.1.1 Disc Diffusion Assay
16.9.1.2 Flask Incubation Assay
16.9.1.3 QS Inhibiting EOs Evaluated by Other Assays
16.9.2 Negatively Tested Oils
16.9.3 EOs Tested on the Inhibition of QS-Related Processes
16.10 Discussion
16.10.1 The QS Activity of EOs Can Only Be Valued Individually for the Respective Assay
16.10.2 An EO Being an Active Inhibitor of AHL-Based QS in One Strain is Not Necessarily Active in Another Strain Also Producing AHLs
16.10.3 EOs Often Not Only Inhibit QS but Also Inhibit Growth
16.10.4 Extraction Method and Collecting Site of the EOs Influences Test Results
16.11 Conclusion
16.12 Abbreviations
References
Chapter 17: Functions of Essential Oils and Natural Volatiles in Plant-Insect Interactions
17.1 Introduction
17.1.1 Brief Historical Overview of Essential Oil (EO) Functional Ecology
17.1.2 Challenges in Identifying EO Functional Roles and Selective Pressures
17.2 Aromatic Medicinal Herbs with Multiple EO Functions
17.2.1 Glandular Plants of Mediterranean Biomes
17.2.2 Focal Studies on Thymus vulgaris EO Chemotypes in the Garrigue
17.2.2.1 Biotic Interactions in a Geographic Mosaic
17.2.2.2 Geographic Variation and Abiotic Stress
17.2.2.3 Thymol in Nectar, Impacts on Bee Health
17.3 Functional Ecological Links between EOs and Resins
17.3.1 Oleoresin in Conifers and Multi-product TPS Enzymes
17.3.2 EOs, Resins, and TPS Enzymes in Angiosperms
17.3.3 Resins as Resources for Nest-Making Bees
17.3.3.1 Resin Collection from Aromatic Tropical Trees
17.3.3.2 Resin Collection from Dalechampia and Clusia Flowers
17.3.4 Cistus and Lavandula: Pollinator Hubs in the Phrygana
17.4 EO-Mediated Pollinator Specialization: Lessons from Aroids
17.4.1 Perfume-Collecting Male Orchid Bees as a Pollination Niche
17.4.2 Scent-Driven Pollinator Modes in the Araceae
References
Chapter 18: Essential Oils as Lures for Invasive Ambrosia Beetles
18.1 Introduction
18.2 Redbay Ambrosia Beetle
18.2.1 Background
18.2.2 Host Attractants
18.2.3 Field Lures
18.2.3.1 Manuka and Phoebe Oils
18.2.3.2 Cubeb Oil
18.2.3.3 Copaiba Oil and Enriched a-Copaene Oil
18.3 Chemical Analysis of the a-Copaene Lure
18.3.1 Method
18.3.2 Results
18.4 Euwallacea Shot-Hole Borers
18.4.1 Background
18.4.2 Host Attractants
18.4.3 Field Lures
18.4.3.1 Quercivorol
18.4.3.2 Enriched a-Copaene Lure
18.5 Other Applications
References
Chapter 19: Adverse Effects and Intoxication with Essential Oils
19.1 Introduction
19.1.1 General Side Effects
19.2 Camphor and Camphor-Containing Essential Oils
19.2.1 Case Reports
19.2.1.1 Dose Range for Oral Intoxication
19.2.1.2 Neurotoxic Effects
19.2.1.3 Effects Following Inhalative Application
19.3 Eucalyptus Oil
19.3.1 Case Reports
19.3.1.1 Dose Range for Oral Intoxication
19.3.1.2 Intoxication after Ingestion
19.3.1.3 Intoxication after Topical Application and Inhalation
19.3.2 Eucalyptol (1,8-Cineole)
19.4 Thujone-Containing Essential Oils
19.4.1 Case Reports
19.5 Peppermint Oil
19.5.1 Adulterations
19.5.2 Case Reports
19.5.3 Menthol
19.5.3.1 Cooling vs. Irritating Effect
19.5.3.2 Menthol-Induced Analgesia
19.5.3.3 Menthol and Tobacco-Related Chemicals
19.5.3.4 Menthol and Dermal Penetration
19.6 Pennyroyal Oil
19.6.1 Case Reports
19.6.2 Pulegone and Menthofuran
19.6.3 Precautions
19.7 Wintergreen Oil
19.7.1 Methyl Salicylate
19.8 Tea Tree Oil (Melaleuca Oil)
19.8.1 Toxicity Following Oral Exposure
19.8.2 Toxicity Following Dermal Exposure
19.8.3 Systemic Reactions
19.8.4 Ototoxicity
19.8.5 Developmental Toxicity
19.8.6 In Vitro. Toxicity
19.9 Sassafras Oil
19.9.1 Safrole
19.10 Clove Oil (Oleum Caryophylli., Caryophylli floris aetheroleum)
19.10.1 Case Reports
19.10.2 Eugenol, Isoeugenol
19.10.2.1 Repeated Dose Toxicity
19.10.2.2 Developmental and Reproductive Toxicity
19.10.2.3 Skin Sensitization
19.10.3 Methyleugenol
19.11 Bergamot Oil
19.12 Essential Oils of Nutmeg and Other Spices
19.13 Conclusion
References
Chapter 20: Adulteration of Essential Oils
20.1 Introduction
20.1.1 General Remarks
20.2 Definition and History
20.3 Adulteration
20.3.1 Unintended Adulteration
20.3.2 Intentional Adulteration
20.3.3 Prices
20.3.4 Availability
20.3.5 Demand of Clients
20.3.6 Regulations
20.3.7 Aging
20.3.8 Cupidity
20.3.9 Simple Sports?
20.4 Possible Adulterations for Essential Oils
20.4.1 Water
20.4.2 Ethanol
20.4.3 Fatty Oils or Mineral Oils
20.4.4 High Boiling Glycols
20.4.5 Oils from Other Parts of the Same Species or Other Species with Similar Essential Oil Composition
20.4.6 Related Botanical Species
20.4.7 Fractions of Essential Oils
20.4.8 Natural Isolates
20.4.9 Chemically Derived Synthetic Compounds, Which Are Proved to Appear in Nature
20.4.10 Steam Distilled Residues from Expression
20.4.11 Enzymatically Produced Chemicals (Natural by Law)
20.5 Methods to Detect Adulterations
20.5.1 Organoleptic Methods
20.5.1.1 Appearance and Color
20.5.1.2 Odor
20.5.1.3 Physical–Chemical Methods
20.5.1.4 Calculation of Relationship Coefficient
20.5.2 Analytical Methods
20.5.2.1 General Tests
20.5.2.2 Thin-Layer Chromatography
20.5.2.3 Gas Chromatography (GC, GLC, HRGC, GC-FID, GC-MS)
20.5.2.4 Chiral Analysis
20.5.2.5 GC-GC and GC.×.GC (Two-Dimensional Gas Chromatography, 2D GC)
20.5.2.6 13C NMR (Nuclear Magnetic Resonance)
20.6 Important Essential Oils and Their Possible Adulteration
20.6.1 Ambrette Seed Oil
20.6.2 Amyris Oil
20.6.3 Angelica Oils
20.6.4 Anise Fruit Oil
20.6.5 Armoise Oil
20.6.6 Basil Oils
20.6.7 Bergamot Oil
20.6.8 Bitter Orange Oil
20.6.9 Bitter Orange Petitgrain Oil
20.6.10 Cajeput Oil
20.6.11 Camphor Oil
20.6.12 Cananga Oil
20.6.13 Caraway Oil
20.6.14 Cardamom Oil
20.6.15 Cassia Oil
20.6.16 Cedar Leaf Oil
20.6.17 Cedarwood Oils
20.6.18 Celery Seed Oil
20.6.19 Chamomile Oil Blue
20.6.20 Chamomile Oil Roman
20.6.21 Cinnamon Bark Oil
20.6.22 Cinnamon Leaf Oil
20.6.23 Citronella Oil
20.6.24 Clary Sage Oil
20.6.25 Clove Oils
20.6.26 Coriander Fruit Oil
20.6.27 Corymbia Citriodora Oil
20.6.28 Corn Mint Oil
20.6.29 Cumin Fruit Oil
20.6.30 Cypress Oil
20.6.31 Dill Oils
20.6.32 Dwarf Pine Oil
20.6.33 Elemi Oil
20.6.34 Eucalyptus Oil
20.6.35 Fennel Oil Sweet
20.6.36 Fennel Oil Bitter
20.6.37 Geranium Oils
20.6.38 Grapefruit Oil
20.6.39 Juniper Berry Oil
20.6.40 Lavandin Oils
20.6.41 Lavender Oil
20.6.42 Lemon Oil
20.6.43 Lemongrass Oil
20.6.44 Lime Oil Distilled
20.6.45 Lime Oil Expressed
20.6.46 Litsea cubeba Oil
20.6.47 Mandarin Oil
20.6.48 Melissa Oil (Lemon Balm)
20.6.49 Mentha citrata Oil
20.6.50 Mountain Pine Oil
20.6.51 Neroli Oil
20.6.52 Nutmeg Oil
20.6.53 Orange Oil Sweet
20.6.54 Origanum Oil
20.6.55 Palmarosa Oil
20.6.56 Parsley Oil
20.6.57 Pine Oil Siberian
20.6.58 Patchouli Oil
20.6.59 Pepper Oil
20.6.60 Peppermint Oil
20.6.61 Petitgrain Oil Paraguay Type
20.6.62 Pimento Oils
20.6.63 Rose Oil
20.6.64 Rosemary Oil
20.6.65 Rosewood Oil
20.6.66 Sage Oil (Salvia officinalis)
20.6.67 Sage Oil Spanish Type
20.6.68 Sandalwood Oil
20.6.69 Spearmint Oils
20.6.70 Spike Lavender Oil
20.6.71 Star Anise Oil
20.6.72 Tarragon Oil
20.6.73 Tea Tree Oil
20.6.74 Thyme Oil
20.6.75 Turpentine Oil
20.6.76 Vetiver Oil
20.6.77 Ylang-Ylang Oils
References
Chapter 21: Essential Oils and Volatiles in Bryophytes
21.1 Introduction
21.2 Terpenoids and Other Volatiles from Bryophytes
21.2.1 Chemical Diversity of Natural Volatiles in Bryophytes
21.2.1.1 Liverwort Components
21.2.1.2 Volatile Components in Mosses and Hornworts
21.2.2 Microbial Terpene Synthase-Like Genes in Bryophytes
21.3 Significance for Chemotaxonomic Studies
21.3.1 Chemotaxonomic Value of Essential Oils
21.3.2 Chemotaxonomic Value of Volatile Extracts
21.4 Biological Functions
21.4.1 Chemical Defense
21.4.2 Allelopathic Activity
21.4.3 Medical Uses
21.5 Conclusions
References
Chapter 22: Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals
22.1 Introduction
22.2 Metabolic Pathways of Acyclic Monoterpenoids
22.2.1 Acyclic Monoterpene Hydrocarbons
22.2.1.1 Myrcene
22.2.1.2 Citronellene
22.2.2 Acyclic Monoterpene Alcohols and Aldehydes
22.2.2.1 Geraniol, Nerol, (+)- and (-)-Citronellol, Citral, and (+)- and (-)-Citronellal
22.2.2.2 Linalool and Linalyl Acetate
22.2.2.3 Dihydromyrcenol
22.3 Metabolic Pathways of Cyclic Monoterpenoids
22.3.1 Monocyclic Monoterpene Hydrocarbon
22.3.1.1 Limonene
22.3.1.2 Isolimonene
22.3.1.3 p-Menthane
22.3.1.4 1-p-Menthene
22.3.1.5 3-p-Menthene
22.3.1.6 a-Terpinene
22.3.1.7 -Terpinene
22.3.1.8 Terpinolene
22.3.1.9 a-Phellandrene
22.3.1.10 p-Cymene
22.3.2 Monocyclic Monoterpene Aldehyde
22.3.2.1 Perillaldehyde
22.3.2.2 Phellandral and 1,2-Dihydrophellandral
22.3.2.3 Cuminaldehyde
22.3.3 Monocyclic Monoterpene Alcohol
22.3.3.1 Menthol
22.3.3.2 Neomenthol
22.3.3.3 (+)-Isomenthol
22.3.3.4 Isopulegol
22.3.3.5 a-Terpineol
22.3.3.6 (-)-Terpinen-4-ol
22.3.3.7 Thymol and Thymol Methyl Ether
22.3.3.8 Carvacrol and Carvacrol Methyl Ether
22.3.3.9 Carveol
22.3.3.10 Dihydrocarveol
22.3.3.11 Piperitenol
22.3.3.12 Isopiperitenol
22.3.3.13 Perillyl Alcohol
22.3.3.14 Carvomenthol
22.3.4 Monocyclic Monoterpene Ketone
22.3.4.1 a, ß-Unsaturated Ketone
22.3.4.2 Saturated Ketone
22.3.4.3 Cyclic Monoterpene Epoxide
22.4 Metabolic Pathways of Bicyclic Monoterpenoids
22.4.1 Bicyclic Monoterpene
22.4.1.1 a-Pinene
22.4.1.2 ß-Pinene
22.4.1.3 (±)-Camphene
22.4.1.4 3-Carene and Carane
22.4.2 Bicyclic Monoterpene Aldehyde
22.4.2.1 Myrtenal and Myrtanal
22.4.3 Bicyclic Monoterpene Alcohol
22.4.3.1 Myrtenol
22.4.3.2 Myrtanol
22.4.3.3 Pinocarveol
22.4.3.4 Pinane-2,3-Diol
22.4.3.5 Isopinocampheol (3-Pinanol)
22.4.3.6 Borneol and Isoborneol
22.4.3.7 Fenchol and Fenchyl Acetate
22.4.3.8 Verbenol
22.4.3.9 Nopol and Nopol Benzyl Ether
22.4.4 Bicyclic Monoterpene Ketones
22.4.4.1 a-, ß-Unsaturated Ketone
22.4.4.2 Saturated Ketone
22.5 Summary
22.5.1 Metabolic Pathways of Monoterpenoids by Microorganisms
22.5.2 Microbial Transformation of Terpenoids as Unit Reaction
References
Chapter 23: Biotransformation of Sesquiterpenoids, Ionones, Damascones, Adamantanes, and Aromatic Compounds by Green Algae, Fungi, and Mammals
23.1 Introduction
23.2 Biotransformation of Sesquiterpenoids by Microorganisms
23.2.1 Highly Efficient Production of Nootkatone (2) from Valencene (1)
23.2.2 Biotransformation of Valencene (1) by .Aspergillus niger and Aspergillus wentii
23.2.3 Biotransformation of Nootkatone (2) by .Aspergillus niger
23.2.4 Biotransformation of Nootkatone (2) by Fusarium culmorum and Botryosphaeria dothidea
23.2.5 Biotransformation of (+)-1(10)-Aristolene (36) from the Crude Drug Nardostachys chinensis by Chlorella fusca., Mucor species, and .Aspergillus niger
23.2.6 Biotransformation of Various Sesquiterpenoids by Microorganisms
23.3 Biotransformation of Sesquiterpenoids by Mammals, Insects, and Cytochrome P-450
23.3.1 Animals (Rabbits) and Dosing
23.3.2 Sesquiterpenoids
23.4 Biotransformation of Ionones, Damascones, and Adamantanes
23.5 Biotransformation of Aromatic Compounds
References
Chapter 24: Use of Essential Oils in Agriculture
24.1 Introduction
24.1.1 Essential Oils: Very Complex Natural Mixes
24.2 Essential Oils as Antipests
24.2.1 Health and Environmental Impact of Botanical Antipests
24.2.2 Pesticidal and Repellent Action of Essential Oils
24.2.2.1 Insecticidal Activities of Essential Oils
24.2.3 Development and Commercialization of Botanical
24.2.3.1 Examples of Essential Oils Used as Antipests
24.3 Essential Oils as Herbicides
24.3.1 Phytotoxicity
24.3.2 Prospects of Organic Weed Control
24.3.3 Examples of Essential Oils in Weed Control
24.3.3.1 Thymus vulgaris
24.3.3.2 Mentha sp.
24.3.3.3 Cymbopogon sp.
24.3.3.4 Eucalyptus sp.
24.3.3.5 Lavandula sp.
24.3.3.6 Origanum sp.
24.3.3.7 Artemisia scoparia
24.3.3.8 Zataria multiflora
24.3.3.9 Tanacetum sp.
24.4 Essential Oils as Inhibitors of Various Pests
24.4.1 Effect on Bacteria
24.4.2 Effect on Fungi
24.4.3 Effect on Viruses
24.4.4 Effect on Nematodes
24.5 Effect of Essential Oils on the Condition of the Soil
24.5.1 Effects of Essential Oils on Microorganisms and Soil
24.5.2 Examples of Essential Oils with an Effect on Soil Conditions
24.5.2.1 Mentha spicata
24.5.2.2 Lavandula sp.
24.5.2.3 Salvia sp.
24.5.2.4 Myrtus communis
24.5.2.5 Laurus nobilis
24.5.2.6 Cymbopogon sp.
24.6 EOs Used in Post-Harvest Disease Control
24.6.1 Effects of Essential Oils on Stored-Product Pests
24.6.2 Examples of Essential Oils Used on Stored Products
24.6.2.1 Thymus zygis
24.6.2.2 Cinnamomum. sp
24.6.2.3 Cymbopogon citratus
24.6.2.4 Laurus nobilis
24.7 Conclusion
References
Chapter 25: Essential Oils Used in Veterinary Medicine
25.1 Introduction
25.2 Oils Attracting Animals
25.3 Oils Repelling Animals
25.4 Oils against Pests
25.4.1 Insecticidal, Pest Repellent, and Antiparasitic Oils
25.4.2 Fleas and Ticks
25.4.3 Mosquitoes
25.4.4 Moths
25.4.5 Aphids, Caterpillars, and Whiteflies
25.4.5.1 Garlic Oil
25.4.6 Ear Mites
25.4.7 Antiparasitic
25.5 Essential Oils Used in Animal Feed
25.5.1 Ruminants
25.5.2 Poultry
25.5.2.1 Studies with CRINA® Poultry
25.5.2.2 Studies with Herbromix®
25.5.3 Pigs
25.6 Essential Oils Used in Treating Diseases in Animals
References
Chapter 26: Encapsulation and Other Programmed/Sustained-Release Techniques for Essential Oils and Volatile Terpenes
26.1 Introduction
26.2 Controlled Release of Volatiles
26.3 Use of Hydrophilic Polymers
26.4 Alginate
26.5 Stabilization of Essential Oil Constituents
26.6 Controlled Release of Volatiles from Nonvolatile Precursors
26.7 Cyclodextrin Complexation of Volatiles
26.8 Enhanced Biological Effect by Prolonged Delivery of Volatiles and Essential Oils
26.9 Methods for Producing Prolonged Delivery Units of Volatiles
26.10 Presenting Volatiles in Nano-Emulsions
26.11 Concluding Remarks
References
Chapter 27: Essential Oils as Carrier Oils
27.1 Introduction
27.2 Essential Oils in General
27.3 Essential Oils as Penetration Enhancers
27.4 Terpenes
27.4.1 Classes and Complexes
27.4.2 Hydrophilic, Lipophilic, Amphiphilic Terpenes
27.4.3 Structure of Terpenes
27.4.4 Boiling Point of Terpenes
27.4.5 Concentration of Terpenes
27.4.6 Increase of Terpenes
27.4.7 Vehicles of Terpenes
27.5 Skin
27.5.1 Layers
27.5.2 Stratum Corneum (SC)
27.5.3 Skin Models
27.5.4 Franz Cell
27.6 Advantages
27.6.1 Advantages of Natural Penetration Enhancers
27.6.2 Advantages of Transdermal Drug Delivery
27.7 Side Effects of Natural Penetration Enhancers
27.7.1 Skin Irritancy and Toxicity
27.7.2 Transepidermal Water Loss (TEWL)
27.8 Mechanism of Action
27.8.1 Effect on Stratum Corneum Lipids
27.8.2 Effect on Hydrogen Bond Networks
27.8.3 Effect on SC Partition of Drugs
27.8.4 Affecting Factors
27.8.5 Screening-Techniques
27.9 Combination of Penetration Enhancers and Drugs
27.9.1 Essential Oils and Anti-Inflammatory Drugs
27.9.1.1 Chuanxiong Oil and Ibuprofen
27.9.1.2 Rosemary Essential Oil and Na-Diclofenac
27.9.1.3 Alpinia oxyphylla Essential Oil and Indomethacin
27.9.1.4 Lippia sidoides Essential Oil and Salicylic Acid
27.9.1.5 Sweet Basil Oil and Indomethacin
27.9.1.6 Zanthoxylum bungeanum Essential Oil and Indomethacin/5-Fluorouracil
27.9.2 Essential Oils and Antiseptic Drugs
27.9.2.1 1,8-Cineole and Chlorhexidine
27.9.3 Essential Oils and Vesicular Carriers
27.9.3.1 Terpenes and Ultra-Deformable Liposomes of Sodium Fluorescein
27.9.3.2 Terpenes and Liposomes of Antisense Oligonucleotide
27.9.3.3 Eucalyptus Oil and Transferosomes of Ketoconazole
27.9.3.4 Speranskia tuberculata Essential Oil and Glycerosomes of Paeoniflorin
27.9.4 Essential Oils and Cytostatic Drugs
27.9.4.1 Myrica rubra Essential Oil and Doxorubicin
27.9.4.2 Mentha x villosa Essential Oil and 5-Fluorouracil
27.9.4.3 Zanthoxylum bungeanum Essential Oil and 5-Fluorouracil (5-FU)/Indomethacin
27.9.5 Essential Oils and Cardiovascular Drugs
27.9.5.1 Zanthoxylum bungeanum Essential Oil and Osthole/Tetramethylpyrazine/Ferulic Acid/Puerarin/Geniposide
27.9.5.2 Eucalyptus Oil and Tetramethylpyrazine
27.9.5.3 Basil Oil and Labetolol Hydrochloride
27.9.5.4 Anethol, Menthone, Eugenol, and Valsartan
27.9.5.5 Basil Oil, Petit Grain Oil, Thyme Oil, and Nitrendipine
27.10 Essential Oils and the Influence of Temperature
27.10.1 Borneol, Osthole, and Increasing Temperature
27.11 Essential Oils and the Effect on Cytochrome P450
27.11.1 Zataria multiflora. Essential Oils with Cancer Chemopreventive Effect
27.12 Essential Oils and their Synergistic Effects
27.12.1 1,8-Cineole and Camphor
27.13 Conclusion
References
Chapter 28: Influence of Light on Essential Oil Constituents
28.1 Introduction
28.2 Mechanisms of Photodegradation
28.2.1 Double-Bond Isomerization
28.2.2 Photooxidation and Epoxidation Reactions
28.2.3 Polymerization Reactions
28.2.4 Other Reaction Mechanisms
28.2.5 Influence of Reaction Conditions
28.2.5.1 Effect of Solvent
28.2.5.2 Effect of Duration and Intensity
28.3 Phototoxicity
28.3.1 Methods for the in vitro Assessment of Phototoxicity
28.3.1.1 3T3 NRU Assay
28.3.1.2 Photohemolysis Test and RBC PT
28.3.1.3 Reconstituted 3D Human Skin Models
28.3.1.4 Photoirritation Factor (PIF) and Mean Photo Effect (MPE)
28.3.2 Phototoxic Essential Oils and Essential Oil Constituents
28.4 Conclusion
References
Chapter 29: Influence of Air on Essential Oil Constituents
29.1 Introduction
29.2 Essential Oils and Terpenoids
29.3 Reaction Mechanisms of Terpenoid Oxidation Patterns
29.3.1 Linalool
29.3.1.1 The Ene-Type Mechanism
29.3.1.2 The Free-Radical Chain Reaction
29.3.1.3 The Direct Reaction Pathway
29.3.2 Geraniol
29.3.2.1 Initiation Phase
29.3.2.2 Propagation Step 1
29.3.2.3 Propagation Step 2
29.4 Chemistry of Fragrance Terpenoid Autoxidation
29.4.1 Linalool
29.4.2 Linalyl Acetate
29.4.3 ß-Caryophyllene
29.4.4 Lavender Oil
29.4.5 Limonene
29.4.6 Geraniol
29.4.7 Geranial
29.4.8 Tea Tree Oil and a-Terpinene
29.5 Conclusion
References
Chapter 30: The Essential Oil Trade
Chapter 31: Industrial Uses of Essential Oils
Acknowledgments
References
Further Reading
Web Sites
Chapter 32: Storage, Labeling, and Transport of Essential Oils
32.1 Marketing of Essential Oils: The Fragrant Gold of Nature Postulates Passion, Experience, and Knowledge
32.2 Impact and Consequences on the Classification of Essential Oils as Natural but Chemical Substances IN REACH
32.3 Dangerous Substances and Dangerous Goods
32.3.1 Material Safety Data Sheet
32.4 Packaging of Dangerous Goods
32.5 Labeling
32.6 List of Regulations for the Consideration of Doing Business in the EU
Acronyms
References
Chapter 33: Recent EU Legislation on Flavours and Fragrances and Its Impact on Essential Oils
33.1 Introduction
33.2 Former Flavouring Directive and Current Flavouring Regulation: Impact on Essential Oils
33.2.1 Maximum Levels of “Restricted Substances”
33.2.1.1 (Restricted Substances under Former) Flavouring Directive 88/388/EC
33.2.1.2 (Restricted Substances under) Current Flavouring Regulation 1334/2008/EC
33.2.2 Definition of “Natural”
33.2.2.1 (Definition of “Natural” under) Former Flavouring Directive 88/388/EC
33.2.2.2 (Definition of “Natural” under) Current Flavouring Regulation 1334/2008/EC
33.3 Conclusion
33.A Appendix
References
Index

Citation preview

Handbook of Essential Oils

Handbook of Essential Oils Science, Technology, and Applications Third Edition

Edited by

K. Hüsnü Can Başer Gerhard Buchbauer

Third edition published 2020 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 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 p ­ ublishers 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 [email protected] 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: Başer, K. H. C. (Kemal Hüsnü Can), editor. | Buchbauer, Gerhard, editor. Title: Handbook of essential oils : science, technology, and applications / [edited by] K. Hüsnü Can Baser, Gerhard Buchbauer. Description: Third edition. | Boca Raton : CRC Press, [2020] | Includes bibliographical references and index. Identifiers: LCCN 2020014781 | ISBN 9780815370963 (hardback) | ISBN 9781351246460 (ebook) Subjects: LCSH: Essences and essential oils--Handbooks, manuals, etc. Classification: LCC QD416.7 .H36 2020 | DDC 661/.806--dc23 LC record available at https://lccn.loc.gov/2020014781 ISBN: 9780815370963 (hbk) ISBN: 9781351246460 (ebk) Typeset in Times LT Std by Nova Techset Private Limited, Bengaluru & Chennai, India

Contents Editors................................................................................................................................................ix Contributors.......................................................................................................................................xi Chapter 1 Introduction...................................................................................................................1 K. Hüsnü Can Başer and Gerhard Buchbauer Chapter 2 History and Sources of Essential Oil Research............................................................3 Karl-Heinz Kubeczka Chapter 3 Sources of Essential Oils............................................................................................. 41 Chlodwig Franz and Johannes Novak Chapter 4 Natural Variability of Essential Oil Components....................................................... 85 Éva Németh-Zámbori Chapter 5 Production of Essential Oils...................................................................................... 125 Erich Schmidt Chapter 6 Chemistry of Essential Oils...................................................................................... 161 Charles Sell Chapter 7 Analysis of Essential Oils......................................................................................... 191 Adriana Arigò, Mariosimone Zoccali, Danilo Sciarrone, Peter Q. Tranchida, Paola Dugo, and Luigi Mondello Chapter 8 Use of Linear Retention Indices in GC-MS Libraries for Essential Oil Analysis....... 229 Emanuela Trovato, Giuseppe Micalizzi, Paola Dugo, Margita Utczás, and Luigi Mondello Chapter 9 Safety Evaluation of Essential Oils: Constituent-Based Approach Utilized for Flavor Ingredients—An Update................................................................................ 253 Sean V. Taylor Chapter 10 Metabolism of Terpenoids in Animal Models and Humans..................................... 275 Walter Jäger and Martina Höferl Chapter 11 Central Nervous System Effects of Essential Oil Compounds................................. 303 Elaine Elisabetsky and Domingos S. Nunes v

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Chapter 12 Effects of Essential Oils on Human Cognition......................................................... 345 Eva Heuberger Chapter 13 Aromatherapy: An Overview and Global Perspectives............................................ 373 Rhiannon Lewis Chapter 14 Essential Oils in Cancer Therapy.............................................................................. 395 Carmen Trummer and Gerhard Buchbauer Chapter 15 Antimicrobial Activity of Selected Essential Oils and Aroma Compounds against Airborne Microbes........................................................................................ 415 Sabine Krist Chapter 16 Quorum Sensing and Essential Oils.......................................................................... 427 Isabel Charlotte Soede and Gerhard Buchbauer Chapter 17 Functions of Essential Oils and Natural Volatiles in Plant-Insect Interactions........ 481 Robert A. Raguso Chapter 18 Essential Oils as Lures for Invasive Ambrosia Beetles............................................. 497 Paul E. Kendra, Nurhayat Tabanca, Wayne S. Montgomery, Jerome Niogret, David Owens, and Daniel Carrillo Chapter 19 Adverse Effects and Intoxication with Essential Oils............................................... 517 Rosa Lemmens-Gruber Chapter 20 Adulteration of Essential Oils................................................................................... 543 Erich Schmidt and Jürgen Wanner Chapter 21 Essential Oils and Volatiles in Bryophytes............................................................... 581 Agnieszka Ludwiczuk and Yoshinori Asakawa Chapter 22 Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals................................................................................................................... 613 Yoshiaki Noma and Yoshinori Asakawa Chapter 23 Biotransformation of Sesquiterpenoids, Ionones, Damascones, Adamantanes, and Aromatic Compounds by Green Algae, Fungi, and Mammals.......................... 769 Yoshinori Asakawa and Yoshiaki Noma

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Chapter 24 Use of Essential Oils in Agriculture......................................................................... 873 Catherine Regnault-Roger, Susanne Hemetsberger, and Gerhard Buchbauer Chapter 25 Essential Oils Used in Veterinary Medicine............................................................. 919 K. Hüsnü Can Başer and Chlodwig Franz Chapter 26 Encapsulation and Other Programmed/Sustained-Release Techniques for Essential Oils and Volatile Terpenes......................................................................... 933 Jan Karlsen Chapter 27 Essential Oils as Carrier Oils.................................................................................... 943 Romana Aichinger and Gerhard Buchbauer Chapter 28 Influence of Light on Essential Oil Constituents...................................................... 961 Marie-Christine Cudlik and Gerhard Buchbauer Chapter 29 Influence of Air on Essential Oil Constituents......................................................... 989 Darija Gajić and Gerhard Buchbauer Chapter 30 The Essential Oil Trade........................................................................................... 1023 Hugo Bovill Chapter 31 Industrial Uses of Essential Oils............................................................................. 1029 W. S. Brud Chapter 32 Storage, Labeling, and Transport of Essential Oils................................................. 1041 Jens Jankowski, Jens-Achim Protzen, and Klaus-Dieter Protzen Chapter 33 Recent EU Legislation on Flavours and Fragrances and Its Impact on Essential Oils...................................................................................................... 1055 Jan C. R. Demyttenaere Index............................................................................................................................................. 1067

Editors K. Hüsnü Can Başer was born on July 15, 1949, in Çankırı, Turkey. He graduated from the Eskisehir I.T.I.A. School of Pharmacy with diploma number 1 in 1972 and became a research assistant in the Pharmacognosy Department of the same school. He did his PhD in pharmacognosy between 1974 and 1978 at Chelsea College of the University of London. Upon returning home, he worked as a lecturer in pharmacognosy at the school from which he had earlier graduated and served as director of Eskisehir I.T.I.A. School of Chemical Engineering between 1978 and 1980. He was promoted to associate professorship in pharmacognosy in 1981. He served as dean of the Faculty of Pharmacy at Anadolu University (1993–2001), vice-dean of the Faculty of Pharmacy (1982–1993), head of the Department of Professional Pharmaceutical Sciences (1982–1993), head of the Pharmacognosy Department (1982–2011), member of the University Board and Senate (1982–2001; 2007), and director of the Medicinal and Aromatic Plant and Drug Research Centre (TBAM) (1980–2002) in Anadolu University. During 1984–1994, he was appointed as the national project coordinator of Phase I and Phase II of the UNDP/UNIDO projects of the government of Turkey titled “Production of Pharmaceutical Materials from Medicinal and Aromatic Plants,” through which TBAM had been strengthened. He was promoted to full professorship in pharmacognosy in 1987. After his early retirement from Anadolu University in 2011, he served as visiting professor in King Saud University in Riyadh, Saudi Arabia (2011–2015). He is currently working as head of the Pharmacognosy Department in the Faculty of Pharmacy of Near East University in Nicosia, N. Cyprus, and director of the Graduate Institute of Health Sciences of the same university. His major areas of research include essential oils, alkaloids, and biological, chemical, pharmacological, technological, and biological activity research into natural products. He is the 1995 Recipient of the Distinguished Service Medal of IFEAT (International Federation of Essential Oils and Aroma Trades) based in London, United Kingdom, and the 2005 recipient of “Science Award” (Health Sciences) of the Scientific and Technological Research Council of Turkey (TUBITAK), which are among the 18 awards he has been bestowed so far. He is listed among the 100 Turks Leading Science. He has published 827 research papers in international refereed journals, 184 papers in Turkish journals, and 141 papers in conference proceedings. He has published altogether 1212 scientific contributions as papers, books, or book chapters. According to SCI, his 594 papers were cited 10,453 times. His H-index is 47. According to Google Scholar, his publications were cited 26,540 times. His H-index is 70; i10 index is 552. His 5 books were published in Turkey, Japan, the UK, and the US (2). More information can be found at http://www.khcbaser.com. Gerhard Buchbauer was born in 1943 in Vienna, Austria. He studied pharmacy at the University of Vienna, from where he received his master’s degree (Mag. pharm.) in May 1966. In September 1966, he assumed the duties of university assistant at the Institute of Pharmaceutical Chemistry and received his doctorate (PhD) in pharmacy and philosophy in October 1971, with a thesis on synthetic fragrance compounds. Further scientific education was practiced postdoc in the team of Professor C. H. Eugster at the Institute of Organic Chemistry, University of Zurich (1977–1978), followed by the habilitation (postdoctoral lecture qualification) in pharmaceutical chemistry with the inaugural dissertation entitled “Synthesis of Analogies of Drugs and Fragrance Compounds with Contributions to Structure-Activity-Relationships” (1979) and appointment as permanent staff of the University of Vienna and head of the first department of the Institute of Pharmaceutical Chemistry. In November 1991, he was appointed as a full professor of pharmaceutical chemistry, University of Vienna; in 2002, he was elected as head of this institute. He retired in October 2008. He has been married since 1973 and had a son in 1974.

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Among others, he is still a member of the permanent scientific committee of International Symposium on Essential Oils (ISEO); a member of the scientific committee of Forum Cosmeticum (1990, 1996, 2002, and 2008); a member of numerous editorial boards (e.g., Journal of Essential Oil Research, the International Journal of Essential Oil Therapeutics, Scientia Pharmaceutica); assistant editor of Flavour and Fragrance Journal; regional editor of Eurocosmetics; a member of many scientific societies (e.g., Society of Austrian Chemists, head of its working group “Food Chemistry, Cosmetics, and Tensides” [2000–2004]; Austrian Pharmaceutical Society; Austrian Phytochemical Society; vice head of the Austrian Society of Scientific Aromatherapy; and so on); technical advisor of IFEAT (1992–2008); and organizer of the 27th ISEO (September 2006, in Vienna) together with Professor Dr. Ch. Franz and senior advisor at the 50th ISEO (September 2019), organized in Vienna again. Based on the sound interdisciplinary education of pharmacists, it was possible to establish an almost completely neglected area of fragrance and avor chemistry as a new research discipline within the pharmaceutical sciences. Our research team is the only one that conducts fragrance research in its entirety and covers synthesis, computer-aided fragrance design, analysis, and pharmaceutical/ medicinal aspects. Because of our efforts, it is possible to show and to prove that these small molecules possess more properties than merely emitting a good odor. Now, this research team has gained a worldwide scientific reputation documented by more than 450 scientific publications, about 100 invited lectures, and about 200 contributions to symposia, meetings, and congresses, as short lectures and poster presentations.

Contributors Romana Aichinger Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Adriana Arigò Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy

Paola Dugo Chromaleont SRL, c/o Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy and Unit of Food Science and Nutrition Department of Medicine University Campus Bio-Medico of Rome Rome, Italy

Yoshinori Asakawa Institute of Pharmacognosy Tokushima Bunri University Tokushima, Japan

Elaine Elisabetsky Department of Biochemistry Universidade Federal Do Rio Grande do Sul Porto Alegre, Rio Grande do Sul, Brazil

Hugo Bovill Ajowan Consulting Bury St Edmunds, Suffolk, United Kingdom

Chlodwig Franz Institute of Animal Nutrition and Functional Plant Compounds University of Veterinary Medicine Vienna, Austria

W. S. Brud Polskie Towarzystwo Aromaterapeutyczne Warszawa, Poland Daniel Carrillo Tropical Research and Education Center University of Florida Homestead, Florida Marie-Christine Cudlik Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Jan C. R. Demyttenaere Director, Scientific & Regulatory Affairs EFFA/IOFI Brussels, Belgium

Darija Gajić Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Susanne Hemetsberger Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Eva Heuberger Pfarrer-Lauer-Strasse St Ingbert, Germany

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Martina Höferl Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Walter Jäger Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Jens Jankowski (deceased) Joh. Vögele KG Lauffen a.N., Germany Jan Karlsen Department of Pharmaceutics University of Oslo Oslo, Norway Paul E. Kendra United States Department of Agriculture, Agricultural Research Service (USDA-ARS) Subtropical Horticulture Research Station (SHRS) Miami, Florida Sabine Krist Lehrstuhl für Medizinische Chemie Medizinische Fakultät Sigmund Freud Privat-Universität and Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Karl-Heinz Kubeczka Untere Steigstraße Margetshöchheim, Germany Rosa Lemmens-Gruber Department of Pharmacology Center of Pharmacy University of Vienna Vienna, Austria

Contributors

Rhiannon Lewis Chemin Les Achaps La Martre, France Agnieszka Ludwiczuk Independent Laboratory of Natural Products Chemistry, Chair and Department of Pharmacognosy Medical University of Lublin Lublin, Poland Giuseppe Micalizzi Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italyand and Center of Sports Nutrition Science University of Physical Education Budapest, Hungary Luigi Mondello Chromaleont SRL, c/o Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy and Unit of Food Science and Nutrition Department of Medicine University Campus Bio-Medico of Rome Rome, Italy and BeSep SRL, c/o Department of Chemistry, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy Wayne S. Montgomery United States Department of Agriculture, Agricultural Research Service (USDA-ARS) Subtropical Horticulture Research Station Miami, Florida Éva Németh-Zámbori Department of Medicinal and Aromatic Plants Szent István University Budapest, Hungary

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Contributors

Jerome Niogret Niogret Ecology Consulting LLC Miami, Florida Yoshiaki Noma Shinkirai, Kitajima-cho Tokushima, Japan Johannes Novak Institute of Animal Nutrition and Functional Plant Compounds University of Veterinary Medicine Vienna, Austria Domingos S. Nunes Department of Chemistry Universidade Estadual de Ponta Grossa, Ponta Grossa, Paraná, Brazil David Owens Carvel Research & Education Center University of Delaware Georgetown, Delaware Jens-Achim Protzen Paul Kaders GmbH Hamburg, Germany Klaus-Dieter Protzen Paul Kaders GmbH Hamburg, Germany Robert A. Raguso Department of Neurobiology and Behavior Cornell University Ithaca, New York Catherine Regnault-Roger Professor emeritus Institute of Interdisciplinary Research on Environment and Materials UPPA University of Pau et des pays de l’Adour Pau, France Erich Schmidt Consultant Essential oils Nördlingen, Germany Danilo Sciarrone Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy

Charles Sell Parsonage Farm Church Lane Kent, England Isabel Charlotte Soede Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria Nurhayat Tabanca United States Department of Agriculture, Agricultural Research Service (USDA-ARS) Subtropical Horticulture Research Station (SHRS) Miami, Florida Sean V. Taylor Flavor & Extract Manufacturers Association and The International Organization of the Flavor Industry Washington, DC Peter Q. Tranchida Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy Emanuela Trovato Chromaleont SRL, c/o Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy Carmen Trummer Department of Pharmaceutical Chemistry Division of Clinical Pharmacy and Diagnostics Center of Pharmacy University of Vienna Vienna, Austria

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Margita Utczás Department of Chemical, Biological, Pharmaceutical and Environmental Sciences University of Messina Messina, Italy and Center of Sports Nutrition Science Universityof Physical Education Budapest, Hungary

Contributors

Jürgen Wanner Kurt Kitzing GmbH Wallerstein, Germany Mariosimone Zoccali Unit of Food Science and Nutrition, Department of Medicine University Campus Bio-Medico of Rome Rome, Italy

1

Introduction K. Hüsnü Can Başer and Gerhard Buchbauer

The overwhelming success of the first edition of the Handbook of Essential Oils: Science, Technology, and Applications had urged the publication of the second edition which was bestowed, in 2016, the ABC James A. Duke Excellence in Botanical Literature Award for the excellent contribution to the vast field of essential oils. This prestigious award by the American Botanical Council for the best book in botanical literature has prompted us to prepare a third edition of this Handbook. As in the previous edition, updated chapters as well as completely new chapters have been included in the third edition. Some important chapters remained as such. Thus, we kept the contributions of the current Chapters 2, 5, 6, 9, 22, 23, 25, 32, and 33 as in the second edition. We skipped Chapters 15, 16, and 26 in the second edition of the Handbook, whereby the former Part Chapter 16, “Aromatherapy with Essential Oils”, has been substituted by Rhiannon Lewis (Chapter 13). In this edition, Chapters 4, 7, 10, 26, 30, and 31 have been updated, and many new contributions have been added, covering the commonly entitled “Biological activities of…” chapters in the form of six chapters. These are “Essential Oils in Cancer Therapy” (Chapter 14), then “Antimicrobial Activity of Selected Essential Oils and Aromas” (Chapter 15), followed by “Quorum Sensing and Essential Oils” (Chapter 16), then “Essential Oils as Carrier Oils” (Chapter 27), and then two new (more chemically written) overviews, namely “Influence of Light on Essential Oil Constituents” and “Influence of Air on Essential Oil Constituents” (now Chapters 28 and 29). The new Chapter 19, entitled “Adverse Effects and Intoxication with Essential Oils” is an overview written by a pharmacologist of the University of Vienna. The former Chapter 12 now has been substituted by the updated chapter “Central Nervous System Effects of Essential Oil Compounds” (now Chapter 11) and another, newly entitled, treatise, namely “Effects of Essential Oils on Human Cognition” (now Chapter 12). “Essential Oils and Volatiles in Bryophytes” is a new chapter (Chapter 21) by Agnieszka Ludwiczuk and Yoshinori Asakawa. “Functions of Essential Oils and Natural Volatiles in Plant–Insect Interactions” (Chapter 17) was contributed by R. Raguso. “Essential Oils as Lures for Invasive Ambrosia Beetles” (Chapter 18) is yet another new contribution. A useful new chapter for GC/MS analysts is entitled “Use of Linear Retention Indices in GC/MS Libraries for Essential Oil Analysis” (Chapter 8). Also with this third edition, we hope that many scientists, especially in the fields of essential oils in botany, chemistry, pharmacognosy, medicine, clinical aromatherapy, and other relevant aspects of these natural products, will find these contributions not only alluring for their own research but also interesting to read and to find out what manifold properties essential oils have. Especially, also in this third edition, we want to provide a strong scientific basis for essential oils and to prevent any trace of esoteric ignorance.

1

2

History and Sources of Essential Oil Research Karl-Heinz Kubeczka

CONTENTS 2.1 Ancient Historical Background.................................................................................................3 2.2 First Systematic Investigations.................................................................................................. 5 2.3 Research during the Last Half Century.....................................................................................6 2.3.1 Essential Oil Preparation Techniques............................................................................6 2.3.1.1 Industrial Processes........................................................................................ 6 2.3.1.2 Laboratory-Scale Techniques......................................................................... 6 2.3.1.3 Microsampling Techniques.............................................................................7 2.3.2 Chromatographic Separation Techniques.................................................................... 12 2.3.2.1 Thin-Layer Chromatography........................................................................ 13 2.3.2.2 GC................................................................................................................. 13 2.3.2.3 Liquid Column Chromatography.................................................................. 19 2.3.2.4 Supercritical Fluid Chromatography............................................................20 2.3.2.5 Countercurrent Chromatography.................................................................. 21 2.3.3 Hyphenated Techniques............................................................................................... 22 2.3.3.1 Gas Chromatography-Mass Spectrometry.................................................... 22 2.3.3.2 High-Resolution GC-FTIR Spectroscopy..................................................... 23 2.3.3.3 GC-UV Spectroscopy...................................................................................24 2.3.3.4 Gas Chromatography-Atomic Emission Spectroscopy................................24 2.3.3.5 Gas Chromatography-Isotope Ratio Mass Spectrometry.............................25 2.3.3.6 High-Performance Liquid Chromatography-Gas Chromatography.............25 2.3.3.7 HPLC-MS, HPLC-NMR Spectroscopy........................................................26 2.3.3.8 Supercritical Fluid Extraction–Gas Chromatography.................................. 27 2.3.3.9 Supercritical Fluid Chromatography-Gas Chromatography........................ 27 2.3.3.10 Couplings of SFC-MS and SFC-FTIR Spectroscopy...................................28 2.3.4 Identification of Multicomponent Samples without Previous Separation...................28 2.3.4.1 UV Spectroscopy..........................................................................................28 2.3.4.2 IR Spectroscopy............................................................................................28 2.3.4.3 Mass Spectrometry....................................................................................... 29 2.3.4.4 13C-NMR Spectroscopy................................................................................ 30 References......................................................................................................................................... 31

2.1  ANCIENT HISTORICAL BACKGROUND Plants containing essential oils have been used since furthest antiquities as spices and remedies for the treatment of diseases and in religious ceremonies because of their healing properties and their pleasant odors. In spite of the obscured beginning of the use of aromatic plants in prehistoric times to prevent, palliate, or heal sicknesses, pollen analyses of Stone Age settlements indicate the use of aromatic plants that may be dated to 10,000 bc. One of the most important medical documents of ancient Egypt is the so-called Papyrus Ebers of about 1550 bc, a 20 m long papyrus, which was purchased in 1872 by the German Egyptologist 3

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G. Ebers, for whom it is named, containing some 700 formulas and remedies, including aromatic plants and plant products like anise, fennel, coriander, thyme, frankincense, and myrrh. Much later, the ancient Greek physician Hippocrates (460–377 bc), who is referred to as the father of medicine, mentioned in his treatise Corpus Hippocratium approximately 200 medicinal plants inclusive of aromatic plants and described their efficacies. One of the most important herbal books in history is the five-volume book De Materia Medica, written by the Greek physician and botanist Pedanius Dioscorides (ca. 40–90), who practiced in ancient Rome. In the course of his numerous travels all over the Roman and Greek world seeking for medicinal plants, he described more than 500 medicinal plants and respective remedies. His treatise, which may be considered a precursor of modern pharmacopoeias, was later translated into a variety of languages. Dioscorides, as well as his contemporary Pliny the Elder (23–79), a Roman natural historian, mention besides other facts turpentine oil and give some limited information on the methods in its preparation. Many new medicines and ointments were brought from the east during the Crusades from the eleventh to the thirteenth centuries, and many herbals, whose contents included recipes for the use and manufacture of essential oil, were written during the fourteenth to the sixteenth centuries. Theophrastus von Hohenheim, known under the name Paracelsus (1493–1541), a physician and alchemist of the fifteenth century, defined the role of alchemy by developing medicines and extracts from healing plants. He believed distillation released the most desirable part of the plant, the Quinta essentia or quintessence by a means of separating the “essential” part from the “nonessential” containing its subtle and essential constituents. The currently used term “essential oil” still refers to the theory of Quinta essentia of Paracelsus. The roots of distillation methods are attributed to Arabian Alchemists centuries with Avicenna (980–1037) describing the process of steam distillation, who is credited with inventing a coiled cooling pipe to prepare essential oils and aromatic waters. The first description of distilling essential oils is generally attributed to the Spanish physician Arnaldus de Villa Nova (1235–1311) in the thirteenth century. However, in 1975, a perfectly preserved terracotta apparatus was found in the Indus Valley, which is dated to about 3000 bc and which is now displayed in a museum in Taxila, Pakistan. It looks like a primitive still and was presumable used to prepare aromatic waters. Further findings indicate that distillation has also been practiced in ancient Turkey, Persia, and India as far back as 3000 bc. At the beginning of the sixteenth century appeared a comprehensive treatise on distillation by Hieronymus Brunschwig (ca. 1450–1512), a physician of Strasbourg. He described the process of distillation and the different types of stills in his book Liber de arte Distillandi de compositis (Strasbourg 1500 and 1507) with numerous block prints. Although obviously endeavoring to cover the entire field of distillation techniques, he mentions in his book only the four essential oils from rosemary, spike lavender, juniper wood, and the turpentine oil. Just before, until the Middle Ages, the art of distillation was used mainly for the preparation of aromatic waters, and the essential oil appearing on the surface of the distilled water was regarded as an undesirable by-product. In 1551 appeared at Frankfurt on the Main the Kräuterbuch, written by Adam Lonicer (1528– 1586), which can be regarded as a significant turning point in the understanding of the nature and the importance of essential oils. He stresses that the art of distillation is a quite recent invention and not an ancient invention and has not been used earlier. In the Dispensatorium Pharmacopolarum of Valerius Cordus, published in Nuremberg in 1546, only three essential were listed; however, the second official edition of the Dispensatorium Valerii Cordi issued in 1592, 61 distilled oils were listed illustrating the rapid development and acceptance of essential oils. In that time, the so-called Florentine flask has already been used for separating the essential oil from the water phase. The German J.R. Glauber (1604–1670), who can be regarded as one of the first great industrial chemists, was born in the little town Karlstadt close to Wuerzburg. His improvements in chemistry, for example, the production of sodium sulfate, as a safe laxative brought him the honor of being named Glauber’s salt. In addition, he improved numerous different other chemical processes and especially new distillation devices also for the preparation of essential oils from aromatic plants. However, it lasted until the nineteenth century to get any real understanding of the composition of true essential oils.

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2.2  FIRST SYSTEMATIC INVESTIGATIONS The first systematic investigations of constituents from essential oils may be attributed to the French chemist M.J. Dumas (1800–1884) who analyzed some hydrocarbons and oxygen as well as sulfurand nitrogen-containing constituents. He published his results in 1833. The French researcher M. Berthelot (1859) characterized several natural substances and their rearrangement products by optical rotation. However, the most important investigations have been performed by O. Wallach, an assistant of Kekule. He realized that several terpenes described under different names according to their botanical sources were often, in fact, chemically identical. He, therefore, tried to isolate the individual oil constituents and to study their basic properties. He employed together with his highly qualified coworkers Hesse, Gildemeister, Betram, Walbaum, Wienhaus, and others fractional distillation to separate essential oils and performed reactions with inorganic reagents to characterize the obtained individual fractions. The reagents he used were hydrochloric acid, oxides of nitrogen, bromine, and nitrosyl chloride—which was used for the first time by W.A. Tilden (1875)—by which frequently crystalline products had been obtained. At that time, hydrocarbons occurring in essential oils with the molecular formula C10H16 were known, which had been named by Kekule terpenes because of their occurrence in turpentine oil. Constituents with the molecular formulas C10H16O and C10H18O were also known at that time under the generic name camphor and were obviously related to terpenes. The prototype of this group was camphor itself, which was known since antiquity. In 1891, Wallach characterized the terpenes pinene, camphene, limonene, dipentene, phellandrene, terpinolene, fenchene, and sylvestrene, which has later been recognized to be an artifact. During 1884–1914, Wallach wrote about 180 articles that are summarized in his book Terpene und Campher (Wallach, 1914) compiling all the knowledge on terpenes at that time, and already in 1887, he suggested that the terpenes must be constructed from isoprene units. In 1910, he was honored with the Nobel Prize for Chemistry “in recognition of his outstanding research in organic chemistry and especially in the field of alicyclic compounds” (Laylin, 1993). In addition to Wallach, the German chemist A. von Baeyer, who also had been trained in Kekule’s laboratory, was one of the first chemists to become convinced of the achievements of structural chemistry and who developed and applied it to all of his work covering a broad scope of organic chemistry. Since 1893, he devoted considerable work to the structures and properties of cyclic terpenes (von Baeyer and Seuffert, 1901). Besides his contributions to several dyes, the investigations of polyacetylenes, and so on, his contributions to theoretical chemistry including the strain theory of triple bonds and small carbon cycles have to be mentioned. In 1905, he was awarded the Nobel Prize for Chemistry “in recognition of his contributions to the development of Organic Chemistry and Industrial Chemistry, by his work on organic dyes and hydroaromatic compounds” (Laylin, 1993). The frequently occurring acyclic monoterpenes geraniol, linalool, citral, and so on have been investigated by F.W. Semmler and the Russian chemist G. Wagner (1899), who recognized the importance of rearrangements for the elucidation of chemical constitution, especially the carbonto-carbon migration of alkyl, aryl, or hydride ions, a type of reaction that was later generalized by H. Meerwein (1914) as Wagner–Meerwein rearrangement. More recent investigations of J. Read, W. Hückel, H. Schmidt, W. Treibs, and V. Prelog were mainly devoted to disentangle the stereochemical structures of menthols, carvomenthols, borneols, fenchols, and pinocampheols, as well as the related ketones (see Gildemeister and Hoffmann, 1956). A significant improvement in structure elucidation was the application of dehydrogenation of sesquiand diterpenes with sulfur and later with selenium to give aromatic compounds as a major method, and the application of the isoprene rule to terpene chemistry, which have been very efficiently used by L. Ruzicka (1953) in Zurich, Switzerland. In 1939, he was honored in recognition of his outstanding investigations with the Nobel Prize in chemistry for his work on “polymethylenes and higher terpenes.” The structure of the frequently occurring bicyclic sesquiterpene ß-caryophyllene was for many years a matter of doubt. After numerous investigations, W. Treibs (1952) has been able to isolate the crystalline caryophyllene epoxide from the autoxidation products of clove oil, and F. Šorm et al.

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(1950) suggested caryophyllene to have a four- and nine-membered ring on bases of infrared (IR) investigations. This suggestion was later confirmed by the English chemist D.H.R. Barton (Barton and Lindsay, 1951), who was awarded the Nobel Prize in Chemistry in 1969. The application of ultraviolet (UV) spectroscopy in the elucidation of the structure of terpenes and other natural products was extensively used by R.B. Woodward in the early forties of the last century. On the basis of his large collection of empirical data, he developed a series of rules (later called the Woodward rules), which could be applied to finding out the structures of new natural substances by correlations between the position of UV maximum absorption and the substitution pattern of a diene or an α,β-unsaturated ketone (Woodward, 1941). He was awarded the Nobel Prize in Chemistry in 1965. However, it was not until the introduction of chromatographic separation methods and nuclear magnetic resonance (NMR) spectroscopy into organic chemistry that a lot of further structures of terpenes were elucidated. The almost exponential growth in our knowledge in that field and other essential oil constituents is essentially due to the considerable advances in analytical methods in the course of the last half century.

2.3  RESEARCH DURING THE LAST HALF CENTURY 2.3.1  Essential Oil Preparation Techniques 2.3.1.1  Industrial Processes The vast majority of essential oils are produced from plant material in which they occur by different kinds of distillation or by cold pressing in the case of the peel oils from citrus fruits. In water or hydrodistillation, the chopped plant material is submerged and in direct contact with boiling water. In steam distillation, the steam is produced in a boiler separate of the still and blown through a pipe into the bottom of the still, where the plant material rests on a perforated tray or in a basket for quick removal after exhaustive extraction. In addition to the aforementioned distillation at atmospheric pressure, high-pressure steam distillation is most often applied in European and American field stills, and the applied increased temperature significantly reduces the time of distillation. The high-pressure steam-type distillation is often applied for peppermint, spearmint, lavandin, and the like. The condensed distillate, consisting of a mixture of water and oil, is usually separated in a so-called Florentine flask, a glass jar, or more recently in a receptacle made of stainless steel with one outlet near the base and another near the top. There, the distillate separates into two layers from which the oil and the water can be separately withdrawn. Generally, the process of steam distillation is the most widely accepted method for the production of essential oils on a large scale. Expression or cold pressing is a process in which the oil glands within the peels of citrus fruits are mechanically crushed to release their content. There are several different processes used for the isolation of citrus oils; however, there are four major currently used processes. Those are pellatrice and sfumatrice—most often used in Italy—and the Brown peel shaver as well as the FMC extractor, which are used predominantly in North and South America. For more details, see, for example, Lawrence 1995. All these processes lead to products that are not entirely volatile because they may contain coumarins, plant pigments, and so on; however, they are nevertheless acknowledged as essential oils by the International Organization for Standardization, the different pharmacopoeias, and so on. In contrast, extracts obtained by solvent extraction with different organic solvents, with liquid carbon dioxide or by supercritical fluid extraction (SFE) may not be considered as true essential oils; however, they possess most often aroma profiles that are almost identical to the raw material from which they have been extracted. They are therefore often used in the flavor and fragrance industry and in addition in food industry, if the chosen solvents are acceptable for food and do not leave any harmful residue in food products. 2.3.1.2  Laboratory-Scale Techniques The following techniques are used mainly for trapping small amounts of volatiles from aromatic plants in research laboratories and partly for determination of the essential oil content in plant material.

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The most often used device is the circulatory distillation apparatus, basing on the publication of Clevenger in 1928 and which has later found various modifications. One of those modified apparatus described by Cocking and Middleton (1935) has been introduced in the European pharmacopoeia and several other pharmacopoeias. This device consists of a heated round-bottom flask into which the chopped plant material and water are placed and which is connected to a vertical condenser and a graduated tube, for the volumetric determination of the oil. At the bottom of the tube, a three-way valve permits to direct the water back to the flask, since it is a continuous closed-circuit distillation device, and at the end of the distillation process to separate the essential oil from the water phase for further investigations. The length of distillation depends on the plant material to be investigated; however, it is usually fixed to 3–4 h. For the volumetric determination of the essential oil content in plants according to most of the pharmacopoeias, a certain amount of xylene—usually 0.5 mL—has to be placed over the water before running distillation to separate even small droplets of essential oil during distillation from the water. The volume of essential oil can be determined in the graduated tube after subtracting the volume of the applied xylene. Improved constructions with regard to the cooling system of the aforementioned distillation apparatus have been published by Stahl (1953) and Sprecher (1963) and, in publications of Kaiser and Lang (1951) and Mechler and Kovar (1977), various apparatus used for the determination of essential oils in plant material are discussed and depicted. A further improvement was the development of a simultaneous distillation–solvent extraction device by Likens and Nickerson in 1964 (see Nickerson and Likens, 1966). The device permits continuous concentration of volatiles during hydrodistillation in one step using a closed-circuit distillation system. The water distillate is continuously extracted with a small amount of an organic- and water-immiscible solvent. Although there are two versions described, one for highdensity and one for low-density solvents, the high-density solvent version using dichloromethane is mostly applied in essential oil research. It has found numerous applications, and several modified versions including different microdistillation devices have been described (e.g., Bicchi et al., 1987; Chaintreau, 2001). A sample preparation technique basing on Soxhlet extraction in a pressurized container using liquid carbon dioxide as extractant has been published by Jennings (1979). This device produces solvent-free extracts especially suitable for high-resolution gas chromatography (GC). As a less time-consuming alternative, the application of microwave-assisted extraction has been proposed by several researchers, for example, by Craveiro et al. (1989), using a round-bottom flask containing the fresh plant material. This flask was placed into a microwave oven and passed by a flow of air. The oven was heated for 5 min and the obtained mixture of water and oil collected in a small and cooled flask. After extraction with dichloromethane, the solution was submitted to GC–mass spectrometry (GC-MS) analysis. The obtained analytical results have been compared with the results obtained by conventional distillation and exhibited no qualitative differences; however, the percentages of the individual components varied significantly. A different approach yielding solvent-free extracts from aromatic herbs by means of microwave heating has been presented by Lucchesi et al. (2004). The potential of the applied technique has been compared with conventional hydrodistillation showing substantially higher amounts of oxygenated compounds at the expense of monoterpene hydrocarbons. 2.3.1.3  Microsampling Techniques 2.3.1.3.1 Microdistillation Preparation of very small amounts of essential oils may be necessary if only very small amounts of plant material are available and can be fundamental in chemotaxonomic investigations and control analysis but also for medicinal and spice plant breeding. In the past, numerous attempts have been made to minimize conventional distillation devices. As an example, the modified Marcusson device may be quoted (Bicchi et al., 1983) by which 0.2–3 g plant material suspended in 50 mL water can be distilled and collected in 100 µL analytical grade pentane or hexane. The analytical results proved to be identical with those obtained by conventional distillation.

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Microversions of the distillation–extraction apparatus, described by Likens and Nickerson, have also been developed as well for high-density (Godefroot et al., 1981) and low-density solvents (Godefroot et al., 1982). The main advantage of these techniques is that no further enrichment by evaporation is required for subsequent gas chromatographic investigation. A different approach has been presented by Gießelmann and Kubeczka (1993) and Kubeczka and Gießelmann (1995). By means of a new developed micro-hydrodistillation device, the volatile constituents of very small amounts of plant material have been separated. The microscale hydrodistillation of the sample is performed using a 20 mL crimp-cap glass vial with a Teflon®-lined rubber septum containing 10 mL water and 200–250 mg of the material to be investigated. This vial, which is placed in a heating block, is connected with a cooled receiver vial by a 0.32 mm ID fused silica capillary. By temperature-programmed heating of the sample vial, the water and the volatile constituents are vaporized and passed through the capillary into the cooled receiver vial. There, the volatiles as well as water are condensed and the essential oil collected in pentane for further analysis. The received analytical results have been compared to results from identical samples obtained by conventional hydrodistillation showing a good correlation of the qualitative and quantitative composition. Further applications with the commercially available Eppendorf MicroDistiller® have been published in several papers, for example, by Briechle et al. (1997) and Baser et al. (2001). A simple device for rapid extraction of volatiles from natural plant drugs and the direct transfer of these substances to the starting point of a thin-layer chromatographic plate has been described by Stahl (1969a) and in his subsequent publications. A small amount of the sample (ca. 100 mg) is introduced into a glass cartridge with a conical tip together with 100 mg silica gel, containing 20% of water, and heated rapidly in a heating block for a short time at a preset temperature. The tip of the glass tube projects ca. 1 mm from the furnace and points to the starting point of the thin-layer plate, which is positioned 1 mm in front of the tip. Before introducing the glass tube, it is sealed with a silicone rubber membrane. This simple technique has proven useful for many years in numerous investigations, especially in quality control, identification of plant drugs, and rapid screening of chemical races. In addition to the aforementioned micro-hydrodistillation with the so-called TAS procedure (T, thermomicro and transfer; A, application; S, substance), several further applications, for example, in structure elucidation of isolated natural compounds such as zinc dust distillation, sulfur and selenium dehydrogenation, and catalytic dehydrogenation with palladium, have been described in the microgram range (Stahl, 1976). 2.3.1.3.2  Direct Sampling from Secretory Structures The investigation of the essential oils by direct sampling from secretory glands is of fundamental importance in studying the true essential oil composition of aromatic plants, since the usual applied techniques such as hydrodistillation and extraction are known to produce in some cases several artifacts. Therefore, only direct sampling from secretory cavities and glandular trichomes and properly performed successive analysis may furnish reliable results. One of the first investigations with a kind of direct sampling has been performed by Hefendehl (1966), who isolated the glandular hairs from the surfaces of Mentha piperita and Mentha aquatica leaves by means of a thin film of polyvinyl alcohol, which was removed after drying and extracted with diethyl ether. The composition of this product was in good agreement with the essential oils obtained by hydrodistillation. In contrast to these results, Malingré et al. (1969) observed some qualitative differences in the course of their study on M. aquatica leaves after isolation of the essential oil from individual glandular hairs by means of a micromanipulator and a stereomicroscope. In the same year, Amelunxen et al. (1969) published results on M. piperita, who separately isolated glandular hairs and glandular trichomes with glass capillaries. They found identical qualitative composition of the oil in both types of hairs, but differing concentrations of the individual components. Further studies have been performed by Henderson et al. (1970) on Pogostemon cablin leaves and by Fischer et al. (1987) on Majorana hortensis leaves. In the latter study, significant differences regarding the oil composition of the hydrodistilled oil and the oil extracted by means of glass capillaries from the trichomes were

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observed. Their final conclusion was that the analysis of the respective essential oil is mainly an analysis of artifacts, formed during distillation, and the gas chromatographic analysis. Even if the investigations are performed very carefully and the successive GC has been performed by cold­ on-column injection to avoid thermal stress in the injection port, significant differences of the GC pattern of directly sampled oils versus the microdistilled samples have been observed in several cases (Bicchi et al., 1985). 2.3.1.3.3  HS Techniques Headspace (HS) analysis has become one of the very frequently used sampling techniques in the investigation of aromatic plants, fragrances, and spices. It is a means of separating the volatiles from a liquid or solid prior to gas chromatographic analysis and is preferably used for samples that cannot be directly injected into a gas chromatograph. The applied techniques are usually classified according to the different sampling principles in static HS analysis and dynamic HS analysis. 2.3.1.3.3.1  Static HS Methods  In static HS analysis, the liquid or solid sample is placed into a vial, which is heated to a predetermined temperature after sealing. After the sample has reached equilibrium with its vapor (in equilibrium, the distribution of the analytes between the two phases depends on their partition coefficients at the preselected temperature, the time, and the pressure), an aliquot of the vapor phase can be withdrawn with a gas-tight syringe and subjected to gas chromatographic analysis. A simple method for the HS investigation of herbs and spices was described by Chialva et al. (1982), using a blender equipped with a special gas-tight valve. After grinding the herb and until thermodynamic equilibrium is reached, the HS sample can be withdrawn through the valve and injected into a gas chromatograph. Eight of the obtained capillary gas chromatograms are depicted in the paper of Chialva and compared with those of the respective essential oils exhibiting significant higher amounts of the more volatile oil constituents. However, one of the major problems with static HS analyses is the need for sample enrichment with regard to trace components. Therefore, a concentration step such as cryogenic trapping, liquid absorption, or adsorption on a suitable solid has to be inserted for volatiles occurring only in small amounts. A versatile and often-used technique in the last decade is solid-phase microextraction (SPME) for sampling volatiles, which will be discussed in more detail in a separate paragraph. Since different other trapping procedures are a fundamental prerequisite for dynamic HS methods, they will be considered in the succeeding text. A comprehensive treatment of the theoretical basis of static HS analysis including numerous applications has been published by Kolb and Ettre (1997, 2006). 2.3.1.3.3.2   Dynamic HS Methods  The sensitivity of HS analysis can be improved considerably by stripping the volatiles from the material to be investigated with a stream of purified air or inert gas and trapping the released compounds. However, care has to be taken if grinded plant material has to be investigated, since disruption of tissues may initiate enzymatic reactions that may lead to formation of volatile artifacts. After stripping the plant material with gas in a closed vessel, the released volatile compounds are passed through a trap to collect and enrich the sample. This must be done because sample injection of fairly large sample volumes results in band broadening causing peak distortion and poor resolution. The following three techniques are advisable for collecting the highly diluted volatile sample according to Schaefer (1981) and Schreier (1984) with numerous references. Cryogenic trapping can be achieved by passing the gas containing the stripped volatiles through a cooled vessel or a capillary in which the volatile compounds are condensed (Kolb and Liebhardt, 1986). The most convenient way for trapping the volatiles is to utilize part of the capillary column as a cryogenic trap. A simple device for cryofocusing of HS volatiles by using the first part of capillary column as a cryogenic trap has been shown in the aforementioned reference inclusive of a discussion of the theoretical background of cryogenic trapping. A similar on-column cold trapping device, suitable for extended period vapor sampling, has been published by Jennings (1981).

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A different approach can be used if large volumes of stripped volatiles have to be trapped using collection in organic liquid phases. In this case, the volatiles distribute between the gas and the liquid, and efficient collection will be achieved, if the distribution factor K is favorable for solving the stripped compounds in the liquid. A serious drawback, however, is the necessity to concentrate the obtained solution prior to GC with the risk to lose highly volatile compounds. This can be overcome if a shortpacked GC column is used containing a solid support coated with a suitable liquid. Novak et al. (1965) have used Celite coated with 30% silicone elastomer E-301 and the absorbed compounds were introduced into a gas chromatograph after thermal desorption. Coating with 15% silicone rubber SE 30 has been successfully used by Kubeczka (1967) with a similar device and the application of a wallcoated tubing with methyl silicone oil SF 96 has been described by Teranishi et al. (1972). A different technique has been used by Bergström (1973) and Bergström et al. (1980). They trapped the scent of flowers on Chromosorb® W coated with 10% silicon high-vacuum grease and filled a small portion of the sorbent containing the volatiles into a precolumn, which was placed in the splitless injection port of a gas chromatograph. There, the volatiles were desorbed under heating and flushed onto the GC column. In 1987, Bichi et al. applied up to 50 cm pieces of thick-film fused silica capillaries coated with a 15 µm dimethyl silicone film for trapping the volatiles in the atmosphere surrounding living plants. The plants under investigation were placed in a glass bell into which the trapping capillary was introduced through a rubber septum, while the other end of the capillary has been connected to pocket sampler. In order to trap even volatile monoterpene hydrocarbons, a capillary length of at least 50 cm and sample volume of maximum 100 mL have to be applied to avoid loss of components through breakthrough. The trapped compounds have been subsequently online thermally desorbed, cold trapped, and analyzed. Finally, a type of enfleurage especially designed for field experiments has been described by Joulain (1987) to trap the scents of freshly picked flowers. Around 100 g flowers were spread on the grid of a specially designed stainless steel device and passed by a stream of ambient air, supplied by an unheated portable air drier. The stripped volatiles are trapped on a layer of purified fat placed above the grid. After 2 h, the fat was collected and the volatiles recovered in the laboratory by means of vacuum distillation at low temperature. With a third often applied procedure, the stripped volatiles from the HS of plant material and especially from flowers are passed through a tube filled with a solid adsorbent on which the volatile compounds are adsorbed. Common adsorbents most often used in investigations of plant volatiles are above all charcoal and different types of synthetic porous polymers. Activated charcoal is an adsorbent with a high adsorption capacity, thermal and chemical stability, and which is not deactivated by water, an important feature, if freshly collected plant material has to be investigated. The adsorbed volatiles can easily be recovered by elution with small amounts (10–50 µL) of carbon disulfide avoiding further concentration of the sample prior to GC analysis. The occasionally observed incomplete recovery of sample components after solvent extraction and artifact formation after thermal desorption has been largely solved by application of small amounts of special type of activated charcoal as described by Grob and Zürcher (1976). Numerous applications have been described using this special type of activated charcoal, for example, by Kaiser (1993) in a great number of field experiments on the scent of orchids. In addition to charcoal, the following synthetic porous polymers have been applied to collect volatile compounds from the HS from flowers and different other plant materials according to Schaefer (1981): Tenax® GC, different Porapak® types (e.g., Porapak P, Q, R, and T), and several Chromosorb types belonging to the 100 series. More recent developed adsorbents are the carbonaceous adsorbents such as Ambersorb®, Carboxen®, and Carbopak®, and their adsorbent properties lie between activated charcoal and the porous polymers. Especially the porous polymers have to be washed repeatedly, for example, with diethyl ether, and conditioned before use in a stream of oxygen-free nitrogen at 200°C–280°C, depending on the sort of adsorbent. The trapped components can be recovered either by thermal desorption or by solvent elution, and the recoveries can be different depending on the applied adsorbent (Cole, 1980). Another very important criterion for the selection of a suitable adsorbent for collecting HS samples is the breakthrough volume limiting the amount of gas passing through the trap.

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A comprehensive review concerning HS gas chromatographic analysis of medicinal and aromatic plants and flowers with 137 references, covering the period from 1982 to 1988 has been published by Bicchi and Joulain in 1990, thoroughly describing and explaining the different methodological approaches and applications. Among other things, most of the important contributions of the Finnish research group of Hiltunen and coworkers on the HS of medicinal plants and the optimization of the HS parameters have been cited in the mentioned review. 2.3.1.3.4  Solid-Phase Microextraction SPME is an easy-to-handle sampling technique, initially developed for the determination of volatile organic compounds in environmental samples (Arthur and Pawliszyn, 1990), and has gained, in the last years, acceptance in numerous fields and has been applied to the analysis of a wide range of analytes in various matrices. Sample preparation is based on sorption of analytes from a sample onto a coated fused silica fiber, which is mounted in a modified GC syringe. After introducing the coated fiber into a liquid or gaseous sample, the compounds to be analyzed are enriched according to their distribution coefficients and can be subsequently thermally desorbed from the coating after introducing the fiber into the hot injector of a gas chromatograph. The commercially available SPME device (Supelco Inc.) consists of a 1 cm length fused silica fiber of ca. 100 µm diameter coated on the outer surface with a stationary phase fixed to a stainless steel plunger and a holder that looks like a modified microliter syringe (Supelco, 2007). The fiber can be drawn into the syringe needle to prevent damage. To use the device, the needle is pierced through the septum that seals the sample vial. Then, the plunger is depressed lowering the coated fiber into the liquid sample or the HS above the sample. After sorption of the sample, which takes some minutes, the fiber has to be drawn back into the needle and withdrawn from the sample vial. By the same procedure, the fiber can be introduced into the gas chromatograph injector where the adsorbed substances are thermally desorbed and flushed by the carrier gas into the capillary GC column. SPME fibers can be coated with polymer liquid (e.g., polydimethylsiloxane [PDMS]) or a mixed solid and liquid coating (e.g., Carboxen®/PDMS). The selectivity and capacity of the fiber coating can be adjusted by changing the phase type or thickness of the coating on the fiber according to the properties of the compounds to be analyzed. Commercially available are coatings of 7, 30, and 100 µm of PDMS, an 85 µm polyacrylate, and several mixed coatings for different polar components. The influence of fiber coatings on the recovery of plant volatiles was thoroughly investigated by Bicchi et al. (2000a,b). Details concerning the theory of SPME, technology, its application, and specific topics have been described by Pawliszyn (1997) and references cited therein. A number of different applications of SPME in the field of essential oil analysis have been presented by Kubeczka (1997a). An overview on publications of the period 2000–2005 with regard to HS-SPME has been recently published by Belliardo et al. (2006) covering the analysis of volatiles from aromatic and medicinal plants, selection of the most effective fibers and sampling conditions, and discussing its advantages and limitations. The most comprehensive collection of references with regard to the different application of SPME can be obtained from Supelco on CD. 2.3.1.3.5  Stir Bar Sorptive Extraction and HS Sorptive Extraction Despite the indisputable simplicity and rapidity of SPME, its applicability is limited by the small amount of sorbent on the needle (1000 t/a); MQ, medium quantities (100–1000 t/a); LQ, low quantities ( ΔP3), while no restriction is present on the other side. In the standby mode, a lower pressure is generated on the side of the first dimension respect to the second dimension branch, where it passes through the solenoid valve without any restriction. In such a configuration, analytes eluting from the first column are directed to FID1 simulating a monodimensional analysis. Once the solenoid valve is activated, the transfer device passes to the cutting mode: the pressure on the first-dimension side of the interface remains unchanged, while the pressure on the second-dimension side becomes lower than that of the first dimension. Under such conditions, the primary-column eluate is free to reach the second capillary. The same system has been used for the determination of the enantiomeric distribution of monoterpene hydrocarbons and monoterpene alcohols in the essential oils of mandarin (Sciarrone et al., 2010a), pistachio (Lo Presti et  al., 2008), and sandalwood (Sciarrone et  al., 2011). The quali/quantitative assessment with respect to normative limits was also carried out in tea tree (Sciarrone et al., 2010b). In Figure 7.5, the monodimensional chiral analysis of mandarin oil is compared to the second dimension chiral separation achieved with the MDGC approach (Sciarrone et al., 2010a). Well-resolved peaks of components present in large amounts, and also of the minor compounds, were attained through the partial transfer of the major concentrated components. For most of the enantiomers, the ee values achieved in eGC and eMDGC were in good agreement, with slight variations probably due to the use of different instruments and columns. However, considerable variations were observed for camphene and linalool, due to the presence of coelutions, while the ee values of camphor and citronellal were calculated only in the multidimensional applications. The latter measurements were achieved thanks to the possibility to overload the primary apolar column by injection of a higher sample amount. Next, only selected slices of the more abundant components were transferred to the secondary chiral column, while in the case of trace components, the entire peaks were selected for the heart-cut. An MDGC approach with a polar and chiral column set, coupled to mass spectrometry and olfactometry port, was applied for the analysis of citrus hybrid peel extract (Casilli et al., 2014). An MDGC-MS method for the

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FIGURE 7.5  Comparison of eGC and eMDGC chiral analysis of mandarin essential oil. (Reprinted from J. Chromatogr. A, 1217. Sciarrone, D. et al., Thorough evaluation of the validity of conventional enantio-gas chromatography in the analysis of volatile chiral compounds in mandarin essential oil: A comparative investigation with multidimensional gas chromatography. 1101–1105. Copyright 2010b, with permission from Elsevier.)

determination of the enantiomeric distribution of selected compounds in lime oils was also reported, namely α-thujene, camphene, β-pinene, sabinene, α-phellandrene, β-phellandrene, limonene, linalool, terpinen-4-ol, and α-terpineol (Bonaccorsi et al., 2012). Gas chromatography–combustion–isotope ratio mass spectrometry and enantioselective multidimensional gas chromatography were exploited complementarily for petitgrain oils landmark and lime essential oil assessment (Schipilliti et al., 2012; Bonaccorsi et al., 2012).

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7.3.5.2  Multidimensional Preparative Gas Chromatography When the isolation of specific volatile components is required because of the lack of standard components or to allow the structure elucidation of unknown molecules, preparative gas chromatography could be considered as an option. Nevertheless, monodimensional prep-GC is commonly considered a time-consuming and “tricky” technique since the isolation of volatile components from the gas stream is not an easy task. Furthermore, a series of shortcomings related to the unsatisfactory degree of purity of the fraction collected have prevented the widespread use of this technique. When the sample is characterized by a high level of complexity (as is often the case of natural samples), the presence of coeluting interferences is very common and requires particular precautions, which affect the time requested for the collection of a certain amount. The first issue is related to the reduced amount of neat, or diluted sample, which can be injected in each run, in order not to impair the efficiency of the system by overloading the GC column. Moreover, the collection of sufficient amounts of chemicals for further evaluations would require a long time. The use of conventional 0.25 mm ID columns requires hundreds of injections to afford the collection of a sufficient amount (µg-mg) due to their reduced sample capacity. For such a reason, megabore columns (0.32–0.53 mm ID) are generally employed in prep-GC systems since they are able to manage higher sample amounts, even at the price of a reduced efficiency compared to microbore capillaries (0.1–0.25 mm ID). Thanks to the improvement in separation power, heart-cutting multidimensional preparative chromatography (MDGC-prep) has demonstrated effectiveness to allow the collection of target compounds from complex samples (Sciarrone et al., 2015a). With such an approach, by the coupling of mega-bore columns with different selectivity in the heart-cut mode, it is possible to achieve a separation system able to afford higher purity of the compounds purification, with higher sample amounts, and finally with higher throughput. Furthermore, the multidimensional approach can be effectively used to obtain increased productivity, thanks to the possibility to overload the first chromatographic dimension by the injection of a high sample amount. After a 1D pre-purification step, the 2D column (or even 3D) would allow to purify the fraction transferred before the collection, reducing the number of runs required. Recently, MDGC systems have been successfully coupled to prep-GC (Ochiai and Sasamoto, 2011; Ball et al., 2012; Sciarrone et al., 2012, 2013, 2014, 2016, 2017, 2019). Sciarrone et al. reported a three-dimensional MDGCprep system, exploiting an apolar 1D column, a medium-polarity 2D wax column, and an ionic liquid stationary phase as 3D, with similar polarity but different in selectivity, each one equipped with a three-restrictors Deans switch device (Figure 7.6). The collection of components present in a range between 10% and 30% in the sample was successfully reported, allowing for further investigations of unknown molecules by NMR, MS, and fourier transform infrared (FTIR). The same group described a further improvement of the system, enabling the collection of low-level components in the sample (1%–10%), obtained including an LC pre-separation step before the three-dimensional GC system. Due to the higher sample capacity of the packed LC column, it was possible to inject 10–20-fold sample amounts than a direct-GC injection (actually limited by the injector liner internal volume). The use of an LC system operated in normal phase allowed the isolation of fractions of eluate containing the concentrated components of interest. The latter were on-line transferred to the GC large-volume injector in order to evaporate the mobile phase (organic) before the start of the GC run. The procedure demonstrated the capability to reduce to about 1/10–1/20 the time to collect a milligram of components. 7.3.5.3 Multidimensional Gas Chromatography Coupled to Isotope Ratio Mass Spectrometry (MDGC-IRMS) Enantioselectivity and isotope discrimination during biosynthesis are well recognized principles of authenticity evaluation. As above discussed, enantioselective multidimensional gas chromatography (MDGC) with the combination of a non-chiral pre-separation column and a chiral main column has proved to be the method of choice for the enantioselective analysis of chiral compounds from complex matrices. In the same way, isotope ratio mass spectrometry (IRMS) is well established and

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FIGURE 7.6  Triple Deans switch multidimensional prep-GC system scheme oil. (Reprinted from Anal. Chim. Acta, 785. Sciarrone, D. et al., Rapid collection and identification of a novel component from Clausena lansium Skeels leaves by means of three-dimensional preparative gas chromatography and nuclear magnetic resonance/infrared/mass spectrometric analysis. 119–125. Copyright 2013, with permission from Elsevier.)

widespread in different research fields for the determination of the isotopic ratio of GC-separated compounds. In the case of carbon, the molecules are oxidized to CO2 into a combustion furnace (C) and transferred to Faraday cups for m/z 44, 45, and 46, by using a magnetic sector. The results can be often related to specific geographical origin and easily discriminate natural from synthetic compounds, allowing highlighting of possible adulterations or frauds, especially in the field of essential oils (Do et al., 2015). Obviously, calibration is of great importance in isotope ratio measurements. Two calibration procedures are commonly used in GC-C-IRMS measurements, the first dealing with the use of standard compounds co-injected and combusted along with the analytes. The second procedure uses standard CO2 gas directly introduced into the ion source. The complexity of natural samples often precludes to find even a few seconds retention time window free of peaks for the selected standards, which in turn must be selected for each matrix. Thus the use of standard CO2 gas, directly introduced into the ion source before and after each sample, has become the most practiced procedure in such cases. Monodimensional gas chromatographic separation is often unable to afford the complete separation of minor components from complex matrices, while MDGC, thanks to its higher peak capacity, gives the highest performance in direct sample cleanup. When using an IRMS system coupled to GC, in mono- or multidimensional systems, the chromatographic conditions have to be carefully optimized in order to avoid a multitude of erroneous judgements (Nitz et al., 1992). Unlike what happens when using a GC hyphenated to a quadrupole mass spectrometer (GC-QMS), where coeluting compounds can still be identified and quantified in the selected ion monitoring (SIM) or extracted ion mode, excellent chromatographic resolution is mandatory for GC-C-IRMS analysis, since the isotopic ratio of the CO2 generated from coeluted peaks cannot be treated in the same way. Baseline-to-baseline integration over an entire peak is moreover required for accurate measurement of its isotopic composition, since column fractionation would retain in a slightly different way the heavier isotopes, eluted in the first part of the peak, and the lighter ones, eluted in the tail (Ricci et al., 1994). As a consequence, the use of MDGC approaches coupled to IRMS detection appears to be the best choice. The hyphenation of a multidimensional GC system in heart-cut mode to IRMS is not a novel concept even if, probably due to technical difficulties, only few papers have appeared in the literature (Juchelka et al., 1998). An MDGC-C-IRMS system was presented for the origin-specific analysis of flavor and fragrance compounds, exploiting the 13C/12C ratio of the detected enantiomers. The authors used a double-oven system equipped with a “live switching” coupling piece (live T-piece) as the transfer system between an apolar-chiral column set. The results showed no significant variations in the 13C/12C ratio of the standard injected, measured

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in mono- and multidimensional conditions. Nowadays, the use of Deans switch devices based on a pressure balancing effect, generally affected by retention time shifts of the next eluting components (see Section 7.3.5.1), exposes to the risk of isotopic fractionation resulting from the incomplete transfer of peaks during consecutive heart-cuts. The recent Deans switch systems, able to maintain a constant 1D backpressure, represent a new frontier to be explored for MDGC-C-IRMS users, and the development of new applications in the field of essential oils analysis is foreseeable.

7.3.6 Comprehensive Two-Dimensional Gas Chromatography and Multidimensional Liquid-Gas Chromatography 7.3.6.1 Analysis of Essential Oils through Comprehensive Two-Dimensional Gas Chromatography (GC × GC) Comprehensive two-dimensional gas chromatography (GC × GC) was first described in 1991 ( Liu and Phillips, 1991 ), with it taking nearly 10 years to be applied to the analysis of an essential oil (Dimandja et al., 2000). Since that first research, various investigations have been published, also regarding complex samples related to essential oils, such as perfumes. In many cases, the use of such a powerful GC methodology appeared to be justified, while for others its use seemed to be an excess in analytical terms. Briefly, a GC × GC analysis is performed by using two capillary columns, each coated with a different type of stationary phase. Most commonly, a conventional column (e.g., 30 m × 0.25 mm ID) with a non-polar stationary phase is used in the first dimension (1D), while a short mid-polarity microbore one (e.g., 1–2 m × 0.10 mm ID) is used in the second dimension (2D). A special transfer device, defined modulator, is situated between the two analytical dimensions and works in a continuous manner throughout the analysis. Cryogenic modulators (a form of thermal modulation) are by far the most popular GC × GC transfer devices. The modulator first accumulates fractions from the 1D and then releases them onto the 2D, where rapid separations occur. The accumulation-release process occurs in a sequential manner, with its time duration defined as the modulation period, usually lasting in the range of 4–8 s. The modulation period is also equivalent to the timeframe of each rapid separation on the second column. A raw GC × GC chromatogram is nothing more than a continuous stream of rapid 2D separations, positioned side-by-side along the retention-time axis. For the fundamental objective of two-dimensional (2D) visualization, peak integration, and identification, the use of dedicated software is necessary. Compared to GC, thermal modulation GC × GC offers enhanced resolving power, selectivity, and sensitivity. The reader is directed to the literature for more details on this powerful GC technology (Marriot et al., 2012). In general, an essential oil can be subjected to a GC analysis for basically three analytical objectives: (a) untargeted analysis, (b) pre-targeted analysis and (c) fingerprinting. A series of examples will be shown, illustrating the need (or not) of comprehensive 2D GC in such types of investigations. 7.3.6.1.1  Untargeted Analyses The untargeted analysis of an essential oil, by using a conventional low-polarity or mid-polarity capillary column, is performed for objectives which may relate to pure research curiosity, or to define the state of preservation. The use of mass spectrometry (MS) is mandatory for peak identification, advisably with the support of linear retention index (LRI) data. The use of relative retention information is important because many terpenes are characterized by a similar structure, a factor that hinders reliable MS differentiation. The generation of two sets of LRI data, by using two parallel columns with a chemically different stationary phase, has been reported for a long time (Bicchi et al., 1988). Such an option, in itself, highlights the requirement of an increased GC resolving power in untargeted analyses. Gas chromatography-olfactometry (GC-O) can also be considered a method used for untargeted analyses, even though the information provided is different with respect to GC-MS. For such a reason, it is a common occurrence to split the column effluent between a mass spectrometer and an olfactometry port to attain complementary types of information (Delahunty et al., 2006). Again,

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the non-sufficient resolving power of a single GC column can hinder the attainment of reliable GC-O-MS essential oil information, calling for the use of a multidimensional (MD) GC method. However, the enhanced separation power of heart-cutting MDGC, and not GC × GC, is the preferred choice in such instances (Delahunty et al., 2006). In general, the separation power of GC × GC is most impressive in untargeted investigations, with essential oils making no exception. The first GC × GC separations involving essential oils appeared in 2000: Dimandja et al. used a thermal sweeper (a now obsolete modulator), and flame ionization detection (FID), for the analysis of peppermint and spearmint oils (Dimandja et al., 2000). A rather curious set of columns was used (the first column was shorter than the second): non-polar, 1 m × 0.1 mm ID × 3.5 µm df + medium-polarity 2 m × 0.1 mm ID × 0.5 µm df. A side-by-side comparison was made with GC-MS results, attained by using a non-polar 30 m column: three times more peaks were detected in the GC × GC-FID analysis of peppermint oil compared to GC-MS (89 vs. 30). Even though not fully expressed, the analytical power of GC × GC was evident in this initial untargeted experiment. In a recent study, Filippi et al. carried out an in-depth qualitative and quantitative investigation of four samples of vetiver oil (Haiti, Indonesia, Brazil, La Réunion) by using GC × GC combined with single-quadrupole (Q) MS and GC × GC-FID, as well as GC-QMS (Filippi et al., 2013). A total ion current (TIC) GC-QMS chromatogram, relative to Haitian vetiver oil, is shown in Figure 7.7A, with a total number of 101 peaks detected with a signal-to-noise ratio (s/n) > 10. The same sample was analyzed by using GC × GC-QMS, with the number of detected peaks (s/n > 10) reaching 535, even though the injector split ratio was doubled compared to the GC-QMS analysis (Figure 7.7B). It is noteworthy, however, that only 117 compounds were identified and then subjected to quantification by using GC × GC-FID. As a consequence, there was still a great deal of concealed information, despite the high number of detected essential oil constituents. In general, such a misbalance is often observed when using GC × GC-MS. The research of Filippi et  al. highlighted the enhanced separation power and sensitivity of GC × GC. An additional emphasized advantage was the formation of elution patterns of specific compound classes (e.g., ketones, hydrocarbons, alcohols, aldehydes, etc.), increasing the reliability of identification. Finally, as will be discussed later, GC × GC generates a true analytical fingerprint possessing a wealth of exploitable information. The on-line multidimensional combination of high-performance liquid chromatography and gas chromatography (LC-GC) has a proven usefulness within the context of essential oil analysis (Marriott et al., 2001). The LC dimension is used to perform the separation of chemical classes of compounds on the basis of polarity. Fractions with a reduced complexity can then be subjected to GC analysis, usually by injecting large sample volumes. The LC-GC concept was extended to the off-line combination of LC and GC × GC-QMS (LC// GC × GC-QMS) by Tranchida et al. (Tranchida et al., 2013b), who exploited the first analytical dimension for the separation of hydrocarbons and O-containing compounds in sweet orange and bergamot essential oils. In the bergamot oil analysis, for instance, approximately 350 peaks were detected with 195 analytes tentatively identified (53 hydrocarbons and 142 oxygenated constituents), against 64 compounds (31 hydrocarbons and 33 oxygenated constituents) using direct GC-QMS. Flow modulation (FM) is an interesting alternative to cryogenic modulation due to the limited hardware and operational costs, and to the capability to modulate with efficiency compounds with a high volatility (e.g., ≤ C6), as well as a low one (e.g., ≥ C30). On the other hand, FM cannot match the higher peak capacity and sensitivity provided by cryogenic modulation (CM). A further recognized disadvantage of FM is the presumed requirement of very high 2D gas flows (e.g., ≥20 mL min−1), a problematic issue when using mass spectrometry (Edwards et al., 2011). However, it has also been demonstrated that through fine method optimization, it is possible to carry out FM GC × GC applications at gas flows within the range 6–8 mL min−1 (Tranchida et al., 2014). Several commercially available MS systems can handle such gas flows. Costa et al. used FM GC × GC-QMS for the untargeted analysis of Artemisia arborescens L. leaf essential oil (Costa et al., 2016). The column set consisted of a low-polarity 20 m × 0.18 mm

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FIGURE 7.7  Analysis of Haitian vetiver oil by using (A) GC-QMS and (B) GC × GC-QMS. (Reprinted from J. Chromatogr. A., 1288. Filippi, J. J. et al., Qualitative and quantitative analysis of vetiver essential oils by comprehensive two-dimensional gas chromatography and comprehensive two-dimensional gas chromatography/ mass spectrometry. 127–148. Copyright 2013, with permission from Elsevier.)

ID × 0.18 µm df 1D and a mid-polarity 10 m × 0.32 mm ID × 0.20 µm df 2D. The use of rather long wide-bore 2D columns in FM analyses is an obliged choice, related not only to the high gas flows (in this case 8 mL min−1), but also to the wide chromatography bands released from the modulator. In fact, differently from CM no solute re-concentration effects occur. Figure 7.8 illustrates three

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FIGURE 7.8  Three chromatogram expansions (A-C) relative to the FM GC × GC-QMS analysis of Artemisia arborescens L. leaf essential oil. (Reprinted from J. Pharm. Biomed. Anal., 117. Costa, R. et al., 2016. Phytochemical screening of Artemisia arborescens L. by means of advanced chromatographic techniques for identification of health-promoting compounds. 499–509. Copyright 2016, with permission from Elsevier.)

2D expansions, extrapolated from the entire chromatogram, relative to the FM GC × GC-QMS analysis of Artemisia arborescens L. leaf essential oil. Mass spectral database searching, with the support of LRI information, enabled the tentative identification of 92 compounds (about two times more were detected).

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7.3.6.1.2  Pre-targeted Analyses With regard to the pre-targeted analysis of essential oils, there are a series of different possibilities, with the use of GC × GC often questionable. For example, pre-targeted essential oil analysis can relate to the determination of phytosanitary compounds, plasticizers, antioxidants, etc. (Di Bella et al., 2006; Garland et al., 1999). It is the specificity and sensitivity of MS which play a fundamental role in such an applicational field. Even so, the use of GC × GC has been described: Tranchida et al. exploited FM GC × GC, hyphenated with triple quadrupole (QqQ) MS, for the determination of the antioxidants butylated hydroxyanisole (BHA) and butylated hydroxytoluene (BHT), as well as the fungicide o-phenylphenol (OPP), in mandarin essential oil (Tranchida et al., 2013a). The QqQ mass spectrometer was a novel device and was capable of rapid switching between the untargeted (scan) and targeted (MSMS) modes during the same analysis. The main aim of the study was to evaluate the QqQMS instrument under challenging analytical conditions. However, the capability of the mass spectrometer to provide two distinct types of information, practically simultaneously, does potentially justify the use of GC × GC in this type of application. A further example of pre-targeted analysis, very useful to confirm (or not) genuineness, is that involving the determination of the enantiomeric composition of specific chiral constituents. Target analyte elution on shape-selective GC stationary phases (enantio-GC), in particular those containing cyclodextrins, represents in this case the most important step of the analytical chain. Enantio-GC essential oil separations are usually carried out by using a single column or, more preferably, on the second dimension of a heart-cutting MDGC instrument (Marriot et al., 2001). Even though both enantio-GC × GC and GC × enantio-GC studies have been reported (Shellie et al., 2001; Shellie and Marriott, 2002), such approaches have not found popularity. On one hand, the modulation process can degrade 1D enantiomer resolution, while on the other, column lengths in the 2D are usually too short to provide satisfactory degrees of separation. The combination of either conventional GC or heart-cutting MDGC, with isotope ratio mass spectrometry (IRMS), has a proven usefulness for the measurement of target analyte isotopic ratios (Schipilliti et al., 2011; Juchelka et al., 1998). Again, MDGC-IRMS represents the preferred choice for the pre-targeted analysis of essential oils, also because the IRMS combustion chamber generally leads to extensive band broadening, causing a resolution loss. Isotopic ratio determinations are performed to guarantee authenticity or to define a geographical origin. To the best of the present authors’ knowledge, the use of GC × GC-IRMS for the analysis of essential oils has not yet been reported. 7.3.6.1.3 Fingerprinting Fingerprinting is a form of untargeted analysis, inasmuch that interest is devoted to the chromatography profile, as a whole or in part, rather than to peak-to-peak assignment. The concept of fingerprinting is two-dimensional in its nature and so it matches well with GC × GC. In fact, an intimate bond can be created between a highly detailed 2D chromatogram and a specific type of sample, preferably by using adequate data treatment software tools. Fine and reliable GC × GC between-sample differentiation can be achieved on the basis of a specific industrial process (e.g., type of roasting), of geographical origin, or of sensorial properties (e.g., oxidized vegetable oils) (Cordero et al., 2010; Purcaro et al., 2014). The GC × GC fingerprinting of essential oils has been rarely reported; in one of such rare instances, Cordero et al. used FM GC × GC with dual detection (FID/QMS) for the fingerprinting of four vetiver oils, namely Java, Brazil, Bourbon, and Haiti (Cordero et  al., 2015). Prior to visualization, thresholds were set for s/n values (>25) and peak volumes (>30,000). Figure 7.9A illustrates the result for the Haitian sample, highlighting also the formation of chemical-class elution patterns. Following alignment, a comparative visualization was made between the Bourbon and Haitian vetiver oils. Regions with a red and green color highlight positive and negative detector response differences, respectively, for the Haitian oil (Figure 7.9B). A further software function allowed a more clear between-oil differentiation: 315 peaks were aligned, with those characterized by the largest peak volume difference (CV% >50%) pinpointed by a yellow circle (Figure 7.9C).

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FIGURE 7.9  (A) Two-dimensional chromatogram of Haitian vetiver oil; (B) visual comparison between Bourbon and Haitian vetiver oils; yellow circles indicate compounds with the largest quantitative differences between the four vetiver oils. (Reprinted from J. Chromatogr. A, 1417. Cordero, C. et al., Potential of the reversed-inject differential flow modulator for comprehensive two-dimensional gas chromatography in the quantitative profiling and fingerprinting of essential oils of different complexity. 79–95. Copyright 2015, with permission from Elsevier.)

For instance, β-vetivenene, a sesquiterpene hydrocarbon, was contained in the highest amounts in the Java and Brazilian oils, while the Haitian oil was characterized, by far, by the highest concentration of (E)-isovalencenol, an oxygenated sesquiterpene. The availability of such detailed information, accompanied by the use of suitable statistical processes, results in a powerful sample classification approach.

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7.3.6.2  On-Line and Off-Line Coupled Liquid Chromatography-Gas Chromatography The analysis of very complex mixtures is often troublesome due to the variety of chemical classes to which the sample components belong to and to their wide range of concentrations. As such, several compounds cannot be resolved by monodimensional GC. In this respect, less complex and more homogeneous mixtures can be attained by the fractionation of the matrix by means of LC prior to GC separation. The multidimensional LC-GC approach, combines the selectivity of the LC separation with the high efficiency and sensitivity of GC separation, enabling the separation of compounds with similar physicochemical properties in samples characterized by a great number of chemical classes. For the highly volatile components, commonly present in essential oils, the most adequate transfer technique is partially concurrent eluent evaporation (Grob, 1987). In this technique, a retention gap is installed, followed by a few meters of pre-column and the analytical capillary GC column, both with an identical stationary phase, for the separation of the LC fractionated components. A vapor exit is placed between the pre-column and the analytical column, allowing partial evaporation of the solvent. Hence, column and detector overloading are avoided. This transfer technique can be applied to the analysis of GC components with a boiling point of at least 50°C higher than the solvent. The interface proposed by Grob was improved in 2009 by Biedermann and Grob, introducing the so-called Y-interface in order to reduce the memory effect (Biedermann and Grob, 2009). In the last few years, different transfer devices have been developed to transfer the fractions to the GC system (Purcaro et al., 2013; Biedermann and Grob, 2012). The composition of citrus essential oils has been elucidated by means of LC-GC, and the development of new methods for the study of single classes of components has been well reported. The aldehyde composition in sweet orange oil has been investigated (Mondello et al., 1994a), as also industrial citrus oil mono- and sesquiterpene hydrocarbons (Mondello et al., 1995) and the enantiomeric distribution of monoterpene alcohols in lemon, mandarin, sweet orange, and bitter orange oils (Dugo et al., 1994a,b). The hyphenation of LC-GC systems to mass spectrometric detectors has also been reported for the analyses of neroli (Mondello et al., 1994b), bitter and sweet oranges, lemon, and petitgrain mandarin oils (Mondello et al., 1996). It has to be highlighted, that the preliminary LC separation, which reduces mutual component interference, greatly simplifies MS identification.

7.3.7 Liquid Chromatography, Liquid Chromatography Hyphenated to Mass Spectrometry, and Multidimensional Liquid Chromatographic Technique in the Analysis of Essential Oils Essential oils generally contain only volatile components, since their preparation is performed by steam distillation. Citrus oils, extracted by cold-pressing machines, are an exception, containing more than 200 volatile and non-volatile components. The non-volatile fraction, constituting 1%–10% of the oil, is represented mainly by hydrocarbons, fatty acids, sterols, carotenoids, waxes, and oxygen heterocyclic compounds (coumarins, psoralens, and polymethoxylated flavones) (Di Giacomo and Mincione, 1994). The latter can have an important role in the identification of a cold-pressed oil and in the control of both quality and authenticity (Di Giacomo and Mincione, 1994; Dugo et al., 1996; Mc Hale and Sheridan, 1989; Mc Hale and Sheridan, 1988), when the information attained by means of GC is not sufficient. The analysis of these compounds is usually performed by means of liquid chromatography (LC), also referred to as high performance liquid chromatography (HPLC), in normal-phase (NP-HPLC) or reversed-phase (RP-HPLC) applications (Dugo and Di Giacomo, 2002). Nowadays, the analysis of oxygen heterocyclic compounds plays a fundamental role in the characterization and quality control of citrus essential oils; therefore, rapid analytical strategies have been developed. Fast methods were validated in order to perform a rapid quality control and determine furocoumarins in essences, according to the European Regulation EC No. 1223/2009, which sets the maximum amount of furocoumarins admitted in finished cosmetic products containing citrus essential oils (EC No. 1223/2009). New HPLC methods are based on the use of

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short columns, which ensure fast separations and guarantee a high separation efficiency thanks to the small dimension of the packaging particles. Fused-core columns are the most employed; the superficially porous structure of their particles reduce the analytes path, and consequently the molecule dispersion and retention times. These columns were employed to study the chemical composition variability of citrus essences depending on seasoning and production procedure (Dugo et al., 2010; Dugo et al., 2011; Dugo et al., 2012; Russo et al., 2012). A satisfactory separation of 38 targets was achieved by using an Ascentis Express C18 column (50 × 4.6 mm ID with particle size of 2.7 µm) in ten minutes, or also in shorter time (three minutes), maintaining a good compromise between chromatographic resolution and analysis time in RP-HPLC-photo dyode array detector PDA Figure 7.10 (Russo et al., 2015). This method was applied to analyze seven citrus essential oils (lime, lemon, bergamot, bitter orange, grapefruit, mandarin, and sweet orange). The method developed was further improved by combining the use of LRIs (using alkyl aryl ketones) and UV spectral libraries for reliable identification of oxygen heterocyclic components in citrus essential oils (Arigò et al., 2019). In some cases, the oxygen heterocyclic components proved to be more effective as markers in adulteration detection than the volatile components (Fan et al., 2015). More recent methods are based on HPLC systems coupled to MS detectors (Mehl et al., 2014; Mehl et al., 2015; Donato et al., 2014; Masson et al., 2016). This approach allows the rapid identification and characterization of oxygen heterocyclic compounds in citrus oils, and detection of some minor components for the first time in some oils was reported (Dugo et al., 1999). In addition, markers to monitor authenticity and possible adulteration were identified. Furthermore, the high selectivity and sensitivity of MS detection, mainly atmospheric pressure chemical ionization (APCI)-positive ionization in Multiple Reaction Monitoring acquisition mode, allows the determination of oxygen heterocyclic compounds in finished cosmetic products, as imposed by the European regulation cited above. A new ionization approach was employed to analyze polymethoxyflavones isolated from the non-volatile residue of mandarin essential oil. This technique consists of a nano-LC-EI-MS configuration made of a nano prominence HPLC system coupled to a GC-QMS, working in electron impact ionization mode (Russo et al., 2016).

FIGURE 7.10  Reverse-phase high-performance liquid chromatography (RP-HPLC) chromatograms of oxygen heterocyclic compounds present in Citrus essential oils. (Reprinted with permission of Taylor & Francis Group, from Russo, M. et al., 2015. J. Essent. Oil Res., 27: 307–315.)

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Apart from citrus oils, other essential oils have also been analyzed by means of liquid chromatography, such as the blackcurrant bud (Píry and Pribela, 1994), Nigella sativa essential oil (Abdel-Reheem et al., 2014); cinnamon, caraway, and cardamom fruit oils (Porel et al., 2014); Zanthoxylum zanthoxyloides Lam. fruit oil (Tine et al., 2017), and mint (Hawryt et al., 2017) and rice leaf essential oils (Minh et al., 2019). The most recent applications of liquid chromatography in the analysis of essential oils are mainly related to the isolation of interesting compounds (Russo et al., 2016; Gu et al., 2016; Padhan et al., 2017; Piochon-Gauthier et al., 2014), to the analysis of thermolabile components (Quassinti et al., 2014; Maggi et al., 2015), or to characterize the oils obtained through innovative and green techniques (Khajenoori et al., 2015). Recent applications regard the determination of several pesticides in essential oils by HPLC coupled to tandem mass spectrometry (Fillatre et al., 2014; Fillatre et al., 2016). High performance liquid chromatography (HPLC) has acquired a role of great importance in food analysis, as demonstrated by the wide variety of applications reported. Single LC column chromatographic processes have been widely applied for sample profile elucidation, providing satisfactory degrees of resolving power; however, whenever highly complex samples require analysis, a monodimensional HPLC system can prove to be inadequate. Moreover, peak overlapping may occur even in the case of relatively simple samples, containing components with similar properties. The basic principles of MDGC are also valid for multidimensional LC (MDLC). The most common use of MDLC separation is the pre-treatment of a complex matrix in an off-line mode. The off-line approach is very easy, but presents several disadvantages: it is time-consuming, operationally intensive, and difficult to automate and to reproduce. Moreover, sample contamination or formation of artefacts can occur. On the other hand, on-line MDLC, though requiring specific interfaces, offers the advantages of ease of automation and greater reproducibility in a shorter analysis time. In the on-line heart-cutting system, the two columns are connected by means of an interface, usually a switching valve, which allows the transfer of fractions of the first column effluent onto the second column. In contrast to comprehensive gas chromatography (GC × GC), the number of comprehensive liquid chromatography (LC × LC) applications reported in the literature are much less. It can be affirmed that LC × LC presents a greater flexibility when compared to GC × GC since the mobile phase composition can be adjusted in order to obtain enhanced resolution (Dugo et  al., 2006a). Comprehensive HPLC systems, developed and applied to the analysis of food matrices, have employed the combination of either NP  ×  RP or RP  ×  RP separation modes. However, it is worthy of note that the two separation mechanisms exploited should be as orthogonal as possible, so that no or little correlation exists between the retention of compounds in both dimensions. A typical comprehensive two-dimensional HPLC separation is attained through the connection of two columns by means of an interface (usually a high pressure switching valve), which entraps specific quantities of first-dimension eluate and directs it onto a secondary column. This means that the first column effluent is divided into “cuts” which are transferred continuously to the second dimension by the interface. The type of interface depends on the methods used, although multiport valve arrangements have been the most frequently employed. Various comprehensive HPLC systems have been developed and proved to be effective for the separation of complex sample components, and in the resolution of a number of practical problems. In fact, the very different selectivities of the various LC modes enable the analysis of complex mixtures with minimal sample preparation. However, comprehensive HPLC techniques are complicated by the operational aspects of switching effectively from one operation step to another, by data acquisition and interpretation issues. Therefore, careful method optimization and several related practical aspects should be considered. In the most common approach, a micro-bore LC column in the first and a conventional column in the second dimension are used. In this case, an 8-, 10-, or 12-port valve equipped with two sample loops (or trapping columns) is used as an interface. A further approach foresees the use of a conventional LC column in the first, and two conventional columns in the second, dimension. One

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or two valves that allow transfers from the first column to two parallel secondary columns (without the use of storage loops) are used as interface. One of the best examples of the application of comprehensive NPLC × RPLC in essential oil analysis is represented by the analysis of oxygen heterocyclic components in cold-pressed lemon oil, by using normal-phase with a micro-bore silica column in the first dimension and a monolithic C18 column in the second dimension with a 10-port switching valve as interface (Dugo et al., 2004). In Figure 7.11, an NPLC × RPLC separation of the oxygen heterocyclic fraction of a lemon oil sample is presented. Oxygen heterocyclic components (coumarins, psoralens, and polymethoxylated flavones) represent the main part of the non-volatile fraction of cold-pressed citrus oils. Their structures and substituents have an important role in the characterization of these oils. Positive peak identification of these compounds was obtained by both the relative location of the peaks in the 2D plane, which varied in relation to their chemical structure and by characteristic UV spectra. In a later experiment, a similar setup was used for a citrus oil extract composed of lemon and orange oil (François et al., 2006). The main difference with respect to the earlier published work (Dugo et al., 2004) was the employment of a bonded phase (diol) column in the first dimension. Under optimized LC conditions, the high degree of orthogonality between the NP and RP systems tested resulted in increased 2D peak capacity. The NPLC × RPLC approach has been also developed for the analysis of carotenoids, pigments mainly distributed in plant-derived foods, especially in orange and mandarin essential oils (Dugo et al., 2006b; Dugo et al., 2008). In terms of structures, food carotenoids are polyene hydrocarbons, characterized by a C40 skeleton that derives from eight isoprene units. They present an extended conjugated double bond (DB) system that is responsible for the yellow, orange, or red colors in plants and are notable for their wide distribution, structural diversity, and various functions. Carotenoids are usually classified in two main groups: hydrocarbon carotenoids, known as carotenes (e.g., β-carotene and lycopene), and oxygenated carotenoids, known as xanthophylls (e.g., β-cryptoxanthin and lutein). The elucidation of carotenoid patterns is particularly challenging because of the complex composition of carotenoids in natural matrices, their great structural diversity, and their extreme instability. An innovative comprehensive dual-gradient elution HPLC system was employed using

FIGURE 7.11  Comprehensive NP (adsorption)-LC × RPLC separation of the oxygen heterocyclic fraction of a lemon oil sample (for peak identification see Dugo et al., 2004). (Reprinted with permission from Dugo, P. et al., 2004. Comprehensive two-dimensional normal-phase (adsorption)-reversed-phase liquid chromatography. Anal. Chem., 76: 2525–2530. Copyright 2004, American Chemical Society.)

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FIGURE 7.12  Contour plot of the comprehensive HPLC analyses of carotenoids present in sweet orange essential oil with peaks and compound classes indicated (for peak identification see Dugo et  al., 2006b). (Reprinted with permission from Dugo, P. et al., 2006b. Elucidation of carotenoid patterns in citrus products by means of comprehensive normal-phase × reversed-phase liquid chromatography. Anal. Chem., 78: 7743–7750. Copyright 2006b, American Chemical Society.)

an NPLC × RPLC setup, composed of silica and C18 columns in the first and second dimensions, respectively. Free carotenoids in orange essential oil and juice (after saponification), were identified by combining the two-dimensional retention data with UV-visible spectra (Dugo et  al., 2006b) obtained by using a PDA detector (Figure 7.12). The carotenoid fraction of a saponified mandarin oil has been studied by means of comprehensive LC, in which a 1D micro-bore silica column was applied for the determination of free carotenoids and a cyanopropyl column for the separation of esters; a monolithic column was used in the 2D (Dugo et al., 2008). Detection was performed by connecting a photodiode array detection (DAD) system in parallel with an MS detection system operated in the APCI positive-ion mode. Thus, the identification of free carotenoid and carotenoid esters was carried out by combining the information provided by the DAD and MS systems and the peak positions in the 2D chromatograms.

7.4  GENERAL CONSIDERATIONS ON ESSENTIAL OIL ANALYSIS As evidenced by the numerous techniques described in the present contribution, chromatography, especially gas chromatography, has evolved into the dominant method for essential oil analysis. This is to be expected because the complexity of the samples must be unraveled by some type of separation, before the sample constituents can be measured and characterized; in this respect, gas chromatography provides the greatest resolving power for most of these volatile mixtures. In the past, a vast number of investigations have been carried out on essential oils, and many of these natural ingredients have been investigated following the introduction of GC-MS, which marked a real turning point in the study of volatile molecules. Es-GC also represented a landmark in the detection of adulterations, and in the cases where the latter technique could fail, gas chromatography hyphenated to isotope ratio mass spectrometry by means of a combustion interface (GC-IRMS) has proved to be a valuable method to evaluate the genuineness of natural

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product components. In addition, the introduction of GC-O was a breakthrough in analytical aroma research, enabling the differentiation of a multitude of volatiles in odor-active and nonodor-active compounds, according to their existing concentrations in a matrix. The investigation of the non-volatile fraction of essential oils, by means of LC and its related hyphenated techniques, contributed greatly toward the progress of the knowledge on essential oils. Many extraction techniques have also been developed, boosting the attained results. Moreover, the continuous demand for new synthetic compounds reproducing the sensations elicited by natural flavors triggered analytical investigations toward the attainment of information on scarcely known properties of well-known matrices.

REFERENCES Abdel-Reheem, M. A. T., M. M. Oraby, and S. K. Alanazi, 2014. Nigella sativa essential oil constituents and its antimicrobial, cytotoxic and necrotic replies. J. Pure Appl. Microbiol., 8: 389–398. Acree, T. E., J. Barnard, and D. Cunningham, 1984. A procedure for the sensory analysis of gas chromatographic effluents. Food Chem., 14: 273–286. AFNOR- Association Française de Normalisation, Saint-Denis, http://www.afnor.org/portail.asp (accessed January 11, 2008). Arigò, A., F. Rigano, G. Micalizzi, P. Dugo, and L. Mondello, 2019. Oxygen heterocyclic compound screening in Citrus essential oils by linear retention index approach applied to liquid chromatography coupled to photodiode array detector. Flav. Fragr. J., 34: 349–364. in press. Ball, G. I., L. Xu, A. P. Mc Nichol, and L. I. Aluwihare, 2012. A two-dimensional, heart-cutting preparative gas chromatograph facilitates highly resolved single-compound isolations with utility towards compoundspecific natural abundance radiocarbon (14C) analyses. J. Chromatogr. A., 1220: 122–131. Beilstein, F., 1872. Ueber den Nachweis von Chlor, Brom und Jod in organischen Substanzen. Ber. Dtscg. Chem. Ges., 5: 620–621. Bicchi, C., C. Brunelli, C. Cordero, P. Rubiolo, M. Galli, and A. Sironi, 2004. Direct resistively heated column gas chromatography (ultrafast module-GC) for high-speed analysis of essential oils of differing complexities. J. Chromatogr. A, 1024: 195–207. Bicchi, C., C. Brunelli, C. Cordero, P. Rubiolo, M. Galli, and A. Sironi, 2005. High-speed gas chromatography with direct resistively-heated column (ultra fast module-GC)-separation measure (S) and other chromatographic parameters under different analysis conditions for samples of different complexities and volatilities. J. Chromatogr. A, 1071: 3–12. Bicchi, C., C. Brunelli, M. Galli, and A. Sironi, 2001. Conventional inner diameter short capillary columns: An approach to speeding up gas chromatographic analysis of medium complexity samples. J. Chromatogr. A, 931: 129–140. Bicchi, C., A. D’Amato, V. Manzin, and P. Rubiolo, 1997. Cyclodextrin derivatives in GC separation of racemic mixtures of volatiles. Part XI. Some applications of cyclodextrin derivatives in GC Enantioseparations of essential oil components. Flav. Fragr. J., 12: 55–61. Bicchi, C., A. D’Amato, and P. Rubiolo, 1999. Cyclodextrin derivatives as chiral selectors for direct gas chromatographic separation of enantiomers in the essential oil, aroma and flavour fields. J. Chromatogr. A, 843: 99–121. Bicchi, C., C. Frattini, G. M. Nano, and A. D’Amato, 1988. On column injection-dual channel analysis of essential oil. J. High Resol. Chromatogr., 11: 56–60. Biedermann, M., and K. Grob, 2009. Memory effects with the on-column interface for on-line coupled high performance liquid chromatography-gas chromatography: The Y-interface. J. Chromatogr. A, 1216: 8652–8658. Biedermann, M., and K. Grob, 2012. On-line coupled high performance liquid chromatography-gas chromatography for the analysis of contamination by mineral oil. Part 1: Method of analysis. J. Chromatogr. A, 1255: 56–75. Boelens, M. H., H. Boelens, and L. J. van Gemert, 1993. Sensory properties of optical isomers. Perf. Flav., 18: 2–16. Bonaccorsi, I., D. Sciarrone, L. Schipilliti, P. Dugo, L. Mondello, and G. Dugo, 2012. Multidimensional enantio gas chromatography/mass spectrometry and gas chromatography-combustion-isotopic ratio mass spectrometry for the authenticity assessment of lime essential oils (C. aurantifolia Swingle and C. latifolia Tanaka). J. Chromatogr. A, 1226: 87–95. Bosart, L. W. 1937. Perfumery Essential Oil Record, 28, 95.

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British Pharmacopoeia BP 2008, 2007. The Stationery Office (TSO), Norwich. Casilli, A., E. Decorzant, A. Jaquier, and E. Delort, 2014. Multidimensional gas chromatography hyphenated to mass spectrometry and olfactometry for the volatile analysis of citrus hybrid peel extract. J. Chromatogr. A, 1373: 169–178. Casimir, D. J., and F. B. Whitfield, 1978. Flavour impact values, a new concept for assigning numerical values for the potency of individual flavour components and their contribution to the overall flavour profile, Ber. Int. Fruchtsaftunion., 15: 325. Chin, S. T., and P. J. Marriott, 2015. Review of the role and methodology of high resolution approaches in aroma analysis. Anal. Chim. Acta, 854: 1–12. Cordero, C., E. Liberto, C. Bicchi et al., 2010. Profiling food volatiles by comprehensive two-dimensional gas chromatography coupled with mass spectrometry: Advanced fingerprinting approaches for comparative analysis of the volatile fraction of roasted hazelnuts (Corylus avellana L.) from different origins. J. Chromatogr. A, 1217: 5848–5858. Cordero, C., P. Rubiolo, L. Cobelli et al., 2015. Potential of the reversed-inject differential flow modulator for comprehensive two-dimensional gas chromatography in the quantitative profiling and fingerprinting of essential oils of different complexity. J. Chromatogr. A, 1417: 79–95. Costa, R., S. Ragusa, M. Russo et al., 2016. Phytochemical screening of Artemisia arborescens L. by means of advanced chromatographic techniques for identification of health-promoting compounds. J. Pharm. Biomed. Anal., 117: 499–509. Cramers, C. A., H. G. Janssen, M. M. van Deursen, and P. A. Leclercq, 1999. High-speed gas chromatography: An overview of various concepts. J. Chromatogr. A, 856: 315–329. Cramers, C. A., and P. A. Leclercq, 1999. Strategies for speed optimisation in gas chromatography: An overview, J. Chromatogr. A, 842: 3–13. Croteau R., Kutchan, T. M., and N. G. Lewis, 2000. Natural Products (Secondary Metabolites). In Biochemistry & Molecular Biology of Plants, 1st ed., B. Buchanan, W. Gruissen, and R. Jones (eds.). New Jersey: ASPB and Wiley, chap. 24. Curtis, T., and Williams, D. G., 2001. Introduction to Perfumery, 2nd ed. New York: Micelle Press, chap. 3. d’Acampora Zellner, B., M. Lo Presti, L. E. Barata, P. Dugo, G. Dugo, and L. Mondello, 2006. Evaluation of leaf-derived extracts as an environmentally sustainable source of essential oils by using gas chromatography-mass spectrometry and enantioselective gas chromatography-olfactometry. Anal. Chem., 78: 883–890. Deans, D. R. 1968. A new technique for heart cutting in gas chromatography. Chromatographia, 1: 18–22. Delahunty, C. M., G. Eyres, and J.-P. Dufour, 2006. Gas chromatography-olfactometry. J. Sep. Sci., 29: 2107–2125. Di Bella, G., L. Serrao, F. Salvo, V. Lo Turco, M. Croce, and G. Dugo, 2006. Pesticide and plasticizer residues in biological citrus essential oils from 2003–2004. Flav. Fragr. J. 21: 497–501. Di Giacomo, A., and B. Mincione, 1994. Gli Olii Essenziali Agrumari in Italia. Baruffa editore. Dimandja, J-M. D., S. B. Stanfill, J. Grainger, and D. G. Patterson, 2000. Application of comprehensive twodimensional gas chromatography (GC × GC) to the qualitative analysis of essential oils. J. High Resolut. Chromatogr., 23: 208–214. Do, T. K. T., F. Hadjji-Minaglou, S. Antoniotti, and X. Fernandez, 2015. Authenticity of essential oils. Trends Anal. Chem., 66: 146–157. Donato, P., I. Bonaccorsi, M. Russo, and P. Dugo, 2014. Determination of new bioflavonoids in bergamot (Citrus bergamia) peel oil by liquid chromatography coupled to tandem ion trap-time of flight mass spectrometry. Flav Fragr. J., 29, 131–136. Dugo, G., Lamonica, G., Cotroneo, A., Stagno d’Alcontres, I., Verzera, A., Donato, M.G., Dugo, P., and G. Licandro, 1992. High resolution gas chromatography for detection of adulterations of citrus cold-pressed essential oils. Perf. & Flav., 17: 57–74. Dugo, G., I. Bonaccorsi, D. Sciarrone et al., 2012. Characterization of cold-pressed and processed bergamot oils by using GC-FID, GC-MS, GC-C-IRMS, enantio-GC, MDGC, HPLC and HPLC-MS-IT-TOF. J. Essent. Oil Res., 24: 93–117. Dugo G., and A. Di Giacomo, 2002. Citrus: The Genus Citrus, 1st ed. Book Chapter, Taylor & Francis Group. Dugo, P., I. Bonaccorsi, C. Ragonese et al., 2011. Analytical characterization of mandarin (Citrus deliciosa Ten.) essential oil. Flav. Frag. J., 26: 34–46. Dugo, P., O. Favoino, R. Luppino, G. Dugo, and L. Mondello, 2004. Comprehensive two-dimensional normalphase (adsorption)-reversed-phase liquid chromatography. Anal. Chem., 76: 2525–2530. Dugo, P., M. M. Fernandez, A. Cotroneo, and L. Mondello, 2006a. Optimization of a comprehensive two-dimensional normal-phase and reversed phase-liquid chromatography system. J. Chromatogr. Sci., 44: 561–565.

Analysis of Essential Oils

223

Dugo, P., M. Herrero, T. Kumm, D. Giuffrida, G. Dugo, and L. Mondello, 2008. Comprehensive normalphase × reversed-phase liquid chromatography coupled to photodiode array and mass spectrometry detectors for the analysis of free carotenoids and carotenoid esters from Mandarin. J. Chromatogr. A, 1189: 196–206. Dugo, P., L. Mondello, E. Cogliandro, A. Verzera, and G. Dugo, 1996. On the genuineness of citrus essential oils. 51. Oxygen heterocyclic compounds of bitter orange oil (Citrus aurantium L.). J. Agric. Food Chem., 44: 544–549. Dugo, P., L. Mondello, E. Sebastiani, R. Ottanà, G. Errante, and G. Dugo, 1999. Identification of minor oxygen heterocyclic compounds of citrus essential oils by liquid chromatography-atmospheric pressure chemical ionisation mass spectrometry. J. Liquid Chrom. & Rel. Tech., 22: 2991–3005. Dugo, P., C. Ragonese, M. Russo et al., 2010. Sicilian lemon oil: Composition of volatile and oxygen heterocyclic fractions and enantiomeric distribution of volatile components. J. Sep. Sci., 33: 3374–3385. Dugo, P., V. Škeřiková, T. Kumm et al., 2006b. Elucidation of carotenoid patterns in citrus products by means of comprehensive normal-phase × reversed-phase liquid chromatography. Anal. Chem., 78: 7743–7750. Dugo, G., A. Verzera, A. Cotroneo, I. Stagno d’Alcontres, L. Mondello, and K. D. Bartle, 1994a. Automated HPLC-HRGC: A powerful method for essential oil analysis. Part II. Determination of the enantiomeric distribution of Linalol in sweet orange, bitter orange and mandarin essential oils. Flav. Fragr. J., 9: 99–104. Dugo, G., A. Verzera, A. Trozzi, A. Cotroneo, L. Mondello, and K. D. Bartle, 1994b. Automated HPLCHRGC: A powerful method for essential oils analysis. Part I. Investigation on enantiomeric distribution of monoterpene alcohols of lemon and mandarin essential oils. Essenz. Deriv. Agrum., 44: 35–44. Edwards, M., A. Mostafa, and T. Górecki, 2011. Modulation in comprehensive two-dimensional gas chromatography: 20 years of innovation. Anal. Bioanal. Chem., 401: 2335–2349. Elbashir, A. A., and H. Y. Aboul-Enein, 2018. Multidimensional gas chromatography for chiral analysis. Critical Rev. Anal. Chem., 48: 416–427. Ettre, L. S., and J. V. Hinshaw, 1993. Basic relationships of Gas Chromatography, 1st ed. Cleveland: Advanstar Data, chap. 4. Regulation (ec) no 1223/2009 of the European Parliament and of the Council of 30 November 2009 on Cosmetic products (recast) (text with eea relevance). European Parliament, Off. J. Eur. Commun. L 342 (2009) 59. European Pharmacopoeia, 6th ed. Strasburg: European Directorate for the Quality of Medicines & Healthcare (Edqm), 2007. Falkenberg, M. B., R. I. Santos, and C. M. O. Simões, 1999. Introdução à Análise Fitoquímica. In Farmacognosia: Da planta ao medicamento, C. M. O. Simões et al. (ed.). 1999. Editora da UFSC and Editora da Universidade/UFRGS: Florianópolis, chap. 10. Fan, H., Q. Wu, J. E. Simon, S. Lou, and C. Ho, 2015. Authenticity analysis of citrus essential oils by HPLCUV-MS on oxygenated heterocyclic components. J. Food and Drug Anal., 23: 30–39. Filippi, J. J., E. Belhassen, N. Baldovini, H. Brevard, and U. J. Meierhenrich, 2013. Qualitative and quantitative analysis of vetiver essential oils by comprehensive two-dimensional gas chromatography and comprehensive two-dimensional gas chromatography/mass spectrometry. J. Chromatogr. A., 1288: 127–148. Fillâtre, Y., D. Rondeau, A. Daguin, and P. Y. Communal, 2016. A workflow for multiclass determination of 256 pesticides in essential oils by liquid chromatography tandem mass spectrometry using evaporation and dilution approaches: Application to lavandin, lemon and cypress essential oils. Talanta, 149: 178–186. Fillâtre, Y., D. Rondeau, A. Daguin, A. Jadas-Hecart, and P. Y. Communal, 2014. Multiresidue determination of 256 pesticides in lavandin essential oil by LC/ESI/sSRM: Advantages and drawbacks of a sampling method involving evaporation under nitrogen. Anal. & Bioanal. Chem., 406: 1541–1550. Finnemore, H., 1926. The Essential Oils, 1st ed. London: Ernest Benn. Firestein, S., 1992. Electrical signals in olfactory transduction. Curr. Opin. Neurobiol., 2: 444–448. Firestein, S., 2001. How the olfactory system makes sense of scents. Nature, 413: 211–218. François, I., A. de Villiers, and P. Sandra, 2006. Considerations on the possibilities and limitations of comprehensive normal phase-reversed phase liquid chromatography (NPLC × RPLC). J. Sep. Sci., 29: 492–498. Friedrich J. E., T. E. Acree, and E. H. Lavin, 2001. Selecting standards for gas chromatography-olfactometry. In Gas Chromatography-Olfactometry: The State of the Art, J. V. Leland et al. (eds.). Washington, D.C.: American Chemical Society, 2001, chap. 13. Fuller, G. H., R. Seltenkamp, and G. A. Tisserand, 1964. The gas chromatograph with human sensor: Perfumer model. Ann. NY Acad. Sci. 116: 711–724. Gaffney, B. M., M. Havekotte, B. Jacobs, L. Costa, 1996. Charm analysis of two Citrus sinensis peel oil volatiles. Perf. Flav., 21: 1.

224

Handbook of Essential Oils

Garland, S. M., R. C. Menary, and N. W. Davies, 1999. Dissipation of propiconazole and tebuconazole in peppermint crops (Mentha piperita (Labiatae)) and their residues in distilled oils. J. Agric. Food Chem., 47: 294–298. Gildemeister, E., and F. R. Hoffman, 1950. Die Ätherischen Öle, Volume One, 3rd ed. Leipzig: Verlag Von Schimmel & Co. Giungato, P., A. Di Gilio, J. Palmisani et al. 2018. Synergistic approaches for odor active compounds monitoring and identification: State of the art, integration, limits and potentialities of analytical and sensorial techniques. Trends Anal. Chem., 107: 116–129. Grob, K., 1987. On-line coupled HPLC-HRGC. In Proc. 8th International Symposium on Capillary Chromatography, Riva del Garda, Italy. Grosch, W., 1994. Determination of potent odourants in foods by aroma extract dilution analysis (AEDA) and calculation of odour activity values (OAVs). Flav. Fragr. J., 9: 147–158. Guenther, E., 1972. The Essential Oils - Volume One: History-Origin in Plants Production-Analysis, Reprint of 1st ed. (1948). Florida: Krieger Publishing Company. Gu, X., Y. Zhao, K. Li et al., 2016. Differentiation of volatile aromatic isomers and structural elucidation of volatile compounds in essential oils by combination of HPLC separation and crystalline sponge method. J. Chromatogr. A, 1474: 130–137. Hahn-Deinstrop, E., 2000. Applied Thin Layer Chromatography: Best Practice and Avoidance of Mistakes, 2nd ed. Weinheim: Wiley-VCH, chap. 1. Hawrył, H. M., M. A. Hawrył, R. Świeboda, K. Stępak, M. Niemiec, and M. Waksmundzka-Hajnos, 2017. Characterization of mint essential oils by high-performance liquid chromatography. Anal. Lett., 50: 2078–2089. Hong, J. H., N. Khan, N. Jamila et al., 2017. Determination of Volatile Flavour Profiles of Citrus spp. Fruits by SDE-GC–MS and Enantiomeric composition of chiral compounds by MDGC–MS. Phytochem. Anal., 28: 392–403. ISO 3761–1997, 1997. International Organization for Standardization, Geneva, http://www.iso.org/iso/iso_ catalogue/catalogue_tc/catalogue_tc_browse.htm?commid=48956 (accessed December 15, 2007). James, A. T., and A. J. P. Martin, 1952. Gas-liquid partition chromatography: The separation and microestimation of volatile fatty acids from formic acid to dodecanoic acid. Biochem. J., 50: 679–690. Japanese Pharmacopoeia JP XV. 2007, 15th ed., Yakuji Nippo, Ltd., Tokyo, 2007. Jirovetz, L., G. Buchbauer, M. B. Ngassoum, and M. Geissler, 2002. Aroma compound analysis of Piper nigrum and Piper guineense essential oils from Cameroon using solid-phase microextraction-gas chromatography, solid-phase microextraction-gas chromatography-mass spectrometry and olfactometry. J. Chromatogr. A, 976: 265–275. Jirovetz, L., and M. B. Ngassoum, 1999. Olfactory evaluation and CG/MS analysis of the essential oil of leaves and flowers of Hyptis pectinata (L.) Poit. From Cameroon. SoFW J., 125: 35. Juchelka, D., T. Beck, U. Hener, F. Dettmar, and A. Mosandl, 1998. Multidimensional gas chromatography coupled on-line with isotope ratio mass spectrometry (MDGC-IRMS): Progress in the analytical authentication of genuine flavor components. J. High Resolut. Chromatogr., 21: 145–151. Khajejenori, M., A. H. Asl, and M. H. Eikani, 2015. Optimization of subcritical water extraction of pimpinella anisum Seeds. Journal of Essential Oils Bearing Plants, 18: 1310–1320. König, W. A., 1991. Gas Chromatographic Enantiomer Separation with Modified Cyclodextrins, 1st ed. Heidelberg: Hüthig. König, W. A., 1998. Enantioselective capillary gas chromatography in the investigation of stereochemical correlations of terpenoids. Chirality, 10: 499–504. Korytár, P., H. G. Janssen, E. Matisová, and U. A. T. Brinkman, 2002. Practical fast gas chromatography: Methods, instrumentation and applications, TrAC, 21, 558–572. Kováts, E., 1958. Gas-chromatographische charakterisierung organischer verbindungen. teil 1: Retentionsindices aliphatischer halogenide, alkohole, aldehyde und ketone, Helv. Chim. Acta, 41, 1915–1932. Lee, M. L., F. J. Yang, and K. D. Bartle, 1984. Open Tubular Column Gas Chromatography, 1st ed. New York: Wiley & Sons, chap. 2. Lehmann, D., A. Dietrich, U. Hener, and A. Mosandl, 1995. Stereoisomeric flavour compounds LXX, 1-p-menthene-8-thiol: Separation and sensory evaluation of the enantiomers by enantioselective gas chromatography/olfactometry. Phytochem. Anal., 6: 255–257. Liberti, A., and G. Conti, 1956. Possibilità di applicazione della cromatografia in fase gassosa allo studio della essenza. In Proc. 1° Convegno Internazionale di Studi e Ricerche sulle Essenze, Reggio Calabria, Italy. Lin, J., and R. L. Rouseff, 2001. Characterization of aroma-impact compounds in cold-pressed grapefruit oil using time–intensity GC–olfactometry and GC–MS. Flav. Fragr. J., 16: 457–463.

Analysis of Essential Oils

225

Linssen, J. P. H., J. L. G. M. Janssens, J. P. Roozen, M. A. Posthumus, 1993. Combined gas chromatography and sniffing port analysis of volatile compounds of mineral water packed in polyethylene laminated packages. Food Chem., 46: 367–371. Liu, Z., and J. B. Phillips, 1991. Comprehensive two-dimensional gas chromatography using an on-column thermal modulator interface. J. Chromatogr. Sci., 29: 227–231. Lo Presti, M., D. Sciarrone, M. L. Crupi et al., 2008. Evaluation of the volatile and chiral composition in Pistacia lentiscus L. essential oil. Flav. Fragr. J., 23: 249–257. Maggi, F., F. Papa, C. Giuliani et al. 2015. Essential oil chemotypification secretory structures of the neglected vegetable Smyrnium olusatrum L. (Apiaceae) growing in central Italy. Flavour Fragr. J., 30: 139–159. Malnic, B., J. Hirono, T. Sato, and L. B. Buck, 1999. Combinatorial receptor codes for odors. Cell, 96: 713–723. Marriott, P. J., S.-T. Chin, B. Maikhunthod, H. G. Schmarr, and S. Bieri, 2012. Multidimensional gas chromatography. TrAC, 34: 1–21. Marriott, P. J., R. Shellie, and C. Cornwall, 2001. Gas chromatographic technologies for the analysis of essential oils. J. Chromatogr. A., 936: 1–22. Martindale: The Complete Drug Reference, 2007. 35th ed., Sweetman, S., Ed., Pharmaceutical Press, London. Masson, J., E. Liberto, J. C. Beolor, H. Brevard, C. Bicchi, and P. Rubiolo, 2016. Oxygenated heterocyclic compounds to differentiate Citrus spp. essential oils through metabolomics strategies. Food Chem., 206: 223–233. McDaniel, M. R., R. Miranda-Lopez, B. T. Watson, N. J. Micheals, and L. M. Libbey, 1992. Pinot noir aroma: A sensory/gas chromatographic approach. In Flavors and Off-Flavors (Developments in Food Science Vol. 24), G. Charalambous (ed.). Amsterdam: Elsevier Science Publishers, pp. 23–26. McGorrin, R. J., 2002. Character impact compounds: Flavors and off-flavors in foods. In Flavor, Fragrance, and Odor Analysis, R. Marsili (ed.), 1st ed. New York: Marcel Dekker, chap. 14. Mc Hale, D., and J. B. Sheridan, 1988. Detection of adulteration of cold-pressed lemon oil. Flav. Fragr. J., 3: 127–133. Mc Hale, D., and J. B. Sheridan, 1989. The oxygen heterocyclic compounds of citrus peel oils. J. Essent. Oil Res., 1: 139–149. Mehl F., G. Marti, J. Boccard et al., 2014. Differentiation of lemon essential oil based on volatile and nonvolatile fractions with various analytical techniques: A metabolomics approach. Food Chem., 143: 325–335. Mehl F., G. Marti, P. Merle et al., 2015. Integrating metabolomics data from multiple analytical platforms for a comprehensive characterization of lemon essential oils. Flav. Fragr. J., 30: 131–138. Minh, T. N., T. D. Xuan, T. M. Van, Y. Andriana, T. D. Khanh, and H. D. Tran, 2019. Phytochemical analysis of potential biological activities of essential oil from rice leaf. Molecules, 24: 546–558. Mondello, L., 2004. Determinazione della Composizione e Individuazione delle Adulterazioni degli Olii Essenziali mediante Ultrafast-GC. In Qualità e sicurezza degli Alimenti. Milan: Morgan Edizioni Scientifiche, 113–116. Mondello, L., K. D. Bartle, G. Dugo, and P. Dugo, 1994a. Automated HPLC-HRGC: A powerful method for essential oil analysis. Part III. Aliphatic and terpene aldehydes of orange oil. J. High Resol. Chromatogr., 17: 312–314. Mondello, L., A. Casilli, P. Q. Tanchida, L. Cicero, P. Dugo, and G. Dugo. 2003. Comparison of fast and conventional GC analysis for citrus essential oils, J. Agric. Food Chem., 51: 5602–5606. Mondello, L., A. Casilli, P. Q. Tranchida, D. Sciarrone, P. Dugo, and G. Dugo, 2008. Analysis of allergens in fragrances by using multiple heart-cut multidimensional gas chromatography-mass spectrometry. LCGC Europe, 21: 130–137. Mondello, L., R. Costa, D. Sciarrone, and G. Dugo, 2010. The chiral compound of citrus oils. In Citrus Oils: Composition, Advanced Analytical Techniques, Contaminants, and Biological Activity, Giovanni Dugo, and Luigi Mondello (eds.). Taylor & Francis Group, LLC, CRC Press. Mondello, L., P. Dugo, K. D. Bartle, G. Dugo, and A. Cotroneo, 1995. Automated HPLC-HRGC: A powerful method for essential oils analysis. Part V. Identification of terpene hydrocarbons of bergamot, lemon, mandarin, sweet orange, bitter orange, grapefruit, clementine and Mexican lime oils by coupled HPLCHRGC-MS(ITD). Flav. Fragr. J., 10: 33–42. Mondello, L., P. Dugo, K. D. Bartle, B. Frere, and G. Dugo, 1994b. On-line high performance liquid chromatography coupled with high resolution gas chromatography and mass spectrometry (HPLC-HRGC-MS) for the analysis of complex mixtures containing highly volatile compounds. Chromatog., 39: 529–538. Mondello, L., P. Dugo, G. Dugo and K. D. Bartle, 1996. On-line HPLC-HRGC-MS for the analysis of natural complex mixtures. J. Chromatogr. Sci., 34: 174–181. Mondello, L., R. Shellie, A. Casilli, P. Q. Tranchida, P. Marriot, and G. Dugo, 2004. Ultra-fast essential oil characterization by capillary GC on a 50 µm ID column, J. Sep. Sci., 27: 699–702.

226

Handbook of Essential Oils

Nishimura, O., 1995. Identification of the characteristic odorants in fresh rhizomes of ginger (Zingiber officinale Roscoe) using aroma extract dilution analysis and modified multidimensional gas chromatography-mass spectroscopy, J. Agric. Food Chem., 43: 2941–2945. Nitz, S., B. Weinreich, and F. Drawert, 1992. Multidimensional gas chromatography-isotope ratio massspectrometry (MDGC-IRMS) A. System description and technical requirements. J. High Resol. Chromatogr., 15: 387–391. Nowotny, H. P., D. Schmalzing, D. Wistuba, and V. Schurig, 1989. Extending the scope of enantiomer separation on diluted methylated β-cyclodextrin derivatives by high-resolution gas chromatography. J. High Res. Chromatogr., 12: 383–393. Ochiai, N., and K. J. Sasamoto, 2011. Selectable one-dimensional or two-dimensional gas chromatographyolfactometry/mass spectrometry with preparative fraction collection for analysis of ultra-trace amounts of odor compounds. J. Chromatogr. A., 1218: 3180–3185. Padhan, D., S. Pattnaik, A. K. Behera, 2017. Growth-arresting activity of acmella essential oil and its isolated component D-limonene (1,8 P-Mentha diene) against Trichophyton rubrum (microbial type culture collection 296). Pharmacognosy Magazine, 13: 555–560. Panda, H., 2003. Essential Oils Handbook, 1st ed. New Delhi: National Institute of Industrial Research, chap. 110. Pantò, S., D. Sciarrone, M. Maimone et al., 2015. Performance evaluation of a versatile multidimensional chromatographic preparative system based on three-dimensional gas chromatography and liquid chromatography–two-dimensional gas chromatography for the collection of volatile constituents. J. Chromatogr. A, 1417: 96–103. Parry, E. J., 1908. The Chemistry of Essential Oils and Artificial Perfumes, 2nd ed. London: Scott, Greenwood & Son. Piochon-Gauthier, M., J. Legault, M. Sylvestre, and A. Pichette, 2014. The essential oil of Populus balsamifera buds: Its chemical composition and cytotoxic activity. Natural Prod. Comm., 9: 257–260. Píry, J., and A. Príbela, 1994. Application of high-performance liquid chromatography to the analysis of the complex volatile mixture of blackcurrant buds (Ribes nigrum L.). J. Chromatogr. A, 665: 105–109. Pollien P., A. Ott, F. Montigon, M. Baumgartner, R. Muñoz-Box, and A. Chaintreau, 1997. Hyphenated headspace-gas chromatography-sniffing technique: Screening of impact odorants and quantitative aromagram comparisons. J. Agric. Food Chem., 45: 2630–2637. Poole, C. F., 2003. The Essence of Chromatography, 1st ed. Amsterdam: Elsevier. Porel, A., Y. Sanyal, and A. Kundu, 2014. Simultaneous HPLC determination of 22 components of essential oils; method robustness with experimental design. Indian J. Pharm. Sci, 76: 19–30. Pragadheesh, V. S., A. Yadav, and C. S. Chanotiya, 2015. Role of substituents in cyclodextrin derivatives for enantioselective gas chromatographic separation of chiral terpenoids in the essential oils of Mentha spicata. J. Chromatogr. B, 1002: 30–41. Proot, M., P. Sandra, and E. Geeraert, 1986. Resolution of triglycerides in capillary SFC as a function of column temperature, J. High Res. Chromatogr. Commun., 9: 189–192. Purcaro, G., C. Cordero, E. Liberto, C. Bicchi, and L. S. Conte, 2014. Toward a definition of blueprint of virgin olive oil by comprehensive two-dimensional gas chromatography. J. Chromatogr. A, 1334: 101–111. Purcaro, G., M. Zoccali, and P. Q. Tranchida et al., 2013. Comparison of two different multidimensional liquidgas chromatography interfaces for determination of mineral oil saturated hydrocarbons in foodstuffs. Anal. Bioanal. Chem., 405: 1077–1084. Quassinti, L., F. Maggi, L. Barboni et al., 2014. Wild celery (Smyrnium olusatrum L.) oil and isofuranodiene induce apoptosis in human colon carcinoma cells. Fitoterapia, 97: 133–141. Ricci, M. P., D. A. Merritt, K. H. Freeman, J. M. Hayes, 1994. Acquisition and processing of data for isotoperatio-monitoring mass spectrometry. Org. Geochem., 21: 561–571. Richardson, A., 1999. Measurement of fragrance perception. In The Chemistry of Fragrances, D. H. Pybus and C. S. Sell (eds.). Cambridge: Royal Society of Chemistry, chap. 8. Rowe, D. J., 2005. Introduction. In Chemistry and Technology of Flavors and Fragrances, D. J. Rowe (ed.). Oxford: The Blackwell Publishing, chap. 1. Rubiolo, P., F. Belliardo, C. Cordero, E. Liberto, B. Sgorbini, and C. Bicchi, 2006. Headspace-solid-phase microextraction fast GC in combination with principal component analysis as a tool to classify different chemotypes of chamomile flower-heads (Matricaria recutita L.). Phytochem. Anal., 17: 217–225. Rubiolo, P., E. Liberto, C. Cagliero et  al., 2007. Linear retention indices in enantioselective GC-Mass Spectrometry (Es-GC-MS) as a tool to identify enantiomers in flavour and fragrance fields. In 38th International Symposium on Essential Oils, Graz. Russo, M., I. Bonaccorsi, R. Costa, R. Trozzi, P., Dugo, and L. Mondello, 2015. Reduced time HPLC analyses for fast quality control of citrus essential oil. J. Essent. Oil Res., 27: 307–315.

Analysis of Essential Oils

227

Russo, M., F. Rigano, A. Arigò et al., 2016. Rapid isolation, reliable characterization, and water solubility improvement of polymethoxyflavones from cold-pressed mandarin essential oil. J. Sep. Sci. 2016, 39, 2018–2027. Russo, M., G. Torre, C. Carnovale, I. Bonaccorsi, L. Mondello, and P. Dugo, 2012. A new HPLC method developed for the analysis of oxygen heterocyclic compounds in Citrus essential oils. J. Essent. Oil Res., 24, 119–129. Sale, J. W., 1953. Analysis of lemon oils. J. Assoc. Offic. Agr. Chemists, 36: 112–119. Schipilliti, L., I. Bonaccorsi, D. Sciarrone, L. Dugo, L. Mondello, and G. Dugo, 2012. Determination of petitgrain oils landmark parameters by using gas chromatography–combustion–isotope ratio mass spectrometry and enantioselective multidimensional gas chromatography. Anal. Bioanal. Chem., 405: 679–690. Schipilliti, L., G. Dugo, L. Santi, P. Dugo, and L. Mondello, 2011. Authentication of bergamot essential oil by gas chromatography-combustion-isotope ratio mass spectrometer (GC-C-IRMS). J. Essent. Oil Res., 23(2): 60–71. Schurig, V., and H. P. Nowotny, 1988. Separation of enantiomers on diluted permetylated β-cyclodextrin by high-resolution gas chromatography. J. Chromatogr., 441: 155–163. Sciarrone, D., R. Costa, C. Ragonese et  al., 2011. Application of a multidimensional GC system with simultaneous mass spectrometric and FID detection to the analysis of sandalwood oil. J. Chromatogr. A, 1218: 137–142. Sciarrone, D., D. Giuffrida, A. Rotondo et  al., 2017. Quali-quantitative characterization of the volatile constituents in Cordia verbenacea D.C. essential oil exploiting advanced chromatographic approaches and nuclear magnetic resonance analysis. J. Chromatogr. A. 1524: 246–253. Sciarrone, D., S. Pantò, F. Cacciola, R. Costa, P. Dugo, and L. Mondello, 2015a. Advanced preparative techniques for the collection of pure components from essential oils. Nat. Volatiles & Essent. Oils, 2: 1–15. Sciarrone, D., S. Pantò, P. Donato, and L. Mondello, 2016. Improving the productivity of a multidimensional chromatographic preparative system by collecting pure chemicals after each chromatographic dimension. J. Chromatogr. A, 1475: 80–85. Sciarrone, D., S. Pantò, C. Ragonese, P. Dugo, and L. Mondello, 2015b. Evolution and status of preparative gas chromatography as a green sample-preparation technique. TrAC, 71: 65–73. Sciarrone, D., S. Pantò, C. Ragonese, P. Q. Tranchida, P. Dugo, and L. Mondello, 2012. Increasing the isolated quantities and purities of volatile compounds by using a triple deans-switch multi-dimensional preparative gas chromatographic system with an apolar-wax-ionic liquid stationary-phase combination. Anal. Chem., 84: 7092–7098. Sciarrone, D., S. Pantò, A. Rotondo et al., 2013. Rapid collection and identification of a novel component from Clausena lansium Skeels leaves by means of three-dimensional preparative gas chromatography and nuclear magnetic resonance/infrared/mass spectrometric analysis. Anal. Chim. Acta, 785: 119–125. Sciarrone, D., S. Pantò, P. Q. Tranchida, P. Dugo, and L. Mondello, 2014. Rapid isolation of high solute amounts using an online four-dimensional preparative system: Normal phase-liquid chromatography coupled to methyl siloxane–ionic liquid–wax phase gas chromatography. Anal. Chem., 86: 4295–4301. Sciarrone, D., C. Ragonese, C. Carnovale et al., 2010a. Evaluation of tea tree oil quality and ascaridole: A deep study by means of chiral and multi heart-cuts multidimensional gas chromatography system coupled to MS Detection. J. Chromatogr. A, 1217: 6422–6427. Sciarrone, D., A. Schepis, G. De Grazia et al., 2019. Collection and identification of an unknown component from Eugenia uniflora essential oil exploiting a multidimensional preparative three-GC system employing apolar, mid-polar and ionic liquid stationary phases. Faraday Discuss. DOI:10.1039/C8FD00234G. Sciarrone, D., L. Schipilliti, C. Ragonese et al., 2010b. Thorough evaluation of the validity of conventional enantio-gas chromatography in the analysis of volatile chiral compounds in mandarin essential oil: A comparative investigation with multidimensional gas chromatography. J. Chromatogr. A, 1217: 1101–1105. Schomburg, G., H. Husmann, E. Hübinger, and W. A. König, 1984. Multidimensional capillary gas chromatography-enantiomeric separation of selected cuts using a chiral second column. J. High Resolut. Chromatogr., 7: 404–410. Scott, R. P. W., 2001. Gas Chromatography, Chrom Ed. Series, http://www.chromatography-online.org/ (accessed December 15, 2007). Semmler, F. W., 1906–1907. Die Ätherischen Öle, Vol. I to IV, 3rd ed. Leipzig: Verlag von Veit. Shellie, R., and P. J. Marriott, 2002. Comprehensive two-dimensional gas chromatography with fast enantioseparation. Anal. Chem., 74: 5426–5430. Shellie, R., P. Marriott, and C. Cornwell, 2001. Application of comprehensive two-dimensional gas chromatography (GC × GC) to the enantioselective analysis of essential oils. J. Sep. Sci., 24: 823–830.

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Sherma, J., 2000. Thin-layer chromatography in food and agricultural analysis, J. Chromatogr. A, 880: 129–147. Simões, C. M. O., and V. Spitzer, 1999. Óleos voláteis. In Farmacognosia: Da planta ao medicamento, C.M.O. Simões et al. (ed.), 1999. Florianópolis: Editora da UFSC and Editora da Universidade/UFRGS, chap. 18. Song, H. S., M. Sawamura, T. Ito, A. Ido, and H. Ukeda, 2000a. Quantitative determination and characteristic flavour of daidai (Citrus aurantium L. var. cyathifera Y. Tanaka) peel oil. Flav. Fragr. J., 15: 323–328. Song, H. S., M. Sawamura, T. Ito, K. Kawashimo, and H. Ukeda, 2000b. Quantitative determination and characteristic flavour of Citrus junos (yuzu) peel oil. Flav. Fragr. J., 15: 245–250. Stillman, R. C., and R. M. Reed, 1932. Perfumery Essential Oil Record, 23: 278. Sybilska, D., and T. Koscielski, 1983. β-cyclodextrin as a selective agent for the separation of o-, m- and p-xylene and ethylbenzene mixtures in gas-liquid chromatography. J. Chromatogr. A., 261: 357–362. TC 57 Essential Oils, International Organization for Standardization, Geneva, http://www.iso.org/iso/iso_ catalogue/catalogue_tc/catalogue_tc_browse.htm?commid=48956, (accessed December 15, 2007). The International Pharmacopoeia (IntPh), 2007. Pharmacopoeia, 4th ed. Geneva: World Health Organization Press. The United States Pharmacopoeia USP/NF 2008, 2007. Maryland: United States Pharmacopoeia Convention Inc. Tine, Y., A. Diop, W. Diatta et al., 2017. Chemical diversity and antimicrobial activity of volatile compounds from Zanthoxylum zanthoxyloides Lam. According to compounds classes, plant organs and Senegalese sample locations. Chem. Biodivers., 14. Todd, J. F. J., 1995. Recommendations for nomenclature and symbolism for mass spectroscopy. Int. J. Mass Spectrom. Ion Process, 142: 211–240. Tranchida, P. Q., A. Casilli, G. Dugo, L. Mondello, and P. Dugo. 2005. Fast gas chromatographic analysis with a 0.05 mm ID micro-bore capillary column, G.I.T. Lab. J., 9: 22. Tranchida, P. Q., Franchina, F. A., Dugo, P., and Mondello, L. 2014. Use of greatly-reduced gas flows in flowmodulated comprehensive two-dimensional gas chromatography-mass spectrometry. J. Chromatogr. A, 1359: 271–276. Tranchida, P. Q., F. A. Franchina, M. Zoccali et  al., 2013a. Untargeted and targeted comprehensive twodimensional GC analysis using a novel unified high-speed triple quadrupole mass spectrometer. J. Chromatogr. A, 1278: 153–159. Tranchida, P. Q., D. Sciarrone, P. Dugo, and L. Mondello, 2012. Heart-cutting multidimensional gas chromatography: Recent evolution, applications, and future prospects. Anal. Chim. Acta, 716: 66–75. Tranchida, P. Q., M. Zoccali, I. Bonaccorsi, P. Dugo, L. Mondello, and G. Dugo, 2013b. The off-line combination of high performance liquid chromatography and comprehensive two-dimensional gas chromatography– mass spectrometry: A powerful approach for highly detailed essential oil analysis. J. Chromatogr. A, 1305: 276–284. Ullrich, F., and W. Grosch, 1987. Identification of the most intense volatile flavour compounds formed during autoxidation of linoleic acid, Z. Lebensm. Unters. Forsch., 184: 277–282. van den Dool, H., and P. D. Kratz, 1963. A generalization of the retention index system including linear temperature programmed gas-liquid chromatography. J. Chromatogr. A, 11, 463–471. van Ruth, S. M., 2001. Methods for gas chromatography-olfactometry: A review. Biomolec. Eng., 17: 121–128. Wagner, H., S. Bladt, and V. Rickl, 2003. Plant Drug Analysis: A Thin Layer Chromatography Atlas, 2nd ed. Heidelberg: Springer Verlag, p. 1. Wong, Y. F., R. N. West, S. T. Chin, and P. J. Marriot, 2015. Evaluation of fast enantioselective multidimensional gas chromatography methods for monoterpenic compounds: Authenticity control of Australian tea tree oil. J. Chromatogr. A, 1406: 307–315. Wong Y. F., D. Yan, R. A. Shellie, D. Sciarrone, and P. J. Marriot, 2019. Rapid plant volatiles screening using headspace SPME and person-portable gas chromatography–mass spectrometry, Chromatographia, 82: 297–305. Wright, D. W., 1997. Application of multidimensional gas chromatography techniques to aroma analysis. In Techniques for Analyzing Food Aroma (Food Science and Technology), R. Marsili (ed.), 1st ed. New York: Marcel Dekker, chap. 5.

8

Use of Linear Retention Indices in GC-MS Libraries for Essential Oil Analysis Emanuela Trovato, Giuseppe Micalizzi, Paola Dugo, Margita Utczás, and Luigi Mondello

CONTENTS 8.1 Introduction........................................................................................................................... 229 8.2 Retention Index Theories....................................................................................................... 230 8.3 Linear Retention Indices Present in the Databases............................................................... 235 8.4 Accuracy of Retention Indices............................................................................................... 235 8.5 Retention Index Compilations...............................................................................................240 8.6 Research Software................................................................................................................. 241 8.7 Application of Retention Indices in GC-FID and GC-MS Essential Oil Analyses.............. 243 8.8 Gas Chromatographic Enantiomer Characterization Supported by Retention Indices.........248 8.9 Retention Indices Applied to Multidimensional Gas Chromatographic Analysis................248 8.10 Concluding Remarks............................................................................................................. 249 References....................................................................................................................................... 249

8.1 INTRODUCTION The continuous fundamental and difficult challenge of the modern science of separation consists of providing adequate chromatographic techniques that allow resolving mixtures of compounds into less complex mixtures or ultimately into pure components with rapid and effective separations, improving at the same time the resolving power, and allowing highly selective or sensitive detections. Essential oils are very complex mixtures consisting of monoterpene and sesquiterpene hydrocarbons, their oxygenated derivatives, and aliphatic oxygenated compounds. The technique most used for essential oil analysis and characterization is GC-MS. Many terpenes have identical mass spectra due to similarities in the initial molecule, or in the fragmentation patterns and rearrangements after ionization, and this raises difficulties in the GC-MS peak identification of these samples. The combining of retention time information with mass spectra results may support the identification by MS library search. In a GC separation, each compound is characterized by a different retention behavior on a specific column, expressed by absolute retention time, retention factor, and the relative retention time. The absolute retention time (tR) varies depending on mobile phase flow, temperature program, column phase ratio and length, and also between consecutive runs on the same column and under the same analytical conditions, and for these reasons, it is not so useful for identification purposes. The need to express gas chromatographic retention data in a standardized system has long been recognized and an enormous effort has been devoted to this purpose. James and Martin (James and Martin, 1952), in their first publication on gas–liquid chromatography in 1952 mentioned that the retention volume of a pure substance on a certain gas chromatographic column is a characteristic value, which could be used for the identification of sample components. 229

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In 1955, Littlewood et al. (1955) attempted to make retention data independent of temperature, stationary phase film thickness, carrier gas flow rate, and pressure drop across the column, introducing the specific retention volume; but this parameter was not practically applied. Instead, from the many proposals, the use of relative retention data expressed with respect to the retention of a standard substance has found consensus. Since gas chromatographic separations are performed over a wide range of temperatures and on stationary phases of different polarities, no single substance would fulfill the role of a universal standard. In general, the most thoroughly studied, diffused, and accepted retention index calculation methods are calculated applying the equation developed by Kováts in 1958 (Kováts, 1958) for isothermal analysis conditions and the equation propounded by van den Dool and Kratz in 1963, (van den Dool and Kratz, 1963) in the case of temperature programming analysis conditions, while values derived from the former equation are usually called retention index (I) or Kováts index (KI). Values calculated using the latter approach are commonly denominated in literature as retention index (I), linear retention index (LRI) or programmed-temperature retention index (PTRI). This chapter provides an overview of the theoretical aspects and applications of retention indices to gas chromatographic analysis of essential oils. Gas chromatographic retention parameters and retention index calculations are described; furthermore, a critical viewpoint is discussed with respect to the indiscriminate use of retention indices from different sources.

8.2  RETENTION INDEX THEORIES As is well known and reported by Poole, the retention of analytes in gas chromatographic capillary columns results from the differential distribution (partition) of the solutes between the stationary liquid and the mobile gas phases (Poole, 2003). A compound’s retention behavior on a specific column is characterized by three parameters: retention time (tR), retention factor (k), and relative retention (r). Retention time (tR) is expressed as the time that the solute spends in the mobile phase, that is the unretained peak time (tM ), summed to the adjusted retention time (t′R), which corresponds to the time the solute spends distributed into the stationary phase (see Equation 8.1):

t R = t M + t R′

(8.1)

The magnitude of retention depends on the partition coefficient, or distribution constant (K), which is defined as the ratio of the equilibrium concentrations of a solute in the stationary (CS) and mobile phases (Cm) during partitioning in the column:

K D = C S /C m

(8.2)

Consequently, the greater is the K value for a sample component, the higher is its solubility and the longer its retention in the stationary phase. Generally, the affinity of a solute for the stationary phase depends on its vapor pressure and the activity coefficient of the solute in that phase. The column’s ability to differentiate two solutes, eluting them with different retention times depends from differences between those two properties. Retention factor (k) quantifies the ratio of the time spent in the stationary phase and that spent in the mobile phase, as expressed by the following equation:

k = t R′ /t M

(8.3)

where t′R is the adjusted retention time and tM the unretained peak time. Relative retention (r) expresses the degree of separation between two peaks, a standard (st) and the solute of interest, not necessarily in adjacent positions. Relative retention can be expressed using

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standard solute retention factors or adjusted retention times. The latter are used in Equation 8.4, where i refers to an individual solute. The relative retention of solutes eluting after the standard will be >1, while that of compounds eluting prior to the standard 98% of the composition of oil of Mentha piperita (peppermint oil), greater than 95% of the oil is accounted for by three chemical groups: (1) terpene aliphatic and aromatic hydrocarbons; (2) terpene alicyclic secondary alcohols, ketones, and related esters; and (3) terpene 2-isopropylidene–substituted cyclohexanone derivatives and related substances. The formation and members of a congeneric group are chosen based on a combination of structural features and known biochemical fate. Substances with a common carbon skeletal structure and functional

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O O Menthyl acetate

OH

OH Menthol

O

OH p-Menthane-3,8-diol

Menthone

FIGURE 9.1  Congeneric groups are formed by members sharing common structural and metabolic features, such as the group of 2-isopropylidene-substituted cyclohexanone derivatives and related substances.

groups that participate in common pathways of metabolism are assigned to the same congeneric group. For instance, menthyl acetate hydrolyzes prior to absorption to yield menthol, which is absorbed and is interconvertible with menthone in fluid compartments (e.g., the blood). Menthol is either conjugated with glucuronic acid and excreted in the urine or undergoes further hydroxylation mainly at C8 to yield a diol that is also excreted, either free or conjugated. Despite the fact that menthyl acetate is an ester, menthol is an alcohol, menthone a ketone, and 3,8-menthanediol a diol, they are structurally and metabolically related (Figure 9.1). Therefore, all are members of the same congeneric group. In the case of M. piperita, the three principal congeneric groups listed earlier have different metabolic options and possess different organ-specific toxic potential. The congeneric group of terpene aliphatic and aromatic hydrocarbons is represented mainly by limonene and myrcene. The second and most predominant congeneric group is the alicyclic secondary alcohols, ketones, and related esters that include d-menthol, menthone, isomenthone, and menthyl acetate. Although the third congeneric group contains alicyclic ketones similar in structure to menthone, it is metabolically quite different in that it contains an exocyclic isopropylidene substituent that undergoes hydroxylation principally at the C9 position, followed by ring closure and dehydration to yield a heteroaromatic furan ring of increased toxic potential. In the absence of a C4–C8 double bond, neither menthone nor isomenthone can participate in this intoxication pathway. Hence, they are assigned to a different congeneric group. The presence of a limited number of congeneric groups in an essential oil is critical to the organization of constituents and subsequent safety evaluation of the oil itself. Members of each congeneric group exhibit common structural features and participate in common pathways of pharmacokinetics and metabolism and exhibit similar toxicologic potential. Recent guidance on chemical grouping has been published (OECD, 2014). If the mass of the essential oil (>95%) can be adequately characterized chemically and constituents assigned to well-defined congeneric groups, the safety evaluation of the essential oil can be reduced to (1) a safety evaluation of each of the congeneric groups comprising the essential oil and (2) a sum of the parts evaluation of the all congeneric groups to account for any chemical or biological interactions between congeneric groups in the essential oil under conditions of intended use. Validation of such an approach lies in the stepwise comparison of the dose and toxic effects for each key congeneric group with similar equivalent doses and toxic effects exhibited by the entire essential oil. Using such an approach, the scientifically independent but industry-sponsored Expert Panel of the U.S. Flavor and Extract Manufacturers Association (FEMA) has conducted safety evaluations on a number of natural flavoring complexes, many of which are essential oils and extracts, under the auspices of the GRAS concept (Smith et al., 2003, 2004). Without question, other approaches have been developed (Meek et al., 2011).

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Potential interactions between congeneric groups can, to some extent, be analyzed by an in-depth comparison of the biochemical and toxicologic properties of different congeneric groups in the essential oil. For some representative essential oils that have been the subject of toxicology studies, a comparison of data for the congeneric groups in the essential oil with data on the essential oil itself (congeneric groups together) is a basis for analyzing for the presence or absence of interactions. Therefore, the impact of interaction between congeneric groups is minimal if the levels of and endpoints for toxicity of congeneric groups (e.g., tertiary terpene alcohols) are similar to those of the essential oil (e.g., coriander oil). Since composition plays such a critical role in the evaluation, analytical identification requirements are also critical to the evaluation. Complete chemical characterization of the essential oil may be difficult or economically unfeasible based on the small volume of essential oil used as a flavor ingredient. In these few cases, mainly for low-volume essential oils, the unknown fraction may be appreciable and a large number of chemical constituents will not be identified. However, if the intake of the essential oil is low or significantly less than its intake from consumption of food (e.g., thyme) from which the essential oil is derived (e.g., thyme oil), there should be no significant concern for safety under conditions of intended use. For those cases in which chemical characterization of the essential oil is limited but the volume of intake is more significant, it may be necessary to perform additional analytical work to decrease the number of unidentified constituents or, in other cases, to perform selected toxicity studies on the essential oil itself. A principal goal of the safety evaluation of essential oils is that no congeneric groups that have significant human intakes should go unevaluated.

9.3.4 Chemical Assay Requirements and Chemical Description of Essential Oil The safety evaluation of an essential oil initially involves specifying the biological origin, physical and chemical properties, and any other relevant identifying characteristics. An essential oil produced under good manufacturing practices should be of an appropriate purity (quality), and chemical characterization should be complete enough to guarantee a sufficient basis for a thorough safety evaluation of the essential oil under conditions of intended use. Because the evaluation is based primarily on the actual chemical composition of the essential oil, full specifications used in a safety evaluation will necessarily include not only information on the origin of the essential oil (commercial botanical sources, geographical sources, plant parts used, degree of maturity, and methods of isolation) and physical properties (specific gravity, refractive index, optical rotation, solubility, etc.) but also chemical assays for a range of essential oils currently in commerce. 9.3.4.1  Intake of the Essential Oil Based on current analytical methodology, it is possible to identify literally hundreds of constituents in an essential oil and quantify the constituents to part per million levels. But is this necessary or desirable? From a practical point of view, the level of analysis for constituents should be directly related to the level of exposure to the essential oil. The requirements to identify and quantify constituents for use of 2,000,000 kg of peppermint oil annually should be far greater than that for use of 2000kg of coriander oil or 50 kg of myrrh oil annually. Also, there is a level at which exposure to each constituent is so low that there is no significant risk associated with intake of that substance. A conservative no significant risk level of 1.5 µg/day (0.0015 mg/day or 0.000025 mg/kg/day) has been adopted by regulatory authorities as a level at which the human cancer risk is below one in one million – this is commonly referred to as the “threshold of regulation” (FDA, 2005). Therefore, if consumption of an essential oil results in an intake of a constituent that is less than 1.5 µg/day, there should be no requirement to identify and quantify that constituent. Determining the level to which the constituents of an essential oil should be identified depends upon estimates of intake of food or flavor additives. These estimates are traditionally calculated using a volume-based or a menu–census approach. A volume-based approach assumes that the total annual volume of use of a substance reported by an industry is distributed over a portion of the population consuming that substance. A menu–census approach is based on the concentration of the substance (essential oil) added to each flavor, the amount of flavor added to each food category, the portion

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of food consumed daily, and the total of all exposures across all food types. Although the latter is quite accurate for food additives consumed at higher levels in a wide variety of food such as food emulsifiers, the former method provides an efficient and conservative approximation of intake, if a fraction of the total population is assumed to consume all of the substance. For the World Health Organization (WHO) and the U.S. FDA, intake is calculated using a method known as the per capita intake (PCI × 10) method, or alternatively known as the maximized surveyderived intake (MSDI) (Rulis et al., 1984; Woods and Doull, 1991). The MSDI method assumes that only 10% of the population consumes the total annual reported volume of use of a flavor ingredient. This approximation provides a practical and cost-effective approach to the estimation of intake for flavoring substances. The annual volumes of flavoring agents are relatively easy to obtain by industry-wide surveys, which can be performed on a regular basis to account for changes in food trends and flavor consumption. A recent poundage survey of U.S. flavor producers was collected in 2005 and published by FEMA in 2007 (Adams et al., 2007). Similar surveys were conducted in recent years in Europe (EFFA, 2005) and Japan (JFFMA, 2002). Calculation of intake using the MSDI method has been shown to result in conservative estimates of intake and thus is appropriate for safety evaluation. Over the last three decades, two comprehensive studies of flavor intake have been undertaken. One involved a detailed dietary analysis (DDA) of a panel of 12,000 consumers who recorded all foods that they consumed over a 14-day period, and the flavoring ingredients in each food were estimated by experience flavorists to estimate intake of each flavoring substance (Hall, 1976; Hall and Ford, 1999). The other study utilized a robust full stochastic model (FSM) to estimate intake of flavoring ingredients by typical consumers in the United Kingdom (Lambe et al., 2002). The results of the data-intensive DDA method and the modelbased FSM support the use of PCI data as a conservative estimate of intake. With regard to essential oils, the MSDI method provides overestimates of intake for oils that are widely distributed in food. The large annual volume of use reported for essential oils such as orange oil, lemon oil, and peppermint oil indicate widespread use in a large variety of foods resulting in consumption of these oils by significantly more than 10% of the population. Citrus flavor is pervasive in a multitude of foods and beverages. Therefore, for selected high-volume essential oils, a simple PCI rather than a per capita × 10 intake may be more appropriate. However, the intake of the congeneric groups and the group of unidentified constituents for these high-volume oils is still estimated by the MSDI method. An alternative approach to MSDI now employed in the evaluation of flavoring substances by the WHO/UN Food and Agricultural Organization Joint Expert Committee on Food Additives (JECFA) relies on a modification of the traditional menu–census approach. The JECFA single portion exposure technique (SPET) identifies the highest intake from a single food category and assigns it as the overall intake for a flavoring substance. This is calculated from the concentration of the flavoring substance within a food category multiplied by the daily portion size for that food category. At JECFA, the highest intake from either MSDI or SPET is now used to assign an intake to a flavoring substance. JECFA has concluded that both methods, MSDI and SPET, can provide complementary and useful information regarding the uses of flavoring substances. To date, SPET has not been used for the safety evaluation of natural complex mixtures at JECFA, as the flavoring agent evaluations at JECFA have focused thus far on chemically defined substances. 9.3.4.2  Analytical Limits on Constituent Identification As described earlier, the analytical requirements for detection and identification of the constituents of an essential oil are set by the intake of the oil and by the conservative assumption that constituents with intakes less than 1.5 µg/day will not need to be identified. For instance, if the annual volume of use of coriander oil in the United States is 10,000 kg, then the estimated daily PCI of the oil is



10, 000 kg/year ×109 µg/kg = 883 µg coriander oil/person/day 365 days/year ×31, 000, 000 persons

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Based on the intake of coriander oil (883 µg/day), any constituent present at greater than 0.17% would need to be chemically characterized and quantified:



1.5 µg/day ×100 = 0.17% 978 µg/day

For the vast majority of essential oils, meeting these characterization requirements does not require exotic analytical techniques and the identification of the constituents is of a routine nature. However, what would the requirements be for very high-volume essential oils, such as orange oil, cold pressed (567,000 kg), or peppermint oil (1,229,000 kg)? In these cases, a practical limit must be applied and can be justified based on the concept that the intake of these oils is widespread and far exceeds the 10% assumption of MSDI. Based on current analytical capabilities, 0.10% or 0.05% could be used as a reasonable limit of detection, with the lower level used for an essential oil that is known or suspected to contain constituents of higher toxic potential (e.g., methyl eugenol in basil). 9.3.4.3  Intake of Congeneric Groups Once the analytical limits for identification of constituents have been met, it is key to evaluate the intake of each congeneric group from consumption of the essential oil. A range of concentration of each congeneric group is determined from multiple analyses of different lots of the essential oil used in flavorings. The intake of each congeneric group is determined from mean concentrations (%) of constituents recorded for each congeneric group. For instance, for peppermint oil the alicyclic secondary alcohol/ketone/related ester group may contain (−)-menthol, (−)-menthone, (−)-menthyl acetate, and isomenthone in mean concentrations of 43.0%, 20.3%, 4.4%, and 0.40%, respectively, with the result that that congeneric group accounts for 68.1% of the oil. It should be emphasized that although members in a congeneric group may vary among the different lots of oil, the variation in concentration of congeneric groups in the oil is relatively small. Routinely, the daily PCI of the essential oil is derived from the annual volumes reported in industry surveys (NAS, 1965, 1970, 1975, 1982, 1987; Lucas et al., 1999; JFFMA, 2002; EFFA, 2005). If a conservative estimate of intake of the essential oil is made using a volume-based approach such that a defined group of constituents are set for each essential oil, target constituents can be monitored in an ongoing quality control program, and the composition of the essential oil can become one of the key specifications linking the product that is distributed in the marketplace to the chemically based safety evaluation. Limited specifications for the chemical composition of some essential oils to be used as food flavorings are currently listed in the Food Chemicals Codex (FCC, 2013). For instance, the chemical assay for cinnamon oil is given as “not less than 80%, by volume, as total aldehydes.” Any specification developed related to this safety evaluation procedure should be consistent with already published specifications including FCC and ISO standards. However, based on chemical analyses for the commercially available oil, the chemical specification or assay can and should be expanded to 1. Specify the mean of concentrations for congeneric groups with confidence limits that ­constitute a sufficient number of commercial lots constituting the vast majority of the oil. 2. Identify key constituents of intake >1.5 µg/day in these groups that can be used to e­ fficiently monitor the quality of the oil placed into commerce over time. 3. Provide information on trace constituents that may be of a safety concern. For example, given its most recent reported annual volume (1060 kg) (Harman et al., 2013), it is anticipated that a chemical specification for lemongrass oil would include (1) greater than 98.7% of the composition chemically identified; (2) not more than 92% aliphatic terpene primary alcohols, aldehydes, acids, and related esters, typically measured as citral; and (3) not more than 15%

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aliphatic terpene hydrocarbons, typically measured as myrcene. The principal goal of a chemical specification is to provide sufficient chemical characterization to ensure safety of the essential oil from use as a flavoring. From an industry standpoint, the specification should be sufficiently descriptive as to allow timely quality control monitoring for constituents that are responsible for the technical flavor function. These constituents should also be representative of the major congeneric group or groups in the essential oil. Also, monitored constituents should include those that may be of a safety concern at sufficiently high levels of intake of the essential oil (e.g., pulegone). The scope of a specification should be sufficient to ensure safety in use but not impose an unnecessary burden on industry to perform ongoing analyses for constituents unrelated to the safety or flavor of the essential oil.

9.4 SAFETY CONSIDERATIONS FOR ESSENTIAL OILS, CONSTITUENTS, AND CONGENERIC GROUPS 9.4.1  Essential Oils 9.4.1.1  Safety of Essential Oils: Relationship to Food The close relationship of natural flavor complexes to food itself has made it difficult to evaluate the safety and regulate the use of essential oils. In the United States, the Federal Food Drug and Cosmetic Act recognizes that a different, lower standard of safety must apply to naturally occurring substances in food than applies to the same ingredient intentionally added to food. For a substance occurring naturally in food, the act applies a realistic standard that the substance must “… not ordinarily render it [the food] injurious to health” (21 CFR 172.30). For added substances, a much higher standard applies. The food is considered to be adulterated if the added substance “… may render it [the food] injurious to health” (21 CFR 172.20). Essential oils used as flavoring substances occupy an intermediate position in that they are composed of naturally occurring substances, many of which are intentionally added to food as individual chemical substances. Because they are considered neither a direct food additive nor a food itself, no current standard can be easily applied to the safety evaluation of essential oils. The evaluation of the safety of essential oils that have a documented history of use in foods starts with the presumption that they are safe based on their long history of use over a wide range of human exposures without known adverse effects. With a high degree of confidence, one may presume that essential oils derived from food are likely to be safe. Annual surveys of the use of flavoring substances in the United States (NAS, 1965, 1970, 1975, 1981, 1987; Lucas et al., 1999; Harman et al., 2013; 21 CFR 172.510) in part, document the history of use of many essential oils. Conversely, confidence in the presumption of safety decreases for natural complexes that exhibit a significant change in the pattern of use or when novel natural complexes with unique flavor properties enter the food supply. Recent consumer trends that have changed the typical consumer diet have also changed the exposure levels to essential oils in a variety of ways. As one example, changes in the use of cinnamon oil in low-fat cinnamon pastries would alter intake for a specialized population of eaters. Also, increased international trade has coupled with a reduction in cultural cuisine barriers, leading to the introduction of novel plants and plant extracts from previously remote geographical locations. Osmanthus absolute (FEMA No. 3750) and Jambu oleoresin (FEMA No. 3783) are examples of natural complexes recently used as flavoring substances that are derived from plants not indigenous to the United States and not commonly consumed as part of a Western diet. Furthermore, the consumption of some essential oils may not occur solely from intake as flavoring substances; rather, they may be regularly consumed as dietary supplements with advertised functional benefits. These impacts have brought renewed interest in the safety evaluation of essential oils. Although the safety evaluation of essential oil must still rely heavily on knowledge of the history of use, a flexible science-based approach would allow for rigorous safety evaluation of different uses for the same essential oil.

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9.4.2 Safety of Constituents and Congeneric Groups in Essential Oils It is well established that when consumed in high quantities, some plants do indeed exhibit toxicity. Historically, humans have used plants as poisons (e.g., hemlock), and many of the intended medicinal uses of plants (pennyroyal oil as an abortifacient) have produced undesirable toxic side effects. High levels of exposure to selected constituents in the plant or essential oil (i.e., pulegone in pennyroyal oil) have been associated with the observed toxicity. However, with regard to flavor use, experience through long-term use and the predominant self-limiting impact of flavorings on our senses have restricted the amount of a plant or plant part that we use in or on food. Extensive scientific data on the most commonly occurring major constituents in essential oils have not revealed any results that would give rise to safety concerns at low levels of exposure. Chronic studies have been performed on more 30 major chemical constituents (menthol, carvone, limonene, citral, cinnamaldehyde, benzaldehyde, benzyl acetate, 2-ethyl-1-hexanol, methyl anthranilate, geranyl acetate, furfural, eugenol, isoeugenol, etc.) found in many essential oils. The majority of these studies were hazard determinations that were sponsored by the National Toxicology Program, and they were normally performed at dose levels many orders of magnitude greater than the daily intakes of these constituents from consumption of the essential oil. Even at these high intake levels, the majority of the constituents show no carcinogenic potential (Smith et a., 2005b). In addition to dose/exposure, for some flavor ingredients, the carcinogenic potential that was assessed in the study is related to several additional factors including the mode of administration, species and sex of the animal model, and target organ specificity. In the vast majority of studies, the carcinogenic effect occurs through a nongenotoxic mechanism in which tumors form secondary to preexisting high-dose, chronic organ toxicity, typically to the liver or kidneys. Selected subgroups of structurally related substances (e.g., aldehydes, terpene hydrocarbons) are associated with a single-target organ and tumor type in a specific species and sex of rodent (i.e., male rat kidney tumors secondary to alpha-2u-globulin neoplasms with limonene in male rats) or using a single mode of administration (i.e., forestomach tumors that arise due to high doses of benzaldehyde and hexadienal given by gavage). Given their long history of use, it appears unlikely that there are essential oils consumed by humans that contain constituents not yet studied that are weak nongenotoxic carcinogens at chronic high-dose levels. Even if there are such cases, because of the relatively low intake (Lucas et al., 1999) as constituents of essential oils, these yet-to-be-discovered constituents would be many orders of magnitude less potent than similar levels of aflatoxins (found in peanut butter), the polycyclic heterocyclic amines (found in cooked foods), or the polynuclear aromatic hydrocarbons (also found in cooked foods). There is nothing to suggest that the major biosynthetic pathways available to higher plants are capable of producing substances such that low levels of exposure to the substance would result in a high level of toxicity or carcinogenicity. The toxic and carcinogenic potentials exhibited by constituent chemicals in essential oils can largely be equated with the toxic potential of the congeneric group to which that chemical belongs. A comparison of the oral toxicity data (JECFA, 2004) for limonene, myrcene, pinene, and other members of the congeneric group of terpene hydrocarbons shows similar low levels of toxicity with the same high-dose target organ endpoint (kidney) in animal studies, of which the relevance to humans is unlikely. Likewise, dietary toxicity and carcinogenicity data (JECFA, 2001) for cinnamyl alcohol, cinnamaldehyde, cinnamyl acetate, and other members of the congeneric group of 3-phenyl-1propanol derivatives show similar toxic and carcinogenic endpoints. The safety data for the congeneric chemical groups that are found in vast majority of essential oils have been reviewed (Adams et al., 1996, 1997, 1998, 2004; JECFA 1997, 1998, 1999, 2000a, b, 2001, 2003, 2004; Newberne et al., 1999; Smith et al., 2002a, b). Available data for different representative members in each of these congeneric groups support the conclusion that the toxic and carcinogenic potential of individual constituents adequately represent similar potentials for the corresponding congeneric group. The second key factor in the determination of safety is the level of intake of the congeneric group from consumption of the essential oil. Intake of the congeneric group will, in turn, depend

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upon the variability of the chemical composition of the essential oil in the marketplace and on the conditions of use. As discussed earlier, chemical analysis of the different batches of oil obtained from the same and different manufacturers will produce a range of concentrations for individual constituents in each congeneric group of the essential oil. The mean concentration values (%) for constituents are then summed for all members of the congeneric group. The total % determined for the congeneric group is multiplied by the estimated daily intake (PCI × 10) of the essential oil to provide a conservative estimate of exposure to each congeneric group from consumption of the essential oil. In some essential oils, the intake of one constituent, and therefore, one congeneric group, may account for essentially all of the oil (e.g., linalool in coriander oil, citral in lemongrass oil, benzaldehyde in bitter almond oil). In other oils, exposure to a variety of congeneric groups over a broad concentration range may occur. As noted earlier, cardamom oil is an example of such an essential oil. Ultimately, it is the relative intake and the toxic potential of each congeneric group that is the basis of the congeneric group-based safety evaluation. The combination of relative intake and toxic potential will prioritize congeneric groups for the safety evaluation. Hypothetically, a congeneric group of increased toxic potential that accounts for only 5% of the essential oil may be prioritized higher than a congeneric group of lower toxic potential accounting for 95%. The following guide and examples therein are intended to more fully illustrate the principles described earlier that are involved in the safety evaluation of essential oils. Fermentation products, process flavors, substances derived from fungi, microorganisms, or animals, and direct food additives are explicitly excluded. The guide is designed primarily for application to essential oils and extracts for use as flavoring substances. The guide is a tool to organize and prioritize the chemical constituents and congeneric groups in an essential oil in such a way as to allow a detailed analysis of their chemical and biological properties. This analysis as well as consideration of other relevant scientific data provides the basis for a safety evaluation of the essential oil under conditions of intended use. Validation of the approach is provided, in large part, by a detailed comparison of the doses and toxic effects exhibited by constituents of the congeneric group with the equivalent doses and effects provided by the essential oil. This methodology, with some variations based on expert judgment as appropriate, has been in use for more than 10 years through the FEMA GRAS program, in safety evaluations conducted by the FEMA Expert Panel (Smith et al., 2004).

9.5  GUIDE AND EXAMPLE FOR THE SAFETY EVALUATION OF ESSENTIAL OILS 9.5.1 Introduction The guide is a procedure involving a comprehensive evaluation of the chemical and biological properties of the constituents and congeneric groups of an essential oil. Constituents in, for instance, an essential oil that are of known structure are organized into congeneric groups that exhibit similar metabolic and toxicologic properties. The congeneric groups are further classified according to levels (Structural Classes I, II, and III) of toxicologic concern using a decision tree approach (Cramer et al., 1978; Munro et al., 1996b). Based on intake data for the essential oil and constituent concentrations, the congeneric groups are prioritized according to intake and toxicity potential. The procedure ultimately focuses on those congeneric groups that, due to their structural features and intake, may pose some significant risk from the consumption of the essential oil. Key elements used to evaluate congeneric groups include exposure, structural analogy, metabolism, and toxicology, which includes toxicity, carcinogenicity, and genotoxic potential (Oser and Hall, 1977; Oser and Ford, 1991; Woods and Doull, 1991; Adams et al., 1996, 1997, 1998, 2004; Newberne et al., 1999; Smith et al., 2002a,b, 2004). Throughout the analysis of these data, it is essential that professional judgment and expertise be applied to complete the safety evaluation of the essential oil. As an example of how a typical evaluation process for an essential oil is carried out according to this guide, the safety evaluation for flavor use of corn mint oil (Mentha arvensis) is outlined in the following text.

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9.5.2  Elements of the Guide for the Safety Evaluation of the Essential Oil 9.5.2.1 Introduction In Step 1 of the guide, the evaluation procedure estimates intake based on industry survey data for each essential oil. It then organizes the chemically identified constituents that have an intake >1.5 µg/ day into congeneric groups that participate in common pathways of metabolism and exhibit a similar toxic potential. In Steps 2 and 3, each identified chemical constituent is broadly classified according to toxic potential (Cramer et al., 1978) and then assigned to a congeneric group of structurally related substances that exhibit similar pathways of metabolism and toxicologic potential. Before the formal evaluation begins, it is necessary to specify the data (e.g., botanical, physical, chemical) required to completely describe the product being evaluated. In order to effectively evaluate an essential oil, attempted complete analyses must be available for the product intended for the marketplace from a number of flavor manufacturers. Additional quality control data are useful, as they demonstrate consistency in the chemical composition of the product being marketed. A Technical Information Paper drafted for the particular essential oil under consideration organizes and prioritizes these data for efficient sequential evaluation of the essential oil. In Steps 8 and 9, the safety of the essential oil is evaluated in the context of all congeneric groups and any other related data (e.g., data on the essential oil itself or for an essential oil of similar composition). The procedure organizes the extensive database of information on the essential oil constituents in order to efficiently evaluate the safety of the essential oil under conditions of use. It is important to stress, however, that the guide is not intended to be nor in practice operates as a rigid checklist. Each essential oil that undergoes evaluation is different, and different data will be available for each. The overriding objective of the guide and subsequent evaluation is to ensure that no significant portion of the essential oil should go unevaluated. 9.5.2.2  Prioritization of Essential Oil According to Presence in Food In Step 1, essential oils are prioritized according to their presence or absence as components of commonly consumed foods (Step 1). This question evaluates the relative intake of the essential oil as an intentionally added flavoring substance versus its intake as a component part of food. Many essential oils are isolated from plants that are commonly consumed as a food. Little or no safety concerns should exist for the intentional addition of the essential oil to the diet, if intake of the oil from consumption of traditional foods (garlic) substantially exceeds intake as an intentionally added flavoring substance (garlic oil). In many ways, the first step applies the concept of “long history of safe use” to essential oils. That is, if exposure to the essential oil occurs predominantly from consumption of a normal diet, a conclusion of safety is straightforward. Step 1 of the guide clearly places essential oils that are consumed as part of a traditional diet on a lower level of concern than those oils derived from plants that are either not part of the traditional diet or whose intake is not predominantly from the diet. The first step also mitigates the need to perform comprehensive chemical analysis for essential oils in those cases where intake is low and occurs predominantly from consumption of food. An estimate of the intake of the essential oil is based on the most recent poundage available from flavor industry surveys and the assumption that the essential oil is consumed by only 10% of the population for an oil having a survey volume 50,000 kg/year. In addition, the detection limit for constituents is determined based on the daily PCI of the essential oil. 9.5.2.2.1  Corn Mint Oil To illustrate the type of data considered in Step 1, consider corn mint oil. Corn mint oil is produced by the steam distillation of the flowering herb of M. arvensis. The crude oil contains upward of 70% (−)-menthol, some of which is isolated by crystallization at low temperature. The resulting dementholized oil is corn mint oil. Although produced mainly in Brazil during the 1970s and 1980s, corn mint oil is now produced predominantly in China and India. Corn mint has a more stringent

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taste compared to that of peppermint oil, M. piperita, but can be efficiently produced and is used as a more cost-effective substitute. Corn mint oil isolated from various crops undergoes subsequent clean up, further distillation, and blending to produce the finished commercial oil. Although there may be significant variability in the concentrations of individual constituents in different samples of crude essential oil, there is far less variability in the concentration of constituents and congeneric groups in the finished commercial oil. The volume of corn mint oil reported in the most recent U.S. poundage survey is 446,000 kg/year (Harman et al., 2013), which is approximately 25% of the potential market of peppermint oil. Because corn mint oil is a high-volume essential oil, it is highly likely that the entire population consumes the annual reported volume, and therefore, the daily PCI is calculated based on 100% of the population (310,000,000). This results in a daily PCI of approximately 3.9 mg/ person/day (0.066 mg/kg bw/day) of corn mint oil:



446, 000 kg/year ×109 µg/kg = 3942 µg/person/day 365 days/year ×310 ×106 persons

Based on the intake of corn mint oil (3942 µg/day), any constituent present at greater than 0.038% would need to be chemically characterized and quantified:



1.5 µg/day ×100 = 0.038% 3942 µg/day

9.5.2.3  Organization of Chemical Data: Congeneric Groups and Classes of Toxicity In Step 2, constituents are assigned to one of three structural classes (I, II, or III) based on toxic potential (Cramer et al., 1978). Class I substances contain structural features that suggest a low order of oral toxicity. Class II substances are clearly less innocuous than Class I substances but do not contain structural features that provide a positive indication of toxicity. Class III substances contain structural features (e.g., an epoxide functional group, unsubstituted heteroaromatic derivatives) that permit no strong presumption of safety and in some cases may even suggest significant toxicity. For instance, the simple aliphatic hydrocarbon, limonene, is assigned to structural Class I, while elemicin, which is an allyl-substituted benzene derivative with a reactive benzylic/allylic position, is assigned to Class III. Likewise, chemically unidentified constituents of the essential oil are automatically placed in Structural Class III, since no presumption of safety can be made. The toxic potential of each of the three structural classes has been quantified (Munro et al., 1996a). An extensive toxicity database has been compiled for substances in each structural class. The database covers a wide range of chemical structures, including food additives, naturally occurring substances, pesticides, drugs, antioxidants, industrial chemicals, flavors, and fragrances. Conservative no observable effect levels (fifth percentile NOELs) have been determined for each class. These fifth percentile NOELs for each structural class are converted to human exposure threshold levels by applying a 100-fold safety factor and correcting for mean bodyweight (60/100). The human exposure threshold levels are referred to as thresholds of toxicological concern (TTC). With regard to flavoring substances, the TTCs are even more conservative, given that the vast majority of NOELs for flavoring substances are above the 90th percentile. These conservative TTCs have since been adopted by the WHO and Commission of the European Communities for use in the evaluation of chemically identified flavoring agents by JECFA and the European Food Safety Authority (EFSA) (JECFA, 1997; EC, 1999). Step 3 is a key step in the guide. It organizes the chemical constituents into congeneric groups that exhibit common chemical and biological properties. Based on the well-recognized biochemical pathways operating in plants, essentially all of the volatile constituents found in essential oils, extracts, and oleoresins belong to well-recognized congeneric groups. Recent reports (Maarse et al., 1992, 1994, 2000; Njissen et al., 2003) of the identification of new naturally occurring constituents indicate

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that newly identified substances fall into existing congeneric groups. The Expert Panel, JECFA, and the EC have acknowledged that individual chemical substances can be evaluated in the context of their respective congeneric group (JECFA, 1997; EC, 1999; Smith et al., 2005a, b). The congeneric group approach provides the basis for understanding the relationship between the biochemical fate of members of a chemical group and their toxicologic potential. Within this framework, the objective is to continuously build a more complete understanding of the absorption, distribution, metabolism, and excretion of members of the congeneric group and their potential to cause systemic toxicity. Within the guidelines, the structural class of each congeneric group is assigned based on the highest structural class of any member of the group. Therefore, if an essential oil contained a group of furanone derivatives that were variously assigned to Structural Classes II and III, in the evaluation of the oil, the congeneric group would, in a conservative manner, be assigned to Class III. The types and numbers of congeneric groups in a safety evaluation program are, by no means, static. As new scientific data and information become available, some congeneric groups are combined while others are subdivided. This has been the case for the group of alicyclic secondary alcohols and ketones that were the subject of a comprehensive scientific literature review in 1975 (FEMA, 1975). Over the last two decades, experimental data have become available indicating that a few members of this group exhibit biochemical fate and toxicologic potential inconsistent with that for other members of the same group. These inconsistencies, almost without exception, arise at highdose levels that are irrelevant to the safety evaluation of low levels of exposure to flavor use of the substance. However, given the importance of the congeneric group approach in the safety assessment program, it is critical to resolve these inconsistencies. Additional metabolic and toxicologic studies may be required to distinguish the factors that determine these differences. Often the effect of dose and a unique structural feature results in utilization of a metabolic activation pathway not utilized by other members of a congeneric group. Currently, evaluating bodies including JECFA, EFSA, and the FEMA Expert Panel have classified flavoring substances into the same congeneric groups for the purpose of safety evaluation. In Steps 5, 6, and 7, each congeneric group in the essential oil is evaluated for safety in use. In Step 5, an evaluation of the metabolism and disposition is performed to determine, under current conditions of intake, whether the group of congeneric constituents is metabolized by well-established detoxication pathways to yield innocuous products. That is, such pathways exist for the congeneric group of constituents in an essential oil, and safety concerns will arise only if intake of the congeneric group is sufficient to saturate these pathways potentially leading to toxicity. If a significant intoxication pathway exists (e.g., pulegone), this should be reflected in a higher decision tree class and lower TTC threshold. At Step 6 of the procedure, the intake of the congeneric group relative to the respective TTC for one of the three structural classes (1800 µg/day for Class I; 540 µg/day for Class II; 90 µg/day for Class III; see Table 9.1) is evaluated. If the intake of the congeneric group is less than the threshold for the respective structural class, the intake of the congeneric group presents no significant safety concerns. The group passes the first phase of the evaluation and is then referred to Step 8, the step in which the safety of the congeneric group is evaluated in the context of all congeneric groups in the essential oil. If, at Step 5, no sufficient metabolic data exist to establish safe excretion of the product or if activation pathways have been identified for a particular congeneric group, then the group moves to Step 7, and toxicity data are required to establish safe use under current conditions of intake. There are examples where low levels of xenobiotic substances can be metabolized to reactive substances. In the event that reactive metabolites are formed at low levels of intake of naturally occurring substances, a detailed analysis of dose-dependent toxicity data must be performed. Also, if the intake of the congeneric group is greater than the human exposure threshold (suggesting metabolic saturation may occur), then toxicity data are also required. If, at Step 7, a database of relevant toxicological data for a representative member or members of the congeneric group indicates that a sufficient margin of safety exists for the intake of the congeneric group, the members of that congeneric group are concluded to be safe under conditions of use of the essential oil. The congeneric group then moves to Step 8.

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TABLE 9.1 Structural Class Definitions and Their Human Intake Thresholds Class I

II

III

a

Description Structure and related data suggest a low order of toxicity. If combined with low human exposure, they should enjoy an extremely low priority for investigation. The criteria for adequate evidence of safety would also be minimal. Greater exposures would require proportionately higher priority for more exhaustive study. Intermediate substances. They are less clearly innocuous than those of Class I, but do not offer the basis either of the positive indication of toxicity or of the lack of knowledge characteristic of those in Class III. Permit no strong initial presumptions of safety or that may even suggest significant toxicity. They thus deserve the highest priority for investigation. Particularly when per capita intake is high of a significant subsection of the population that has a high intake, the implied hazard would then require the most extensive evidence for safety in use.

Fifth Percentile NOEL (mg/kg/day)

Human Exposure Threshold (TTC)a (µg/day)

3.0

1800

0.91

540

0.15

90

The human exposure threshold was calculated by multiplying the fifth percentile NOEL by 60 (assuming an individual weighs 60 kg) and dividing by a safety factor of 100.

In the event that insufficient data are available to evaluate a congeneric group at Step 7, or the currently available data result in margins of safety that are not sufficient, the essential oil cannot be further evaluated by this guide and must be set aside for further considerations. Use of the guide requires scientific judgment at each step of the sequence. For instance, if a congeneric group that accounted for 20% of a high-volume essential oil was previously evaluated and found to be safe under intended conditions of use, the same congeneric group found at less than 2% of a low-volume essential oil does not need to be further evaluated. Step 8 considers additivity or synergistic interactions between individual substances and between the different congeneric groups in the essential oil. As for all other toxicological concerns, the level of exposure to congeneric groups is relevant to whether additive or synergistic effects present a significant health hazard. The vast majority of essential oils are used in food in extremely low concentrations, which therefore results in very low intake levels of the different congeneric groups within that oil. Moreover, major representative constituents of each congeneric group have been tested individually and pose no toxicological threat even at dose levels that are orders of magnitude greater than normal levels of intake of essential oils from use in traditional foods. Based on the results of toxicity studies both on major constituents of different congeneric groups in the essential oil and on the essential oil itself, it can be concluded that the toxic potential of these major constituents is representative of that of the oil itself, indicating the likely absence of additivity and synergistic interaction. In general, the margin of safety is so wide and the possibility of additivity or synergistic interaction so remote that combined exposure to the different congeneric groups and the unknowns are considered of no health concern, even if expert judgment cannot fully rule out additivity or synergism. However, case-by-case considerations are appropriate. Where possible combined effects might be considered to have toxicological relevance, additional data may be needed for an adequate safety evaluation of the essential oil. Additivity of toxicologic effect or synergistic interaction is a conservative default assumption that may be applied whenever the available metabolic data do not clearly suggest otherwise. The

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extensive database of metabolic information on congeneric groups (JECFA, 1997, 1998, 1999, 2000a, b, 2001, 2003, 2004) that are found in essential oils suggests that the potential for additive effects and synergistic interactions among congeneric groups in essential oils is extremely low. Although additivity of effect is the approach recommended by NAS/NRC committees (NRC, 1988, 1994) and regulatory agencies (EPA, 1988), the Presidential Commission of Risk Assessment and Risk Management recommended (Presidential Commission, 1996) that “For risk assessments involving multiple chemical exposures at low concentrations, without information on mechanisms, risks should be added. If the chemicals act through separate mechanisms, their attendant risks should not be added but should be considered separately.” Thus, the risks of chemicals that act through different mechanisms, that act on different target systems, or that are toxicologically dissimilar in some other way should be considered to be independent of each other. The congeneric groups in essential oils are therefore considered separately. Further, the majority of individual constituents that comprise essential oils are themselves used as flavoring substances that pose no toxicological threat at doses that are magnitudes greater than their level of intake from the essential oil. Rulis (1987) reported that “The overwhelming majority of additives present a high likelihood of having safety assurance margins in excess of 105.” He points out that this is particularly true for additives used in the United States at less than 100,000 lb/year. Because more than 90% of all flavoring ingredients are used at less than 10,000 lb/year (Hall and Oser, 1968), this alone implies intakes commonly many orders of magnitude below the no-effect level. Nonadditivity thus can often be assumed. As is customary in the evaluation of any substance, high-end data for exposure (consumption) are used, and multiple other conservatisms are employed to guard against underestimation of possible risk. All of these apply to complex mixtures as well as to individual substances. 9.5.2.3.1  Corn Mint Oil Congeneric Groups In corn mint oil, the principal congeneric group is composed of terpene alicyclic secondary alcohols, ketones, and related esters, as represented by the presence of (−)-menthol, (−)-menthone, (+)-isomenthone, (−)-menthyl acetate, and other related substances. Samples of triple-distilled commercial corn mint oil may contain up to 95% of this congeneric group. The biochemical and biological fate of this group of substances has been previously reviewed (Adams et al., 1996; JECFA, 1999). Key data on metabolism, toxicity, and carcinogenicity are cited in the following text (Table 9.2) in order to complete the evaluation. Although constituents in this group are effectively detoxicated via conjugation of the corresponding alcohol or ω-oxidation followed by conjugation and excretion (Williams, 1940; Madyastha and Srivatsan, 1988; Yamaguchi et al., 1994), the intake of the congeneric group (3745 µg/person/day or 3.75 mg/person/day, see Table 9.2) is higher than the exposure threshold of (540 µg/person/day or 0.540 mg/person/day for Structural Class II. Therefore, toxicity data are required for this congeneric group. In both short- and long-term studies (NCI, 1979; Madsen et al., 1986), menthol, menthone, and other members of the group exhibit NOAELs at least 1000 times the daily PCI (eaters only) (3.75 mg/person/day or 0.062 mg/kg bw/day) of this congeneric group resulting from intake of the essential oil. For members of this group, numerous in vitro and in vivo genotoxicity assays are consistently negative (Florin et al., 1980; Heck et al., 1989; Sasaki et al., 1989; Muller, 1993; Zamith et al., 1993; Rivedal et al., 2000; NTP Draft, 2003a). Therefore, the intake of this congeneric group from consumption of M. arvensis is not a safety concern. Although it is a constituent of corn mint oil and is also a terpene alicyclic ketone structurally related to the aforementioned congeneric group, pulegone exhibits a unique structure (i.e., 2-isopropylidenecyclohexanone) that participates in a well-recognized intoxication pathway (see Figure 9.2) (McClanahan et al., 1989; Thomassen et al., 1992; Adams et al., 1996; Chen et al., 2001) that leads to the formation of menthofuran. This metabolite subsequently oxidizes and ring opens to yield a highly reactive 2-ene-1,4-dicarbonyl intermediate that reacts readily with proteins resulting in hepatotoxicity at intake levels at least two orders of magnitude less than no observable effect levels for structurally related alicyclic ketones and secondary alcohols (menthone, carvone, and menthol). Therefore, pulegone

III (90)

2-Isopropylidene cyclohexanone and metabolites (e.g., pulegone)

2

8

95

Step 3. High% from Multiple Commercial Samples

79

315

3745

Step 4. Intake, µg/ Person/Day

Based on daily per capita intake of 3942 µg/person/day for corn mint oil.

I (1800)

Aliphatic terpene hydrocarbon (e.g., limonene, pinene)

a

II (540)

Secondary alicyclic saturated and unsaturated alcohol/ketone/ketal/ ester (e.g., menthol, menthone, isomenthone, menthyl acetate)

Congeneric Group

 Step 2. Decision Tree Class (TTC, µg/Person/ Day)

TABLE 9.2 Safety Evaluation of Corn Mint Oil, Mentha arvensisa

3. Hydroxylation catalyzed by cytochrome P-450 to yield a series of ring- and side-chain-hydroxylated pulegone metabolites, one of which is a reactive 2-ene-1,4-dicarbonyl derivative. This intermediate is known to form protein adducts leading to enhanced toxicity. (Austin et al., 1988)

Michael-type addition leading to mercapturic acid conjugates that are excreted or further hydroxylated and excreted.

1. Reduction to yield menthone or isomenthone, followed by hydroxylation of ring or side-chain positions and then conjugation with glucuronic acid. 2. Conjugation with glutathione in a

carboxy metabolites excreted as glucuronic acid conjugates.

1. Glucuronic acid conjugation of the alcohol followed by excretion in the urine.2. ω-Oxidation of the side-chain substituents to yield various polyols and hydroxyacids and excreted as glucuronic acid conjugates. 1. ω-Oxidation to yield polar hydroxy and

Step 5. Metabolism Pathways

Yes, 79  TTC

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H Reduction

OAc Isopulegyl acetate

H

H

Hydrolysis

Isomerization O

OH Isopulegol

Isopulegone

O Pulegone

9-Hydroxylation Glucuronic acid conjugate

H

OH 9-Hydroxyisopulegone

H

O Menthofuran

FIGURE 9.2  Metabolism of isopulegone, pulegone, and isopulegyl acetate.

and its metabolite (menthofuran), which account for 2% of the composition of corn mint oil is a congeneric group of terpene hydrocarbons ((+) and (−)-pinene, (+) limonene, etc.). Although these may contribute up to 8% of the oil, upon multiple redistillations during processing, the hydrocarbon content can be significantly reduced (80% and are both hepatotoxic and pneumotoxic in mice (Engel, 2003). Though pulegone has been used for flavoring food and oral hygiene products, the pulegone content in foods and beverages is restricted by EU law. At nontoxic concentrations, pulegone is oxidized selectively at the 10-position and further metabolized into menthofuran. In vitro data show that this metabolic pathway is mainly catalyzed by human CYP2E1, and to a lesser extent by CYP1A2 and CYP 2C19 (Khojasteh-Bakht et al., 1999). Alternatively, it may be reduced to menthone, which has been detected in trace levels in urine samples. It might be possible that pulegone is also reduced at the carbonyl group first. Consequently, pulegol is either reduced very efficiently to menthol or rearranged to 3-p-menthen-8-ol (Engel, 2003) (Figure 10.23). In rats, three major pathways have been identified: (a) hydroxylation followed by glucuronidation, (b) reduction to menthone and hydroxylation, and (c) conjugation with glutathione and further metabolism (Chen et al., 2001; Ferguson et al., 2007).

10.2.23  α-Terpineol α-Terpineol, a monocyclic monoterpene alcohol, is found in the essential oil sources such as neroli, petitgrain, Melaleuca alternifolia, and Origanum vulgare (Bornscheuer et al., 2014). Based on its

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OH

OH

COOH 2-(2-Hydroxy-4-methylphenyl)propionic acid

8,9-Dehydrothymol

OH HO OH O

4-Hydroxypulegone

O

O

5-Hydroxypulegone Thymol

Rat in vivo

HO

O

O

OH

Rat in vivo

7α-Hydroxymintlactone Rat in vivo Human in vivo

Human, rat in vivo O

O

O

OH

3-Hydroxypulegone

10-Hydroxypulegone

Pulegone Human, rat in vivo

Rat in vivo

OH

O

O O

OH

O

OH

HO 8-Hydroxy-4-p-menthen-3-one

10-Hydroxymenthone Pulegol Human in vivo

Rat in vivo

Menthone

OH

Human in vivo

OH

5-Hydroxymenthone

OH

O 7-Hydroxymenthone

HO Menthol

3,9-Epoxy-pmenthan-3-ol

O

O p-Menth-3-en-8-ol

Menthofuran

Piperitone

FIGURE 10.23  Proposed metabolism of pulegone in humans and rats. (Adapted from Engel 2003. J. Agric. Food Chem. 51:6589–6597; Chen et al. 2001. Drug Metab. Dispos. 29:1567–1577.)

pleasant lilac-like odor, α-terpineol is widely used in the manufacture of perfumes, cosmetics, soaps, and antiseptic agents. After oral administration to rats (600 mg/kg body weight), α-terpineol is metabolized to p-menthane-1,2,8-triol, probably formed from the epoxide intermediate. Notably, allylic methyl oxidation and the reduction of the 1,2-double bond are the major routes for the biotransformation of α-terpineol in rat (Figure 10.24). Although allylic oxidation of C-1 methyl

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O

Rat in vitro

HO

HO HO α-Terpineol

OH

p-Menthane-1,2,8-triol OH

COOH

COOH

HO

HO

HO Dihydrooleuropeic acid

Oleuropeic acid

FIGURE 10.24  Proposed metabolism of α-terpineol in rats. (Adapted from Madyastha and Srivatsan 1988. Environ. Contam. Toxicol. 41:17–25.)

seems to be the major pathway, the alcohol p-ment-1-ene-7,8-diol could not be isolated from the urine samples. Probably, this compound is accumulated and is readily further oxidized to oleuropeic acid (Madyastha and Srivatsan, 1988a).

10.2.24 Terpinen-4-ol (–)-Terpinen-4-ol is the main compound of Melaleuca alternifolia essential oil, which is widely used in cosmetic and pharmaceutical products due to its antiseptic properties (Bornscheuer et al., 2014). As shown in Figure 10.25, terpinen-4-ol is oxidized to p-menth-1-en-4,8-diol and 1,2-epoxyp-menthan-4-ol. Further experiments with human recombinant CYPs identified CYP3A4, CYP 2A6, and to a lesser extent CYP1A2 as the main enzymes involved in the formation of terpinene-4-ol (Haigou and Miyazawa, 2012).

10.2.25  α- and β-Thujone α-Thujone and β-thujone are bicyclic monoterpenes that differ in the stereochemistry of the C-4 methyl group. The isomer ratio depends on the plant source, with high content of α-thujone in Thuja occidentalis and β-thujone in the essential oils of Tanacetum vulgare, Salvia officinalis, and Artemisia absinthium. Due to their neurotoxicity, maximum permissible amounts for thujones in O Human in vitro OH OH p-Menth-1-en-4,8-diol

Human in vitro OH Terpinen-4-ol

OH 1,2-Epoxy-p-menthan-4-ol

FIGURE 10.25  Proposed metabolism of terpinen-4-ol in human liver microsomes. (Adapted from Haigou and Miyazawa 2012. J. Oleo Sci. 61:35–43.)

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OH

O Mouse, rat in vivo

OH

OH

O O Thujol (R) Neothujol (S)

4,10-Dehydro-α-thujone Mouse, rat in vivo HO O

10-Hydroxy-α-thujone β-thujone

Carvacrol

Mouse, rabbit in vitro/in vivo

Human, rat, mouse in vitro

Human in vitro

O

Mouse, rat in vivo 4-Hydroxy-α-thujone β-thujone Glucuronide

OH

Mouse, rat α-Thujone β-Thujone O in vivo OH

Mouse, rat in vivo Human, rat, mouse in vitro

Mouse, rat Mouse, rat Glucuronide in vivo 2-Hydroxy-α-thujone in vivo Glucuronide β-thujone

8-Hydroxy-α-thujone O

OH 7-Hydroxy-α-thujone β-thujone

O Mouse, rat in vivo

7,8-Dehydro-α-thujone β-thujone

FIGURE 10.26  Proposed in vitro and in vivo metabolism of α- and β-thujone in rats. (Adapted from Ishida et al. 1989. Xenobiotica 19:843–855; Höld et al. 2000. Environ. Chem. Toxicol. 97:3826–3831; Höld et al. 2001. Chem. Res. Toxicol. 14:589–595.)

foods and beverages have been defined in the EU (O’Neil, 2006; Bornscheuer et al., 2014). α-Thujone is best known as the active ingredient of the alcoholic beverage absinthe, which was a very popular European drink in the 1800s. In vivo rabbit, rat, and mouse models conformed to the in vitro data using rat, mice, and human liver microsomes where α- and β-thujone are extensively metabolized to six hydroxy-thujones and three dehydrothujones (Figure 10.26). In Phase II reactions, several metabolites are further conjugated with glucuronic acid (Ishida et al., 1989; Höld et al., 2000, 2001). Recent studies have shown that the main human metabolites 4- and 7-hydroxy-a-thujone are formed by CYP2A, CYP2B6, CYP2D6, and CYP3A4 (Abass et al., 2011). CYP3A4 seems to be involved in the formation of most of the thujone metabolites in human models (Jiang and Ortiz de Montellano, 2009). Based on the in vitro and in vivo data, biotransformation should also be pronounced in humans after the intake of α- and β-thujone.

10.2.26 Thymol Thymol is the major constituent of the essential oils of Thymus vulgaris and Origanum vulgare. It is medically used in preparations against bronchitis and oral infections. Due to its antiseptic properties, it is also an ingredient of mouthwashes and toothpastes (O’Neil, 2006; Bornscheuer et al., 2014). Although after oral application to rats, large quantities were excreted unchanged or as their glucuronide and sulfate conjugates, extensive oxidation of the methyl and isopropyl groups also occurred (Austgulen et al., 1987), resulting in the formation of derivatives of benzyl alcohol and 2-phenylpropanol and their corresponding carboxylic acids. Ring hydroxylation was only a minor reaction in rats (Figure 10.27), whereas in humans, several ring-hydroxylated products were found

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OH OH

OH

OH

p-Cymene-2,3-diol

HO p-Cymene-3,8-diol

p-Cymen-8-en-3-ol Human in vivo

OH COOH 2-(2-Hydroxy-4-methylphenyl)propionic acid OH

OH OH

OH Thymol Rat in vivo Human in vivo

O

OH

Rat in vivo Human in vivo

9-Hydroxythymol OH

OH 7,9-Dihydroxythymol COOH

HO

O

Thymoquinone

OH

Thymoquinol

OH

7-Hydroxythymol

OH

3-Hydroxy-4-(1-methylethyl)benzoic acid

FIGURE 10.27  Proposed metabolism of thymol in humans and rats. (Adapted from Austgulen et al. 1987. Pharmacol. Toxicol. 61:98–102; Thalhamer et al. 2011. J. Pharm. Biomed. Anal. 56:64–69; Pisarčíková et al. 2017. Biomed. Chromatogr. 31:e3881.)

after oral application. All metabolites could be detected in human urine samples that were treated with glucuronidase prior to analysis (Thalhamer et al., 2011). CYP1A2, CYP2A6, and CYP2D6 were shown to be the main enzymes responsible for metabolizing thymol in human liver microsomes (Dong et al., 2012b). Thymol is excreted as glucuronide and sulfate into human urine which can be also assumed for its metabolites (Kohlert et al., 2002). After feeding broiler chickens various amounts of thymol, high concentrations of thymol sulfate and thymol glucuronide could be detected in the duodenal wall; decreasing concentrations of thymol sulfate were also found in plasma and liver samples (Pisarčíková et al., 2017).

10.3  METABOLISM OF SESQUITERPENES 10.3.1 Caryophyllene (–)-β-Caryophyllene is a common sesquiterpene with a clove- or turpentine-like odor. It can be found in the essential oils of Syzygium aromaticum, Piper nigrum, and Humulus lupulus. It is used as a flavoring substance; for example, for chewing gums (O’Neil, 2006; Bornscheuer et al., 2014). The biotransformation of (–)-β-caryophyllene in rabbits yielded the main metabolite (10S)-(– )-14-hydroxycaryophyllen-5,6-oxide and the minor biotransformation product caryophyllene-5,6oxide-2,12-diol. The formation of the minor metabolites is easily explained via the regioselective

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O

O OH H

H

H

H

OH

Caryophyllen-5,6-oxide-2,12-diol

Rabbit in vivo

(–)-Caryophyllen-5,6-oxide O

(–)-β-Caryophyllene

H

HO

HO

H

(10S)-(–)14-Hydroxy caryophyllen-5,6-oxide

FIGURE 10.28  Proposed metabolism of β-caryophyllene in rabbits. (Adapted from Asakawa et al. 1981. J. Pharm. Sci. 70:710–711; Asakawa et al. 1986. Xenobiotica 16:753–767.)

hydroxylation of the diepoxide intermediate (Asakawa et al., 1981, 1986). Based on in vivo data from rabbits, extensive biotransformation in humans is highly suggested after oral administration (Figure 10.28).

10.3.2 Farnesol Farnesol is present in many essential oils such as Cymbopogon species and neroli. It is used in perfumery and soaps to emphasize the odors of sweet floral perfumes and due to its fixative and antibacterial properties (O’Neil, 2006; Bornscheuer et al., 2014). Interestingly, it is also produced in humans where it acts on numerous nuclear receptors (Joo and Jetten, 2010). In vitro studies using recombinant drug metabolizing enzymes and human liver microsomes have shown that CYP isoenzymes participate in the metabolism of farnesol to 12-hydroxyfarnesol (Figure 10.29). Subsequently, farnesol and its metabolite are glucuronidated (DeBarber et al., 2004; Staines et al., 2004).

10.3.3 Longifolene Longifolene is primarily found in Indian turpentine oil, which is commercially extracted from Pinus roxburghii (chir pine) (Bornscheuer et al., 2014). In rabbits, longifolene is metabolized as follows: OH

Human in vitro

Farnesol Human, rabbit rat in vitro HO Hydroxyfarnesol

O

Glu

Farnesylglucuronide Human OH in vitro HO

O

Glu

Hydroxyfarnesylglucuronide

FIGURE 10.29  Proposed metabolism of farnesol in human liver microsomes. (Adapted from DeBarber et al. 2004. Biochim. Biophys. Acta 1682:18–27; Staines et al. 2004. Biochem. J. 384:637–645.)

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FIGURE 10.30  Proposed metabolism of (+)-longifolene in rabbits. (Adapted from Asakawa et al. 1986. Xenobiotica 16:753–767.) HO

Rabbit in vivo

HO

HO

HO O

Patchoulol

OH

OH

15-Hydroxypatchoulol

A hydroxy acid

Norpatchoulenol

FIGURE 10.31  Proposed metabolism of patchoulol in rabbits and dogs. (Adapted from Bang et al. 1975. Tetrahedron Lett. 26:2211–2214; Ishida 2005. Chem. Biodivers. 2:569–590.)

attack on the exo-methylene group from the endo-face to form its epoxide followed by isomerization of the epoxide to a stable endo-aldehyde. Then, rapid CYP-catalyzed hydroxylation of this endoaldehyde occurs (Asakawa et al., 1986) (Figure 10.30).

10.3.4  Patchoulol (–)-Patchoulol is the major active ingredient and the most odor-intensive component of patchouli oil, the volatile oil of Pogostemon cablin, one of the most important raw materials of perfumery, which is also used for its insect repellent activity (Bornscheuer et al., 2014). In rabbits and dogs, patchoulol is hydroxylated at the C-15 position, yielding a diol that is subsequently oxidized to a carboxylic acid. After decarboxylation and oxidation, the 3,4-unsaturated norpatchoulen-1-ol is formed, which also has a characteristic odor (Figure 10.31). All these urinary metabolites are also found as glucuronides, explaining their excellent water solubility (Bang et al., 1975; Ishida, 2005).

REFERENCES Abass, K., P. Reponen, S. Mattila et al. 2011. Metabolism of α-thujone in human hepatic preparations in vitro. Xenobiotica, 41: 101–111. Asakawa, Y., T. Ishida, M. Toyota et al. 1986. Terpenoid biotransformation in mammals. IV Biotransformation of (+)-longifolene, (-)-caryophyllene, (-)-caryophyllene oxide, (-)-cyclocolorenone, (+)-nootkatone, (-)-elemol, (-)-abietic acid and (+)-dehydroabietic acid in rabbits. Xenobiotica, 16: 753–767. Asakawa, Y., and Z. Taira, T. Takemoto et al. 1981. X-ray crystal structure analysis of 14-hydroxycaryophyllene oxide, a new metabolite of (-)-caryophyllene, in rabbits. Journal of Pharmaceutical Sciences, 70: 710–711. Asakawa, Y., M. Toyota, and T. Ishida. 1988. Biotransformation of 1,4-cineole, a monoterpene ether. Xenobiotica, 18: 1129–1134. Austgulen, L. T., E. Solheim, and R. R. Scheline. 1987. Metabolism in rats of p-cymene derivatives: Carvacrol and thymol. Pharmacology & Toxicology, 61: 98–102. Bang, L., G. Ourisson, and P. Teisseire. 1975. Hydroxylation of patchoulol by rabbits. Hemi-synthesis of norpatchoulenol, the odour carrier of patchouli Oil. Tetrahedron Letters, 16: 2211–2214. Bickers, D., P. Calow, H. Greim et al. 2003. A toxicologic and dermatologic assessment of linalool and related esters when used as fragrance ingredients. Food and Chemical Toxicology: An International Journal Published for the British Industrial Biological Research Association, 41: 919–942.

Metabolism of Terpenoids in Animal Models and Humans

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Bornscheuer, U., W. Streit, B. Dill et al., (eds.). 2014. Römpp online 4.0. Stuttgart, Germany: Georg Thieme Verlag KG. Boyle, R., S. McLean, W. Foley et al. 2001. Metabolites of dietary 1,8-cineole in the male koala (Phascolarctos cinereus). Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology, 129: 385–395. Chadha, A., and K. M. Madyastha. 1984. Metabolism of geraniol and linalool in the rat and effects on liver and lung microsomal enzymes. Xenobiotica, 14: 365–374. Chen, L.-J., E. H. Lebetkin, and L. T. Burka. 2001. Metabolism of (R)-(+)-pulegone in F344 rats. Drug Metabolism and Disposition, 29: 1567–1577. Crowell, P. L., S. Lin, E. Vedejs et al. 1992. Identification of metabolites of the antitumor agent d-limonene capable of inhibiting protein isoprenylation and cell growth. Cancer Chemotherapy and Pharmacology, 31: 205–212. DeBarber, A. E., L. A. Bleyle, J.-B. O. Roullet et al. 2004. Omega-hydroxylation of farnesol by mammalian cytochromes p450. Biochimica et Biophysica Acta, 1682: 18–27. Diliberto, J. J., P. Srinivas, D. Overstreet et al. 1990. Metabolism of citral, an alpha,beta-unsaturated aldehyde, in male F344 rats. Drug Metabolism and Disposition: The Biological Fate of Chemicals, 18: 866–875. Dong, R.-H., Z.-Z. Fang, L.-L. Zhu et al. 2012a. Identification of UDP-glucuronosyltransferase isoforms involved in hepatic and intestinal glucuronidation of phytochemical carvacrol. Xenobiotica, 42: 1009–1016. Dong, R.-H., Z.-Z. Fang, L.-L. Zhu et al. 2012b. Identification of CYP isoforms involved in the metabolism of thymol and carvacrol in human liver microsomes (HLMs). Die Pharmazie, 67: 1002–1006. Duisken, M., D. Benz, T. H. Peiffer et al. 2005. Metabolism of delta(3)-carene by human cytochrome p450 enzymes: Identification and characterization of two new metabolites. Current Drug Metabolism, 6: 593–601. Engel, W. 2001. In vivo studies on the metabolism of the monoterpenes S-(+)- and R-(-)-carvone in humans using the metabolism of ingestion-correlated amounts (MICA) approach. Journal of Agricultural and Food Chemistry, 49: 4069–4075. Engel, W. 2003. In vivo studies on the metabolism of the monoterpene pulegone in humans using the metabolism of ingestion-correlated amounts (MICA) approach: explanation for the toxicity differences between (S)(−)- and (R)-(+)-pulegone. Journal of Agricultural and Food Chemistry, 51: 6589–6597. Eriksson, K., and J. O. Levin. 1996. Gas chromatographic-mass spectrometric identification of metabolites from alpha-pinene in human urine after occupational exposure to sawing fumes. Journal of Chromatography. B, Biomedical Applications, 677: 85–98. Ferguson, L.-J. C., E. H. Lebetkin, F. B. Lih et al. 2007. 14C-labeled pulegone and metabolites binding to α2u-globulin in kidneys of male F-344 rats. Journal of Toxicology and Environmental Health, Part A, 70: 1416–1423. Gelal, A., P. Jacob 3rd, L. Yu et al. 1999. Disposition kinetics and effects of menthol. Clinical Pharmacology and Therapeutics, 66: 128–135. Gyoubu, K., and M. Miyazawa. 2007. In vitro metabolism of (-)-camphor using human liver microsomes and CYP2A6. Biological and Pharmaceutical Bulletin, 30: 230–233. Hagvall, L., J. M. Baron, A. Börje et al. 2008. Cytochrome P450-mediated activation of the fragrance compound geraniol forms potent contact allergens. Toxicology and Applied Pharmacology, 233: 308–313. Haigou, R., and M. Miyazawa. 2012. Metabolism of (+)-terpinen-4-ol by cytochrome P450 enzymes in human liver microsomes. Journal of Oleo Science, 61: 35–43. Höld, K. M., N. S. Sirisoma, and J. E. Casida. 2001. Detoxification of alpha- and beta-Thujones (the active ingredients of absinthe): Site specificity and species differences in cytochrome P450 oxidation in vitro and in vivo. Chemical Research in Toxicology, 14: 589–595. Höld, K. M., N. S. Sirisoma, T. Ikeda et al. 2000. Alpha-thujone (the active component of absinthe): Gammaaminobutyric acid type A receptor modulation and metabolic detoxification. Proceedings of the National Academy of Sciences of the United States of America, 97: 3826–3831. Horst, K., and M. Rychlik. 2010. Quantification of 1,8-cineole and of its metabolites in humans using stable isotope dilution assays. Molecular Nutrition & Food Research, 54: 1515–1529. Ishida, T. 2005. Biotransformation of terpenoids by mammals, microorganisms, and plant-cultured cells. Chemistry & Biodiversity, 2: 569–590. Ishida, T., Y. Asakawa, T. Takemoto et al. 1979. Terpenoid biotransformation in mammals. II: Biotransformation of dl-camphene in rabbits. Journal of Pharmaceutical Sciences, 68: 928–930. Ishida, T., Y. Asakawa, T. Takemoto et al. 1981. Terpenoids biotransformation in mammals III: Biotransformation of alpha-pinene, beta-pinene, pinane, 3-carene, carane, myrcene, and p-cymene in rabbits. Journal of Pharmaceutical Sciences, 70: 406–415. Ishida, T., M. Toyota, and Y. Asakawa. 1989. Terpenoid biotransformation in mammals. V. Metabolism of (+)-citronellal, (+-)-7–hydroxycitronellal, citral, (-)-perillaldehyde, (-)-myrtenal, cuminaldehyde, thujone, and (+-)-carvone in rabbits. Xenobiotica, 19: 843–855.

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Jäger, W., M. Mayer, P. Platzer et al. 2000. Stereoselective metabolism of the monoterpene carvone by rat and human liver microsomes. The Journal of Pharmacy and Pharmacology, 52: 191–197. Jäger, W., M. Mayer, G. Reznicek et  al. 2001. Percutaneous absorption of the montoterperne carvone: Implication of stereoselective metabolism on blood levels. The Journal of Pharmacy and Pharmacology, 53: 637–642. Jiang, Y., and P. R. Ortiz de Montellano. 2009. Cooperative effects on radical recombination in CYP3A4catalyzed oxidation of the radical clock β-thujone. ChemBioChem, 10: 650–653. Joo, J. H., and A. M. Jetten. 2010. Molecular mechanisms involved in farnesol-induced apoptosis. Cancer Letters, 287: 123–135. Kaffenberger, R. M., and M. J. Doyle. 1990. Determination of menthol and menthol glucuronide in human urine by gas chromatography using an enzyme-sensitive internal standard and flame ionization detection. Journal of Chromatography, 527: 59–66. Khojasteh, S. C., S. Oishi, and S. D. Nelson. 2010. Metabolism and toxicity of menthofuran in rat liver slices and in rats. Chemical Research in Toxicology, 23: 1824–1832. Khojasteh-Bakht, S. C., W. Chen, L. L. Koenigs et al. 1999. Metabolism of (R)-(+)-pulegone and (R)-(+)menthofuran by human liver cytochrome P-450s: Evidence for formation of a furan epoxide. Drug Metabolism and Disposition, 27: 574–580. Kohlert, C., G. Schindler, R. W. März et al. 2002. Systemic availability and pharmacokinetics of thymol in humans. The Journal of Clinical Pharmacology, 42: 731–737. Lassila, T., S. Mattila, M. Turpeinen et al. 2016. Tandem mass spectrometric analysis of S- and N-linked glutathione conjugates of pulegone and menthofuran and identification of P450 enzymes mediating their formation. Rapid Communications in Mass Spectrometry: RCM, 30: 917–926. Leibman, K. C., and E. Ortiz. 1973. Mammalian metabolism of terpenoids. I. Reduction and hydroxylation of camphor and related compounds. Drug Metabolism and Disposition: The Biological Fate of Chemicals, 1: 543–551. Madyastha, K. M., and V. Srivatsan. 1988a. Biotransformations of alpha-terpineol in the rat: Its effects on the liver microsomal cytochrome P-450 system. Bulletin of Environmental Contamination and Toxicology, 41: 17–25. Madyastha, K. M., and V. Srivatsan. 1988b. Studies on the metabolism of l-menthol in rats. Drug Metabolism and Disposition: The Biological Fate of Chemicals, 16: 765–772. Matsumoto, T., T. Ishida, T. Yoshida et al. 1992. The enantioselective metabolism of p-cymene in rabbits. Chemical & Pharmaceutical Bulletin, 40: 1721–1726. Meesters, R. J. W., M. Duisken, and J. Hollender. 2007. Study on the cytochrome P450-mediated oxidative metabolism of the terpene alcohol linalool: Indication of biological epoxidation. Xenobiotica, 37: 604–617. Meesters, R. J. W., M. Duisken, and J. Hollender. 2009. Cytochrome P450-catalysed arene-epoxidation of the bioactive tea tree oil ingredient p-cymene: Indication for the formation of a reactive allergenic intermediate? Xenobiotica, 39: 663–671. Miyazawa, M., and K. Gyoubu. 2006. Metabolism of (+)-fenchone by CYP2A6 and CYP2B6 in human liver microsomes. Biological & Pharmaceutical Bulletin, 29: 2354–2358. Miyazawa, M., and K. Gyoubu. 2007. Metabolism of (-)-fenchone by CYP2A6 and CYP2B6 in human liver microsomes. Xenobiotica, 37: 194–204. Miyazawa, M., S. Marumoto, T. Takahashi et al. 2011. Metabolism of (+)- and (-)-menthols by CYP2A6 in human liver microsomes. Journal of Oleo Science, 60: 127–132. Miyazawa, M., and K. Nakanishi. 2006. Biotransformation of (-)-menthone by human liver microsomes. Bioscience, Biotechnology, and Biochemistry, 70: 1259–1261. Miyazawa, M., and M. Shindo. 2001. Biotransformation of 1,8-cineole by human liver microsomes. Natural Product Letters, 15: 49–53. Miyazawa, M., M. Shindo, and T. Shimada. 2001a. Oxidation of 1,8-cineole, the monoterpene cyclic ether originated from Eucalyptus polybractea, by cytochrome P450 3A enzymes in rat and human liver microsomes. Drug Metabolism and Disposition, 29: 200–205. Miyazawa, M., M. Shindo, and T. Shimada. 2001b. Roles of cytochrome P450 3A enzymes in the 2-hydroxylation of 1,4-cineole, a monoterpene cyclic ether, by rat and human liver microsomes. Xenobiotica, 31: 713–723. Miyazawa, M., M. Shindo, and T. Shimada. 2002. Metabolism of (+)- and (−)-limonenes to respective carveols and perillyl alcohols by CYP2C9 and CYP2C19 in human liver microsomes. Drug Metabolism and Disposition, 30: 602–607.

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Nakahashi, H., and M. Miyazawa. 2011. Biotransformation of (-)-camphor by Salmonella typhimurium OY1002/2A6 expressing human CYP2A6 and NADPH-P450 reductase. Journal of Oleo Science, 60: 545–548. Nakahashi, H., N. Yagi, and M. Miyazawa. 2013. Biotransformation of (+)-fenchone by Salmonella typhimurium OY1002/2A6 expressing human CYP2A6 and NADPH-P450 reductase. Journal of Oleo Science, 62: 293–296. O’Neil, M. J., (eds.). 2006. The Merck Index: An Encyclopedia of Chemicals, Drugs, and Biologicals. Whitehouse Station, NJ: Wiley. Pass et al. 2002. Microsomal metabolism and enyzme kinetics of the terpene p-cymene in the common brushtail possum (Trichosurus vulpecula), koala (Phascolarctos cinereus) and rat. Xenobiotica 32: 383–397. Pisarčíková, J., V. Oceľová, Š. Faix et al. 2017. Identification and quantification of thymol metabolites in plasma, liver and duodenal wall of broiler chickens using UHPLC-ESI-QTOF-MS. Biomedical Chromatography, 31: e3881. Schmidt, L., V. N. Belov, and T. Göen. 2013. Sensitive monitoring of monoterpene metabolites in human urine using two-step derivatisation and positive chemical ionisation-tandem mass spectrometry. Analytica Chimica Acta, 793: 26–36. Schmidt, L., V. N. Belov, and T. Göen. 2015. Human metabolism of Δ3-carene and renal elimination of Δ3caren-10-carboxylic acid (chaminic acid) after oral administration. Archives of Toxicology, 89: 381–392. Schmidt, L., and T. Göen. 2017a. Human metabolism of α-pinene and metabolite kinetics after oral administration. Archives of Toxicology, 91: 677–687. Schmidt, L., and T. Göen. 2017b. R-Limonene metabolism in humans and metabolite kinetics after oral administration. Archives of Toxicology, 91: 1175–1185. Shimada, T., M. Shindo, and M. Miyazawa. 2002. Species differences in the metabolism of (+)- and (-)-limonenes and their metabolites, carveols and carvones, by cytochrome P450 enzymes in liver microsomes of mice, rats, guinea pigs, rabbits, dogs, monkeys, and humans. Drug Metabolism and Pharmacokinetics, 17: 507–515. Spatzenegger, M., and W. Jäger. 1995. Clinical importance of hepatic cytochrome P450 in drug metabolism. Drug Metabolism Reviews, 27: 397–417. Spichiger, M., R. C. Mühlbauer, and R. Brenneisen. 2004. Determination of menthol in plasma and urine of rats and humans by headspace solid phase microextraction and gas chromatography–mass spectrometry. Journal of Chromatography B, 799: 111–117. Staines, A. G., P. Sindelar, M. W. H. Coughtrie et al. 2004. Farnesol is glucuronidated in human liver, kidney and intestine in vitro, and is a novel substrate for UGT2B7 and UGT1A1. The Biochemical Journal, 384: 637–645. Sun, X.-M., Q.-F. Liao, Y.-T. Zhou et al. 2014. Simultaneous determination of borneol and its metabolite in rat plasma by GC-MS and its application to pharmacokinetics study. Journal of Pharmaceutical Analysis, 4:345–350. Thalhamer, B., W. Buchberger, and M. Waser. 2011. Identification of thymol phase I metabolites in human urine by headspace sorptive extraction combined with thermal desorption and gas chromatography mass spectrometry. Journal of Pharmaceutical and Biomedical Analysis, 56: 64–69. Zhang, R., C. Liu, T. Huang et al. 2008. In vitro characterization of borneol metabolites by GC-MS upon incubation with rat liver microsomes. Journal of Chromatographic Science, 46: 419–423. Zhao, J., Y. Lu, S. Du et al. 2012. Comparative pharmacokinetic studies of borneol in mouse plasma and brain by different administrations. Journal of Zhejiang University. Science B, 13: 990–996.

11

Central Nervous System Effects of Essential Oil Compounds Elaine Elisabetsky and Domingos S. Nunes

CONTENTS 11.1 Overview..............................................................................................................................304 11.2 Introduction.........................................................................................................................304 11.2.1 Translatability and Reproducibility.........................................................................304 11.3 Review Methodology........................................................................................................... 305 11.3.1 Literature Review..................................................................................................... 305 11.3.2 Identification of EOs and/or their Compounds with Psychopharmacologic Action.....306 11.4 Compounds from EOs with Psychopharmacology Potential..............................................306 11.4.1 Alcohols................................................................................................................... 310 11.4.2 Phenols and Aromatic Methyl Ethers...................................................................... 312 11.4.3 Hydrocarbons........................................................................................................... 313 11.4.4 Carbonyl Compounds.............................................................................................. 313 11.4.5 Monoterpene Epoxides and Furanes........................................................................ 314 11.4.6 Nitrogenated Compounds........................................................................................ 314 11.5 Compounds from EOs with Anxiolytic Properties............................................................. 315 11.5.1 Assessing Anxiolytic Properties.............................................................................. 315 11.5.1.1 Methods..................................................................................................... 316 11.5.1.2 Mechanisms of Action............................................................................... 316 11.5.2 Anxiolytic Properties of EO Compounds................................................................ 316 11.6 Hypnotic Properties of Compounds from EOs.................................................................... 319 11.6.1 Assessing Hypnotic Properties................................................................................ 319 11.6.1.1 Methods..................................................................................................... 320 11.6.1.2 Mechanisms of Action............................................................................... 320 11.6.2 Hypnotic Properties of EO Compounds.................................................................. 320 11.7 Compounds Reported to Possess Anticonvulsant Effects................................................... 322 11.7.1 Assessing Antiepileptic Properties.......................................................................... 323 11.7.1.1 Methods..................................................................................................... 323 11.7.1.2 Mechanisms of Action............................................................................... 323 11.7.2 Anticonvulsant Properties of EO Compounds......................................................... 323 11.8 Antidepressant Properties.................................................................................................... 327 11.8.1 Assessing Antidepressant Properties....................................................................... 328 11.8.1.1 Methods..................................................................................................... 328 11.8.1.2 Mechanisms of Action............................................................................... 328 11.8.2 Antidepressant Properties of OE Compounds......................................................... 328 11.9 Neuroprotector Properties of Compounds from EOs.......................................................... 330 11.9.1 Assessing Neuroprotective Properties..................................................................... 330 11.9.1.1 Methods..................................................................................................... 330 11.9.1.2 Mechanisms of Action............................................................................... 331 11.9.2 Neuroprotective Properties of EO Compounds....................................................... 331 11.10 Concluding Remarks and Perspectives................................................................................ 334 References....................................................................................................................................... 334 303

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11.1 OVERVIEW The purpose of this chapter is to provide a critical review of the scientific product in the area of psychopharmacology of essential oils (EOs). The chapter is based on the literature review of EOs reported to act as anxiolytics, hypnotics, anticonvulsants, antidepressants, and neuroprotectors, especially in experimental models. Other psychopharmacology and/or neurological effects (including analgesia, antipsychotic, hallucinogenic, and other effects), as well as pharmacokinetic and toxicology aspects are beyond the scope of this chapter and have been discussed in Chapters 10, 17, and 26. Clinical observations focusing on the effects of EOs* are informative and should be encouraged (Höferl et al., 2016; Soto-Vásquez and Alvarado-Garcia, 2018; Wang and Heinbockel, 2018), but wellsubstantiated conclusions require accurately characterized samples to allow comparisons among laboratories and sound clinical trials. In this context, it is disappointing that hundreds of clinical observations by aromatherapists and naturopaths are not well documented to facilitate the design of observational clinical studies. Although not conclusive, observational studies are of great value to inform on specific therapies and/or the design of more rigorous clinical trials (Graz et al., 2007; Graz et al., 2010; Nunes et al., 2010; Peers et al., 2014). Because EOs are complex mixtures of dozens of compounds (the very feature that makes them unique), the sample variability makes biological experiments with EOs difficult to reproduce. Without reproducibility, the robustness of results is weakened or at least questionable (Janero, 2016). Moreover, the understanding of reliable effects and its underlying mechanism of action are key features to translational research. Hoping that this review and constructive critics will optimize pharmacological innovation in EOs research, we chose to focus on the effects of compounds isolated from EOs.

11.2 INTRODUCTION 11.2.1 Translatability and Reproducibility Translatability refers to the translation of experimental results in the field of biomedicine to safe and effective therapies. Despite the academic efforts and the substantial investment of capital in drug development by the pharmaceutical industry, innovations in this area are rare (Wehling, 2009). The question of translational research is complex and has many dimensions (Mullane et al., 2014), but unavoidable issues include hypothesis-driven research, the suitability of the methods employed to answer the question in place (Goodman et al., 2016), and the proper execution of validated methods (Kimmelman et al., 2014). As in all fields of biomedical inquiry, the question of translatability is applicable EO research, and an overall evaluation of research designs and methods is needed to attain meaningful improvements in the area (Ioannidis, 2018). This review shows key features to translatability (e.g., kinetics, validated target, mechanism of action, toxicity) (Wehling, 2009) are often not consider, ultimately weakening drug development in the area. Replicability† in science has been abundantly discussed (Ioannidis, 2018). Alarming data show that, despite the rigorous methodologies common to science and scientific methods, 70% of the researchers interviewed responded that they could not replicate colleagues’ experiments (Baker, 2016). It is not uncommon that even the laboratory that originated the data cannot reproduce its own results. The crisis sets in when it was estimated that as much as 75% of biomedical research may be lost (unreproducible), which severely decreases its value and leads to huge investments by the pharmaceutical industry on projects that fail (Goodman et al., 2016). Underlying the lack of experimental reproducibility, among others, are sample size (N), false positives, biases of * Pharmacological activities of EOs and associated plant species have been reviewed elsewhere (Nunes et al., 2010; Löscher and Schmidt, 2011; Sousa, 2012; Peers et al., 2014; Baker, 2016). † Replicability has been defined as “the ability of a researcher to duplicate the results of a prior study if the same procedures are followed but new data are collected”; reproducibility “refers to the ability of a researcher to duplicate results of a prior study using the same materials as were used by the original investigator.” (Bollen et al., 2015)

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various natures, misuse of statistics, absence of adequate randomization methods, lack of blinded observation, and observations made by a single observer (Kimmelman et al., 2014; Goodman et al., 2016). Considering the time, financial resources, and number of animals involved in biomedical research, it is fair to claim ethical and moral obligations to improve the quality and reliability of preclinical studies (Peers et al., 2014). As stated by Ioannidis (Ioannidis, 2005), “… the high rate of non-replication (lack of confirmation) of research discoveries is a consequence of the convenient, yet ill-founded strategy of claiming conclusive research findings solely on the basis of a single study assessed by formal statistical significance, typically for a p-value less than 0.05.” The literature review shows that many compounds from EOs have been subjected to just one study; though they have been included in the review, its conclusions must be viewed with caution until replication is available. It is also noteworthy that some studies evaluate a given property of a compound by using one experimental model, whereas others use a battery of models. Another common limitation is the postulation of a mechanism of action based on the reversal of a given effect by an antagonist in a single dose experiment (Kenakin et al., 2014). Single dose experiments, while perfectly accepted to suggest the involvement of a target in the mechanism of action, must be further corroborated with proper evidence for a conclusive definition. As in other areas of biomedical research, more often than not, conclusion on promising results or therapeutic claims are fairly unsubstantiated.

11.3  REVIEW METHODOLOGY 11.3.1 Literature Review For this chapter, bibliographical references were collected in the online database Web of Science, since 1945 up to now (December 2018). An initial survey was done by searching in the field “issue” the following association of the keywords: “essential oils” AND “antidepressant OR antidepressive OR anxyolitic OR anxiolytic OR anticonvulsivant OR anticonvulsive OR anticonvulsant OR neuroprotector OR neuroprotection OR hypnotic OR epilepsy.” The resulting 785 reference abstracts were acquired and read to establish an initial list of 61 substances isolated from EOs reported to have pharmacological studies using experimental models for anxiolytic, hypnotic, anticonvulsant, antidepressant, and/or neuroprotector effects. A second survey followed on each of these 61 substances, this time searching in the field “issue” the association of the keywords “compound name” AND “antidepressant OR antidepressive OR anxyolitic OR anxiolytic OR anticonvulsivant OR anticonvulsive OR anticonvulsant OR neuroprotector OR neuroprotection OR hypnotic OR epilepsy.” From the resulting 875 abstracts, about 300 were selected for in-depth analysis. This analysis led us to reduce the potential initial 61 compounds to the 34 compounds discussed in this chapter. Reports of activity published as abstracts at meetings, online bulletins, or devoid of DOI and/or another main stream identification were excluded. Compounds with reported activity in the areas of interest of this chapter but reporting frail data (e.g., unusual methods and/or known methods improperly conducted, inadequate samples sizes, lacking positive and/or negative controls, unvalidated endpoint observations, unacceptable statistics) were also left out.* A brief description of the pathology, the mechanism of action of major marketed drugs, and a brief description of the mostly used methods used for evaluation are given for each pharmacology topic in order to facilitate the reader appraisal on the aspects reported for each compound.

* 1-(4-(3,5-Dibutyl-4-hydroxybenzyl) piperazin-1-yl)-2-methoxyethan-1-one, 1-nitro-2-phenylethane, 3-butylidene-4,5dihydrophtalide, α-atlantone, benzylalcohol, carvone, cedrol, citral, citronellal, cyano-carvone, dimethoxytoluene, eucalyptol, geranial, hydroxycarvone, N-isopropylanthranilate, N-methylanthranilate, methylisoeugenol, myrcene, neral, α-phellandrene, sarisan, sulcatylacetate, α-terpineol, terpinen-4-ol, α-turmerone, β-turmerone, (-)-verbenone.

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11.3.2 Identification of EOs and/or their Compounds with Psychopharmacologic Action The need for hypothesis-driven research, instead of fishing expeditions in search for a hypothesis (often based on the available results) has been thoroughly discussed (Ioannidis, 2018), as well as its implications for pharmacology and drug development (Kenakin et al., 2014). One approach in the search for psychoactive compounds from EOs is to formulate hypotheses on pharmacological properties based on the medicinal use of aromatic species in traditional systems of medicine. Buchbauer and colleagues (Buchbauer et  al., 1993) showed the effects of inhalation of several EOs and their compounds in mice mobility. Since sedative-like effects (decreased mobility) were observed in low concentrations, the data justify the efficacy of pure fragrance compounds and/or their EOs used as mild sedatives in folk medicine. Examples include coumarin for the soporific property of hay and linalool and linalyl acetate for sandalwood oil, neroli oil, and especially lavender oil. By correlating the effects on mobility with compounds concentrations retrieved in plasma, their lipophilic characters, and detection thresholds, the authors concluded that chemical structures and functional groups of the compounds are likely to determine their biological effects. Another example is the ethnopharmacology approach to search for anticonvulsants in Amazonian traditional medicine (Elisabetsky and Brum, 2003). Based on the descriptions of symptoms and diseases by healers and users among caboclo or riberinhos in the State of Pará (Brazil), a list of conditions that could refer to epilepsy-like conditions and associated home medicines was generated. The most frequently mentioned formula was the combination of the juices (obtained by mechanical pressure) from arruda (Ruta graveolens L., Rutaceae), cipó pucá (Cissus sicyoides L., Vitaceae), catinga de mulata (Aeollanthus suaveolens Mart. Ex Spreng., Lamiaceae) leaves, and a teaspoon of gergelim preto (Sesamun indicum L., Pedaliaceae) seeds. The formula, prepared as reported, slowed (but did not inhibit) pentylenetetrazole (PTZ)-induced seizures in mice. A. suaveolens, an African species never studied before, as the popular name suggests, is strongly aromatic, and its EO proved to have anticonvulsant properties (Coelho de Souza et al., 1997). The main components in the EO were (E)-βfarnesene (37.75%), δ-decene-2-lactone (20.6%–44.3%), linalyl acetate (11.32%), linalool 10.49%), and δ-decanolactone (0.37%–3.02%) (Elisabetsky and Brum, 2003). Although the study of (E)-β-farnesene is limited, we found that linalyl acetate was devoid of activity (Coelho de Souza et al., 1997) and that linalool and γ-decanolactone were active in several hypno-sedative and seizure animal models (Pereira et al., 1997). Several linalool-producing plants are or have been used in traditional medical systems for purposes evocative of central sedative effects, most notably lavender baths or inhalation. Because neurotransmitter systems are present in neuronal circuitries implicated in more than a neuropsychiatric condition, plants used traditionally for one purpose may act through a pharmacological mechanism applicable to other pharmacological areas. A plant species used to “improve sleep” generates the hypothesis of hypnotic, anxiolytic and anticonvulsant properties, as the same mechanism of action may apply to all. Antidepressants are used for analgesics and/or anxiolytic purposes, and insults to central nervous system (CNS) apparently share the commonality of generating oxidative stress and neuroinflammation (Hurley and Tizabi, 2013). Unfortunately, one finds in the literature reference to “a plant traditionally used” without detailed use or references to original field data, or even statements unsubstantiated by the original report (often the wrong information is thereafter used repetitively after the first misquotation). An accurate ethnopharmacology is obviously necessary for this approach.

11.4  COMPOUNDS FROM EOs WITH PSYCHOPHARMACOLOGY POTENTIAL The components isolated from EOs belong to several chemical classes, such as monoterpenes, sesquiterpenes, and phenylpropanoids. There are not enough studies on structure-activity relationships that would facilitate the search for new psychoactive EO compounds. In the absence of ethnopharmacological information, the major advantage of using EOs in the search for new drugs with central activity would be the availability of a wide variety of natural substances and their molecular characteristics.

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There are enormous difficulties in the way to find pharmacologically useful drugs (Lipinski, 2000), but absorption, distribution, metabolism, and excretion were proposed as simple filters in the search of active substances in large databases (Clark, 1999; Lipinski et al., 2012). Of particular relevance to this chapter is the estimative that over 98% of all small-molecule drugs do not cross the blood–brain barrier (BBB) (Pardridge, 2005). Lipinsky’s “rule of five” describes a set of requirements for a drug to be orally absorbed (Lipinski et al., 2012): (1) molecular weight, MW 0.5 0.1

Inula graveolens (L.) Desf.

0.3 ± 0.4

3.3 ± 1.1

0.5

Juniperus communis L. (1)

3.3 ± 0.4

1.3 ± 1.1

>0.5 0.5

Juniperus communis L. (2)

2.3 ± 0.4

0.5

0.3

Juniperus oxycedrus L.

6.3 ± 1.1

3.0 ± 0.7 0.0

nd

Lavandula buchii var. tolpidifolia (Svent.)(1)

7.8 ± 0.4

0.0

>0.5 0.5

Lavandula buchii var. tolpidifolia (Svent)(2)

5.7 ± 0.4

1.0 ± 0.0

0.5

0.1

Lavandula buchii var. tolpidifolia (Svent.)(3)

0.3 ± 0.4

0.5 ± 0.0 0.0

0.3

0.1

Ledum groenlandicum Oeder

6.8 ± 0.4

0.1 0.3

nd

0.5

nd

Lippia citriodora (Palau) Kunth

6.5 ± 0.7

0.5 ± 0.0

Mentha arvensis L.

0.5 ± 0.7

1.0 ± 0

>0.5 0.1

0.5

Mentha pulegium L.

5.3 ± 0.4

2.3 ± 0.4

0.3

0.1

Mentha × citrata Ehrh

2.8 ± 0.4

4.0 ± 0.7

0.5

0.5

Mentha × piperita L. Myrtus communis L. (1)

3.8 ± 0.4

1.0 ± 0.0

0.5

0.3

0.5 ± 0.7

4.0 ± 1.4

0.3

0.1

Nardostachys jatamansi (D.Don) DC.

7.0 ± 0.7

1.3 ± 0.4

Origanum compactum Benth.

0.3 ± 0.4

2.5 ± 0.0

>0.5 0.1

0.1

0.1

nd

Origanum majorana L. (1)

6.5 ± 0.7

0.5 ± 0.0

4.5 ± 0.7

0.8 ± 0.4

>0.5 0.1

0.5

Origanum majorana L. (2) Pinus ponderosa Douglas ex C.Lawson

1.3 ± 0.4

3.0 ± 0.0

0.3

0.1

Rosmarinus officinalis L.

6.0 ± 0.7

0.5 ± 0.0

0.5

Satureja montana L.

0.4 ± 0.4

2.3 ± 0.4

>0.5 0.1

Styrax benzoin Dryand

0.8 ± 0.4

3.5 ± 0.7

Thymus vulgaris L. (1) Thymus vulgaris L. (2)

3.3 ± 0.4 0.0

0.1

0.1 nd

2.3 ± 1.1

>0.5 0.1

0.1

2.3 ± 0.0

0.0

0.1

Source: Adapted from Mokhetho et al., 2018. J. Essent. Oil Res. 30 (6): 399–408. Abbreviation:  nd = not detected.

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MIC-assay revealed good antimicrobial activity with MICs  0.05) This result indicates an anti-biofilm activity of T. daenensis EO independent from hldgene expression (Sharifi et al., 2018). Bacterial strains: S. aureus isolates from respiratory patients, dairy cattle with subclinical mastitis, and milk and food samples. Using culture characteristics, biochemical tests, and PCR, isolates were identified. Results were calculated as mean of 10 isolates. Additionally, S. aureus ATCC 25923 was used as reference strain. Additionally, S. epidermidis ATCC 12228, a non-biofilm-producer strain, was used in all the experiments. EOs: EO components were determined by GC-MS. Plant material was obtained from a university in Iran; EOs were then obtained by hydro-distillation. MIC and MBC (minimal bactericidal concentration) were quantified. Experiments were carried out using subMICs. Performed assays: • Microtiter plate test (mtP) to assess activity against biofilm formation and biofilm eradication as earlier described. (Sieniawska et al., 2013) MIC/2 to MIC/16 concentration was used. • SEM: To assess antibiofilm-activity. • Quantitative real-time RT-polymerase chain reaction (PCR) of hdl (RNAIII transcript) to detect QSI: MIC/2 concentrations of EOs were used, assay was performed as described before. EO treated biofilm formed by S. aureus ATCC 25923 was used to measure hld-gene expression. • Tetrazolium-based colorimetric assay (MTT) to detect cytotoxicity. Inhibition of QS (hdl expression inhibition in S. aureus ATCC 25923) S. hortensis MIC/2. Negatively tested oil: T. daenensis 16.8.3.30  Carum copticum EO Snoussi et al. (2018) tested the effect of Carum copticum EO. C. copticum fruits are widely used in India as a spice in curry powder and are known to have several areas of application due to their antibacterial, antifungal, anti-inflammatory, anti-hypertensive, and antitussive efficacy. The authors investigated the constituents of the EO by GC-EIMS. Additionally, polyphenols of the methanolic extract were analyzed. C. copticum EO, as well as the methanolic extract, was positively tested on antibacterial, antifungal, and antioxidative activity. The main components of the EO included oxygenated monoterpenes (53.0%), with thymol accounting for the greatest share. The monoterpene hydrocarbons p-cymene and γ-terpinene together accounted for about 44% of the oil. Disc diffusion test with CV026 revealed an anti-QS activity of the EO, although the diameter of inhibition is not mentioned. Microplate assay with ATCC12472 revealed an IC50 value for violacein inhibition of 0.23 mg/mL, whereas the MIC was 0.6 mg/mL (Snoussi et al., 2018).

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EOs: Dried seeds were purchased in Jeddah, a city in Saudi Arabia. EO was obtained by hydrodistillation. GC-EIMS (gas chromatography electron ionization mass spectrometry) was performed to identify components of the oil, MIC, MBC, and MFC (minimal fungicidal concentration). Sensor strains: ATCC 12472 (microplate assay) and CV026 (disc diffusion assay) Performed assays: • DPPH assay: To measure antioxidant activity • Disc diffusion assay: As described before, slightly adjusted; 2 µL of the EO was used. • Micro-plate assay: Adjusted flask incubation assay as described before, spectrophotometric quantification of violacein, IC50 was recorded 16.8.3.31  Cinnamomum tamala oil Combined with DNase Banu et al. (2018) tested the antibiofilm and QSI potential of Cinnamomum tamala EO from India, as well as its main compounds cinnamaldehyde and linalool, on the pathogen Vibrio parahaemolyticus, which can cause food-borne infections when consuming raw sea foods. Additionally, synergistic effects of EO together with DNase and marine bacterial DNase on preformed biofilms were investigated. eDNA (environmental DNA) is seen to be as an important component of biofilms to ensure structural integrity in V. parahaemolyticus. Enzymes like DNase can therefore weaken biofilms and, as a result, can improve the efficacy of antibacterial agents by improving biofilm penetration. The authors also tested linalool and cinnamaldehyde on their preservative efficacy in prawns infected with V. parahaemolyticus and proved them to reduce bacterial load and inhibit lipid peroxidation equal to the standard food preservative sodium benzoate. Also, SEM analysis revealed that the compounds do not damage muscle tissue of the tested prawns, which makes them a possible new food preservative. Looking at biofilm disruption, results revealed a higher efficacy of the EO combined with DNases than the compounds alone to disrupt preformed biofilms or biofilm formation. C. tamala EO significantly inhibited EPS and alginate production dose-dependently (subMICs were used). Due to these findings, Banu et al. expected that the application of antibiotics in combination with EOs reducing EPS and alginate to possibly enhance penetration through biofilms and improve their efficacy that way. The similar assumption was made for eDNA inhibition, as again the combination of EO with DNases lead to reduced eDNA accumulation. Swarming and swimming was measured using subMICs (0.5% v/v) of the oil. Both types of motility were significantly inhibited in a dosedependent manner by the oil as well as by the two major compounds of the oil. As swarming motility is QS dependent, this result can be evaluated as QS inhibition. All in all, the work of this author group demonstrated multiple fields of application of C. tamala EO and an efficient way to combine EO/EO compounds in order inhibit biofilms, virulence factor production, and bacterial motility (Banu et al., 2018). Sensor strains: V. parahaemolyticus (ATCC 17802) EO: Oil was obtained C. tamala leaves collected in Almora, Uttarkhand, India. Components were identified by GC-MS. MIC was determined in the study. DNase: Commercially acquired Marine Bacterial DNase: isolated from Vibrio alginolyticus Performed assays: Biofilm assays (test agents: EO, DNases, and combinations) • Crystal violet assay: To detect young biofilm inhibition of EO as well as the effect on preformed biofilms • Visualization of biofilm inhibition by light microscopy, confocal laser scanning microcopy • eDNA staining by fluorescence microscopy: To detect DNA present in a biofilm Other assays: • Inhibition of virulence factors (test agent: EO): EPS, alginate (embedded in the EPS of biofilms) • Swarming and swimming motility assay (test agents: EO, cinnamaldehyde, and linalool)

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• Bacteriological analysis of tiger prawns (P. vannamei), including measurement of the initial bacterial load, bactericidal effect, preservative effect of linalool, cinnamaldehyde, ciprofloxacin, and sodium benzoate • SEM of linalool or cinnamaldehyde combined with DNase to prove that muscle tissue of the prawns is not damaged.

16.9  OVERVIEW OF THE RESULTS This section gives an overview of the results discussed in the Section 16.8 (see Tables 16.9 through 16.13). If oils were tested in different concentrations below their minimal inhibitory concentrations (MICs) of the sensor strains, the label “subMIC” is used. If no MIC was evaluated in the respective study or higher concentrations were used, concentrations or volumes are shown. In the case of a study not clearly stating the concentration, cells are marked by N/A. Methods used are discussed in Section 16.8.2. Results given for disc diffusion (DD) test quantify QSI by measuring colorless (but viable) cells around EO-impregnated paper discs (in mm). It is important to mention that several different concentrations of EOs were used in the studies, therefore values cannot always be compared. Additionally, some publications do not clearly state whether the diameter contains or excludes the antimicrobial diameter (in case of exceeding subMIC concentrations). Results from other assays equal the reduction of violacein/fluorescence/luminescence/β-galactosidase, depending on the used method. Most of the oils were tested in different concentrations; these tables only lists the maximal reductions at the respective concentrations of EOs. If both MQSIC and other inhibition values are published, MQSIC is cited. For cases not indicating exact values, N/A is cited.

16.9.1  Positively Tested Oils 16.9.1.1  Disc Diffusion Assay TABLE 16.9 Essential Oils Inhibiting Quorum Sensing Detected by Disc Diffusion Assay EO

Concentration

Sensor Strain

Diameter of QS Inhibition (mm)

Artemisia annua L.

100%

ATCC12472

70 ± 0.2

Artemisia dracunculus L. Carum copticum L. Chamaemelum nobile (L.) All

1 mg/mL (20 µL) 100% (2 µL) 1 mg/mL (20 µL)

ATCC 12472 CV026 ATCC 12472

0.5 ± 0.7 N/A

Cinnamomum verum J.Presl Cinnamomum verum J.Presl Cinnamomum verum J.Presl

subMICs subMICs 100 mg/mL

CV026 ATCC12472 CV026

11 12 15

Citrus limon (L.) Osbeck (1) Citrus limon (L.) Osbeck (2) Citrus reticulata Blanco (1) Citrus reticulata Blanco (2) Citrus sinensis (L.) Osbeck Coriandrum sativum L. Cuminum cyminum L.

1 mg/mL (20 µL) 1 mg/mL (20 µL) 1 mg/mL (20 µL) 1 mg/mL (20 µL) 1 mg/mL (20 µL) 100% subMICs

ATCC 12472 ATCC 12472 ATCC 12472 ATCC 12472 ATCC 12472 ATCC12472 CV026

5.5 ± 1.1 1.0 ± 0.0 4.0 ± 0.0 3.3 ± 0.4 3.3 ± 0.4 63.50 ±2.12 N/A

Cymbopogon citratus (DC.) Stapf

200 mg/mL

CV026

4.3 ± 0.4

15

Reference Mukherji and Prabhune (2014) Mokhetho et al. (2018) Snoussi et al. (2018) Mokhetho et al. (2018) Khan et al. (2009) Khan et al. (2009) Mukherji and Prabhune. (2014) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Duarte et al. (2016) Venkadesaperumal et al. (2016) Mukherji and Prabhune (2014) (Continued)

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TABLE 16.9 (Continued) Essential Oils Inhibiting Quorum Sensing Detected by Disc Diffusion Assay EO

Concentration

Sensor Strain

Diameter of QS Inhibition (mm)

Reference

Cymbopogon citratus (DC.) Stapf Cymbopogon flexuosus (Nees ex Steud.) W.Watson Cymbopogon giganteus Chiov. Cymbopogon martini (Roxb.) W.Watson Cymbopogon winterianus Jowitt Dorema aucheri Boiss. Eucalyptus citriodora Hook (China) Eucalyptus dives Schauer

1 mg/mL (20 µL)

ATCC 12472

3.0 ± 0.7

Mokhetho et al. (2018)

1 mg/mL (20 µL)

ATCC 12472

3.3 ± 0.4

1 mg/mL (20 µL)

ATCC 12472

2.0 ± 0.7

1 mg/mL (20 µL)

ATCC 12472

3.5 ± 1.4

1 mg/mL (20 µL)

ATCC 12472

2.3 ± 1.1

N/A 10 µL

CV026 ATCC 12472

N/A 8.36 ± 0.93

Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Sepahi et al. (2015) Luís et al. (2017)

1 mg/mL (20 µL)

ATCC 12472

3.0 ± 0.7

Eucalyptus globulus Labill. Eucalyptus globulus Labill.

N/A 1 mg/mL (20 µL)

ATCC12472 ATCC 12472

10 1.0 ± 0.7

Eucalyptus radiata Sieber ex DC. Eugenia caryophyllata Thunb. Ferula asafoetida L. Foeniculum vulgare Mill.

N/A

ATCC12472

20

1 mg/mL (20 µL)

ATCC 12472

3.3 ± 0.4

N/A subMICs

CV026 CV026

N/A N/A

Gaultheria procumbens L.

1 mg/mL (20 µL)

ATCC 12472

3.0 ± 1.4

Geranium robertianum L. Geranium robertianum L. Hyptis dilatata Benth. Inula graveolens (L.) Desf.

100% 100% 30/60 µL (diluted) 1 mg/mL (20 µL)

CV026+ATTC 31301 CV026+Ezf 10–20 CV31592 ATCC 12472

13 15 N/A

Juniperus communis L. Juniperus communis L. (1)

3 µL 1 mg/mL (20 µL)

CV6269 ATCC 12472

20

Juniperus communis L. (2)

1 mg/mL (20 µL)

ATCC 12472

3.0 ± 0.7

Lavandula angustifolia Mill. Lavandula angustifolia Mill. Lavandula angustifolia Mill. (France) Lavandula angustifolia Mill. (High altitude, France) Lavandula buchii Webb var. tolpidifolia Svent. (3) Lavandula buchii Webb var. tolpidifolia (Svent) M. C. León Labill. (2) Lavandula officinalis Chaix ex Vill. (Croatia)

subMICs subMICs 10 µL

CV026 ATCC12472 ATCC 12472

10 11 8.60 ± 0.56

Mokhetho et al. (2018) Sepahi et al. (2015) Venkadesaperumal et al., (2016) Mokhetho et al. (2018) Szabó et al. (2010) Szabó et al. (2010) Álvarez et al. (2015) Mokhetho et al. (2018) Kerekes et al. (2013) Mokhetho et al. (2018) Mokhetho et al. (2018) Khan et al. (2009) Khan et al. (2009) Luís et al. (2017)

10 µL

ATCC 12472

5.01 ± 0.43

Luís et al. (2017)

1 mg/mL (20 µL)

ATCC 12472

0.5 ± 0.0

1 mg/mL (20 µL)

ATCC 12472

1.0 ± 0.0

Mokhetho et al. (2018) Mokhetho et al. (2018)

10 µL

ATCC 12472

3.3 ± 1.1

1.3 ± 1.1

4.84 ± 0.08

Mokhetho et al. (2018) Luís et al. (2016) Mokhetho et al. (2018) Luís et al. (2016)

Luís et al. (2017) (Continued)

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TABLE 16.9 (Continued) Essential Oils Inhibiting Quorum Sensing Detected by Disc Diffusion Assay EO

Concentration

Sensor Strain

Diameter of QS Inhibition (mm)

Reference

Lippia citriodora (Palau) Kunth Lippia graveolens Kunt

1 mg/mL (20 µL)

ATCC 12472

0.5 ± 0.0

25 mg/mL

ATCC12472

Melaleuca alternifolia (Maiden & Betche) Cheel

N/A

ATCC 12472

5.8 ± 1.04 15–19

Mentha × citrata Ehrh

1 mg/mL (20 µL)

ATCC 12472

4.0 ± 0.7

María V. Alvarez et al. (2012) Mokhetho et al. (2018)

Mentha × piperita L. Mentha arvensis L.

1 mg/mL (20 µL)

ATCC 12472

1.0 ± 0.0

Mokhetho et al. (2018)

1 mg/mL (20 µL) subMICs subMICs

ATCC 12472

1.0 ± 0 10 11

Mokhetho et al. (2018)

Mentha piperita L. Mentha piperita L. Mentha pulegium L.

1 mg/mL (20 µL) pure EO/N/A

CV026 ATCC12472 ATCC 12472

2.3 ± 0.4

Mokhetho et al. (2018) Alvarez et al. (2014)

Khan et al. (2009) Khan et al. (2009) Mokhetho et al. (2018) Pellegrini et al. (2014) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Szabó et al. (2010)

ATCC12472

>90 ± 0.1

1 mg/mL (20 µL)

ATCC 12472

4.0 ± 1.4

Nardostachys jatamansi (D.Don) DC. Origanum compactum Benth. Origanum majorana L.

1 mg/mL (20 µL)

ATCC 12472

1.3 ± 0.4

1 mg/mL (20 µL)

ATCC 12472

2.5 ± 0.0

100% 3 µL

CV6269

20

Origanum majorana L. (1)

1 mg/mL (20 µL)

ATCC 12472

0.5 ± 0.0

Mokhetho et al. (2018)

Origanum majorana L. (2)

1 mg/mL (20 µL)

ATCC 12472

0.8 ± 0.4

Mokhetho et al. (2018)

Pinus ponderosa Douglas ex C. Lawson Piper nigrum L.

1 mg/mL (20 µL)

ATCC 12472

3.0 ± 0.0

50 µL/mL

CV026

N/A

Rosa damascena Mill.

100%

CV026+ATTC 31298

18

Mokhetho et al. (2018) Venkadesaperumal et al. (2016) Szabó et al. (2010)

Rosa damascena Mill.

100%

20

Szabó et al. (2010)

Rosmarinus officinalis L. (Tunisia) Rosmarinus officinalis L

10 µL

CV026+Ezf 10–17 ATCC 12472

8.69 ± 0.81

Luís et al. (2017)

100%

CV026+ATTC 31303

18

Szabó et al. (2010)

Rosmarinus officinalis L

100%

18

Rosmarinus officinalis L. Rosmarinus officinalis L. Salvia officinalis L.

1 mg/mL (20 µL) N/A 100%

CV026+Ezf 10–22 ATCC 12472

Salvia sclarea L.

3 µL

CV6269

>90 ± 0.1 25

Satureja montana L.

ATCC 12472

2.3 ± 0.4

Mokhetho et al. (2018)

Satureja odora (Gris.) Epl.

1 mg/mL (20 µL) 100%

ATCC12472

90 ±1

Pellegrini et al. (2014)

Schinus molle L.

100%

ATCC12472

50 ± 1.2

Pellegrini et al. (2014)

Styrax benzoin Dryand

1 mg/mL (20 µL) subMICs

ATCC 12472

Mokhetho et al. (2018)

CV026

3.5 ± 0.7 17

subMICs

ATCC12472

19

1 mg/mL (20 µL) 1 mg/mL (20 µL)

ATCC 12472 ATCC 12472

2.3 ± 1.1 2.3 ± 0.0

Minthostachys mollis (Kunth) Griseb. Myrtus communis L. (1)

Syzigium aromaticum (L.) Merr. & L.M. Perry Syzigium aromaticum (L.) Merr. & L.M. Perry Thymus vulgaris L. (1) Thymus vulgaris L. (2)

ATCC 12472 ATCC12472

0.5 ± 0.0 15–19

Szabó et al. (2010) Mokhetho et al. (2018) Alvarez et al. (2012) Pellegrini et al. (2014) Szabó et al. (2010)

Khan et al. (2009) Khan et al. (2009) Mokhetho et al. (2018) Mokhetho et al. (2018)

463

Quorum Sensing and Essential Oils

16.9.1.2  Flask Incubation Assay TABLE 16.10 Essential Oils Inhibiting Quorum Sensing Detected by Flask Incubation Assay EO Artemisia annua L. Artemisia dracunculus L.

Concentration

Sensor strain

% Inhibition

Reference

MQSIC = 0.0138%

ATCC12472

500%

Pellegrini et al. (2014)

MQSIC = 0.6 mg/mL

ATCC 12472

60.0%

Mokhetho et al. (2018)

Calamintha nepeta (L.) Savi ssp. nepeta Carum copticum L.

MQSIC = 0.2 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

MQSIC = 0.23 mg/ML

ATCC 12472

50.0%

Snoussi et al. (2018)

Cedrus atlantica (Endl.) Manetti ex Carrière Chamaemelum nobile (L.) All Cinnamomum verum J.Presl Citrus clementina Hort ex Tan Citrus limon (L.) Osbeck

MQSIC = 6.0 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

MQSIC = 0.5 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

subMICs

CV026 CV CIP 103350T

78.0% 50.0%

Kalia et al. (2015) Poli et al. (2018)

MQSIC = 0.4 mg/mL Citrus limon (L.) Osbeck MQSIC = 0.1 mg/mL (1) Citrus limon (L.) Osbeck (2) MQSIC = 0.1 mg/mL Citrus reticulata Blanco (1) MQSIC = 0.1 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

ATCC 12472

10.0%

Mokhetho et al. (2018)

ATCC 12472

10.0%

Mokhetho et al. (2018)

ATCC 12472

10.0%

Mokhetho et al. (2018)

Citrus reticulata Blanco (2)

MQSIC = 0.5 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

Citrus sinensis (L.) Osbeck MQSIC = 0.3 mg/mL Citrus sinensis L. 100%

ATCC 12472

30.0%

Mokhetho et al. (2018)

1500.0%

Szabó et al. (2010)

>90%

Duarte et al. (2016)

MQSIC = 0.1 mg/mL

Coriandrum sativum L.

5 µL/mL

CV026+Ezf 10–21 ATCC12472

Cuminum cyminum L.

50 µL/mL

CV026

>70%

Cymbopogon citratus (DC.) Stapf. Cymbopogon citratus (DC.) Stapf Cymbopogon flexuosus (Nees ex Steud.) W. Watson Cymbopogon giganteus Chiov. Cymbopogon martini (Roxb.) W. Watson

MQSIC = 0.2 mg/mL

CV CIP 103350T

50.0%

Venkadesaperumal et al. (2016) Poli et al.

MQSIC = 0.1 mg/mL

ATCC 12472

10.0%

Mokhetho et al. (2018)

MQSIC = 0.1 mg/mL

ATCC 12472

10.0%

Mokhetho et al. (2018)

MQSIC = 0.1 mg/mL

ATCC 12472

10.0%

Mokhetho et al. (2018)

MQSIC (mg/mL) = 0.1 0.5 mg/mL

ATCC 12472

50% 95%

Mokhetho et al. (2018)

Cymbopogon winterianus  MQSIC = 0.1 mg/mL Jowitt Elettaria cardamomum (L.) subMICs Maton

ATCC 12472

50.0%

Mokhetho et al. (2018)

ATCC 12472

≈75%– 80% 50.0%

Abdullah et al. (2017)

>90% 6 mm

Luís et al. (2016)

50.0%

Mokhetho et al. (2018)

50.0%

Poli et al. (2018)

Eucalyptus dives Schauer

MQSIC = 0.1 mg/mL Eucalyptus globulus Labill. 5 µL/mL Eucalyptus globulus Labill. 100% Eucalyptus globulus Labill.  MQSIC = 0.1 mg/mL Eucalyptus polybractea MQSIC = 0.05 mg/mL R.T. Baker

ATCC 12472 ATCC12472 CV026+ATTC 31300 ATCC 12472 CV CIP 103350T

Mokhetho et al. (2018) Szabó et al. (2010)

(Continued)

464

Handbook of Essential Oils

TABLE 16.10 (Continued) Essential Oils Inhibiting Quorum Sensing Detected by Flask Incubation Assay EO Eucalyptus radiata Sieber ex DC. Eugenia caryophyllata Thunb. Eugenia caryophyllata Thunb. Eugenia caryophyllata Thunb. Foeniculum vulgare Mill.

Concentration

Sensor strain

% Inhibition

Reference

5 µL/mL

ATCC12472

>90%

Luís et al. (2016)

50 mL (dil.)

CV026

80.0%

Eris and Ulusoy (2013)

50 mL (dil.)

VIR07

72.0%

Eris and Ulusoy (2013)

50 mL (dil.)

ATCC12472

39.0%

Eris and Ulusoy (2013)

50 µL/mL

75.7%

Sepahi et al. (2015)

50.0%

Poli et al.

Foeniculum vulgare Mill.

MQSIC = 0.2 mg/mL

CV026 + C6HSL CV CIP 103350T

Gaultheria procumbens L.

MQSIC = 0.1 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

Helichrysum italicum (Roth) G. Don fil. Inula graveolens (L.) Desf.

MQSIC = 0.8 mg/mL

CV CIP 103350T

50.0%

Poli et al.

MQSIC = 0.5 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.3 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.3 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

CV026 + C6HSL

50.0%

Olivero-Verbel et al. (2014)

MQSIC = 66.91 µg/mL CV026 + C6HSL

50.0%

Olivero-Verbel et al. (2014)

CV026 + C6HSL

50.0%

Olivero-Verbel et al. (2014)

MQSIC = 30.32 µg/mL CV026 + C6HSL

50.0%

Olivero-Verbel et al. (2014)

Juniperus communis L. (1) Juniperus communis L. (2) Lippia alba (Mill.) N.E.Br. ex Britton & P.Wilson (C) CT1 Lippia alba (Mill.) N.E.Br. ex Britton & P.Wilson (A) CT2 Lippia alba (Mill.) N.E.Br. ex Britton & P.Wilson (D) CT1 Lippia alba (Mill.) N.E.Br. ex Britton & P.Wilson (E) CT2 Lippia alba (Mill.) N.E.Br. ex Britton & P.Wilson (B) CT2 Lavandula angustifolia L.

|MQSIC = 2.24 µg/mL

CV026 + C6HSL

50.0%

Olivero-Verbel et al. (2014)

100%

CV026 + ATTC 31299

10 mm

Szabó et al. (2010)

Lavandula angustifolia L.

100%

CV026 + Ezf 10–18 ATCC 12472

15 mm

Szabó et al. (2010)

50.0%

Mokhetho et al. (2018)

ATCC 12472

50.0%

Mokhetho et al. (2018)

CV CIP 103350T

50.0%

Poli et al. (2018)

ATCC12472

50.0%

Pellegrini et al. (2014)

ATCC12472

>90% 50.0%

Duarte et al. (2016)

ATCC 12472 ATCC12472

>95%

Alvarez et al. (2014)

MQSIC = 0.62 µg/mL

MQSIC = 15.79 µg/mL

Lavandula buchii Webb var. MQSIC = 0.1 mg/mL tolpidifolia (Svent.) M. C. León (3) Lavandula buchii Webb MQSIC = 0.1 mg/mL var. tolpidifolia (Svent) M. C. León (2) Lavandula stoechas L. MQSIC = 0.2 mg/mL Lepechinia floribunda MQSIC = 0.0137% (Benth.) Epling Linalool 5 µL/mL Lippia citriodora (Palau) MQSIC = 0.5 mg/mL Kunth Lippia graveolens Kunt 0.13%

Mokhetho et al. (2018)

(Continued)

465

Quorum Sensing and Essential Oils

TABLE 16.10 (Continued) Essential Oils Inhibiting Quorum Sensing Detected by Flask Incubation Assay EO

Concentration

Matricaria recutita L. Matricaria recutita L. Matricaria recutita L. Melaleuca alternifolia (Maiden & Betche) Cheel

50 mL (dil.) 50 mL (dil.) 50 mL (dil.)

Mentha × citrata Ehrh

Sensor strain

% Inhibition

Reference

CV026 VIR07 ATCC12472 ATCC 12472

67.0% 65.0% 25.0% 80% 50%

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Alvarez et al. (2012)

MQSIC = 0.5 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.3 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.1 mg/mL subMIC

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.1 mg/mL

CV026 ATCC 12472

83.3% 50.0%

Husain et al. (2015) Mokhetho et al. (2018)

Mentha suaveolens Ehrh. ssp. insularis (Req.) Greuter Minthostachys mollis (Kunth) Griseb. Myrtus communis L.

MQSIC = 0.1 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

MQSIC = 0.0649%

ATCC12472

50.0%

Pellegrini et al. (2014)

MQSIC = 0.1 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

Myrtus communis L. (1)

MQSIC = 0.1 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

Origanum compactum Benth. MQSIC = 0.1 mg/mL Origanum majorana L. (1) MQSIC = 0.5 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

ATCC 12472

50.0%

Mokhetho et al. (2018)

Origanum majorana L. (2)

50.0%

Mokhetho et al. (2018)

Mentha × piperita L. Mentha arvensis L. Mentha piperita L. Mentha pulegium L.

0.125 µg/mL MQSIC = 0.21 µg/mL

MQSIC = 0.1 mg/mL

ATCC 12472

Pinus ponderosa Douglas ex C.Lawson Pinus sylvestris L. Pinus sylvestris L. Pinus sylvestris L. Piper bogotense C. DC.

MQSIC = 0.1 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

50 mL (dil.) 50 mL (dil.) 50 mL (dil.)

CV026 VIR07 CV12472

MQSIC = 93.1 µL/mL

CV026 + C6AHL

75.0% 61.0% 32.0% 50.0%

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Olivero et al. (2011)

Piper brachypodon C. DC.

MQSIC = 513.8 µL/ mL

CV026 + C6AHL

50.0%

Olivero et al. (2011)

Piper bredemeyeri J. Jacq.

MQSIC = 45.6 µL/mL subMICs

CV026 + C6AHL

50.0%

Olivero et al. (2011)

CV026 +C6HSL

>55%

CV026 VIR07 CV12472 ATCC 12472

80.0% 70.0% 43.0% 50.0%

Venkadesaperumal et al. (2016) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Mokhetho et al.

Piper nigrum L. Rosa damascena Mill. Rosa damascena Mill. Rosa damascena Mill. Rosmarinus officinalis L.

50 mL (dil.) 50 mL (dil.) 50 mL (dil.)

Rosmarinus officinalis L.

0.125 µg/mL MQSIC = 0.21 µg/mL

ATCC 12472

80% 50%

Alvarez et al. (2012)

Salvia officinalis L.

MQSIC = 0.0259%

ATCC1247

50.0%

Pellegrini et al. (2014)

Satureja montana L.

MQSIC = 0.1 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

Schinus molle L.

MQSIC = 0.005% subMICs

ATCC12472

50.0%

Pellegrini et al. (2014)

ATCC12472

78.4%

Khan et al. (2009)

MQSIC = 0.1 mg/mL

CV026 ATCC 12472

90.0% 50.0%

Myszka et al. (2016) Mokhetho et al. (2018)

MQSIC = 0.1 mg/mL

ATCC 12472

50.0%

Mokhetho et al. (2018)

MQSIC = 0.2 mg/mL

CV CIP 103350T

50.0%

Poli et al. (2018)

Syzigium aromaticum (L.) Merr. & L.M. Perry Thymus vulgaris L. Thymus vulgaris L. (1) Thymus vulgaris L. (2) Xanthoxylum armatum DC.

MQSIC = 0.5 mg/mL

subMICs

ß-Galactosidase Fluorescence Fluorescence QSIS ß-Galactosidase ß-Galactosidase ß-Galactosidase ß-Galactosidase QSIS Fluorescence Fluorescence QSIS QSIS Fluorescence Fluorescence Fluorescence Fluorescence Fluorescence Fluorescence QSIS Luminescence Fluorescence

subMICs subMIC

subMIC

50 mL

4 mg/mL 4 mg/mL 4 mg/mL 4 mg/mL 50 mL 1.2 mg/mL

1.2 mg/mL

50 mL 50 mL 1.2 mg/mL

1.2 mg/mL

1.2 mg/mL

1.2 mg/mL

1.2 mg/mL

1.2 mg/mL

50 mL N/A

1.2 mg/mL

Cinnamomum verum Cinnamomum verum

Cinnamomum verum

Cinnamomum zeylanicum Citrus reticulata Citrus reticulata Citrus reticulata Citrus reticulata Cymbopogon nardus Elettaria cardamomum

Elettaria cardamomum

Eugenia caryophyllata Juniperus communis L.alba (3) CT 2

L.alba (3) CT 2

L.alba (2) CT 2

L.alba (4) CT2

L.alba (5) CT2

L.alba (5) CT2

Lavandula angustfolia Lavandula angustifolia

Lippia alba (1) CT 1

Assay

Concentration

EO

TABLE 16.11 QS Inhibiting EOs Evaluated by Other Assays

16.9.1.3  QS Inhibiting EOs Evaluated by Other Assays

P. putida + pRK12 (C12)

E. coli + pJBA132 (C6) QSIS12 E. Coli [pSB1075] + 3-oxo-C12-HSL

P. putida + pRK12 (C12)

E. coli + pJBA132 (C6)

E.coli + pJBA132 (C6)

E. coli + pJBA132 (C6)

P. putida + pRK12 (C12)

E. coli + pJBA132 (C6) QSIS2 QSIS6

P. putida + pRK12 (C12)

ATCC27853 ATCC27853 HT5 HT5 QSIS5

E. coli [pSB401] +  C6-HSL QSIS4

PAO1 E. Coli [pSB1075] + 3-oxo-C12-HSL

Sensor Strain

65.0%

N/A N/A

44%–72%

9.0%

61%–62%

15%–84%

72%–78%

N/A N/A 1.0%

21%–22%

49.0% 50.0% 38.0% 37.0% N/A 31.0%

N/A

N/A

38.0% N/A

QS Inhibition

(Continued)

Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013) Yap et al. (2014)

Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Luciardi et al. (2016) Luciardi et al. (2016) Luciardi et al. (2016) Luciardi et al. (2016) Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013)

Yap et al. (2014)

M. Kalia et al. (2015) Yap et al. (2014)

Reference

466 Handbook of Essential Oils

QSIS Fluorescence QSIS Fluorescence Fluorescence QSIS QSIS QSIS Hdl expression Fluorescence ß-Galactosidase QSIS QSIS Fluorescence

50 mL 1.2 mg/mL

50 mL 1.2 mg/mL

1.2 mg/mL

50 mL 50 mL 50 mL subMIC 1.2 mg/mL

subMICs 50 mL 50 mL 1.2 mg/mL

Matricaria recutita Myntotachys mollis

Myrtus communis Ocotea sp.

Ocotea sp.

Pinus sylvestris Rosa damascena Salvia officinalis Satureja hortensis Swinglea glutinosa

Syzigium aromaticum Thymus vulgaris Vanilla fragrans Zingiber officinale

Assay Fluorescence

1.2 mg/mL

Concentration

Lippia alba (1) CT 1

EO

TABLE 16.11 (Continued) QS Inhibiting EOs Evaluated by Other Assays Sensor Strain

E. coli + pJBA132 (C6)

E. coli + pJBA132 (C6) PAO1 QSIS10 QSIS11

E. coli + pJBA132 (C6) QSIS8 QSIS1 QSIS9 S. aureus ATCC 25923

P. putida + pRK12 (C12)

E. coli + pJBA132 (C6) QSIS7

E. coli + pJBA132 (C6) QSIS3

56.0% N/A Fluorescence 34%–69%

N/A N/A N/A N/A 28%–73%

71%–76%

N/A 25.0%

N/A 56%–65%

18.0%

QS Inhibition

Reference

Husain et al. (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Sharifi et al. (2018) Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012)

Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012)

Jaramillo-Colorado et al. (2012)

Quorum Sensing and Essential Oils 467

468

Handbook of Essential Oils

16.9.2 Negatively Tested Oils TABLE 16.12 Essential Oils Showing no QSI Effect in the Respective Studies EO

Assay

Cymbopogon martini

DD

Cymbopogon martini Apium graveolens

DD DD

Apium graveolens Boswellia carteri

DD DD

Cananga odorata

DD

Cedrus sp. Citrus aurantium Citrus bergamia Citrus bergamia

QSIS QSIS QSIS DD

Citrus limon

DD

Citrus limon Citrus limon Citrus limonum Citrus paradisi

DD DD QSIS DD

Citrus paradisi Citrus reticulata Citrus sinensis

DD QSIS DD

Citrus sinensis Citrus sinensis

DD DD

Sensor Strain CV026 + C6AHL CV12472 CV026 + C6AHL CV12472 CV026 + C6AHL CV026 + C6AHL QSIS1 QSIS1 QSIS1 CV026 + C6AHL CV026 + C6AHL CV12472 CV6269 QSIS1 CV026 + C6AHL CV12472 QSIS1 CV026 + C6AHL CV12472 CV026 + C6AHL

Reference Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Mukherji and Prabhune (2014) Mukherji and Prabhune (2014) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Mukherji and Prabhune. (2014) Khan et al. (2009) Khan et al. (2009) Szabó et al. (2010) Eris and Ulusoy (2013) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy. (2013) Khan et al. (2009) Khan et al. (2009) Mukherji and Prabhune (2014)

Citrus sinensis

V

CV026 +ATTC 31302

Szabó et al. (2010)

Cymbopogon citratus

DD

Khan et al. (2009)

Cymbopogon citratus Cymbopogon citratus (South Africa) Cymbopogon nardus

DD DD DD

CV026 + C6AHL CV12472 ATCC 12475

Eucalyptus globulus Eucalyptus globulus

QSIS V

Eucalyptus smithii (Australia) Eucalyptus sp

DD DD

Eucalyptus sp.

DD

Eucalyptus sp. Eucalyptus staigeriana (Australia) Eugenia caryophyllus Foeniculum vulgare

DD DD V DD

Foeniculum vulgare

DD

Foeniculum vulgare Foeniculum vulgare Foeniculum vulgare Miller Jasminum officinale Juniperus communis (Bosnia) Juniperus communis

DD DD QSIS QSIS DD V

Juniperus oxycedrus

DD

CV026 + C6AHL QSIS1 CV026 + Ezf 10–19 ATCC 12472 CV026 + C6AHL CV026 + C6AHL CV12472 ATCC 12473 ATCC 12472 CV026 + C6AHL CV026 + C6AHL CV12472 CV12472 QSIS1 QSIS1 ATCC 12476 CV026 + ATTC 31302/Ezf 10–20 ATCC 12472

Khan et al. (2009) Luís et al. (2017) Mukherji and Prabhune (2014) Eris and Ulusoy (2013) Szabó et al. (2010) Luís et al. (2017) Mukherji and Prabhune (2014) Khan et al. (2009) Khan et al. (2009) Luís et al. (2017) Mokhetho et al. (2018) Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Luís et al. (2017) Szabó et al. (2010) Mokhetho et al. (2018) (Continued)

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TABLE 16.12  (Continued) Essential Oils Showing no QSI Effect in the Respective Studies EO

Assay

Juniperus oxycedrus L.alba (2) CT 2

V F

L.alba (4) CT2

F

Laurus nobilis Lavandula angustifolia

QSIS F

Lavandula angustifolia (Croatia) Lavandula buchii var. tolpidifolia (1) Lavandula buchii var. tolpidifolia (1) Ledum groenlandicum Ledum groenlandicum Lilium candium Matricaria recutica L.,

DD DD V DD V QSIS V

Melaleuca alternifolia

DD

Melissa officinalis Mentha piperita

QSIS DD

Mentha piperita Myntotachys mollis

QSIS F

Myristica fragrans

DD

Myristica fragrans Myroxylon pereirae Nardostachys jatamansi Nigella sativa Ocimum basilicum Ocimum basilicum

DD QSIS V QSIS QSIS DD

Olea europa Olea europaea

QSIS DD

Olea europaea Orange

DD V

Petroselinum crispum

DD

Petroselinum crispum Pinus sp. Punica granatum Rosmarinus off.

DD QSIS QSIS DD

Rosmarinus officinalis

DD

Rosmarinus officinalis Rosmarinus officinalis Santalum album

DD QSIS DD

Santalum album Santalum album Styrax benzoin Swinglea glutinosa

DD QSIS V F

T. daenensis

hdl expression

Sensor Strain ATCC 12473 P. putida + pRK12 (C12) P. putida + pRK12 (C12) QSIS1 E. coli [pSB401] + 3-oxo-C6ATCC 12474 ATCC 12472 ATCC 12474 ATCC 12472 ATCC 12475 QSIS1 CV026 + ATTC 31302/Ezf 10–21 CV026 + C6AHL QSIS1 CV026+ C6AHL QSIS1 P. putida + pRK12 (C12) CV026 + C6AHL CV12472 QSIS1 ATCC 12473 QSIS1 QSIS1 CV026 + C6AHL QSIS1 CV026 + C6AHL CV12472 CV026 + ATTC 31302/Ezf 10–19 CV026 + C6AHL CV12472 QSIS1 QSIS1 CV026 + C6AHL CV026 + C6AHL CV12472 QSIS1 CV026 + C6AHL CV12472 QSIS1 ATCC 12474 P. putida + pRK12 (C12) S. aureus ATCC 25923

Reference Mokhetho et al. (2018) Jaramillo-Colorado et al. (2012) Jaramillo-Colorado et al. (2012) Eris and Ulusoy (2013) Yap et al. (2014) Luís et al. (2017) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Mokhetho et al. (2018) Eris and Ulusoy (2013) Szabó et al. (2010) Mukherji and Prabhune (2014) Eris and Ulusoy (2013) Mukherji and Prabhune (2014) Eris and Ulusoy (2013) Jaramillo-Colorado et al. (2012) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Mokhetho et al. (2018) Eris and Ulusoy (2013) Eris and Ulusoy. (2013) Mukherji and Prabhune (2014) Eris and Ulusoy (2013) Khan et al. (2009) Khan et al. (2009) Szabó et al. (2010) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Mukherji and Prabhune (2014) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Mokhetho et al. (2018) Jaramillo-Colorado et al. (2012) Sharifi et al. (2018) (Continued)

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TABLE 16.12  (Continued) Essential Oils Showing no QSI Effect in the Respective Studies EO

Assay DD DD DD DD QSIS QSIS DD DD F DD DD QSIS

Thymus vulgaris Thymus vulgaris Trachyspermum ammi Trachyspermum ammi Viola odorata Vitis vinifera Zea mays Zea mays Zingiber officinale Zingiber officinale Zingiber officinale Zingiber officinale

Sensor Strain

Reference Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Khan et al. (2009) Khan et al. (2009) Jaramillo-Colorado et al. (2012) Khan et al. (2009) Khan et al. (2009) Eris and Ulusoy (2013)

CV026 + C6AHL CV12472 CV026+ C6AHL CV12472 QSIS1 QSIS1 CV026 + C6AHL CV12472 P. putida + pRK12 (C12) CV026 + C6AHL CV12472 QSIS1

Abbreviations: DD = Disc diffusion assay, V = Flask incubation assay including spectrophotometric violacein quantification, QSIS: QS-inhibitor-selector assay, β: β-galactosidase assay, F: Fluorescence-based assay, L: Luminescence-based assay, hdl-expression-assay

16.9.3  EOs Tested on the Inhibition of QS-Related Processes TABLE 16.13 Essential Oils Tested on the Inhibition of QS-Related Processes EO Cinnamomum tamala

Sensor Strain V. parahaemolyticus (ATCC 17802)

Cinnamomum verum

PAO1

Citrus reticulata EOP

ATCC27853

Citrus reticulata EOP

HT5

Citrus reticulata EOPD

ATCC27853

Citrus reticulata EOPD

HT5

Dorema aucheri

PAO1

Eugenia caryophyllata Eugenia caryophyllata Ferula asafoetida

PAO1 PCI PAO1

Positive Biofilm eDNA EPS Alginate, Swarming Swimming Biofilm, protease Pyocyanin Alginate Swarming motility Biofilm Elastase B Biofilm Elastase B Biofilm Elastase B Biofilm Elastase B Biofilm Elastase, Pyoverdine Biofilm Biofilm Elastase Pyoverdine Pyocyanin

Negative

Reference Farisa Banu et al. (2018)

M. Kalia et al. (2015)

Luciardi et al. (2016) Luciardi et al. (2016) Luciardi et al. (2016) Luciardi et al. (2016) Pyocyanin

Sepahi et al. (2015)

Biofilm

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Sepahi et al. (2015)

(Continued)

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Quorum Sensing and Essential Oils

TABLE 16.13 (Continued) Essential Oils Tested on the Inhibition of QS-Related Processes EO

Sensor Strain

Matricaria recutita Matricaria recutita Mentha piperita

PAO1 PCI PAO1

Mentha piperita

Aeromonas hydrophila

Pinus sylvestris Pinus sylvestris Rosa damascena Rosa damascena Satureja hortensis

PAO1 PCI PAO1 PCI Staphyloccus aureus

Syzigium aromaticum

PAO1

Syzigium aromaticum

Aeromonas hydrophila

Syzigium aromaticum

PAO1

Thymus daenensis

Staphylococcus aureus

Thymus vulgare

Pseudomonas fluorescens

Positive

Negative Biofilm

Biofilm Biofilm Elastase Protease Pyoverdine Pyocyanin Chitinase EPS Swarming Biofilm Protease EPS

Husain et al. (2015)

Biofilm Biofilm Biofilm Biofilm Biofilm Planctonic growth Biofilm Swarming Biofilm Rotease EPS Biofilm Elastase Protease Pyocyanin Chitinase EPS Swimming Biofilm Planktonic growth Biofilm Swimming fg1A gene-expression

Reference Eris and Ulusoy (2013) Eris and Ulusoy (2013) Husain et al. (2015)

Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Eris and Ulusoy (2013) Sharifi et al. (2018) Khan et al. (2009) Husain et al. (2013)

Husain et al. (2013)

Sharifi et al. (2018) Myszka et al. (2016)

16.10 DISCUSSION As presented before, several EOs where tested positively on QS inhibition or inhibition of QS-related processes like biofilm formation and virulence-factor production in several different sensor strains. The large variety of assays used to detect QS inhibition of EOs and deviations in their execution makes it difficult to compare the activity of the oils. To illustrate this point, the aspects of the studies complicating comparability are present here.

16.10.1 The QS Activity of EOs Can Only Be Valued Individually for the Respective Assay QS inhibitors detected by the disc diffusion assay using C. violaceum CV026 or ATCC12472 as sensor strains with an inhibition of >20 mm can be accounted as extremely sensitive. EOs belonging

472

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to this group are Artemisia annua, Coriandrum sativum, Eucalyptus radiata, Juniperus communis, Minthostachys mollis, Origanum majorana, Salvia sclarea, Satureja odora, and Schinus molle. 15–19 mm inhibition can still be valued as sensitive and was visible in the following oils: Cinnamomum verum, Coriandrum sativum, Geranium robertianum, Melaleuca alterinifolia, Rosa damascena L., Rosmarinus officinalis, Syzygium aromaticum, Cinnamomum verum, Eucalyptus globulus, Geranium robertianum, Lavandula angustifolia, and Mentha piperata EOs showed a moderate effect of 9–14 mm QSI. Repetitions emerging in different groups are due to diverging results in the studies, which is not surprising as oils often were obtained in different countries resulting in altered constituents, and varying amounts of EOs were used depending on the study. When looking at the results from the flask incubation assay, the introduction of MQSIC (concentration of EO leading to a 50% reduction of violacein production) allows a better comparability of EOs, whereas a low MQSIC reveals high QSI potential. EOs revealing a very low MQSIC of >0.1 mg/mL are different Lippia alba hemotypes, Piper bredemeyeri, Piper bogotense, Schinus molle, and Eucalyptus polybractea. MQSIC of 0.1 mg/mL was measured, for example, by thyme, oregano, myrtle, lavender, eucalyptus, and different mint oils. When comparing results of those different assays, it is visible that in some cases, EOs show a QSI activity in several assays like Cinnamomum verum which was proved to be active in the disc diffusion assay, flask incubation assay (both C. violaceum strains), ß-galactosidase-assay, and fluorescence assay (both E. coli strains). Although some oils show rather conflicting results depending on the assay. An example is oregano oil (Lippia graveolens Kunt) which was highly active in the flask incubation assay but has a rather low activity when being tested by disc diffusion assay (Alvarez et al., 2014).

16.10.2 An EO Being an Active Inhibitor of AHL-Based QS in One Strain is Not Necessarily Active in Another Strain Also Producing AHLs Due to the varying side chains of AHLs in different organisms, EOs show different activities depending on the QS system of the tested organism, which was demonstrated by Yap et al. (2014) when investigating the effect of lavender oil on either short chain or long chain AHL sensitive sensor strains. For one thing, this aspect is important to consider when conducting studies on EOs, as the respective AHLs have to be defined. This ambiguity exists, for example, in the study done by Szabo et al. (2010), as the research team did not specify the AHLs produced by E. coli ATTC 31298 and Ezf 10–17; they just referred to an unpublished work by Szegedi. This point should be considered before assuming that specific EOs already tested positively might inhibit QS in other bacteria with AHL-based QS.

16.10.3  EOs Often Not Only Inhibit QS but Also Inhibit Growth As mentioned before, the disc diffusion assay makes a clear differentiation between QS inhibition and growth inhibition, possible because a strain which is only QS inhibited is colorless and turbid whereas a clear halo indicates cell death. The assays often include concurrent cell viability tests in order to individually determine QS inhibition or EOs are used in subMIC concentrations.

16.10.4  Extraction Method and Collecting Site of the EOs Influences Test Results Constituents of EOs vary significantly depending on their production method, resulting in different QS activity, which was shown, for example, by Jaramillo-Colorado et al. (2012) when investigating clove oil either extracted by microwave-assisted hydro-distillation or merely hydro-distillation. The relevance of the collecting site regarding constituents of the EOs was illustrated when comparing different EOs from Lippia alba (Jaramillo-Colorado et al., 2012). As mentioned in Section 16.8, EOs are defined according to their extraction method, therefore it is quite relevant to include this information in publications, although this specification is often not mentioned in the discussed studies. This aspect should be considered when interpreting study results. An example is a study published by Ganesh and Vittal Rai (2015) in which they discussed the anti-QS activity of EOs. However, they used supercritical CO2 extraction to obtain the oils, which is

Quorum Sensing and Essential Oils

473

an extraction method not in accordance to the ISO definition of EOs (Ganesh and Vittal Rai, 2015). GC-MS-analysis can also give a clue on the extraction method, as oil ingredients may indicate an extraction method not being rule consistent. This was demonstrated by a study conducted by Eris and Ulusoy (2013) as they analyzed the components of the tested oils by GC-MS. The main clove oil components mentioned in the paper where the solvents propylene glycol (73.2%) and diethylene glycol (16.0%). They also found propylene glycol in chamomile oil. The team also mentioned “vanilla EO,” although vanilla does not contain an EO (Eris and Ulusoy, 2013).

16.11 CONCLUSION QS systems in different organisms vary widely; research on those pathways is especially concentrated on AHL-based QS, which may be due to the fact that many pathogenic bacteria possess LUXRIhomologous QS systems. As QS regulates a wide range of bacterial functions such as biofilm formation, virulence-factor production, or swarming motility, QS inhibitors represent a group of active substances, possibly having multifaceted effects on bacterial populations differing markedly from classical antibiotic targets. Research on EOs on QS inhibitors only started a few years ago with in vitro studies on C. violaceum. Given the fact that EOs already are well studied and are safe substances which are easy to gain, they meet important criteria for possible new drugs. In vitro studies on different sensor strains, all based on AHLs as autoinducers revealed a significant QS-inhibitory potential of many different EOs. Not only did several oils inhibit QS itself but also they inhibit QS-related processes. However, methods of detecting QS inhibition are widely, differing from each other, and therefore prevent a transparent comparison of the EOs. In summary, EOs are potent QS inhibitors in the tested sensor strains, but research on this topic is still at its beginning. As step toward clinical studies is using the preclinical infection model Caenorhabditis elegans (Husain et al., 2013; Ganesh and Vittal Rai,2016). Closer investigations are necessary to, for example, understand the exact inhibition mechanisms or to define the effective components of the oils.

16.12 ABBREVIATIONS Abbreviation

Definition

“I” in, e.g., LuxI/LasI “R” in, e.g., LuxR/LasR ABC agr AHL or HSL AI AI-2 AIP ATCC12472 ATCC31532 BHL CAI-1 CGI CqsA CqsS CT CV ATCC31592 CV026 CviIR DD EO(s)

AI synthesis protein AI receptor protein ATP-binding cassette Accessory gene regulator Acyl homoserine lactone Autoinducer V. harveyi autoinducer 2, a furanosyl borate diester Autoinducer peptide C. violaceum wild type strain C. violaceum wild type strain N-butyryl-L-homoserine-lactones V. cholerae autoinducer 1 Cell growth inhibition V. cholerae quorum-sensing autoinducer V. cholerae quorum-sensing sensor Chemotype C. violaceum strain C. violaceum mutant strain of ATCC31532 QS system in C. violaceum Disc diffusion assay Essential oil(s)

474

EPS F GC-EIMS Gfp HAI-1 HD HHL HT5 IC50 IQS IZS L LasRI LB agar LUX lux LUXRI homologous systems MBC MFC MIC MQSIC MWHD OD OdDHL PAO1 PBS PCI PQS PqsR QS QscR QSI QSIS QSIS 1 QSM RhlIR RhlR RT-PCR SEM SPME ß subMIC V VIR07 VqsR ZOI ZOT

Handbook of Essential Oils

Exopolysacharide Flourescence based assay Gas chromatography electron ionization mass spectrometry Green fluorescence protein V. harveyi autoinducer 1, an AHL Hydro-distillation N-hexanoyl-HSL “Hospital Tucuman 5,” a P. aeruginosa strain isolated from a patient Concentration of EO leading to a 50% reduction of violacein production Integrated quorum sensing signal Inhibition zone size Luminescence-based assay P. aeruginosa QS system, designated after ist elastase-regulating properties Luria Bertani agar/lysogeny broth, a growth medium for bacteria Proteins coded by luminescence genes (lux) Luminescence genes (lux = Latin for light) QS systems homologous to LUXRI-QS in e.g. V.fisheri using AHL as Ais Minimal bactericidal concentration Minimal fungicidal concentration Minimal inhibitory concentration Minimum QS inhibitory concentration; the minimal EO concentration, inhibiting violacein production of C. violaceum by at least 50% Microwave assisted hydrodestillation Dptical density N-(3-oxododecanoyl)-homoserine lactone Wild-type strain of P. aeruginosa Phosphate buffered saline, a buffer solution P. aeruginosa clinical isolate pseudomonas quinolone signal PQS receptor Quorum sensing QS-control repressor QS inhibition/QS inhibitor(s) QSI selector, a QS-detection method QS strain constructed via QSI-selector method A Quorum sensing molecule(s) P. aeruginosa QS system, designated after RhlR Protein encoded by a rhamnolipid synthase gene cluster, AI receptor of BHL Real-time polymerase chain reaction Scanning electron microscopy Solid-phase microextraction ß-galactosidase assay Concentrations below MIC Flask incubation assay C. violaceum mutant strain of ATCC12472 virulence and QS regulator Zone of inhibition = antimicrobial activity DD Zone of turbidity = QSI DD

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475

REFERENCES Abdullah, A. A., M. S. Butt, M. Shalid, Q. Huang. 2017. Evaluating the antimicrobial potential of green cardamom essential oil focusing on quorum sensing inhibition of Chromobacterium violaceum. J. Food Sci. Technol. 54 (8): 2306–2315. Adonizio, A. L., K. Downum, B. C. Bennett, K. Mathee. 2006. Anti-quorum sensing activity of medicinal plants in southern Florida. J. Ethnopharmacol. 105 (3): 427–435. Agarwal, V., L. Priyanka, V. Pruthi et  al. 2008. Prevention of Candida albicans biofilm by plant oils. Mycopathologia 165 (1): 13–19. Albuquerque, P., A. Casadevall. 2012. Quorum sensing in fungi--a Review. Med. Mycol. 50 (4): 337–345. Álvarez, F., E. Tello, K. Bauer et al. 2015. Cytotoxic and antimicrobial diterpenes from Hyptis dilatata. Curr. Bioact. Compd. 11 (3): 189–197. Alvarez, M. V., M. R. Moreira, A. Ponce. 2012. Antiquorum sensing and antimicrobial activity of natural agents with potential use in food. J. Food Saf. 32 (3): 379–387. Alvarez, M. V., L. A. Ortega-Ramirez, M. M. Gutierrez-Pacheco et al. 2014. Oregano essential oil-pectin edible films as anti-quorum sensing and food antimicrobial agents. Front. Microbiol 5 (12): 1–7. August, P. R., T. H. Grossman, C. Minor et al. 2000. Sequence analysis and functional characterization of the violacein biosynthetic pathway from Chromobacterium violaceum. J. Mol. Microbiol Biotechnol. 2 (4): 513–519. Bala, A., R. Kumar, K. Harjai. 2011. Inhibition of quorum sensing in Pseudomonas aeruginosa by azithromycin and its effectiveness in urinary tract infections." J Med. Microbiol. 60: 300–306. Banu, S. F., D. Rubini, R. Murugan et al. 2018. Exploring the antivirulent and sea food preservation efficacy of essential oils combined with DNase on Vibrio parahaemolyticus. Lwt-Food Science and Technology 95: 107–115. Bassler, L. Bonnie. 2002. Small talk: Cell-to-cell communication in bacteria. Cell 109 (4): 421–424. Bassler B. L., M. Wright, R. E. Showalter, M. R. Silverman. 1993. Intercellular signalling in Vibrio harveyi: Sequence and function of genes regulating expression of luminescence. Mol. Microbiol. 9 (4): 773–786. Bassler B. L., M. Wright, M. R. Silverman. 1994. Multiple signalling systems controlling expression of luminescence in Vibrio harveyi: Sequence and function of genes encoding a second sensory pathway. Mol. Microbiol. 13 (2): 273–286. Bauer A. W., W. M. Kirby, J. C. Sherris, M. Turck. 1966. Antibiotic susceptibility testing by a standardized single disk method. Tech. Bull. Regist. Med. Technol. 6 (3): 49–52. Blosser, R. S., K. M. Gray. 2000. Extraction of Violacein from Chromobacterium violaceum provides a new quantitative bioassay for N-acyl homoserine lactone autoinducers. J. Microbiol. Methods. 40 (1): 47–55. Bouayed, N., N. Dietrich, C. Lafforgue et al. 2016. Process-oriented review of bacterial quorum quenching for membrane biofouling mitigation in membrane bioreactors (MBRs). Membranes 6 (4): 52–58. Bryers, J. D. 2009. Medial biofilms. Biotechnol. Bioeng. 2008. 100 (1): 1–18. Burt, S. A. 2004. Essential oils: Their antibacterial properties and potential applications in foods - a review. Int. J. Food Microbiol. 94 (3): 223–253; cited from: Gracia-Valenzuela, M. H., C. Orozco-Medina, C. Molina-Maldonado et al. 2012. Antibacterial effect of essential oregano oil (Lippia berlandieri) on pathogenic bacteria of shrimp Litopenaeus vannamei. Hidrobiologica 22(3): 201–206. Campbell, N. A., J. B. Reece J. B. 2009. Biologie, 8. Aktualisierte Auflage. Verlag Pearson Studium, München, pp 280–287. ISBN 978-3-8273-7287-1, 280–87. Cao, J. G., E. A. Meighen. 1989. Purification and structural identification of an autoinducer for the luminescence system of Vibrio harveyi. J. Biol. Chem. 264 (36): 21670–21676. Chen, H., G. R. Fink. 2006. Feedback control of morphogenesis in fungi by aromatic alcohols. Gene Development. 20: 1150–1161. Chen X., S. Schauder, N. Potier et al., 2002. Structural identification of a bacterial quorum-sensing signal containing boron. Nature 415 (6871): 545–549. Cho, T., T. Aoyama, M. Toyoda et al. 2008. Farnesol as a quorum-sensing molecule in Candida albicans. Japanese Journal of Medical Mycology 49 (4): 281–286. Choo, J. H., Y. Rukayadi, J. K. Hwang et al. 2006. Inhibition of bacterial quorum sensing by vanilla extract. Lett. Appl. Microbiol. 42 (6): 637–641. Chugani, S. A., M. Whiteley, K. M. Lee et al. 2001. QscR, a modulator of quorum-sensing signal synthesis and virulence in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. U. S. A. 98 (5): 2752–2757. Cole, S. P., J. Harwood, R. Lee et al. 2004. Characterization of monospecies biofilm formation by Helicobacter pylori. J. Bacteriol. 186 (10): 3124–3132. Costerton, W. J., Z. Lewandowski, D. E. Caldwell et al. 1995. Microbial biofilms. Annu. Rev. Microbiol. 49 (1): 711–745.

476

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Dong, Y. H., L. Y. Wang, L. H. Zhang et al. 2007. Quorum-quenching microbial infections: Mechanisms and implications. Philos. Trans. R. Soc. Lond. Series B, Biol. Sci. 362 (1483): 1201–1211. Duarte, A., A. Luis, M. Oleastro, F. C. Domingues. 2016. Antioxidant properties of coriander essential oils and linalool and their potential to control Campylobacter spp. Food Control 61: 115–122. Eberhard, A. A., L. Burlingame, C. E. Eberhard et  al. 1981. Structural identification of autoinducer of Photobacterium fischeri luciferase. Biochemistry 20 (9): 2444–2449. Engebrecht, J., M. Silverman. 1984. Identification of genes and gene products necessary for bacterial bioluminescence. Proc. Natl. Acad. Sci. U. S. A. 81 (13): 4154–4158. Erez, Z., I. Steinberger-Levy, M. Shamir et al. 2017. Communication between viruses guides lysis-lysogeny decisions. Nature 541 (7638): 488–493. Eris, R., S. Ulusoy. 2013. Rose, clove, chamomile essential oils and pine turpentine inhibit quorum sensing in Chromobacterium violaceum and Pseudomonas aeruginosa. J. EO Bear. Plants 16 (2): 126–135. Feoktistova, M., P. Geserick, M. Leverkus et al. 2016. Crystal violet assay for determining viability of cultured cells. Cold Spring Harb. Protoc. 2016 (4): pdb.prot087379. doi:10.1101/pdb.prot087379. Fuqua, W. C., S. C. Winans, E. P. Greenberg. 1994. Quorum sensing in bacteria: The LuxR-Luxl family of cell density-responsive transcriptional regulators. J. Bacteriol. 176 (2), 269–275. Ganesh, P. S., R. Vittal Rai. 2015. Evaluation of anti-bacterial and anti-quorum sensing potential of essential oils extracted by supercritical CO2 method against Pseudomonas aeruginosa. J. EO Bear. Plants 18 (2): 264–275. Ganesh, P. S., R. Vittal Rai. 2016. Inhibition of quorum-sensing-controlled virulence factors of Pseudomonas aeruginosa by Murraya koenigii essential oil: A study in a Caenorhabditis elegans infectious model. J. Med. Microbiol. 65 (12): 1528–1535. Greenberg, E. P. 1997. Quorum sensing in gram-negative bacteria. ASM News. 63 (7): 371–377. Haque, S., F. Ahmad, S. A. Dar et al. 2018. Developments in strategies for quorum sensing virulence factor inhibition to combat bacterial drug resistance. Microb. Pathog. 121 (March): 293–302. Henke, J. M., B. L. Bassler. 2004. Three parallel quorum-sensing systems regulate gene expression in Vibrio harveyi. J. Bacteriol. 186 (20): 6902–6914. Hentzer, M., H. Wu, J. B. Andersen et al. 2003. Attenuation of Pseudomonas aeruginosa virulence by quorumsensing inhibitors. EMBO J. 22 (15): 3803–3815. Hogan, D. 2006. Talking to themselves: Autoregulation and quorum sensing in fungi. Eukaryot. Cell 5 (4): 613–619. Hornby, J. M., E. C. Jensen, A. D. Lisec et al. 2001. Quorum sensing in the dimorphic fungus Candida albicans is mediated by farnesol. Appl. Environ. Microbiol. 67 (7): 2982–2992. Huan Liu, S. Srinivas, X. He. 2013. Quorum sensing in Vibrio and its relevance to bacterial virulence. J. Bacteriol. Parasitol. 04 (03). doi:10.4172/2155-9597.1000172. Husain, F. M., I. Ahmad, A. Asif et al. 2013. Influence of clove oil on certain quorum-sensing-regulated functions and biofilm of Pseudomonas aeruginosa and Aeromonas hydrophila. J. Biosci. 38 (5): 835–44. Husain, F. M., I. Ahmad, M. S. Khan et al. 2015. Sub-MICs of Mentha piperita EO and menthol Inhibits AHL mediated quorum sensing and biofilm of gram-negative bacteria. Front. Microbiol. 6: 420–432. ISO/9235.2. ISO-Rule 1997, Aromatic Natural Raw Materials-Vocabulary, Geneva, Switzerland: International Standard Organisation, 2013, Standards catalogue, ISO/TC54-Essential Oils, https://www.iso.org/obp/ ui/#iso:std:iso:9235:ed-2:v1:en:term:2.11. Jaramillo-Colorado, B., J. Olivero-Verbel, E. E. Stashenko et al. 2012. Anti-quorum sensing activity of essential oils from Colombian plants. Nat. Prod. Res. 26 (12): 1075–86. Jensen, R. O., K. Winzer, S. R. Clarke et al. 2008. Differential recognition of Staphylococcus aureus quorumsensing signals depends on both extracellular loops1 and 2 of the transmembrane sensor AgrC. J. Mol. Biol., 381: 300–309. Juhas, M., L. Wiehlmann, B. Huber et al. 2004. Global regulation of quorum sensing and virulence by VqsR in Pseudomonas aeruginosa. Microbiology 150 (4): 831–841. Kalia, V. C. 2013. Quorum sensing inhibitors: An overview. Biotechnol. Adv. 31 (2): 224–245. Kalia, M., V. Kumar Yadav, P. Kumar Singh et al. 2015. Effect of cinnamon oil on quorum sensing-controlled virulence factors and biofilm formation in Pseudomonas aeruginosa. PLoS ONE 10 (8): 1–18. Kaplan, H. B., E. P. Greenberg. 1985. Diffusion of autoinducer is involved in regulation of the Vibrio fischeri luminescence system. J. Bacteriol. 163 (3): 1210–1214. Kerekes, E. B., E. Deak, M. Tako et al. 2013. Anti-biofilm forming and anti-quorum sensing activity of selected essential oils and their main components on food-related micro-organisms. J. Appl. Microbiol. 115 (4): 933–942. Khan, M. S. A., M. Zahin, S. Hasan et al. 2009. Inhibition of quorum sensing regulated bacterial functions by plant essential oils with special reference to clove oil. Lett. Appl. Microbiol. 49 (3): 354–360.

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Kim, Y.-G., J. H. Lee, G. Gwon et al. 2016. Essential oils and eugenols inhibit biofilm formation and the virulence of Escherichia coli O157:H7. Sci. Rep. 6: 1–11. Kleerebezem, M., L. E. Quadri, O. P. Kuipers et al. 1997. Quorum sensing by peptide pheromones and twocomponent signal-transduction systems in gram-positive bacteria. Mol. Microbiol. 24 (5): 895–904. Krishnan, T., W.-F. Yin, K.-G. Chan et al. 2012. Inhibition of quorum sensing-controlled virulence factor production in pseudomonas aeruginosa PAO1 by Ayurveda spice clove (Syzygium aromaticum) bud extract. Sensors 12 (4): 4016–4030. Kulkarni, R. K., K. W. Nickerson. 1980. Nutritional control of dimorphism in Ceratocystis ulmi. Exp. Mycol. 5 (2): 149–154. Le Thi, T. T., C. Prigent-Combaret, C. Dorel, P. Le Lejeune. 2001. First stages of biofilm formation: Characterization and quantification of bacterial functions involved in colonization process. Methods Enzymology 336: 152–159. Lee, J., L. Zhang. 2014. The hierarchy quorum sensing network in Pseudomonas aeruginosa. Protein Cell 6 (1): 26–41. Luciardi, M. C., M. Amparo-Blázquez, E. Cartagena et al. 2016. Mandarin essential oils inhibit quorum sensing and virulence factors of Pseudomonas aeruginosa. LWT - Food Sci. Technol. 68: 373–380. Luís, A., A. Duarte, J. Gominho et al. 2016. Chemical composition, antioxidant, antibacterial and antiquorum sensing activities of Eucalyptus globulus and Eucalyptus radiata EOs. Ind. Crops Prod. 79: 274–282. Luís, A., A. Duarte, L. Pereira et al. 2017. Chemical profiling and evaluation of antioxidant and anti-microbial properties of selected commercial essential oils: A comparative study. Medicines 4 (2): 36. Lynch, M. J., S. Swift, D. F. Kirke et al. 2002. The regulation of biofilm development by quorum sensing in Aeromonas hydrophila. Environ. Microbiol. 4 (1): 18–28. Maree, J., G. P. Kamatou, S. Gibbons et al. 2014. The application of GC-MS combined with chemometrics for the identification of antimicrobial compounds from selected commercial EOs. Chemometr. Intell. Lab. Syst. 130: 172–81. McClean, K. H., M. K. Winson, L. Fish et al. 1997. Quorum sensing and Chrornobacteriurn violaceurn: Exploitation of violacein production and inhibition for the detection of N-acyl homoserine lactones. Microbiology 143 (1997): 3703–11. McLean, R. J. C., L. S. Pierson 3rd, C. Fuqua et al. 2004. A simple screening protocol for the identification of quorum signal antagonists. J. Microbiol. Methods 58 (3): 351–360. Meighen, E. A. 1993. Bacterial bioluminescence: Organization regulation and application of the lux genes. FASEB J. 7 (11): 1016–1022. Miller-Keane Encyclopedia and Dictionary of Medicine, Nursing, and Allied Health, Seventh Edition. (2003). Retrieved from http://Medical-Dictionary.Thefreedictionary.Com/Biofilm, Accessed July 22, 2016. Mokhetho, K. C., M. Sandasi, A. Ahmad et al. 2018. Identification of potential anti-quorum sensing compounds in EOs: A gas chromatography-based metabolomics approach. J. Essent. Oil Res. 30 (6): 399–408. Morohoshi, T., M. Kato, K. Fukamachi et al. 2008. N -acylhomoserine lactone regulates violacein production in Chromobacterium violaceum type strain ATCC12472. FEMS Microbiol. Lett. 279: 124–130. Mukherji, R., A. Prabhune. 2014. Novel glycolipids synthesized using plant essential oils and their application in quorum sensing inhibition and as antibiofilm agents. Scientific World J. 2014 (3): 890709. doi:10.1155/2014/890709. Myszka, K., M. T. Schmidt, M. Majcher et  al. 2016. Inhibition of quorum sensing-related biofilm of Pseudomonas fluorescens KM121 by Thymus vulgare essential oils and its major bioactive compounds. Int. Biodeterior. Biodegrad. 114: 252–59. Nazzaro, F., F. Fratianni, L. DeMartino et  al. 2013. Effect of essential oils on pathogenic bacteria. Pharmaceuticals 6 (12): 1451–74. J. W. Hastings. Nealson, K. H., T. Platt et al. 1970. Cellular control of the synthesis and activity of the bacterial luminescent system. J. Bacteriol. 104 (1): 313–322. Oh, K. B., H. Miyazawa, T. Naito et al. 2001. Purification and characterization of an autoregulatory substance capable of regulating the morphological transition in Candida albicans. Proc. Natl. Acad. Sci. U. S. A. 98 (8): 4664–4668. Olivero, J. T., N. P. C. Pajaro, E. Stashenko. 2011. Antiquorum sensing activity of essential oils isolated from different species of the genus Piper. Vitae, 18 (1): 77–82. Olivero-Verbel, J., A. Barreto-Maya, A. Bertel-Sevilla et al. 2014. Composition, anti-quorum sensing and antimicrobial activity of essential oils from Lippia alba. Braz. J. Microbiol. 45 (3): 759–767.

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Packiavathy, I., Pitchaikani Sasikumar, S. K. Pandian, A. Veera Ravi. 2013. Prevention of quorum-sensingmediated biofilm development and virulence factors production in Vibrio spp. by curcumin. Appl. Microbiol. Biotechnol. 97 (23): 10177–10187. Parsek, M. R., E. P. Greenberg. 2005. Sociomicrobiology: The connections between quorum sensing and biofilms. Trends Microbiol. 13 (1): 27–33. Pearson, J. P., K. M. Gray, L. Passador et al. 1994. Structure of the autoinducer required for expression of Pseudomonas aeruginosa virulence genes. Proc. Natl. Acad. Sci. U. S. A. 91 (1): 197–201. Pearson, J. P., C. van Delden, B. H. Iglewski et al. 1999. Active efflux and diffusion are involved in transport of pseudomonas aeruginosa cell-to-cell signals. J. Bacteriol. 181 (4): 1203–1210. Pellegrini, M. C., M. V. Alvarez, A. G. Ponce et al. 2014. Anti-quorum sensing and antimicrobial activity of aromatic species from South America. J. Essent. Oil Res. 26 (6): 458–465. Poli, J.-P., E. Guinoiseau, D. de Rocca Serra et  al. 2018. Anti-quorum sensing activity of 12 EOs on Chromobacterium violaceum and specific action of cis-cis-p-menthenolide from Corsican Mentha Suaveolens ssp. Insularis. Molecules. 23 (9): E2125. Prouty, A. M., W. H., Schwesinger, J. S. Gum. 2002. Biofilm formation and interaction with the surfaces of gallstones by Salmonella spp. Society 70 (5): 2640–2649. Ramage, G., S. P. Saville, B. L. Wickers et al. 2002. Inhibition of Candida albicans biofilm formation by farnesol, a quorum-sensing molecule. Appl. Environ. Microbiol. 68 (11): 5459–5463. Rasmussen, T. B., T. Bjarnsholt, M. E. Skindersoe et al. 2005. Screening for quorum-sensing inhibitors (QSI) by use of a novel genetic system, the QSI selector. J. Bacteriol. 187 (5): 1799–1814. Reimmann, C., N. Ginet, L. Michel et al. 2002. Genetically programmed autoinducer destruction reduces virulence gene expression and swarming motility in Pseudomonas aeruginosa PAO1. Microbiology 148 (4): 923–932. Riedel, K., S. Schoenmann, L. Eberl. 2005. Quorum sensing in plant-associated bacteria. BioSpektrum 11: 1385–1388. Sepahi, E., S. Tarighi, F. S. Ahmadi, A. Bagheri. 2015. Inhibition of quorum sensing in Pseudomonas aeruginosa by two herbal EOs from Apiaceae family. J. Microbiol. 53 (2): 176–180. Sharifi, A., A. Mohammadzadeh, T. Salehi Zahraei, P. Mahmoodi, 2018. Antibacterial, antibiofilm and antiquorum sensing effects of Thymus daenensis and Satureja hortensis essential oils against Staphylococcus aureus isolates. J. Appl. Microbiol. 124 (2): 379–388. Sieniawska, E., R. Los, T. Baj et al. 2013. Antimicrobial efficacy of Mutellina purpurea essential oil and α-pinene against Staphylococcus epidermidis grown in planktonic and biofilm cultures. Ind. Crops Prod. 51. 152–157. Smith, R. S., B. H. Iglewski. 2003a. P. aeruginosa quorum-sensing systems and virulence. Curr. Opin. Microbiol. 6 (1): 56–60. Smith, R. S., Iglewski, B. H. 2003b. Pseudomonas aeruginosa quorum sensing as a potential antimicrobial target. J. Clin. Investig. 112 (10): 1460–1465. Snoussi, M., E. Noumi, R. Punchappady-devasya et al. 2018. Antioxidant properties and anti-quorum sensing potential of Carum copticum essential oil and phenolics against Chromobacterium violaceum. J. Food Sci. Technol. 55 (8): 2824–2832. Solano, C., M. Echeverz. 2014. Biofilm dispersion and quorum sensing. Curr. Opin. Microbiol. 18 (1): 96–104. Stevens, A. M., K. M. Dolan, E. P. Greenberg. 1994. Synergistic binding of the Vibrio fischeri LuxR transcriptional activator domain and RNA polymerase to the Lux promoter region.” Proc. Natl. Acad. Sci. U. S. A. 91 (26): 12619–12623. Surette, M. G., M. B. Miller, B. L. Bassler. 1999. Quorum sensing in Escherichia coli, Salmonella typhimurium, and Vibrio harveyi: A new family of genes responsible for autoinducer production. Proc. Natl. Acad. Sci. U. S. A. 96 (4): 1639–1644. Szabó, M. Á., G. Z. Varga, J. Hohmann et al. 2010. Inhibition of quorum-sensing signals by EOs. Phytother. Res. 24:782–786. Toole, G. O., H. B. Kaplan, R. Kolter. 2000. Biofilm formation as microbial development. Annu. Rev. Microbiol. 54:49–79. Vattem, D. A., K. Mihalik, S. H. Crixell, R. J. McLean. 2007. Dietary phytochemicals as quorum sensing inhibitors. Fitoterapia 78 (4): 302–310. Venkadesaperumal, G., S. Rucha, K. Sundar, P. K. Shetty. 2016. Anti-quorum sensing activity of spice oil nanoemulsions against food borne pathogens. LWT - Food Sci. Technol. 66. Elsevier Ltd: 225–231. Visick, K. L., J. Foster, J. Doino et al. 2000. Vibrio fischeri lux genes play an important role in colonization and development of the host light organ. J. Bacteriol. 182 (16): 4578–4586.

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Wagner-Döbler, I., V. Thie, L. Eberl et al. 2005. Discovery of complex mixtures of novel long-chain quorum sensing signals in free-living and host-associated marine alphaproteobacteria. Chembiochem 6 (12): 2195–2206. Waters, C. M., B. L. Bassler. 2005. Quorum sensing: Cell-to-cell communication in bacteria. Annu. Rev. Cell Dev. Biol. 21 (1): 319–346. Whitehead, N. A., A. M. L. Barnard, H. Slater et al. 2001. Quorum-sensing in gram-negative bacteria. FEMS Microbiol. Rev. 25 (4): 365–404. Whiteley, M., S. P. Diggle, E. P. Greenberg et al. 2017. Progress in and promise of bacterial quorum sensing research. Nature 551 (7680): 313–320. Winson, M. K., S. Swift, L. Fish et al. 1998. Construction and analysis of LuxCDABE-based plasmid sensors for investigating N-acyl homoserine lactone-mediated quorum sensing. FEMS Microbiol. Lett. 163 (2): 185–192. Yap, P. S. X., T. Krishnan, B. C. Yiap et al. 2014. Membrane disruption and anti-quorum sensing effects of synergistic interaction between Lavandula angustifolia (lavender oil) in combination with antibiotic against plasmid-conferred multi-drug-resistant Escherichia coli. J. Appl. Microbiol. 116 (5): 1119–1128. Yap, P. S. X., T. Krishnan, K. G. Khan, S. H. E. Lim. 2015. Antibacterial mode of action of Cinnamomum verum Bark EO, alone and in combination with piperacillin, against a multi-drug-resistant Escherichia coli strain. J. Microbiol. Biotechnol. 25 (8): 1299–1306. Yarwood, J. M., P. M. Schlievert. 2003. Quorum sensing in Staphylococcus infections. J. Clin. Investig. 112 (11): 1620–1625.

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Functions of Essential Oils and Natural Volatiles in Plant-Insect Interactions Robert A. Raguso

CONTENTS 17.1 Introduction........................................................................................................................... 481 17.1.1 Brief Historical Overview of Essential Oil (EO) Functional Ecology...................... 481 17.1.2 Challenges in Identifying EO Functional Roles and Selective Pressures................. 482 17.2 Aromatic Medicinal Herbs with Multiple EO Functions...................................................... 483 17.2.1 Glandular Plants of Mediterranean Biomes.............................................................. 483 17.2.2 Focal Studies on Thymus vulgaris EO Chemotypes in the Garrigue....................... 483 17.2.2.1 Biotic Interactions in a Geographic Mosaic............................................... 483 17.2.2.2 Geographic Variation and Abiotic Stress................................................... 483 17.2.2.3 Thymol in Nectar, Impacts on Bee Health.................................................484 17.3 Functional Ecological Links between EOs and Resins.........................................................484 17.3.1 Oleoresin in Conifers and Multi-product TPS Enzymes...........................................484 17.3.2 EOs, Resins, and TPS Enzymes in Angiosperms..................................................... 485 17.3.3 Resins as Resources for Nest-Making Bees.............................................................. 485 17.3.3.1 Resin Collection from Aromatic Tropical Trees......................................... 485 17.3.3.2 Resin Collection from Dalechampia and Clusia Flowers.......................... 486 17.3.4 Cistus and Lavandula: Pollinator Hubs in the Phrygana.......................................... 487 17.4 EO-Mediated Pollinator Specialization: Lessons from Aroids............................................. 488 17.4.1 Perfume-Collecting Male Orchid Bees as a Pollination Niche................................. 488 17.4.2 Scent-Driven Pollinator Modes in the Araceae......................................................... 489 References....................................................................................................................................... 491

17.1 INTRODUCTION 17.1.1 Brief Historical Overview of Essential Oil (EO) Functional Ecology Essential oils (EOs) provide a glimpse into the hidden world of plant metabolic diversity, revealed through analytical chemistry. EO diversity is manifested at all levels of organization, ranging from the contents of glandular trichomes or floral tissues within a plant, to the physiological plasticity in EO emissions induced by biotic or abiotic stresses, to heritable EO variation (chemotypes) across the geographic distribution of a given species, and (finally), to species-specific variation in EO bouquets between related species. EOs have played important historical roles in establishing the conceptual foundations of chemosystematics (Rodman et al., 1981; Adams, 1998) as well as plant defense and coevolutionary theory (Raguso et al., 2015a), and improvements in analytical and statistical methods have drawn EOs closer to the mainstream of ecological research (Kallenbach et al., 2014; Raguso et al., 2015b). More recent research has led to the rapidly growing field of herbivore-induced plant responses, in which herbivore damage may dramatically alter a plant’s constitutive EO composition, along with non-volatile chemical defenses, physical defenses and extra-floral nectar. Herbivore induction often triggers systemic release of a fundamentally different EO blend, with the potential 481

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to attract carnivores whose activities may reduce herbivore damage and thus (indirectly) constitute a form of defense (Karban and Baldwin, 1997; Dicke and Baldwin, 2010). There is a growing realization that herbivore-induced responses impact flowers and below-ground plant tissues, as well as vegetative shoots and leaves, revealing unexpected richness in chemically mediated ecological interactions across the lives of plants (Johnson et al., 2015; Papadopoulou and van Dam, 2017). The primary goal of this chapter is to outline progress in our understanding of the biological functions of EOs, with an emphasis on complex insect–plant interactions, especially pollination. This outline will focus on natural sources of EO variation, in contrast to the historical emphasis on domesticated and horticultural plants in the voluminous literature on EO chemistry, especially in flowers.

17.1.2 Challenges in Identifying EO Functional Roles and Selective Pressures Chemical ecologists face at least three major challenges in their quest to identify the natural functions of EOs in plants. The first challenge is to determine which volatile compounds are released into the headspace of living plants, where they are most likely to mediate biological interactions. Natural volatile composition and dosage are important because attraction, repellence, toxicity, growth inhibition, and other functions often are determined by quantitative relationships among EO blend components (Dötterl and Vereecken, 2010; Galen et al., 2011). This largely methodological challenge has been addressed using non-invasive headspace sorption techniques and sensitive analytical methods. These approaches now extend beyond the traditional coupled gas chromatography-mass spectrometry (GCMS) to include hyphenated GC systems (Mondello et al., 2008; Mitrevski and Marriott, 2012), direct volatile sampling via proton-transfer-reaction mass spectrometry (PTR-MS; Riffell et al., 2014; FarréArmengol et al., 2016) and tissue surface chemical mapping using matrix-assisted laser desorption/ ionization (MALDI) mass spectrometry (Kaspar et al., 2011; Shroff et al., 2015). The second major challenge for chemical ecologists is to identify the ecological roles played by EOs, which are often complex and unexpected. This challenge has deep roots in the work of early 20th-century scientists (Clements and Long, 1923; Knoll, 1926), whose clever behavioral assays decoupled EOs and other chemical stimuli from floral color, shape, and texture, challenging living pollinators to respond to experimentally modified floral traits (Borg-Karlson, 1990). A major conceptual change has come with the recognition that a full spectrum of organisms (from microbes to browsing ungulates) interact with flowers and, with it, the realization that bioassays should be extended to these organisms as well as to pollinators (Junker et al., 2013). Recent insights on the functional ecology of flowers have been gained through novel genetic tools (e.g., inbred lines, hybrids, gene silencing; Galliot et  al., 2006; Kessler et  al., 2008) and more sophisticated experimental manipulation of floral stimuli (e.g., augmenting 3D-printed flowers with EO extracts; Policha et al., 2016; Nordström et al., 2017). The third major challenge, critical to ecological and evolutionary theory, has been to measure the selective forces acting upon and shaping EOs as phenotypic traits. Progress in this area has been impaired by the difficulty of measuring and experimentally manipulating complex chemical blends as traits, extending their natural variation while gaining a clearer understanding of their inheritance. The genetic tools described above have helped substantially in this regard, when combined with untargeted approaches that measure the impacts of floral antagonists (herbivores, florivores, nectar- and pollen-thieves) and microbes as well as pollinators, typically using fruitor seed-production as the ultimate measure of fitness (Kessler et  al., 2008, 2013). It remains daunting to dissect highly complex chemical traits, such as pine oleoresins or glandular trichome exudates, which may contain tens to hundreds of related compounds resulting from interactions among several genetic loci, through the action of their encoded biosynthetic enzymes (Pichersky and Raguso, 2018). In some plant families (e.g., Asteraceae, Lamiaceae, Verbenaceae), floral and vegetative EOs may be quite similar due to shared classes of glandular trichomes covering the entire plant (Manan et al., 2016; Tissier et al., 2017). This review begins with a discussion of plants in the mint family, well known as sources of culinary spices and herbal medicines (Bozin et al., 2006).

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17.2  AROMATIC MEDICINAL HERBS WITH MULTIPLE EO FUNCTIONS 17.2.1 Glandular Plants of Mediterranean Biomes All of the world’s major biomes contain plants with medicinal properties. Of these, the Mediterranean biome, defined by cool, wet winters and hot, dry summers and bounded between 30°–45° N or S latitudes (Cowling et al., 1996), contains multitudes of aromatic plants, many of which have been cultivated for their medicinal and/or culinary properties from prehistoric times (Kantsa et al., 2015). The maquis shrublands of the Mediterranean basin show a remarkable resemblance to the South African fynbos, Californian chaparral, Chilean matorral, and Australian mallee in their physiognomy and ecology, being open, fire-adapted habitats with unusually high degrees of local endemism. Indeed, these five Mediterranean climate regions combined account for less than 3% of the world’s land surface yet contain nearly almost 20% of the world’s floristic diversity (Rundel et al., 2018). In southern Europe, the maquis transitions into a coastal scrub habitat typified by low-nutrient limestone soils dominated by aromatic, resinous shrubby plants in the Lamiaceae family, including some of the best-known sources of culinary herbs and commercially valuable EOs (e.g., Lavandula, Salvia, Rosmarinus, and Origanum). In southern France, this calcareous coastal scrub, the garrigue, is home to Thymus vulgaris, noteworthy for its qualitative variation in EO composition, manifested in a fine-scale mosaic of population-level differences perceptible to the human nose (Gouyon et al., 1986). Careful genetic crosses and GC-MS analyses led to the assignment of individual plants to six different chemical phenotypes or “chemotypes,” each dominated by a single volatile oxygenated monoterpenoid compound, with epistatic relationships dictated by their biosynthetic pathway dynamics (Vernet et al., 1986). These chemotypes include acyclic monoterpene alcohols (geraniol and linalool), followed by cyclic terpene alcohols (alpha terpineol and 4-thujanol), and phenolic terpene alcohols (thymol and carvacrol; Linhart and Thompson, 1999).

17.2.2 Focal Studies on Thymus vulgaris EO Chemotypes in the Garrigue 17.2.2.1  Biotic Interactions in a Geographic Mosaic Careful experimental studies have revealed a fascinating landscape of ecological functions and selective forces shaping the mosaic of thyme EO chemotypes across southern France. Depending on location, thyme foliage is assailed by a variety of herbivores, ranging from Arima marginata, a chrysomelid beetle specialized on aromatic plants, to more generalized invertebrates (Helix snails and Leptophyes grasshoppers) and vertebrate browsers (domesticated or feral sheep and goats) (Linhart and Thompson, 1999). Interestingly, no single EO chemotype is an ideal herbivore deterrent; linalooldominated plants are least attractive to sheep and grasshoppers but are most vulnerable to snails and beetles, whereas the phenolic chemotypes (thymol and carvacrol) are repellent to most animals and microbes but are vulnerable to grasshopper attack (Linhart and Thompson, 1995, 1999). Local differences in biotic interactions would favor balanced polymorphism in EO chemotypes, especially given that these traits are under relatively simple Mendelian genetic control. An additional level of biotic interactions involves competition between T. vulgaris and neighboring plants, particularly grasses, which could crowd or overtop these short, slowly growing plants in the open landscape of the garrigue (Tarayre et al., 1995; Linhart et al., 2005). The phenolic terpenes emerge as the most effective allelopathic agents against the germination or survival of competitor grasses such as Brachypodium phoenicoides and Bromus spp., whether tested as aqueous leachates from T. vulgaris foliage or as dilute EO distillates (Linhart et al., 2014). However, the impacts of growing beneath or alongside a canopy of thyme leaves are complicated by the positive effects that the plants may have on surrounding soil nutrients, which may counter-balance direct allelopathy due to EOs. 17.2.2.2  Geographic Variation and Abiotic Stress The biotic interactions described above would appear to favor thymol- or carvacrol-dominated EO chemotypes of Thymus vulgaris across the garrigue, with minor exceptions. Instead, one finds a

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patchwork of thyme populations in which non-phenolic chemotypes may be dominant, and the thymol and carvacrol chemotypes appear to partition microhabitat by soil type and slope aspect (Gouyon et al., 1986). Additional studies indicate that abiotic (thermal) factors associated with elevational gradients provide the most powerful check against a selective sweep by the phenolic chemotypes of thyme EOs and may explain why sharp clinal variation is maintained despite considerable gene flow (Linhart and Thompson, 1999). Thymol and carvacrol chemotypes are rare at higher elevations where winter temperatures are coldest and show demonstrably lower ability (as seedlings) to survive and regrow after early winter freezing temperatures (−10°C), in strong contrast to their superior ability to endure hot, dry summers (Amiot et al., 2005). Reciprocal transplant experiments support the association of specific thyme chemotypes with differences in thermal tolerance (Thompson et al., 2007). However, if the climate of southern France continues to warm (without explosive growth of Leptophyes grasshopper populations), the phenolic EO chemotypes of T. vulgaris would be predicted to spread beyond the lowest elevations of the garrigue. 17.2.2.3  Thymol in Nectar, Impacts on Bee Health Thymol and carvacrol are not limited to the essential oil of Thymus vulgaris. Surveys of the EOs of other Thymus species, of related Origanum and other mint-family plants reveal a complex mosaic of terpenoid-dominated EOs from Portugal (Miguel et al., 2004; Figueiredo et al., 2008) to Turkey (Tümen et al., 1995; Bagci and Başer, 2005) and north to the Baltic Sea (Mockute and Bernotiene, 2001). Despite the low solubility of the phenolic terpenes in aqueous medium, thymol has been reported as a constituent of many floral nectars, including those of Thymus capitatus and T. serpyllum, which are important nectar sources for diverse bee groups across the Mediterranean region (Petanidou and Vokou, 1993; Petanidou and Smets, 1996), as well as the honey bee (Apis mellifera) and its associated honey industry (Alissandrakis et al., 2007; Karabagias et al., 2014). Recent research has focused on the non-caloric value that thymol, eugenol, and other EO substances might provide to bees foraging in diverse floral habitats, especially those infected with internal parasites. The antimicrobial potential of thymol was noted in earlier studies of the utility of T. vulgaris EO against a spectrum of microbial agents at low dosages (Hammer et al., 1999; Rota et al., 2008) and in reduction of infection by Nosema ceranae, a microsporidian fungal pathogen of honey bees, along with increased survivorship of bees fed a thymol syrup (Costa et al., 2010). A serious problem for social bees with generalized foraging patterns (Bombus terrestris in Europe, B. impatiens in North America) is the spread of the trypanosome Crithidia bombi, an obligate gut parasite of bumble bees that is transmitted among bees through infected feces (Schmid-Hempel and Durrer, 1991). In addition to direct impacts on bee survival and reproduction, infection by C. bombi also reduces the ability of bees to learn different floral traits in association with sugar rewards (Gegear et al., 2006). One promising development is that thymol was found to inhibit the growth of C. bombi when applied in dosages comparable to their natural occurrence in T. vulgaris floral nectar (5–8 ppm), presumably by disrupting the parasite’s cell and mitochondrial membranes (PalmerYoung et al., 2016). Subsequent research indicates that the prophylactic effects of nectar thymol on C. bombi are synergized when eugenol, another widespread component of floral EOs and nectars, is present, suggesting that diversified nectar meals may help bees to mitigate trypanosome infection (Palmer-Young et al., 2017b). However, strains of thymol-resistant C. bombi may evolve quickly, even when multiple EO components are used in biocontrol (Palmer-Young et al., 2017a). We are just beginning to understand how non-sugar nectar components, including but not limited to volatile EO constituents, impact the health of bees and other flower-visiting animals (Richardson et al., 2015).

17.3  FUNCTIONAL ECOLOGICAL LINKS BETWEEN EOs AND RESINS 17.3.1  Oleoresin in Conifers and Multi-product TPS Enzymes The preceding discussion of EO antimicrobial properties and the evolution of pathogen resistance calls to mind the fundamental relationship between terpenoid EOs and the non-volatile terpenoid

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compounds present in the oleoresins of coniferous trees. In the oleoresin of pines and other conifers, volatile “turpentine” (monoterpene and sesquiterpene EO blends) helps to solubilize and mobilize non-volatile “rosin” (terpenoid resin acids and gums), which acts as both a physical and chemical defense by sealing wounds and overwhelming bark beetles and associated fungal pathogens (Paine et  al., 1997). Oleoresin is an ancient plant defense, whose prevalence in prehistoric forests is evidenced by fossilized amber deposits, whose chemical composition may still be analyzed (Pereira et al., 2009). Early biochemical studies of oleoresin components in grand fir (Abies grandis) trees revealed that individual terpene synthase enzymes and their structural genes (the tps superfamily) often are responsible for the biosynthesis of a single major product and numerous minor products (Steele et al., 1998a). It is thought that the generation of complex and variable EO blends from relatively few enzymes represents a coevolutionary response to the selective pressures exerted by bark beetles (Scolytidae), making it more difficult for them to perceive aggregation pheromones or evolve resistance to a shifting target of toxic oleoresin blends (Phillips and Croteau, 1999; Raffa, 2001). Controlled physiological assays using grand fir saplings demonstrated that wounding induces the expression of C10 (monoterpene), C15 (sesquiterpene), and C20 (diterpene acid)-related tps genes in specialized resin ducts, beginning with almost immediate monoterpene synthase activity, followed by coordinated expression of sesquiterpene and diterpene acid-related tps genes and their associated biosynthetic enzymes (Steele et al., 1998b). Thus, the EOs and non-volatile resin components of oleoresin in coniferous trees are biosynthetically and functionally linked.

17.3.2  EOs, Resins, and TPS Enzymes in Angiosperms Terpene synthases are responsible for terpenoid biosynthesis in angiosperms as well as gymnosperms, and the “one gene, multiple products” pleiotropic relationship first described for tps genes in fir trees is common to many flowering plant lineages (Chen et al., 2011). For example, the cineole synthase enzyme, responsible for the biosynthesis of 1,8-cineole (eucalyptol), produces from 7 to 10 additional monoterpenoids in Salvia officinalis (Lamiaceae), the common sage (Wise et al., 1998), in Arabidopsis thaliana (Cruciferae), a self-pollinating weed and model for genetic research (Chen et al., 2004) and across Nicotiana sect. Alatae (Solanaceae), a group of wild tobacco species from southern Brazil (Fähnrich et al., 2012). In addition to coniferous trees, many angiosperms also demonstrate links between EOs, gums, and resins in ways that have been valued since human antiquity. The family Burseraceae includes aromatic trees whose resins—frankincense, myrrh, and copal—were used as forms of currency, to honor royalty and to invoke the sacred throughout the ancient world (Pichersky and Raguso, 2018). Neotropical members of this family (Bursera and Protium) have become model systems for the study of coevolutionary “arms races,” in which specialized beetles and other herbivores drive the evolution of more complex EO blends and resins by evolving counteradaptations to simpler blends. The diversification of terpenoid EO blends in Bursera and Protium is especially pronounced among related species that share a habitat, presumably under selective pressure to avoid attracting the herbivores using the related species as host plants (Becerra, 2007; Fine et al., 2013). Molecular analyses reveal high levels of tps gene duplication in Protium, providing a likely mechanism for the diversification of EOs and resin components (Zapata and Fine, 2013).

17.3.3 Resins as Resources for Nest-Making Bees 17.3.3.1  Resin Collection from Aromatic Tropical Trees Like humans, bees also value plant resins, but for more practical reasons—to use as antimicrobial and waterproof building materials for their nests. Honey bees mix naturally collected plant waxes and resins with their own glandular secretions to produce propolis, which has been valued for its medicinal and sacred properties from ancient to modern times (Burdock, 1998; Salatino et al., 2011). Beyond honey bees, literally thousands of social and solitary bee species utilize plant resins as nest-building materials. This is especially true in tropical rainforests, where an effective bees’ nest

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should shed the constant rain, repel or exclude predators, and prevent pathogenic microbial growth (Drescher et al., 2014). Leonhardt et al. (2011) studied such a community, calculating food-web networks to explore the resin-foraging relationships between 16 species of meliponine (stingless) bees and 15 tree species in a lowland dipterocarp rainforest in Borneo. Although the resin samples were chemically diverse (1117 total compounds), only 113 of these compounds were incorporated into bee nests, featuring sesquiterpenoids and triterpenes but lacking the monoterpene and diterpene components of the original resins. Behavioral experiments in the same habitat revealed that stingless bees are attracted to the resin of a rainforest conifer, Agathis borneensis, through olfactory responses to the volatile monoterpene and sesquiterpene EO extracts of the resin (Leonhardt et al., 2010). Stingless bees appear to learn specific ratios of these volatile components, just as honey bees are conditioned on quantitative ratios in the scents of rewarding flowers (Wright and Schiestl, 2009). 17.3.3.2  Resin Collection from Dalechampia and Clusia Flowers In Central and South America, the one-sided relationship between bees and their resin sources is leveraged into a plant-pollinator mutualism by two plant genera whose flowers produce resins as floral rewards, instead of nectar (Armbruster, 1984). Many of the approximately 120 species of Dalechampia (Euphorbiaceae) secrete a floral resin of triterpene ketones and alcohols (e.g., 11a,12aEpoxi-3-oxo-D-friedooleanan-14-ene (1); de Araújo et al., 2007), which is collected for nest building by female orchid bees (euglossines), along with stingless bees and megachilid bees that may collect both resin and pollen (Armbruster et al., 2009). Like poinsettia and other spurges, Dalechampia plants combine small monoecious (single sex on the same plant) flowers with showy bracts in a pseudanthial “blossom,” which serves as the unit of pollinator attraction (Figure 17.1A). Phenotypic selection studies revealed that bees primarily utilize visual cues when foraging for Dalechampia resins. In Gabon, West Africa, D. ipomoifolia produces colorless floral resins, and its megachilid bee pollinators show foraging preferences for colorful bracts whose size is correlated with resin gland area and reward quantity, selecting for morphologically integrated flowers that optimize pollen placement onto stigmas by bees while collecting resins (Armbruster et al., 2005; Figure 17.1B). In contrast, the tropical Mexican species D. schottii has small, inconspicuous bracts but produces bluecolored resins. Female orchid bee pollinators (Euglossa sp.) ignored floral bracts of D. schottii as a proxy for reward quality and their foraging preferences resulted in direct selection on resin gland area (Bolstad et al., 2010). Phylogenetic analyses reveal that resin-based pollination is a sharedancestral condition in the genus Dalechampia, with several independent, derived shifts to pollination by EO-collecting male orchid bees (see Section 17.4.1) in species that emit monoterpenes from the stigmas of female flowers (Armbruster, 1997). The coevolutionary patterns of resin deployment in defense of leaves, developing fruits, and male flowers in conjunction with the targeting of these tissues by the larvae of specialized herbivorous butterflies suggests that resin-based plant defenses served as a pre-adaptation for the evolution of resin-based pollination in Dalechampia (Armbruster et al., 2009). The genus Clusia (Guttiferae) contains about 150 dioecious (single-sex flowers on different plants) epiphyte and tree species, in which male flowers offer pollen and resins whereas female flowers only provide resins from sterile staminodes (Armbruster, 1984; Figure 17.1C). As for Dalechampia, the primary pollinators of Clusia flowers are female orchid bees and stingless (meliponine) bees that collect floral resins for nest building across tropical America. However, the floral resins of Clusia are not terpene-based at all, consisting of poly-isoprenylated benzophenones such as grandone (2), which are also found in the fruits (Figure 17.1D), roots and leaves of Clusia trees (de Oliveira et al., 1996). Unsurprisingly, Clusia resins offer antimicrobial as well as structural benefits, with evidence for targeted toxicity against gram-staining Paenobacillus bacteria with pathogenic impacts on honey bees (Lokvam and Braddock, 1999). The phenolic resins of Clusia flowers show no obvious volatile fraction, unlike the tree-produced terpenoid resins and oleoresins described above. Instead, the visually showy flowers of Clusia produce floral EOs with diverse chemical composition (aliphatic

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FIGURE 17.1  Plants with relationships between essential oils, resins, and resin-collecting bees. Panels a–d: Neotropical plants with resin-secreting flowers. (A) Blossom of Dalechampia tiliifolia from Gamboa, Panama, with showy bracts and a glistening resin gland (center). Insert (1) shows an oxygenated triterpene component of Dalechampia resin (11a,12a-Epoxi-3-oxo-D-friedooleanan-14-ene). (B) Female Euglossa orchid bee (arrow) collecting resin—note body placement resulting in pollen transfer. (C) Clusia uvitana from La Selva biological station, Costa Rica, with resin glands at the flower’s center. (D) Maturing fruits of a Clusia tree, Incachaca, Bolivia, with resin darkening at the site of a wound (arrow). Insert (2) shows grandone, a poly-isoprenylated benzophenone component of Clusia resin. Panels e–g: Resinous, purple-flowered Mediterranean plants with sesquiterpene-dominated essential oils and prominent roles as bee-attractants in plant-pollinator networks in Greece. (E,F) Cistus x purpurea, resinous foliage and flower. Insert (3) shows Labd-13-ene-8a,15-diol, an oxygenated diterpene from Cistus creticus resin. (G) Lavandula stoechas, cultivated in central California, with a similar Mediterranean climate. (Photos by R.A. Raguso.)

and aromatic compounds as well as terpenoids), which attract stingless bees in field trials (Nogueira et  al., 2001). As with herbivore-avoidance in Bursera and Protium trees, species-specific floral EOs may enhance the fitness of co-occurring Clusia trees, in this case by promoting reproductive isolation through scent-mediated constancy by bees.

17.3.4  Cistus and Lavandula: Pollinator Hubs in the Phrygana As discussed in Section 17.2, the Mediterranean region is rich in aromatic plants and bee diversity, including numerous flowering as well as non-flowering (e.g., cypress and pine tree) sources of propolis (Potts et al., 2006). Thus, plant-pollinator relationships in this region might be expected to include chemical links to resinous plants. Many of the signature resin components of propolis in Crete and mainland Greece are labdane-type diterpenes (Popova et al., 2010), which typify the commercially valuable labdanum resin produced by Cistus creticus shrubs (Cistaceae). As in grand fir, the expression of tps genes and associated enzymes (e.g., copal-8-ol diphosphate synthase) that mediate the synthesis of labdane-related diterpenes (e.g., Labd-13-ene-8a,15-diol (3)) in Cistus is localized to glandular trichomes (vs. resin ducts) and is induced by mechanical wounding (Falara et al., 2010; Figure 17.1E). Cistus cretica is a characteristic shrub of the phrygana, the Greek equivalent of the French garrigue, where it blooms for several months and is visited by a broad spectrum of bees. Recent studies on the Greek island of Lesvos explored the complex relationships between C. cretica and 40 additional species of co-flowering plants, their “community volatilome” of 351 total

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EO compounds and 168 identified insect-pollinator species. These studies were remarkable for their community-wide reach, their unbiased statistical approach, and the resulting non-random relationships identified between insect visitation, floral EOs, and color, as coded through beeand butterfly-specific visual-perception models (Kantsa et al., 2017). Cistus cretica, along with an aromatic mint-family plant, Lavandula stoechas, emerged as dominant “hubs” in the plantpollinator network (Figure 17.1F,G), combining purple floral color with distinctive sesquiterpene EOs, dense floral displays, and extended blooming periods; all properties that underscored strong statistical relationships with native bees, especially megachilids (Kantsa et al., 2018). Lavandula stoechas provides comparable services as a cornucopian floral resource for solitary bees along the coast of southern Spain (Herrera, 1997), as does the related Lavandula latifolia in upland Iberian habitats (Herrera, 2005). Future studies should explore the connection between solitary bee pollination, resin collection, and floral attraction to sesquiterpene-dominated EOs in Cistus and Lavandula plants (Figure 17.1).

17.4  EO-MEDIATED POLLINATOR SPECIALIZATION: LESSONS FROM AROIDS 17.4.1  Perfume-Collecting Male Orchid Bees as a Pollination Niche The study of floral volatiles as pollinator attractants began over 50 years ago with examinations of highly specialized pollination systems in orchids, the most diverse family of angiosperm plants. The success of these early studies hinged on field experiments that revealed how EOs mediate the behavioral responses of pollinators, including a diverse guild of over 600 species of New World orchids pollinated by male euglossine (orchid) bees (Williams and Whitten, 1983). Neotropical orchids in several genera (e.g., Stanhopea, Gongora, Catasetum) produce floral EOs in such high quantities that they are harvested in the liquid phase by male bees (Gerlach and Schill, 1991; Eltz et al., 1999), which use EO blends collected from flowers and other sources in pheromone-like reproductive displays (Pokorny et al., 2017). Studies of the perfume blends collected by males of 15 sympatric Euglossa bee species in Panama revealed strong divergence of chemical composition, again in species-specific patterns that would enhance reproductive isolation if used by female bees for mate choice and recognition (Zimmermann et al., 2009). Thus, the reproductive biology and behavior of male orchid bees has driven the evolution of an EO-based, specialized pollination system in orchids, just as the reproductive biology of female orchid bees has driven the evolution of resinbased, specialized pollination systems in Dalechampia and Clusia plants. Not all plants pollinated by EO-seeking male orchid bees are orchids. For example, males of several Eulaema species collect trans-carvone oxide, an unusual EO component in flowers, resulting in convergent evolution among several species of Catasetum orchids along with Dalechampia spathulata (Euphorbiaceae; Whitten et  al., 1986), Unonopsis stipitata (Annonaceae) (Teichert et al., 2009), Gloxinia perennis (Gesneriaceae; Gerlach and Schill, 1991), and Anthurium and Spathiphyllum species (Araceae; Schwerdtfeger et al., 2002). These last two families—the Gesneriaceae and Araceae—provide a useful comparison when considering how EOs contribute to plant-pollinator diversification. Both families are pan-tropical and diverse, with growth forms varying from understory plants to epiphytes. Despite its high diversity in floral form and color, pollinator affinities, and a preponderance of trichomes, the Gesneriaceae or “African violet” family (160 genera, 3300 spp.) is noteworthy for its lack of floral scent, outside of Gloxinia and related genera (Möller and Clark, 2013). In contrast, the Araceae (arum family or “aroids”) show comparable diversification (126 genera, 3300 spp.) with a much greater emphasis on EO chemistry, associated with multiple evolutionary shifts between pollinator classes and from rewarding mutualisms to deception (Bröderbauer et al., 2012; Chartier et al., 2014). The following section outlines several of these transitions in the Araceae, as a model for understanding how volatile chemistry can mediate specialized pollination even when floral morphology is relatively conservative.

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17.4.2 Scent-Driven Pollinator Modes in the Araceae Pollinator diversity in aroids is surprising, given the apparent constraints in their floral bauplan. All species in this family place small, single-sex flowers (“florets”) onto a dense spike inflorescence (a “spadix”) subtended by a bract (a “spathe”), which may be colorful and, in some species, may form a chamber (Bröderbauer et al., 2012; Figure 17.2). Only a few aroids provide nectar as a floral reward, and many species present male and female floral functions that are separated in space and/ or time. When the spadix includes sterile tissue (lacking florets), this “appendix” often is involved in EO production and thermogenesis (Skubatz et al., 1996). Recent studies have used phylogenetic trait reconstruction to map the evolution of plant-pollinator interactions (Chartier et al., 2014) and insect-trapping mechanisms (i.e., within floral chambers; Bröderbauer et al., 2012) in aroids. These analyses, combined with a growing body of literature on floral volatiles and pollinator attraction in aroids, paint an intriguing picture of EO diversification associated with distinctive pollinator niches (Gibernau and Vignola, 2016). The primitive condition in aroids is inferred from basal lineages with

FIGURE 17.2  Floral diversification in the Araceae. (A) Lysichiton americanum, with yellow banner spathe, indole (4) and rove beetle pollinators on the spadix, near Seattle, Washington, US. (B) Bee-pollinated Anthurium sp. with ipsdienol (5) and (C) scarab beetle pollinated Philodendron radiatum, with p-methoxystyrene (6), both at La Selva biological station, Costa Rica. (D) The spathe of Xanthosoma mexicana (Barro Colorado Island, Panama) forms a chamber below the spadix, emitting cis-jasmone (7) and providing a mating site for cyclocephaline scarabs. (E) Giant spathe of Amorphophallus titanum opening after dusk, and (F) margin between male florets and sterile appendix (arrow), the source of fetid dimethyltrisulfide (8) and heat in Amorphophallus konjak, both cultivated at Cornell University, Ithaca, NY, US. (Photos by R.A. Raguso.)

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hermaphroditic, pollen-rewarding flowers, including extant “skunk cabbage” genera (Lysichiton and Symplocarpus) from temperate North America and East Asia. Lysichiton americanum produces a bright-yellow spathe scented with C9 and C11 aliphatic alkenes and indole (4), which attracts staphylinid (rove) beetles that mate and eat pollen while aggregating on the spadix (Brodie et al., 2018; Figure 17.2A). Indole is a widespread floral volatile with context-dependent functions, as it is present in herbivore dung and deceptive dung-mimicking flowers (Urru et al., 2011; Schiestl and Dötterl, 2012), in jasmine and other night-blooming, hawkmoth-pollinated flowers (Bischoff et al., 2015), and in Stanhopea orchids pollinated by male orchid bees in the genus Eulema (Williams and Whitten, 1983). However, in the community context of springtime temperate forests in northwestern North America, indole paired with bright yellow attracts one species of rove beetle, Pelecomalium testaceum, the pollinator of Lysichiton americanum (Brodie et al., 2018). Nearly one-third of all aroid species belong to the genus Anthurium, with the full diversity of pollinators that might be expected of a large genus (Croat, 1980). Spathes range from waxy and brilliantly colored (scarlet in the bird-pollinated A. sanguineum; Kraemer and Schmitt, 1999) to flimsy, green, and reflexed in the orchid bee-pollinated A. rubrinervium, from which bees collect EOs directly from the spadix (Hentrich et  al., 2007; Figure 17.2B). Anthurium rubrinervium produces ipsdienol (5), a monoterpene alcohol that attracts male Euglossa bees to orchids in tropical rainforests but is better known as an aggregation pheromone for scolytid bark beetles (Ips pini), a major pest in temperate pine forests (Whitten et al., 1988). Floral EOs remain poorly known for most of the approximately 1000 species of Anthurium, but the examples above illustrate a theme common to aroids and orchids: the olfactory preferences and perfume-gathering behavior of male orchid bees represent a niche dimension that can be exploited by flowers, resulting in convergent evolution of EOs, including carvone oxide for Eulema-pollinated plants and ipsdienol for Euglossa-pollinated plants. The greater antiquity of orchid bee lineages suggests that orchids and aroids evolved more recently, recruiting euglossine bees as specialized pollinators when they gained the biosynthetic capability to produce different EO blends (Ramírez et al., 2011). The evolution of monoecy in aroids is associated with protogyny (maturation of female flowers before male flowers) and the modification of the spathe into a chamber, with the capacity to trap and detain pollinators among female-phase flowers until (hours later) male flowers shed pollen onto departing insects (Chartier et al., 2014). Trap inflorescences have evolved independently in several aroid genera and play a major role in the “mating mutualism” pollination mode of the large (about 400 sp.) genus Philodendron (Bröderbauer et  al., 2012). In the best studied species, P. selloum, the large white, waxy spathe opens at dusk with a burst of fragrance and heat associated with the highest respiratory rate measured for a flowering plant (Seymour, 2001). Large dynastine scarab beetles are attracted by the scent and spathe, enter the warm floral chamber, and spend the evening mating in a warm, protected space (Gottsberger and Silberbauer-Gottsberger, 1991; Figure 17.2C,D). Although the EO blend of P. selloum is complex (over 80 compounds), three abundant and antennally sensitive volatiles (4-methoxystyrene (6), 3,4-dimethoxystyrene, and cis-jasmone (7)) are sufficient to attract the specific beetle pollinator, Erioscelis emarginata (Dötterl et al., 2012). Like orchid bees, scarab beetles are abundant, ancient, species-rich, and have well-developed chemical-communication systems using methoxylated aromatic compounds, aliphatic acyloins, and fatty acid-derived esters (Schiestl and Dötterl, 2012). Beside aroids, the mating mutualism trap flower or “love hotel” pollination syndrome includes some of the most spectacular flowers of the New World tropics, such as the Victoria water lily (Nymphaeaceae), giant Magnolia flowers (Magnoliaceae), Panama hat plants (Cyclanthaceae), and rind-like Annona flowers (Annonaceae), all of which detain scarab beetle pollinators in chambers, produce heat, and emit strong fragrances at nightfall (Gottsberger, 1990; Ervik and Knudsen, 2003). The EO components that attract these beetles are diverse (jasmones, ionones, terpenoids), interesting (methoxylated aromatics and chiral aliphatic acyloins), and sometimes novel in a floral context (4-methyl-5-vinyl thiazole), common to Caladium bicolor and four Annona species pollinated by Cyclocephala scarab beetles (Maia et al., 2012).

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Finally, aroids are well known practitioners of brood-site deception, in which the chamber inflorescence traps saprophytic flies or beetles with fetid odors that mimic feces, carrion, fungi, or rotting fruit, decaying substrates that are used by the larvae of these insects for sustenance (Urru et al., 2011; Jürgens et al., 2013). As with sexual deception, the effectiveness of brood-site deception is thought to hinge on the scarcity of quality resources, resulting in powerful selective pressure to respond to cues that reliably indicate such resources (Schiestl and Dötterl, 2012). Deceptive pollination has evolved at least five times in the Araceae, such that fecal- and urinemimicry dominate in the genus Arum (indicated by indole, skatole, cresol, and 2-heptanone) and carrion mimicry is well represented in Amorphophallus (associated with S-volatiles and thermogenesis; Kite et al., 1998; Figure 17.2E,F). Carrion mimicry in Helicodiceros muscivoros requires calliphorid (blow) flies to enter the floral trap during the female phase, which is accomplished through emitting dimethyl di- and tri-sulfides (8) as fly attractants and producing heat from the hairy, tail-like appendix, which flies land upon before walking into the chamber (Angioy et  al., 2004). Twenty years after their initial studies, Kite and Hetterscheid (2017) expanded their initial GC-MS analyses to a total of 92 species of the Old World tropical genus Amorphophallus (about 200 spp.) cultivated in a common greenhouse, extending our knowledge far beyond the spectacular A. titanum, which has become a global sensation in botanical gardens but remains poorly understood in nature (Barthlott et al., 2009). The patterns emerging from Kite and Hetterscheid’s study serve as a fitting summary of the major themes outlined in this chapter: (1) not all species of Amorphophallus produce fetid odors, as EO composition (and humanperceived odor quality) varies markedly across the genus with geography, rather than presenting variations on a theme of gaseous sulfur malodor; (2) EOs often co-vary with inflorescence color, with carrion mimicry associated with sulfides and darker (blood red) colors; (3) species sharing distinctive odorants (e.g., trimethylamine, the odor of rotting fish) recur in several clades and are not closest relatives; and (4) conversely, closely related species with shared geographic distributions differ markedly in floral scent and other characteristics, suggesting selection to reduce pollinator competition and/or enhance reproductive isolation through pollinator specialization. The lessons learned from the EOs of aroids reinforce the larger themes running through this chapter and highlight that competing selective forces—mating success, reproductive isolation, avoidance of predators and pathogens, abiotic stress, and community context—shape the ecological functions of plant essential oils.

REFERENCES Adams, R. P. 1998. The leaf essential oils and chemotaxonomy of Juniperus sect. Juniperus. Biochem. Syst. Ecol., 26: 637–645. Alissandrakis, E., P. A. Tarantilis, P. C. Harizanis, and M. Polissiou. 2007. Comparison of the volatile composition in thyme honeys from several origins in Greece. J. Agric. Food Chem., 55: 8152–8157. Amiot, J., Y. Salmon, C. Collin, and J. D. Thompson. 2005. Differential resistance to freezing and spatial distribution in a chemically polymorphic plant Thymus vulgaris. Ecol. Lett., 8: 370–377. Angioy, A. M., M. C. Stensmyr, I. Urru, M. Puliafito, I. Collu, and B. S. Hansson. 2004. Function of the heater: The dead horse arum revisited. Proc. R. Soc. B., 271: S13–S15. de Araújo, M. R. S., M. A. S. Lima, and E. R. Silveira. 2007. Triterpenes and steroids of Dalechampia pernambucensis Baill. Biochem. Syst. Ecol., 35: 311–313. Armbruster, W. S. 1984. The role of resin in angiosperm pollination: Ecological and chemical considerations. Am. J. Bot., 71: 1149–1160. Armbruster, W. S. 1997. Exaptations link evolution of plant–herbivore and plant–pollinator interactions: A phylogenetic inquiry. Ecology, 78: 1661–1672. Armbruster, W. S., L. Antonsen, and C. Pélabon. 2005. Phenotypic selection on Dalechampia blossoms: Honest signaling affects pollination success. Ecology, 86: 3323–3333. Armbruster, W. S., J. Lee, and B. G. Baldwin. 2009. Macroevolutionary patterns of defense and pollination in Dalechampia vines: Adaptation, exaptation, and evolutionary novelty. Proc. Natl Acad. Sci. USA, 106: 18085–18090.

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Bagci, E., and K. H. C. Başer. 2005. Study of the essential oils of Thymus haussknechtii Velen and Thymus kotschyanus Boiss. et Hohen var. kotschyanus (Lamiaceae) taxa from the eastern Anatolian region in Turkey. Flavour Fragr. J., 20: 199–202. Barthlott, W., J. Szarzynski, P. Vlek, W. Lobin, and N. Korotkova. 2009. A torch in the rain forest: Thermogenesis of the titan arum (Amorphophallus titanum). Plant Biol., 11: 499–505. Becerra, J. X. 2007. The impact of herbivore–plant coevolution on plant community structure. Proc. Natl Acad. Sci. USA, 104: 7483–7488. Bischoff, M., R. A. Raguso, A. Jürgens, and D. R. Campbell. 2015. Context-dependent reproductive isolation mediated by floral scent and color. Evolution, 69: 1–13. Bolstad, G. H., W. S. Armbruster, C. Pélabon, R. Pérez-Barrales, and T. F. Hansen. 2010. Direct selection at the blossom level on floral reward by pollinators in a natural population of Dalechampia schottii: Fulldisclosure honesty? New Phytol., 188: 370–384. Borg-Karlson, A. K. 1990. Chemical and ethological studies of pollination in the genus Ophrys (Orchidaceae). Phytochemistry, 29: 1359–1387. Bozin, B., N. Mimica-Dukic, N. Simin, and G. Anackov. 2006. Characterization of the volatile composition of essential oils of some Lamiaceae spices and the antimicrobial and antioxidant activities of the entire oils. J. Agric. Food Chem., 54: 1822–1828. Brodie, B. S., A. Renyard, R. Gries, H. Zhai, S. Ogilvie, J. Avery, and G. Gries. 2018. Identification and field testing of floral odorants that attract the rove beetle Pelecomalium testaceum (Mannerheim) to skunk cabbage, Lysichiton americanus (L.). Arthropod-Plant Inte., 12: 591–599. Bröderbauer, D., A. Diaz, and A. Weber. 2012. Reconstructing the origin and elaboration of insect-trapping inflorescences in the Araceae. Am. J. Bot., 99: 1666–1679. Burdock, G. A. 1998. Review of the biological properties and toxicity of bee propolis (propolis). Food Chem. Toxicol., 36: 347–363. Chartier, M., M. Gibernau, and S. S. Renner. 2014. The evolution of pollinator–plant interaction types in the Araceae. Evolution, 68: 1533–1543. Chen, F., D. K. Ro, J. Petri, J. Gershenzon, J. Bohlmann, E. Pichersky, and D. Tholl. 2004. Characterization of a root-specific Arabidopsis terpene synthase responsible for the formation of the volatile monoterpene 1, 8–cineole. Plant Physiol., 135: 1956–1966. Chen, F., D. Tholl, J. Bohlmann, and E. Pichersky. 2011. The family of terpene synthases in plants: A mid-size family of genes for specialized metabolism that is highly diversified throughout the kingdom. Plant J., 66: 212–229. Clements, F. E., and F. L. Long. 1923. Experimental Pollination: An Outline of the Ecology of Flowers and Insects (No. 336). Carnegie Institution of Washington. Costa, C., M. Lodesani, and L. Maistrello. 2010. Effect of thymol and resveratrol administered with candy or syrup on the development of Nosema ceranae and on the longevity of honeybees (Apis mellifera L.) in laboratory conditions. Apidologie, 41: 141–150. Cowling, R. M., P. W. Rundel, B. B. Lamont, M. K. Arroyo, and M. Arianoutsou. 1996. Plant diversity in Mediterranean-climate regions. Trends Ecol. Evol., 11: 362–366. Croat, T. B. 1980. Flowering behavior of the neotropical genus Anthurium (Araceae). Am. J. Bot., 67: 888–904. Dicke, M., and I. T. Baldwin. 2010. The evolutionary context for herbivore-induced plant volatiles: Beyond the “cry for help”. Trends Plant Sci., 15: 167–175. Dötterl, S., A. David, W. Boland, I. Silberbauer-Gottsberger, and G. Gottsberger. 2012. Evidence for behavioral attractiveness of methoxylated aromatics in a dynastid scarab beetle-pollinated Araceae. J. Chem. Ecol., 38: 1539–1543. Dötterl, S., and N. J. Vereecken. 2010. The chemical ecology and evolution of bee–flower interactions: A review and perspectives. Can. J. Zool., 88: 668–697. Drescher, N., H. M. Wallace, M. Katouli, C. F. Massaro, and S. D. Leonhardt. 2014. Diversity matters: How bees benefit from different resin sources. Oecologia, 176: 943–953. Eltz, T., W. M. Whitten, D. W. Roubik, and K. E. Linsenmair. 1999. Fragrance collection, storage, and accumulation by individual male orchid bees. J. Chem. Ecol., 25: 157–176. Ervik, F., and J. T. Knudsen. 2003. Water lilies and scarabs: Faithful partners for 100 million years? Biol. J. Linn. Soc., 80: 539–543. Fähnrich, A., A. Brosemann, L. Teske, M. Neumann, and B. Piechulla. 2012. Synthesis of “cineole cassette” monoterpenes in Nicotiana section Alatae: Gene isolation, expression, functional characterization and phylogenetic analysis. Plant Mol. Biol., 79: 537–553. Falara, V., E. Pichersky, and A. K. Kanellis. 2010. A copal-8-ol diphosphate synthase from the angiosperm Cistus creticus subsp. creticus is a putative key enzyme for the formation of pharmacologically active, oxygen-containing labdane-type diterpenes. Plant Physiol., 154: 301–310.

Functions of Essential Oils and Natural Volatiles in Plant-Insect Interactions

493

Farré-Armengol, G., J. Peñuelas, T. Li, P. Yli-Pirilä, I. Filella, J. Llusia, and J. D. Blande. 2016. Ozone degrades floral scent and reduces pollinator attraction to flowers. New Phytol., 209: 152–160. Figueiredo, A. C., J. G. Barroso, L. G., Pedro, L. Salgueiro, M. G. Miguel, and M. L. Faleiro. 2008. Portuguese Thymbra and Thymus species volatiles: Chemical composition and biological activities. Curr. Pharm. Des., 14: 3120–3140. Fine, P. V. A., M. R. Metz, J. Lokvam et al. 2013. Insect herbivores, chemical innovation, and the evolution of habitat specialization in Amazonian trees. Ecology, 94: 1764–1775. Galen, C., R. Kaczorowski, S. L. Todd, J. Geib, and R. A. Raguso. 2011. Dosage-dependent impacts of a floral volatile compound on pollinators, larcenists, and the potential for floral evolution in the alpine skypilot Polemonium viscosum. Am. Nat., 177: 258–272. Galliot, C., J. Stuurman, and C. Kuhlemeier. 2006. The genetic dissection of floral pollination syndromes. Curr. Opin. Plant Biol., 9: 78–82. Gegear, R. J., M. C. Otterstatter, and J. D. Thompson. 2006. Bumble-bee foragers infected by a gut parasite have an impaired ability to utilize floral information. Proc. R. Soc. B, 273: 1073–1078. Gerlach, G., and R. Schill. 1991. Composition of orchid scents attracting euglossine bees. Bot. Acta, 104: 385–391. Gibernau, M., and R. D. S. Vignola. 2016. Pollinators and visitors of aroid inflorescences III–phylogenetic & chemical insights. Aroideana, 39: 4–22. Gottsberger, G. 1990. Flowers and beetles in the South American tropics. Bot. Acta, 103: 360–365. Gottsberger, G., and I. Silberbauer-Gottsberger. 1991. Olfactory and visual attraction of Erioscelis emarginata (Cyclocephalini, Dynastinae) to the inflorescences of Philodendron selloum (Araceae). Biotropica, 23: 23–28. Gouyon, P. H., P. Vernet, J. L. Guillerm, and G. Valdeyron. 1986. Polymorphisms and environment: The adaptive value of the oil polymorphisms in Thymus vulgaris L. Heredity, 57: 59–66. Hammer, K. A., C. F. Carson, and T. V. Riley. 1999. Antimicrobial activity of essential oils and other plant extracts. J. Appl. Microbiol., 86: 985–990. Hentrich, H., R. Kaiser, and G. Gottsberger. 2007. Floral scent collection at the perfume flowers of Anthurium rubrinervium (Araceae) by the kleptoparasitic orchid bee Aglae caerulea (Euglossini). Ecotropica, 13: 149–155. Herrera, C. M. 2005. Plant generalization on pollinators: Species property or local phenomenon? Am. J. Bot., 92: 13–20. Herrera, J. 1997. The role of colored accessory bracts in the reproductive biology of Lavandula stoechas. Ecologyv 78: 494–504. Johnson, M. T., S. A. Campbell, S. A., and S. C. Barrett. 2015. Evolutionary interactions between plant reproduction and defense against herbivores. Annu. Rev. Ecol. Evol. Syst., 46: 191–213. Junker, R. R., N. Blüthgen, T. Brehm, J. Binkenstein, J. Paulus, H. M. Schaefer, and M. Stang. 2013. Specialization on traits as basis for the niche-breadth of flower visitors and as structuring mechanism of ecological networks. Funct. Ecol., 27: 329–341. Jürgens, A., S. L. Wee, A. Shuttleworth, and S. D. Johnson. 2013. Chemical mimicry of insect oviposition sites: A global analysis of convergence in angiosperms. Ecol. Lett., 16: 1157–1167. Kallenbach, M., Y. Oh, E. J. Eilers, D. Veit, I. T. Baldwin, and M. C. Schuman. 2014. A robust, simple, high-throughput technique for time-resolved plant volatile analysis in field experiments. Plant J., 78: 1060–1072. Kantsa, A., R. A. Raguso, A. G. Dyer, S. P. Sgardelis, J. M. Olesen, and T. Petanidou. 2017. Community-wide integration of floral colour and scent in a Mediterranean scrubland. Nat. Ecol. Evol., 1: 1502–1510. Kantsa, A., R. A. Raguso, A. G. Dyer, J. M. Olesen, T. Tscheulin, and T. Petanidou. 2018. Disentangling the role of floral sensory stimuli in pollination networks. Nat. Commun., 9: 1041. https://www.nature.com/ articles/s41467-018-03448-w Kantsa, A., S. Sotiropoulou, M. Vaitis, and T. Petanidou. 2015. Plant volatilome in Greece: A review on the properties, prospects, and chemogeography. Chem. Biodivers., 12: 1466–1480. Karabagias, I. K., M. V. Vavoura, A. Badeka, S. Kontakos, and M. G. Kontominas. 2014. Differentiation of Greek thyme honeys according to geographical origin based on the combination of phenolic compounds and conventional quality parameters using chemometrics. Food Anal. Methods, 7: 2113–2121. Karban, R., and I. T. Baldwin. 1997. Induced Responses to Herbivory. Chicago: Univ Chicago Press. 319p. Kaspar, S., M. Peukert, A. Svatos, A. Matros, and H. P. Mock. 2011. MALDI-imaging mass spectrometry–an emerging technique in plant biology. Proteomics, 11: 1840–1850. Kessler, D., C. Diezel, D. G. Clark, T. A. Colquhoun, and I. T. Baldwin. 2013. Petunia flowers solve the defence/ apparency dilemma of pollinator attraction by deploying complex floral blends. Ecol. Lett., 16: 299–306.

494

Handbook of Essential Oils

Kessler, D., K. Gase, and I. T. Baldwin. 2008. Field experiments with transformed plants reveal the sense of floral scents. Science, 321: 1200–1202. Kite, G. C., W. L. A. Hetterscheid, M. J. Lewis et al. 1998. Inflorescence odours and pollinators of Arum and Amorphophallus (Araceae). In: S.J. Owens and P.J. Rudall (Editors). Reproductive Biology, pp. 295–315. Royal Botanic Gardens. Kew. Kite, G. C., and W. L. Hetterscheid. 2017. Phylogenetic trends in the evolution of inflorescence odours in Amorphophallus. Phytochemistry, 142: 126–142. Knoll, F. R. 1926. Die Arum-blütenstände und ihre Besucher. Abhandlungen der Zoologisch–Botanischen Gesellschaft Wien, 12: 382–481. Kraemer, M., and U. Schmitt. 1999. Possible pollination by hummingbirds in Anthurium sanguineum Engl. (Araceae). Plant Syst. Evol., 217: 333–335. Leonhardt, S. D., S. Zeilhofer, N. Blüthgen, and T. Schmitt. 2010. Stingless bees use terpenes as olfactory cues to find resin sources. Chem. Senses, 35: 603–611. Leonhardt, S. D., T. Schmitt, and N. Blüthgen, 2011. Tree resin composition, collection behavior and selective filters shape chemical profiles of tropical bees (Apidae: Meliponini). PLOS ONE, 6: e23445. https://doi. org/10.1371/journal.pone.0023445 Linhart, Y. B., and J. D. Thompson. 1995. Terpene-based selective herbivory by Helix aspersa (Mollusca) on Thymus vulgaris (Labiatae). Oecologia, 102: 126–132. Linhart, Y. B., and J. D. Thompson. 1999. Thyme is of the essence: Biochemical polymorphism and multispecies deterrence. Evol. Ecol. Res., 1: 151–171. Linhart, Y. B., K. Keefover-Ring, K. A. Mooney, B. Breland, and J. D. Thompson. 2005. A chemical polymorphism in a multitrophic setting: Thyme monoterpene composition and food web structure. Am. Nat., 166: 517–529. Linhart, Y. B., P. Gauthier, K. Keefover-Ring, and J. D. Thompson. 2014. Variable phytotoxic effects of Thymus vulgaris (Lamiaceae) terpenes on associated species. Int. J. Plant Sci., 176: 20–30. Lokvam, J., and J. F. Braddock. 1999. Anti-bacterial function in the sexually dimorphic pollinator rewards of Clusia grandiflora (Clusiaceae). Oecologia, 119: 534–540. Maia, A. C. D., S. Dötterl, R. Kaiser et al. 2012. The key role of 4-methyl-5-vinylthiazole in the attraction of scarab beetle pollinators: A unique olfactory floral signal shared by Annonaceae and Araceae. J. Chem. Ecol., 38: 1072–1080. Manan, A. A., R. M. Taha, E. E. Mubarak, and H. Elias. 2016. In vitro flowering, glandular trichomes ultrastructure, and essential oil accumulation in micropropagated Ocimum basilicum L. In Vitro Cell. Dev. Biol. Plant, 52: 303–314. Miguel, G., M. Simoes, A. C. Figueiredo, J. G. Barroso, L. G., Pedro, and L. Carvalho. 2004. Composition and antioxidant activities of the essential oils of Thymus caespititius, Thymus camphoratus and Thymus mastichina. Food Chem., 86: 183–188. Mitrevski, B., and P. J. Marriott. 2012. Novel hybrid comprehensive 2D–multidimensional gas chromatography for precise, high-resolution characterization of multicomponent samples. Anal. Chem., 84: 4837–4843. Mockute, D., and G. Bernotiene. 2001. The α-terpenyl acetate chemotype of essential oil of Thymus pulegioides L. Biochem. Syst. Ecol., 29: 69–76. Möller, M., and J. L. Clark. 2013. The state of molecular studies in the family Gesneriaceae: A review. Selbyana, 31: 95–125. Mondello, L., P. Q. Tranchida, P. Dugo, and G. Dugo. 2008. Comprehensive two-dimensional gas chromatography-mass spectrometry: A review. Mass Spectrom. Rev., 27: 101–124. Nogueira, P. C. D. L., V. Bittrich, G. J. Shepherd, A. V. Lopes, and A. J. Marsaioli. 2001. The ecological and taxonomic importance of flower volatiles of Clusia species (Guttiferae). Phytochemistry, 56: 443–452. Nordström, K., J. Dahlbom, V. S. Pragadheesh et al. 2017. In situ modeling of multimodal floral cues attracting wild pollinators across environments. Proc. Natl Acad. Sci. USA, 114: 13218–13223. de Oliveira, C. M., A. Porto, V. Bittrich, I. Vencato, and A. J. Marsaioli. 1996. Floral resins of Clusia spp.: Chemical composition and biological function. Tetrahedron Lett., 37: 6427–6430. Paine, T. D., K. F. Raffa, and T. C. Harrington. 1997. Interactions among scolytid bark beetles, their associated fungi, and live host conifers. Annu. Rev. Entomol., 42: 179–206. Palmer-Young, E. C., B. M. Sadd, and L. S. Adler. 2017a. Evolution of resistance to single and combined floral phytochemicals by a bumble bee parasite. J. Evol. Biol., 30: 300–312. Palmer-Young, E. C., B. M. Sadd, P. C. Stevenson, R. E. Irwin, and L. S. Adler. 2016. Bumble bee parasite strains vary in resistance to phytochemicals. Sci. Rep., 6: 37087. https://www.nature.com/articles/srep37087 Palmer-Young, E. C., B. M. Sadd, R. E. Irwin, and L. S. Adler. 2017b. Synergistic effects of floral phytochemicals against a bumble bee parasite. Ecol. Evol., 7: 1836–1849.

Functions of Essential Oils and Natural Volatiles in Plant-Insect Interactions

495

Papadopoulou, G. V., and N. M. van Dam. 2017. Mechanisms and ecological implications of plant-mediated interactions between belowground and aboveground insect herbivores. herbivores. Ecol. Res., 32: 13–26. Pereira, R., I. de Souza Carvalho, B.R. Simoneit, and D. de Almeida Azevedo. 2009. Molecular composition and chemosystematic aspects of Cretaceous amber from the Amazonas, Araripe and Recôncavo basins, Brazil. Org. Geochem., 40: 863–875. Petanidou, T., and D. Vokou. 1993. Pollination ecology of Labiatae in a phryganic (East Mediterranean) ecosystem. Am. J. Bot., 80: 892–899. Petanidou, T., and E. Smets. 1996. Does temperature stress induce nectar secretion in Mediterranean plants? New Phytol., 133: 513–518. Phillips, M. A., and R. B. Croteau. 1999. Resin-based defenses in conifers. Trends Plant Sci., 4: 184–190. Pichersky, E., and R. A. Raguso. 2018. Why do plants produce so many terpenoid compounds? New Phytol., 220: 692–702. Pokorny, T., I. Vogler, R. Losch et al. 2017. Blown by the wind: The ecology of male courtship display behavior in orchid bees. Ecology, 98: 1140–1152. Policha, T., A. Davis, M. Barnadas, B. T. Dentinger, R. A. Raguso, and B. A. Roy. 2016. Disentangling visual and olfactory signals in mushroom-mimicking Dracula orchids using realistic three-dimensional printed flowers. New Phytol., 210: 1058–1071. Popova, M. P., K. Graikou, I. Chinou, and V. S. Bankova. 2010. GC-MS profiling of diterpene compounds in Mediterranean propolis from Greece. J. Agric. Food Chem., 58: 3167–3176. Potts, S. G., T. Petanidou, S. Roberts, C. O’Toole, A. Hulbert, and P. Willmer. 2006. Plant-pollinator biodiversity and pollination services in a complex Mediterranean landscape. Biol. Conserv., 129: 519–529. Raffa, K. F. 2001. Mixed messages across multiple trophic levels: The ecology of bark beetle chemical communication systems. Chemoecology, 11: 49–65. Raguso, R. A., A. A. Agrawal, A. E. Douglas, G. Jander, A. Kessler, K. Poveda, and J. S. Thaler. 2015a. The raison d’être of chemical ecology. Ecology, 96: 617–630. Raguso, R. A., J. N. Thompson, and D. R. Campbell. 2015b. Improving our chemistry: Challenges and opportunities in the interdisciplinary study of floral volatiles. Nat. Prod. Rep., 32: 893–903. Ramírez, S. R., T. Eltz, M. K. Fujiwara et  al. 2011. Asynchronous diversification in a specialized plantpollinator mutualism. Science, 333: 1742–1746. Richardson, L. L., L. S. Adler, A. S. Leonard et al. 2015. Secondary metabolites in floral nectar reduce parasite infections in bumblebees. Proc. R. Soc. B., 282: 20142471. Riffell, J. A., E. Shlizerman, E. Sanders et al. 2014. Flower discrimination by pollinators in a dynamic chemical environment. Science, 344: 1515–1518. Rodman, J. E., A. R. Kruckeberg, and I. A. Al-Shehbaz. 1981. Chemotaxonomic diversity and complexity in seed glucosinolates of Caulanthus and Streptanthus (Cruciferae). Syst. Bot., 6: 197–222. Rota, M. C., A. Herrera, R. M. Martínez, J. A. Sotomayor, and M. J. Jordán. 2008. Antimicrobial activity and chemical composition of Thymus vulgaris, Thymus zygis and Thymus hyemalis essential oils. Food Control, 19: 681–687. Rundel, P. W., M. T. Arroyo, R. M. Cowling et al. 2018. Fire and plant diversification in Mediterranean-climate regions. Front. Plant Sci., 9: 851. Salatino, A., C. C. Fernandes-Silva, A. A. Righi, and M. L. F. Salatino. 2011. Propolis research and the chemistry of plant products. Nat. Prod. Rep., 28: 925–936. Schiestl, F. P., and S. Dötterl. 2012. The evolution of floral scent and olfactory preferences in pollinators: Coevolution or pre-existing bias? Evolution, 66: 2042–2055. Schmid-Hempel, P., and S. Durrer. 1991. Parasites, floral resources and reproduction in natural populations of bumblebees. Oikos, 62: 342–350. Schwerdtfeger, M., G. Gerlach, and R. Kaiser. 2002. Anthecology in the neotropical genus Anthurium (Araceae): A preliminary report. Selbyana, 23: 258–267. Seymour, R. S. 2001. Diffusion pathway for oxygen into highly thermogenic florets of the arum lily Philodendron selloum. J. Exp. Bot., 52: 1465–1472. Shroff, R., K. Schramm, V. Jeschke et al. 2015. Quantification of plant surface metabolites by matrix-assisted laser desorption–ionization mass spectrometry imaging: Glucosinolates on Arabidopsis thaliana leaves. Plant J., 81: 961–972. Skubatz, H., D. D. Kunkel, W. N. Howald, R. Trenkle, and B. Mookherjee. 1996. The Sauromatum guttatum appendix as an osmophore: Excretory pathways, composition of volatiles and attractiveness to insects. New Phytol., 134: 631–640. Steele, C. L., J. Crock, J. Bohlmann, and R. Croteau. 1998a. Sesquiterpene synthases from grand fir (Abies grandis) comparison of constitutive and wound-induced activities, and cDNA isolation,

496

Handbook of Essential Oils

characterization, and bacterial expression of δ-selinene synthase and γ-humulene synthase. J. Biol. Chem., 273: 2078–2089. Steele, C. L., S. Katoh, J. Bohlmann, and R. Croteau. 1998b. Regulation of oleoresinosis in grand fir (Abies grandis): Differential transcriptional control of monoterpene, sesquiterpene, and diterpene synthase genes in response to wounding. Plant Physiol., 116: 1497–1504. Tarayre, M., J. D. Thompson, J. Escarré, and Y. B. Linhart. 1995. Intra-specific variation in the inhibitory effects of Thymus vulgaris (Labiatae) monoterpenes on seed germination. Oecologia, 101: 110–118. Teichert, H., S. Dötterl, B. Zimma, M. Ayasse, and G. Gottsberger. 2009. Perfume-collecting male euglossine bees as pollinators of a basal angiosperm: The case of Unonopsis stipitata (Annonaceae). Plant Biol., 11: 29–37. Thompson, J. D., P. Gauthier, J. Amiot et al. 2007. Local adaptation in a chemically polymorphic plant: The relative performance of Thymus vulgaris chemotypes in a spatially heterogeneous environment. Ecol. Monogr., 77: 421–439. Tissier, A., J. A. Morgan, and N. Dudareva. 2017. Plant volatiles: Going “in” but not “out” of trichome cavities. Trends Plant Sci., 22: 930–938. Tümen, G., N. Kirimer, and K. H. C. Başer. 1995. Composition of the essential oil of Thymus species growing in Turkey. Chemistry of Natural Compounds, 31: 42–47. Urru, I., M. C. Stensmyr, and B. S. Hansson. 2011. Pollination by brood-site deception. Phytochemistry, 72: 1655–1666. Vernet, P., R. H. Gouyon, and G. Valdeyron. 1986. Genetic control of the oil content in Thymus vulgaris L: A case of polymorphism in a biosynthetic chain. Genetica, 69: 227–231. Whitten, W. M., H. G. Hills, and N. H. Williams. 1988. Occurrence of ipsdienol in floral fragrances. Phytochemistry, 27: 2759–2760. Whitten, W. M., N. H. Williams, W. S. Armbruster, M. A. Battiste, L. Strekowski, L., and N. Lindquist. 1986. Carvone oxide: An example of convergent evolution in euglossine pollinated plants. Syst. Bot., 11: 222–228. Williams, N. H., and W. M. Whitten. 1983. Orchid floral fragrances and male euglossine bees: Methods and advances in the last sesquidecade. Biol. Bull., 164: 355–395. Wise, M. L., T. J. Savage, E. Katahira, and R. Croteau. 1998. Monoterpene synthases from common sage (Salvia officinalis) cDNA isolation, characterization, and functional expression of (+)-sabinene synthase, 1, 8–cineole synthase, and (+)-bornyl diphosphate synthase. J. Biol. Chem., 273: 14891–14899. Wright, G. A., and F. P. Schiestl. 2009. The evolution of floral scent: The influence of olfactory learning by insect pollinators on the honest signaling of floral rewards. Funct. Ecol., 23: 841–851. Zapata, F., and P. V. Fine. 2013. Diversification of the monoterpene synthase gene family (TPSb) in Protium, a highly diverse genus of tropical trees. Mol. Phylogenet. Evol., 68: 432–442. Zimmermann, Y., S. R. Ramírez, and T. Eltz. 2009. Chemical niche differentiation among sympatric species of orchid bees. Ecology, 90: 2994–3008.

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Essential Oils as Lures for Invasive Ambrosia Beetles Paul E. Kendra, Nurhayat Tabanca, Wayne S. Montgomery, Jerome Niogret, David Owens, and Daniel Carrillo

CONTENTS 18.1 Introduction........................................................................................................................... 497 18.2 Redbay Ambrosia Beetle....................................................................................................... 498 18.2.1 Background................................................................................................................ 498 18.2.2 Host Attractants......................................................................................................... 499 18.2.3 Field Lures................................................................................................................. 499 18.2.3.1 Manuka and Phoebe Oils............................................................................500 18.2.3.2 Cubeb Oil....................................................................................................500 18.2.3.3 Copaiba Oil and Enriched α-Copaene Oil................................................. 502 18.3 Chemical Analysis of the α-Copaene Lure........................................................................... 505 18.3.1 Method....................................................................................................................... 505 18.3.2 Results........................................................................................................................ 505 18.4 Euwallacea Shot-Hole Borers............................................................................................... 507 18.4.1 Background................................................................................................................ 507 18.4.2 Host Attractants.........................................................................................................509 18.4.3 Field Lures.................................................................................................................509 18.4.3.1 Quercivorol.................................................................................................509 18.4.3.2 Enriched α-Copaene Lure..........................................................................509 18.5 Other Applications................................................................................................................. 510 References....................................................................................................................................... 512

18.1 INTRODUCTION Ambrosia beetles in the weevil subfamily Scolytinae (Coleoptera: Curculionidae) are wood borers that have developed close symbiotic relationships with fungi (Hulcr and Stelinski, 2017). Upon dispersal, adult females carry fungal conidia (spores) in specialized storage organs known as mycangia. Colonizing females excavate galleries in the xylem of host trees, inoculate the gallery walls with conidia, and subsequently cultivate fungal gardens to provide food for themselves and their progeny. Most species of ambrosia beetle will colonize only physiologically stressed or dying trees, and host-seeking females locate vulnerable trees by their elevated emissions of ethanol (Miller and Rabaglia, 2009; Ranger et al., 2010). As such, these beetles can be considered beneficial insects that initiate decomposition of woody material and contribute to the maintenance of healthy forest ecosystems. Unfortunately, with significant increases in global trade in recent decades, exotic ambrosia beetles have become one of the most frequently intercepted insects at ports of entry (Rabaglia et al., 2008). Although benign or minor pests in their native range, a few species and their fungal symbionts have emerged as serious threats to forests and agriculture following establishment in new environments and exploitation of novel hosts (Hulcr and Dunn, 2011). This is the case with the redbay ambrosia beetle, Xyleborus glabratus Eichhoff (Fraedrich et al., 2008; Kendra et al., 2013a) and a group of 497

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cryptic species morphologically similar to the tea shot-hole borer, Euwallacea fornicatus Eichhoff (Kasson et al., 2013; O’Donnell et al., 2015). These ambrosia beetles are endemic to Southeast Asia, but are now established in portions of the US, where they vector fungal pathogens responsible for vascular wilt diseases in commercially grown avocado (Persea americana Mill.) and numerous native American trees. A critical component for successful pest management is the identification of effective attractants, followed by development of lures for early pest detection. This process is facilitated by an understanding of the unique aspects of the insect’s chemical ecology. First, as with other ambrosia beetles within the tribe Xyleborini, species-specific sex pheromones are not used by X. glabratus or Euwallacea spp.; this is because new females usually mate with their flightless male siblings prior to dispersal from the natal gallery. Second, atypical of ambrosia beetles, neither X. glabratus or Euwallacea spp. are strongly attracted to ethanol. These beetles function ecologically as primary colonizers, capable of attacking healthy unstressed trees. As a result, they are attracted to the volatile terpenoids naturally emitted from the wood of host trees (i.e., kairomones used for host location). Since essential oils consist of concentrated plant terpenoids, they have provided an ideal substrate for development of lures for ambrosia beetle pests (Hanula and Sullivan, 2008; Kendra et al., 2012c, 2014b, 2015b, 2016a, 2018; Owens et al., 2017). This chapter will summarize the succession of essential oil lures used for X. glabratus over the last decade, outline development of the current lure which is highly enriched in (–)-α-copaene, present chemical analysis of the α-copaene lure, and describe its recent applications for detection of Euwallacea in Florida avocado groves.

18.2  REDBAY AMBROSIA BEETLE 18.2.1 Background Females of X. glabratus are the primary vectors of the fungus that causes laurel wilt, a lethal disease of trees and woody shrubs within the family Lauraceae, and a disease not observed in nature until the establishment of X. glabratus in the US (Fraedrich et al., 2008; Hanula et al., 2008) (Figure 18.1). Although multiple fungal species have been isolated from the mycangia of X. glabratus, the beetle’s predominant symbiont, Raffaelea lauricola T. C. Harr., Fraedrich and Aghayeva, is the confirmed pathogen (Harrington et al., 2008, 2010). Introduction of R. lauricola into susceptible hosts triggers secretion of gels and the formation of parenchymal tyloses within xylem vessels (Ploetz et al., 2012).

FIGURE 18.1  Female (A) and male (B) of the redbay ambrosia beetle, Xyleborus glabratus, vector of the fungal pathogen that causes laurel wilt. (C) Cross-section through the trunk of a swampbay tree, Persea palustris, showing extensive galleries excavated by redbay ambrosia beetles. (D) Silkbay tree, Persea humilis, killed by laurel wilt in Highlands County, FL, US.

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This defensive response can be extreme, resulting in decreased water transport, systemic wilt, and ultimately death of the infected tree. Since initially detected near a maritime port in Georgia in 2002, X. glabratus has continued to spread throughout the southeastern US. It is now established in nine states (USDA-FS, 2018) where laurel wilt has reached epidemic levels (reviewed by Kendra et al., 2013a; Hughes et al., 2015; Ploetz et al., 2017b). Throughout this range, primary hosts are native Persea species, including redbay (Persea borbonia [L.] Spreng.), swampbay (P. palustris [Raf.] Sarg.), and silkbay (P. humilis Nash), and as of 2017, an estimated 300 million native bay trees have been killed (Hughes et al., 2017). In southern Florida, avocado trees are also readily attacked. Although X. glabratus clearly transported the R. lauricola pathogen southward into commercial groves, the progression of laurel wilt in this agroecosystem has unfolded quite differently than that in native forests. Apparently avocado is a poor host for X. glabratus, limiting its reproduction (Brar et al., 2013); in contrast, established species of ambrosia beetle successfully colonize avocado trees stressed by laurel wilt. This situation has led to an unprecedented lateral transfer of R. lauricola to at least nine additional beetle species, which now comprise a community of secondary vectors of the laurel-wilt pathogen in avocado groves (Carrillo et al., 2014; Ploetz et al., 2017c). Because of the severe impact of laurel wilt, it has been identified as a high-priority disease by the USDA National Plant Disease Recovery System (USDA-ARS, 2018), and separate recovery plans have been drafted for the disease in forest hosts (Hughes et al., 2015) and in avocado (Ploetz et al., 2017a).

18.2.2 Host Attractants All known hosts of X. glabratus in the US are members of the Lauraceae family; therefore, research on kairomone attractants has focused on this taxonomic group. Kendra et al., (2014a) conducted a comparative study of nine lauraceous species (including avocado cultivars representative of each of the three botanical races) to determine boring preferences and in-flight attraction of X. glabratus as related to phytochemical emissions from host wood. Emissions of α-copaene, α-cubebene, α-humulene, and calamenene (all sesquiterpene hydrocarbons) were positively correlated with attraction to Lauraceae. Of these compounds, α-copaene and α-humulene had been correlated previously with attraction to three additional avocado cultivars, as well as to wood from lychee (Litchi chinensis Sonn.; Sapindaceae) (Kendra et al., 2011a). Lychee is not a reproductive host of X. glabratus, but particular cultivars will attract females and initiate boring behavior owing to sesquiterpene emission profiles similar to those of the Lauraceae (Kendra et al., 2011a, 2011b, 2013b). In addition, independent research identified eucalyptol (1,8 cineole; a monoterpene ether) as another host-based attractant (Kuhns et al., 2014). For all these terpenoids, olfactory chemoreception has been confirmed in female X. glabratus using freshly dissected antennae and newly developed electroantennography (EAG) techniques designed to accommodate the minute antennae of this species (Figure 18.2) (Kendra et  al., 2012a, 2014a). Comparative EAG recordings have been useful for assessing potential attractants, with good correlation between amplitude of antennal responses and relative attraction observed in the field. The current hypothesis is that the initial step in the host-location process consists of in-flight detection (by antennal olfactory receptors) of a mixture of volatile terpenoids characteristic of the Lauraceae (i.e., a long-range “signature bouquet” indicative of an appropriate host) (Kendra et al., 2011a, 2014a; Niogret et al., 2011a). This is followed by a series of cues presented in sequential order, including mid-range visual cues and integration of various short-range cues (olfactory, gustatory, contact chemosensory, tactile) which reinforce the message that a suitable host has been located, triggering a switch from “host-location” behavior to “host-acceptance and boring” behavior (Kendra et al., 2014a and references therein).

18.2.3 Field Lures Although identification of specific attractant chemicals has been useful for understanding the chemical ecology of X. glabratus, production of field lures using synthetic sesquiterpenes poses

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FIGURE 18.2  (A) Scanning electron micrograph of female redbay ambrosia beetle antenna (length = 0.4 mm), showing concentration of olfactory sensilla on the distal end of antennal club. (B) Two-pronged gold electroantennography (EAG) electrode modified to accommodate minute antennae by soldering a flexible gold wire to one prong. (C) Magnified view of the EAG antennal preparation, with head capsule mounted to the prong (indifferent electrode) and back of antennal club in contact with wire (different electrode).

a challenge. Many sesquiterpenes, particularly α-copaene, are difficult to synthesize, not readily available, and prohibitively costly in quantities sufficient for trap deployment (Flath et al., 1994). Thus, development of economical lures has relied on the use of plant-derived essential oils (whole or distilled oil products) naturally high in attractive sesquiterpenes. 18.2.3.1  Manuka and Phoebe Oils Crook et  al., (2008) identified two essential oils attractive to the emerald ash borer, Agrilus planipennis Fairmaire (Coleoptera: Buprestidae). These consisted of manuka oil (extracted from Leptospermum scoparium Forst. and Forst.; Myrtaceae), and phoebe oil (obtained from Phoebe porosa Mez; Lauraceae). When first tested in Georgia and South Carolina, these two oils were also found to be good baits for trapping X. glabratus (Hanula and Sullivan, 2008). However, when commercial manuka and phoebe oil lures were evaluated in Florida in 2009–2010, both were found to be fairly non-specific, capturing a variety of non-target Scolytinae (Kendra et al., 2011b, 2012c). Moreover, the manuka oil lures were not competitive with host Persea wood (Figure 18.3A) and had a field life of only 2–3 weeks due to rapid depletion of sesquiterpenes under Florida field conditions (Figure 18.3C). In contrast, phoebe oil lures were competitive with host wood (Figure 18.3), captured significantly more X. glabratus than manuka oil lures (Figure 18.3B), and despite rapid loss of terpenoids initially, continued to release low levels of attractive sesquiterpenes for at least 10 weeks (Figure 18.3C). In addition, EAG response to phoebe oil was comparable to that of Persea wood, and both were significantly higher than EAG response to manuka oil (Kendra et al., 2012a). Unfortunately, shortly after completion of the field studies by Kendra et al., (2012c), phoebe oil lures were no longer available commercially due to a limited supply of source trees in Brazil. Therefore, the suboptimal manuka oil lure became the standard for detection of X. glabratus in the US. 18.2.3.2  Cubeb Oil The poor performance (and increasing cost) of manuka oil lures prompted research to evaluate additional essential oils as potential alternatives. Field tests in 2012 (Kendra et al., 2013a, 2014b) compared captures of female X. glabratus among seven essential oils. Treatments included whole oil preparations of manuka and phoebe oils, plus five new oils: cubeb, ginger root, angelica seed, tea

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FIGURE 18.3  Captures of female redbay ambrosia beetle in field tests conducted in Florida, US. (A) Mean (±SE) captures in an 8-wk test (Alachua County) with commercial phoebe oil lure (Phoeb), commercial manuka oil lure (Manuk), wood bolts of three avocado cultivars: “Brooks Late,” Guatemalan race (Guat); “Simmonds,” West Indian race (W. Ind); “Seedless Mexican,” Mexican race (Mex), and an unbaited control (Contr). (Adapted from Kendra et al., 2011a. J. Chem. Ecol., 37: 932–942.) (B) Mean (±SE) captures and (C) summed weekly captures in a 12-wk test (Highlands County) with commercial oil lures and an unbaited control. Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al., 2012c. J. Econ. Entomol., 105: 659–669.)

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FIGURE 18.4  Mean (±SE) captures of female redbay ambrosia beetle with seven essential oils deployed in a 4-wk field test conducted in Highlands County, FL, USA. Treatments consisted of manuka (Man), cubeb (Cub), phoebe (Phb), ginger root (Gin), angelica seed (Ang), tea tree (Tea), Valencia orange (Ora) oils, and an unbaited control trap (Ctrl). A membrane-based dispenser was used to prepare the lures, and each lure was loaded with 5 mL of neat oil. Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al., 2013a. Am. J. Plant. Sci., 4: 727–738.)

tree, and Valencia orange oils. These latter oils were chosen based on their sesquiterpene content, which included constituents correlated with captures of X. glabratus previously (Kendra et  al., 2011a, 2012c). In addition, they all had been reported to attract males of the Mediterranean fruit fly, Ceratitis capitata Wied. (Diptera: Tephritidae), another pest that responds positively to substrates containing α-copaene (Flath et al., 1994; Niogret et al., 2011b). Of the new oils evaluated, ginger root and angelica seed oils were moderately attractive, but whole cubeb oil (extracted from berries of tailed pepper, Piper cubeba L.; Piperaceae) was identified as a strong new attractant for X. glabratus (Figure 18.4) (Kendra et al., 2013a, 2014b). In 2013, a commercial formulation of cubeb oil became available, which consisted of a plastic bubble lure containing a proprietary oil product distilled to remove monoterpenoids, thereby enriching the sesquiterpene content (Synergy Semiochemicals Corp., Burnaby, BC, Canada). The cubeb bubble lure captured significantly more X. glabratus than the commercial manuka oil lure in several independent field trials (Hanula et al., 2013; Kendra et al., 2014b). In EAG analyses, olfactory responses to the cubeb lure and phoebe lure were equivalent, and both were significantly greater than response obtained with the manuka oil lure (Kendra et al., 2014b). Further evaluations (Kendra et al., 2015b) indicated that field attraction of cubeb oil lures was significantly better than that of phoebe oil lures (Figure 18.5A), and the bubble lures had a field life of at least 12 weeks (Figure 18.5B). Temporal analysis of cubeb lure emissions revealed that its superior longevity was due to extended low release of sesquiterpenes, primarily α-copaene (Figure 18.6A) and α-cubebene (Figure 18.6B) (Kendra et al., 2015b). The bubble dispenser used for the new cubeb lure has a much lower surface area-to-volume ratio than the flat rectangular design of the manuka oil lure (Figure 18.7). This difference, coupled with a thicker release membrane, has resulted in a much improved delivery system for sustained release of sesquiterpene attractants. 18.2.3.3  Copaiba Oil and Enriched α-Copaene Oil After the cubeb bubble lure replaced the manuka lure for detection of X. glabratus, research efforts (in collaboration with Synergy Semiochemicals Corp.) focused on elucidation of the attractive components found in cubeb oil. Since this essential oil is composed of a complex mixture of

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FIGURE 18.5  Captures of female redbay ambrosia beetle in a 12-wk field test conducted in Highlands County, FL, US. (A) Mean (±SE) captures and (B) summed weekly captures obtained with commercial oil lures and an unbaited control. Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al., 2015b. J. Econ. Entomol., 108: 350–361.)

terpenoids, hydro-distillation was used to separate whole cubeb oil into multiple fractions, based on boiling point of the terpenoid constituents. These fractions were then evaluated through EAG analyses and binary-choice bioassays to quantify olfactory and behavioral responses, respectively. Results indicated that fractions with high α-copaene and α-cubebene content were the most bioactive for female X. glabratus (Kendra et al., 2016a), supporting previous hypotheses regarding the importance of these kairomones (Kendra et al., 2011a, 2012c, 2014a, 2014b, 2015b; Niogret et al., 2013). Based on this observation, two additional essential oil products were evaluated, both formulated in slow-release bubble dispensers. The first lure contained whole copaiba oil (extracted from species of Copaifera L.; Leguminosae), an essential oil low in α-cubebene, but with twice as much α-copaene as whole cubeb oil. In field trials (Figure 18.8A), the prototype copaiba oil lure captured equal numbers of X. glabratus as the commercial cubeb oil lure (Kendra et al., 2016a). The second prototype lure contained a proprietary essential oil product that was highly distilled to achieve 50% α-copaene content. In multiple field tests, this enriched α-copaene lure captured significantly more X. glabratus than either the cubeb oil lure or the copaiba oil lure (Figure 18.8A)

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FIGURE 18.6  Emissions of (A) α-copaene and (B) α-cubebene quantified over time from commercial oil lures field-deployed for 12 wk in Miami-Dade County, FL, US. Inset enhances the scale for emissions beginning at 4 wk, the point at which manuka lures lost efficacy for attraction of redbay ambrosia beetle in field tests. Volatiles were isolated by super-Q collection, analyzed by GC-MS, and identified by comparison of Kovats retention index with synthetic chemicals. (Adapted from Kendra, P. E. et al., 2015b. J. Econ. Entomol., 108: 350–361.)

and displayed field longevity of at least three months (Figure 18.8B) (Kendra et al., 2016a, 2016b). EAG response to the α-copaene lure was also significantly higher than response recorded with either the cubeb oil lure or the copaiba oil lure (Kendra et al., 2016a). This research confirmed the role of α-copaene as a primary host-location cue, and identified the 50% α-copaene lure as an improved tool for more sensitive detection of X. glabratus than that provided by the cubeb oil lure. The α-copaene bubble lure is now produced commercially and has been adopted by state and federal action agencies for use in X. glabratus monitoring programs in the US.

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FIGURE 18.7  Commercially available lures for redbay ambrosia beetle. (A) Cubeb bubble lure (Synergy Semiochemicals Corp.; Burnaby, BC, Canada). (B) Manuka oil lure (ChemTica Internacional; Heredia, Costa Rica).

18.3  CHEMICAL ANALYSIS OF THE α-COPAENE LURE 18.3.1 Method Three replicates of the α-copaene lure (Synergy Semiochemicals Corp., Product # 3302, Lot # 160511) were analyzed by gas chromatography-mass spectrometry (GC-MS) using a 5975B mass-selective detector (Agilent Technologies, Santa Clara, SA, US) and equipped with a DB-5 column (30 m, 0.25 mm ID, film thickness 0.25 µm; Agilent Technologies). The oven temperature was programmed from 60°C (1.3 min) to 246°C at a rate of 3°C/min. Helium was used as the carrier gas at a flow rate of 1.3 mL/ min. The PTV injector and ion source temperature were 200°C and 230°C, respectively. Mass spectra were recorded at 70 eV. The mass range was m/z 35–450 and scan rate was set at 2.8 scans/second. Data acquisition and processing were done using software Mass Hunter B.07.02 (Agilent Technologies). Chemical identification of lure components was based on (a) linear retention indices (LRI), which were calculated using n-alkanes on the DB-5 column according to the calculation method developed by van den Dool and Kratz (1963); (b) comparison of the mass spectra with published literature values (MassFinder, 2004; Adams, 2007; FFNSC-3, 2015; NIST, 2017; Wiley, 2017); (c) comparison with our own library “SHRS Essential Oil Constituents-DB-5,” which was built with authentic standards and components of known essential oils analyzed on the DB-5 column; and (d) comparison with reference compounds purchased from Fluka Chemical Co., Buchs, SG, Switzerland (δ-cadinene, Cas # 483-76-1; α-copaene. Cas # 3856-25-5; β-elemene, Cas # 515-13-9; valencene, Cas # 4630-07-3), Sigma-Aldrich Ltd, St. Louis, MO, US (alloaromadendrene, Cas # 25246-27-9; β-caryophyllene, Cas # 87-44-5; caryophyllene oxide, Cas # 1139-30-6, cyclosativene, Cas # 22469-52-9; and α-humulene, Cas # 6753-98-6), and α-cubebene (Cas # 17699-14-8) from Bedoukian Research, Inc., Danbury, CT, US.

18.3.2 Results Fifteen compounds were identified from the α-copaene enriched oil lure, and relative percentages of the components are shown in Table 18.1. The analysis indicated that sesquiterpene hydrocarbons were

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FIGURE 18.8  Captures of female redbay ambrosia beetle in a 12-wk field test conducted in Highlands County, FL, US. (A) Mean (±SE) captures and (B) summed weekly captures obtained with commercial cubeb oil lure, prototype copaiba oil lure, prototype 50% α-copaene lure (50% cop), and an unbaited control trap. Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al, 2016a. J. Pest Sci., 89: 427–438.)

the dominant constituents, comprising 99.3% of the lure contents; only one oxygenated sesquiterpene was detected, caryophyllene oxide (0.7%). Major sesquiterpene hydrocarbons consisted of α-copaene (60.2%), β-caryophyllene (18.7%), and δ-cadinene (10.3%), with lesser amounts of α-cubebene (2.4%), α-humulene (3.2%), and γ-muurolene (1.1%). These results are consistent with those reported by Owens et al. (2018b) for a different lot of α-copaene lures analyzed on a DB-5MS column. Although α-copaene is the primary attractant, other sesquiterpene components (part of the laurel bouquet, Kendra et al., 2014a) may be contributing to the overall attraction of this lure. Owens et al. (2018b) also evaluated the enantiomeric distribution of α-copaene on an Rt-βDEXse column (2,3-di-O-ethyl-6-O-tert-butyl dimethylsilyl-β-cyclodextrin; Restek Corp., Bellefonte, PA, US), which indicated that the (–)-enantiomer was predominant (99.91%) in the commercial lure, with a very high enantiomeric excess of 99.82% (Figure 18.9). Knowledge of enantiomer distributions is an important consideration when developing management programs for insect pests. Although

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TABLE 18.1 Chemical Composition of α-copaene Enriched Lure (n = 3) #

LRIexpa1

LRILitb2

Compound

%

Identification methodc1

1

1335

1335

δ-Elemene

0.27 ± 0.02

a, b, c

2

1348

1345

2.42 ± 0.11

a, b, c, d

3

1363

1369

α-Cubebene Cyclosativene

0.01 ± 0.00

a, b, c, d

4

1374

1374

α-Copaene

60.21 ± 0.09

a, b, c, d

5

1388

1387

β-Cubebene

0.31 ± 0.01

a, b, c

6

1390

1389

β-Elemene

0.25 ± 0.02

a, b, c, d

7

1407

1409

α-Gurjunene

0.84 ± 0.03

a, b, c

8

1416

1417

β-Caryophyllene

9

1450

1452

10

1456

1458

11

1474

1478

12

1488

13

1498

14 15

18.72 ± 0.10

a, b, c, d

α-Humulene Alloaromadendrene

3.19 ± 0.02

a, b, c, d

0.81 ± 0.05

a, b, c, d

1.07 ± 0.07

a, b, c

1496

γ-Muurolene Valencene

0.43 ± 0.02

a, b, c, d

1500

α-Muurolene

0.54 ± 0.02

a, b, c

1521

1522

1580

1582

δ-Cadinene Caryophyllene oxide

10.26 ± 0.12

a, b, c, d

0.66 ± 0.01

a, b, c, d

LRI was calculated against n-alkanes based on DB-5 column. LRI values from Adams library (2007). c1 Identification method given in Section 18.3.1. a1

b1

X. glabratus and E. nr. fornicatus (see Section 18.4.3.2 below) are highly attracted to (–)-α-copaene in this essential oil lure, the (+)−α-copaene isolated from angelica seed oil (Angelica archangelica L.) was found to be more attractive to males of C. capitata then its (–)- enantiomer (Flath et al., 1994).

18.4  EUWALLACEA SHOT-HOLE BORERS 18.4.1 Background The type species, E. fornicatus, is a polyphagous beetle, but it is best documented as a pest of cultivated tea, Camellia sinensis (L.) Kuntze (Theaceae), in Sri Lanka, India, and elsewhere in Asia (Hazarika et al., 2009) (Figure 18.10). Females typically colonize small-diameter branches which reduces production of tender foliage (flushes), the commodity harvested commercially. In recent years, beetles morphologically indistinguishable from E. fornicatus have emerged as pests of avocado, woody ornamentals, and native trees outside of Asia. Pest populations have become established in the US (Rabaglia et  al., 2006; Eskalen et  al., 2013), Israel (Mendel et  al., 2012), Australia (Campbell and Geering, 2011), Mexico (García-Avila et al., 2016), and, most recently, South Africa (Paap et al., 2018). Molecular investigations have concluded that beetles from different geographic regions possess sufficient genetic diversity to constitute a cryptic species complex; therefore, all members are conservatively referred to as E. near fornicatus currently, pending taxonomic separation (Kasson et al., 2013, O’Donnell et al., 2015). The primary fungal symbionts are species within the genus Fusarium (Hypocreales: Nectriaceae) (Freeman et al., 2012). Unlike the R. lauricola pathogen that systemically colonizes host xylem, Fusarium spp. fungi remain localized and destroy the functional xylem in close proximity to beetle galleries. With avocado trees, beetle attacks tend to be concentrated at the base of branches, so the localized lesions essentially girdle the branch, impede water conduction, and eventually kill the branch. This newly described vascular disease has been termed Fusarium dieback (Freeman et al., 2012).

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FIGURE 18.9  GC-MS chromatogram of the α-copaene enriched lure and enantiomeric distribution of α-copaene using an Rt-βDEXse column.

FIGURE 18.10  Female (A) and male (B) of Euwallacea nr. fornicatus, vector of the fungal pathogen that causes Fusarium dieback. (C) Avocado tree (Persea americana) in Miami-Dade County, FL, US, with multiple attacks by E. nr. fornicatus concentrated at the base of the branch.

In the US, two distinct species of E. nr. fornicatus now exist in California, commonly referred to as the polyphagous shot-hole borer, first detected in the vicinity of Los Angeles, and the Kuroshio shot-hole borer, found farther south near San Diego (Eskalen et al., 2013; Boland, 2016). These beetles are attributed with high mortality of a variety of phylogenetically diverse species, with the

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most severe impact on avocado, box elder (Acer negundo L., Sapindaceae), castor bean (Ricinus communis L., Euphorbiaceae), California live oak (Quercus agrifolia Née, Fagaceae), and red willow (Salix laevigata Bebb, Salicaceae). A third distinct population, more closely aligned with tea shot-hole borer (Stouthamer et al., 2017), is now established in southern Florida where avocado is the preferred host (Carrillo et al., 2016; Kendra et al., 2017; Owens et al., 2018b). That state’s avocado industry, already heavily impacted by laurel wilt, now faces a new threat with Fusarium dieback. In addition to avocado, reported hosts of Florida E. nr. fornicatus include native species like swampbay and wild tamarind (Lysiloma latisiliquum [L.] Benth., Fabaceae), ornamental royal poinciana (Delonix regia [Boj. ex Hook] Raf., Fabaceae), naturalized exotics like woman’s tongue tree (Albizia lebbeck [L.] Bentham, Fabaceae), and fruit crops such as mango (Mangifera indica L., Anacardiaceae) and soursop (Annona muricata L., Annonaceae) (Rabaglia et al., 2006; Carrillo et al., 2012, 2016; Owens et al., 2018a).

18.4.2 Host Attractants Unlike the situation with X. glabratus, little attention has been directed toward identification of specific host-based attractants for the Euwallacea pests. This is not unexpected, given the extensive host range exhibited by members of the E. nr. fornicatus complex. For example, hosts of the tea shothole borer are reported to include species from more than 35 plant families in Asia (Danthanarayana, 1968), and the polyphagous shot-hole borer, as implied by its common name, has even more documented hosts in California, with species represented from more than 50 plant families (Eskalen et al., 2013). To the authors’ knowledge, only one published report has addressed host attractants of E. fornicatus; this was in Sir Lanka and was restricted to tea (Karunaratne et al., 2008). In olfactometer tests, females were attracted to the dominant volatiles known from tea, including eugenol, hexanol, α- and β-pinene, geraniol, and methyl salicylate. In addition, attraction was increased when these volatiles were combined with ethanol. With E. nr. fornicatus in Florida field trials, ethanol appeared to be a weak attractant when combined with quercivorol lures (see Section 18.4.3.1 below) (Carrillo et al., 2015; Kendra et al., 2015a); however, with polyphagous shot-hole borer, ethanol was found to be a repellent in tests conducted in California (Dodge et al., 2017) and in Israel (Byers et al., 2018).

18.4.3 Field Lures 18.4.3.1 Quercivorol Current monitoring for E. nr. fornicatus utilizes a commercial lure containing quercivorol, (1S, 4R)p-menth-2-en-1-ol (Kashiwagi et al., 2006), a proposed fungal volatile emitted by Fusarium spp. (i.e., a food-based attractant) (Cooperband et al., 2017). These lures were first found attractive to host-seeking females in Florida (Carrillo et al., 2015), with comparable results later obtained with Euwallacea populations in California (Dodge et al., 2017) and Israel (Byers et al., 2017). A recent analysis (Owens et al., 2018b) of the commercial lure indicated it contains a mixture of four isomers of p-menth-2-en-1-ol, and the (1S,4R)-enantiomer is not the major component. However, this lure is commonly referred to as the “quercivorol lure” in entomological literature, but it is unclear exactly which isomer(s) is/are responsible for attraction of E. nr. fornicatus. 18.4.3.2 Enriched α-Copaene Lure In 2015, while conducting field tests for X. glabratus in a Florida avocado grove, a serendipitous discovery was made. Females of E. nr. fornicatus also appeared to be attracted to the enriched α-copaene lure (Kendra et al., 2015a). More focused evaluations in 2016 demonstrated that α-copaene is equal in attraction to quercivorol and that the combination of semiochemicals results in additive to synergistic increases in trap capture (Figure 18.11) (Kendra et al., 2017). EAG analyses confirmed olfactory chemoreception of both compounds, with a higher response elicited with the combination of

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FIGURE 18.11  Mean (±SE) captures of female Euwallacea nr. fornicatus in field tests conducted in commercial avocado groves in Miami-Dade County, FL, US. (A) Results obtained at a site free of laurel wilt with high beetle populations. (B) Results from a site with advanced laurel wilt and very low numbers of E. nr. fornicatus. Lure treatments consisted of a quercivorol bubble lure and an α-copaene bubble lure, deployed separately and in combination. Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al, 2017. PLOS ONE, 12: e0179416.)

volatiles (Figure 18.12). This two-component lure, combining a food attractant with a host attractant, is the most effective lure identified to date for E. nr. fornicatus in Florida, with field longevity of 12 weeks, minimal attraction of non-target ambrosia beetles, and an effective sampling range of ∼30 m (Owens et al., 2018b, 2019). In early 2018, this combination lure was adopted by SAGARPA (Secretaría de Agricultura, Ganadería, Desarrollo Rural, Pesca y Alimentación) in Mexico in survey programs for both E. nr. fornicatus and X. glabratus in high-risk areas, including ports, international borders, and avocado-production regions (DGSV-CNRF, 2018).

18.5  OTHER APPLICATIONS Although recognized as valuable tools for early pest detection, essential oils have played another important role in our investigations of ambrosia beetles. In 2011, a method was developed for field

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FIGURE 18.12  Electroantennogram (EAG) responses of female Euwallacea nr. fornicatus to volatiles emitted from commercial lures containing quercivorol and α-copaene. (A) Representative EAG recordings obtained with single excised antennae. (B) Comparative EAG responses (mean ± SE) to fixed 2-mL doses of volatiles. Responses to lure volatiles are expressed as normalized percentages relative to the standard reference compound (ethanol, 2-mL saturated vapor). Bars topped with the same letter are not significantly different. (Adapted from Kendra, P. E. et al, 2017. PLOS ONE, 12: e0179416.)

collection of live, host-seeking female X. glabratus using host wood and essential oils as bait (Kendra et al., 2012b). As an alternative to establishing a laboratory colony of X. glabratus, this method provided an immediate supply of adult beetles for research use, on an “as needed” basis. Furthermore, females obtained in this manner were behaviorally and physiologically in “host-seeking mode,” the perfect cohort for evaluation of host-based attractants in controlled tests. The method consisted of spreading a white cotton sheet in a clearing of a forest exhibiting laurel wilt, placing freshly cut Persea or lychee wood in the center of the sheet, and suspending essential oil lures (initially manuka and/or phoebe) from a tripod positioned above the wood. At intervals of 15–20 minutes, new wood was added to the sheet and lures were fanned to generate a pulsed release of attractive host volatiles. This effectively lured in female ambrosia beetles from the surrounding woods. As they landed, beetles were collected by hand with a soft brush and stored in plastic boxes with moist tissue until needed for experimentation. This simple collection method provided the foundation for numerous projects since 2011, not only advancing our knowledge of X. glabratus, but of many other scolytine beetles, including pest and non-pest species. An immediate benefit of real-time collections was the documentation of species-specific flight windows. This was observed initially for female X. glabratus, which initiated flight much earlier than other species, allowing for selective capture of this target species (Kendra et al., 2012b). Subsequently, flight windows were recorded for Xyleborus affinis Eichhoff and X.

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ferrugineus (Fabricius) (Kendra et al., 2012a), eight species of Hypothenemus (Johnson et al., 2016), E. nr. fornicatus (Kendra et al., 2017), and the community of ambrosia beetles found in Florida avocado groves affected by laurel wilt (Menocal et al., 2018). Field-caught females greatly accelerated comparative evaluations of host species and essential oil attractants through EAG analyses (Kendra et al., 2012a, 2014b, 2016a), choice and no-choice behavioral bioassays (Kendra et  al., 2013b, 2014a, 2016a), and field cage release-recapture experiments (Kendra et al., 2015b, 2016b). In addition, the collection of in-flight ambrosia beetles facilitated collaborations with plant pathologists to study the fungal symbionts transported in female mycangia (Bateman et al., 2015; Campbell et al., 2016; Ploetz et al., 2017c).

REFERENCES Adams, R. P., 2007. Identification of Essential Oil Components by Gas Chromatography/Mass Spectrometry, 4th ed. Carol Stream: Allured Publishing Corp. Bateman, C., P. E. Kendra, R. Rabaglia, and J. Hulcr, 2015. Fungal symbionts in three exotic ambrosia beetles, Xylosandrus amputatus, Xyleborinus andrewesi, and Dryoxylon onoharaense (Coleoptera: Curculionidae: Scolytinae: Xyleborini) in Florida. Symbiosis, 66: 141–148. Boland, J. M., 2016. The impact of an invasive ambrosia beetle on the riparian habitats of the Tijuana River Valley, California. Peer J, 4: e2141. Brar, G. S., J. L. Capinera, P. E. Kendra, S. Mclean, and J. E. Peña, 2013. Life cycle, development, and culture of Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). Florida Entomol, 96: 1158–e1167. Byers, J. A., Y. Maoz, and A. Levi-Zada, 2017. Attraction of the Euwallacea sp. near fornicatus (Coleoptera: Curculionidae) to quercivorol and to infestation in avocado. J. Econ. Entomol., 110: 1512–1517. Byers, J. A., Y. Maoz, D. Wakarchuk, D. Fefer, and A. Levi-Zada, 2018. Inhibitory effects of semiochemicals on the attraction of an ambrosia beetle Euwallacea nr. fornicatus to quercivorol. J. Chem. Ecol., 44: 565–575. Campbell, P., and A. Geering, 2011. Biosecurity capacity building for the Australian avocado industry – Laurel wilt. Proceedings VII World Avocado Congress 2011 (Actas VII Congreso Mundial del Aguacate 2011). Cairns, Australia. 5–9 September 2011. http://www.avocadosource.com/WAC7/Section_03/ CampbellPaul2011.pdf Campbell, A. S., R. C. Ploetz, T. J. Draeden, P. E. Kendra, and W. S. Montgomery, 2016. Geographic variation in mycangial communities of Xyleborus glabratus. Mycologia, 108: 657–667. Carrillo, D., R. E. Duncan, and J. E. Peña, 2012. Ambrosia beetles (Coleoptera: Curculionidae: Scolytinae) that breed in avocado wood in Florida. Florida Entomol., 95: 573–579. Carrillo, D., R. E. Duncan, J. N. Ploetz, A. F. Campbell, R. C. Ploetz, and J. E. Peña, 2014. Lateral transfer of a phytopathogenic symbiont among native and exotic ambrosia beetles. Plant Pathol., 63: 54–62. Carrillo, D., T. Narvaez, A. A. Cossé, R. Stouthamer, and M. Cooperband, 2015. Attraction of Euwallacea nr. fornicatus (Coleoptera: Curculionidae: Scolytinae) to lures containing quercivorol. Florida Entomol., 98: 780–782. Carrillo, D., L. F. Cruz, P. E. Kendra, T. I. Narvaez, W. S. Montgomery, A. Monterroso, C. De Grave, and M. F. Cooperband, 2016. Distribution, pest status, and fungal associates of Euwallacea nr. fornicatus in Florida avocado groves. Special Issue: Invasive Insect Species. Insects, 7: 55. Cooperband, M. F., A. A. Cossé, T. H. Jones, D. Carrillo, K. Cleary, I. Canlas, and R. Stouthamer, 2017. Pheromones of three ambrosia beetles in the Euwallacea fornicatus species complex: Ratios and preferences. Peer J., 5: e3957. Crook, D. J., A. Khirimian, J. A. Francese, I. Fraser, T. M. Poland, A. J. Sawyer, and V. C. Mastro, 2008. Development of a host-based semiochemical lure for trapping emerald ash borer, Agrilus planipennis (Coleoptera: Buprestidae). Environ. Entomol., 37: 356–365. Danthanarayana, W., 1968. The distribution and host-range of the shot-hole borer (Xyleborus fornicatus Eichh.) of tea. Tea Quarterly., 39: 61–69. DGSV-CNRF, 2018. Dirección General de Sanidad Vegetal — Centro Nacional de Referencia Fitosanitaria. Manuales operativos de plagas cuarentenarias. Plan de acción para la vigilancia y aplicación de medidas de control contra complejos ambrosiales reglamentados en México: Xyleborus glabratus- Raffaelea lauricola y Euwallacea sp.—Fusarium euwallaceae. http://www.sinavef.senasica.gob.mx/SIRVEF/ ManualesOperativos.aspx Dodge, C., J. Coolidge, M. Cooperband, A. Cossé, D. Carrillo, and R. Stouthamer, 2017. Quercivorol as a lure for the polyphagous and Kuroshio shot hole borers, Euwallacea spp. nr. fornicatus (Coleoptera: Scolytinae), vectors of Fusarium dieback. Peer J., 5: e3656.

Essential Oils as Lures for Invasive Ambrosia Beetles

513

Eskalen, A., R. Stouthamer, S. C. Lynch, P. F. Rugman-Jones, M. Twizeyimana, A. Gonzalez, and T. Thibault, 2013. Host range of Fusarium dieback and its ambrosia beetle (Coleoptera: Scolytinae) vector in southern California. Plant Dis., 97: 938–951. FFNSC-3, 2015. Flavors and Fragrances of Natural and Synthetic Compounds 3. Hoboken, NJ, USA: Mass Spectral Database, Scientific Instrument Services Inc. Flath, R. A., R. T. Cunningham, T. R. Mon, and J. O. John, 1994. Male lures for Mediterranean fruit fly (Ceratitis capitata Wied.): Structural analogs of α-copaene. J. Chem. Ecol., 20: 2595–2609. Fraedrich, S. W., T. C. Harrington, R. J. Rabaglia, M. D. Ulyshen, A. E. Mayfield III, J. L. Hanula, J. M. Eickwort, and D. R. Miller, 2008. A fungal symbiont of the redbay ambrosia beetle causes a lethal wilt in redbay and other Lauraceae in the southeastern USA. Plant Dis., 92: 215–224. Freeman, S., A. Protasov, M. Sharon, K. Mohotti, M. Eliyahu, N. Okon-Levy, M. Maymon, and Z. Mendel, 2012. Obligate feed requirement of Fusarium sp. Nov., an avocado wilting agent, by the ambrosia beetle Euwallacea aff. fornicata. Symbiosis, 58: 245–251. García-Avila, C. D. J., F. J. Trujillo-Arriaga, J. A. López-Buenfil, R. González-Gómez, D. Carrillo, L. F. Cruz, I. Ruiz-Galván, A. Quezada-Salinas, and N. Acevedo-Reyes, 2016. First report of Euwallacea nr. fornicatus (Coleoptera: Curculionidae) in Mexico. Florida Entomol., 99: 555–556. Hanula, J. L., and B. Sullivan, 2008. Manuka oil and phoebe oil are attractive baits for Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae), the vector of laurel wilt. Environ. Entomol., 37: 1403–1409. Hanula, J. L., A. E. Mayfield III, S. W. Fraedrich, and R. J. Rabaglia, 2008. Biology and host associations of redbay ambrosia beetle, Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae), exotic vector of laurel wilt killing redbay (Persea borbonia) trees in the southeastern United States. J. Econ. Entomol., 101: 1276–1286. Hanula, J. L., B. T. Sullivan, and D. Wakarchuk, 2013. Variation in manuka oil lure efficacy for capturing Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae), and cubeb oil as an alternative attractant. Environ. Entomol., 42: 333–340. Harrington, T. C., S. W. Fraedrich, and D. N. Aghayeva, 2008. Raffaelea lauricola, a new ambrosia beetle symbiont and pathogen on the Lauraceae. Mycotaxon, 104: 399–404. Harrington, T. C., D. N. Aghayeva, and S. W. Fraedrich, 2010. New combinations of Raffaelea, Ambrosiella, and Hyalorhinocladiella, and four new species from the redbay ambrosia beetle, Xyleborus glabratus. Mycotaxon, 111: 337–361. Hazarika, L. K., M. Bhuyan, and B. N. Hazarika, 2009. Insect pests of tea and their management. Ann. Rev. Entomol., 54: 267–284. Hughes, M. A., J. A. Smith, R. C. Ploetz et al., 2015. Recovery plan for laurel wilt on redbay and other forest species caused by Raffaelea lauricola and disseminated by Xyleborus glabratus. Plant Health Prog., 16: 173–210. Hughes, M. A., J. J. Riggins, F. H. Koch, A. I. Cognato, C. Anderson, J. P. Formby, T. J. Dreaden, R. C. Ploetz, and J. A. Smith, 2017. No rest for the laurels: Symbiotic invaders cause unprecedented damage to southern USA forests. Biol. Invasions., 19: 2143–2157. Hulcr, J. and R. R. Dunn, 2011. The sudden emergence of pathogenicity in insect-fungus symbioses threatens naïve forest ecosystems. Proc. R. Soc. B. Biol. Sci., 278: 2866–2873. Hulcr, J. and L. L. Stelinski, 2017. The ambrosia symbiosis: From evolutionary ecology to practical management. Annu. Rev. Entomol., 52: 285–303. Johnson, A. J., P. E. Kendra, J. Skelton, and J. Hulcr, 2016. Species diversity, phenology, and temporal flight patterns of Hypothenemus pygmy borers (Coleoptera: Curculionidae: Scolytinae) in South Florida. Environ. Entomol., 45: 627–632. Karunaratne, W. S., V. Kumar, J. Pettersson, and N. S. Kumar, 2008. Response of the shot-hole borer of tea, Xyleborus fornicatus (Coleoptera: Scolytidae) to conspecifics and plant semiochemicals. Acta Agr. Scand. B—Soil Plant Sci., 58: 345–351. Kashiwagi, T., T. Nakashima, S-I Tebayashi, and C-S Kim, 2006. Determination of the absolute configuration of quercivorol, (1S,4R)-p-menth-2-en-1-ol, an aggregation pheromone of the ambrosia beetle Platypus quercivorus (Coleoptera: Platypodidae). Biosci. Biotech. Biochem., 70: 2544–2546. Kasson, M. T., K. O’Donnell, A. P. Rooney et al., 2013. An inordinate fondness for Fusarium: Phylogenetic diversity of fusaria cultivated by ambrosia beetles in the genus Euwallacea on avocado and other plant hosts. Fungal Genet. Biol., 56: 147–157. Kendra, P. E., W. S. Montgomery, J. Niogret, J. E. Peña, J. L. Capinera, G. Brar, N. D. Epsky, and R. R. Heath, 2011a. Attraction of the redbay ambrosia beetle, Xyleborus glabratus, to avocado, lychee, and essential oil lures. J. Chem. Ecol., 37: 932–942. Kendra, P. E., J. S. Sanchez, W. S. Montgomery, K. E. Okins, J. Niogret, J. E. Peña, N. D. Epsky, and R. R. Heath, 2011b. Diversity of Scolytinae (Coleoptera: Curculionidae) attracted to avocado, lychee, and essential oil lures. Florida Entomol., 94: 123–130.

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Kendra, P. E., W. S. Montgomery, J. Niogret, M. A. Deyrup, L. Guillén, and N. E. Epsky, 2012a. Xyleborus glabratus, X. affinis, and X. ferrugineus (Coleoptera: Curculionidae: Scolytinae): Electroantennogram responses to host-based attractants and temporal patterns in host-seeking flight. Environ. Entomol., 41: 1597–1605. Kendra, P. E., W. S. Montgomery, J. S. Sanchez, M. A. Deyrup, J. Niogret, and N. D. Epsky, 2012b. Method for collection of live redbay ambrosia beetles, Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). Florida Entomol., 95: 513–516. Kendra, P. E., J. Niogret, W. S. Montgomery, J. S. Sanchez, M. A. Deyrup, G. E. Pruett, R. C. Ploetz, N. D. Epsky, and R. R. Heath, 2012c. Temporal analysis of sesquiterpene emissions from manuka and phoebe oil lures and efficacy for attraction of Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). J. Econ. Entomol., 105: 659–669. Kendra, P. E., W. S. Montgomery, J. Niogret, and N. D. Epsky, 2013a. An uncertain future for American Lauraceae: A lethal threat from redbay ambrosia beetle and laurel wilt disease (A review). Special Issue: The Future of Forests. Am. J. Plant. Sci., 4: 727–738. Kendra, P. E., R. C. Ploetz, W. S. Montgomery, J. Niogret, J. E. Peña, G. S. Brar, and N. D. Epsky, 2013b. Evaluation of Litchi chinensis for host status to Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae) and susceptibility to laurel wilt disease. Florida Entomol., 96: 1442–1453. Kendra, P. E., W. S. Montgomery, J. Niogret, G. E. Pruett, A. E. Mayfield III, M. MacKenzie, M. A. Deyrup, G. R. Bauchan, R. C. Ploetz, and N. D. Epsky, 2014a. North American Lauraceae: Terpenoid emissions, relative attraction and boring preferences of redbay ambrosia beetle, Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). PLOS ONE, 9: e102086. Kendra, P. E., W. S. Montgomery, J. Niogret, E. Q. Schnell, M. A. Deyrup, and N. D. Epsky, 2014b. Evaluation of seven essential oils identifies cubeb oil as most effective attractant for detection of Xyleborus glabratus. J. Pest Sci., 87: 681–689. Kendra, P. E., T. I. Narvaez, W. S. Montgomery, and D. Carrillo, 2015a. Ambrosia beetle communities in forest and agricultural ecosystems with laurel wilt disease (D3524). 53rd Annual Meeting of the Entomological Society of America, Minneapolis, MN, USA 15–18 November 2015. https://www.esa.confex.com/ esa/2015/webprogram/Session27045.html Kendra, P. E., J. Niogret, W. S. Montgomery, M. A. Deyrup, and N. D. Epsky, 2015b. Cubeb oil lures: Terpenoid emissions, trapping efficacy, and longevity for attraction of redbay ambrosia beetle (Coleoptera: Curculionidae: Scolytinae). J. Econ. Entomol., 108: 350–361. Kendra, P. E., W. S. Montgomery, M. A. Deyrup, and D. Wakarchuk, 2016a. Improved lure for redbay ambrosia beetle developed by enrichment of α-copaene content. J. Pest Sci., 89: 427–438. Kendra, P. E., W. S. Montgomery, E. Q. Schnell, M. A. Deyrup, and N. D. Epsky, 2016b. Efficacy of α-copaene, cubeb, and eucalyptol lures for detection of redbay ambrosia beetle (Coleoptera: Curculionidae: Scolytinae). J. Econ. Entomol., 109: 2428–2435. Kendra, P. E., D. R. Owens, W. S. Montgomery, T. I. Narvaez, G. R. Bauchan, E. Q. Schnell, N. Tabanca, and D. Carrillo, 2017. α-Copaene is an attractant, synergistic with quercivorol, for improved detection of Euwallacea nr. fornicatus (Coleoptera: Curculionidae: Scolytinae). PLOS ONE, 12: e0179416. Kendra, P. E., W. S. Montgomery, J. Niogret, N. Tabanca, D. Owens, and N. D. Epsky, 2018. Utility of essential oils for development of host-based lures for Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae), vector of laurel wilt. Special Issue: Research for Natural Bioactive Products. Open Chem., 16: 393–400. Kuhns, E. H., X. Martini, Y. Tribuiani, M. Coy, C. Gibbard, J. Peña, J. Hulcr, and L. L. Stelinski, 2014. Eucalyptol is an attractant of the redbay ambrosia beetle, Xyleborus glabratus. J. Chem. Ecol., 40: 355–362. MassFinder, 2004. MassFinder Software, Version 3. Hamburg: Dr. Hochmuth Scientific Consulting. Mendel, Z., A. Protasov, M. Sharon, A. Zveibil, S. Ben Yehuda, K. O’Donnell, R. Rabaglia, M. Wysoki, and S. Freeman, 2012. An Asian ambrosia beetle Euwallacea fornicatus and its novel symbiotic fungus Fusarium sp. pose a serious threat to the Israeli avocado industry. Phytoparasitica, 40: 235–238. Menocal, O., P. E. Kendra, W. S. Montgomery, J. H. Crane, and D. Carrillo, 2018. Vertical distribution and daily flight periodicity of ambrosia beetles (Coleoptera: Curculionidae) in Florida avocado orchards affected by laurel wilt. J. Econ. Entomol., 111: 1190–1196. Miller, D. R., and R. J. Rabaglia, 2009. Ethanol and (-)-α-pinene: Attractant kairomones for bark and ambrosia beetles in the southeastern U.S. J. Chem. Ecol., 35: 435–448. Niogret, J., P. E. Kendra, N. D. Epsky, and R. R. Heath, 2011a. Comparative analysis of terpenoid emissions from Florida host trees of the redbay ambrosia beetle, Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). Florida Entomol., 94: 1010–1017.

Essential Oils as Lures for Invasive Ambrosia Beetles

515

Niogret, J., W. S. Montgomery, P. E. Kendra, R. R. Heath, and N. D. Epsky, 2011b. Attraction and electroantennogram responses of male Mediterranean fruit fly (Diptera: Tephritidae) to volatile chemicals from Persea, Litchi, and Ficus wood. J. Chem. Ecol., 37: 483–491. Niogret, J., N. C. Epsky, R. J. Schnell, E. J. Boza, P. E. Kendra, and R. R. Heath, 2013. Terpenoid variations within and among half-sibling avocado trees, Persea americana Mill. (Lauraceae). PLOS ONE, 8: e73601. NIST, 2017. NIST/EPA/NIH Mass Spectral Library, Version: NIST 17. Mass Spectrometry Data Center. Gaithersburg, MD, USA: National Institute of Standard and Technology. O’Donnell, K., S. Sink, R. Libeskind-Hadas et al., 2015. Discordant phylogenies suggest repeated host shifts in the Fusarium-Euwallacea ambrosia beetle mutualism. Fungal Genet. Biol., 82: 277–290. Owens, D., W. S. Montgomery, T. I. Narvaez, M. A. Deyrup, and P. E. Kendra, 2017. Evaluation of lure combinations containing essential oils and volatile spiroketals for detection of host-seeking Xyleborus glabratus (Coleoptera: Curculionidae: Scolytinae). J. Econ. Entomol., 110: 1596–1602. Owens, D., L. F. Cruz, W. S. Montgomery, T. I. Narvaez, E. Q. Schnell, N. Tabanca, R. E. Duncan, D. Carrillo, and P. E. Kendra, 2018a. Host range expansion and increasing damage potential of Euwallacea nr. fornicatus (Coleoptera: Curculionidae) in Florida. Florida Entomol., 101: 229–e73236. Owens, D., P. E. Kendra, N. Tabanca, T. I. Narvaez, W. S. Montgomery, E. Q. Schnell, and D. Carrillo, 2018b. Quantitative analysis of contents and volatile emissions from α-copaene and quercivorol lures, and longevity for attraction of Euwallacea nr. fornicatus in Florida. Special Issue: Invasive insect pests of forests and urban trees: Pathways, early detection, and management. J. Pest Sci., 91: doi 10.1007/ s10340-018-0960-6. Owens, D., M. Seo, W. S. Montgomery, M. J. Rivera, L. L. Stelinski, and P. E. Kendra, 2019. Dispersal behavior of Euwallacea nr. fornicatus (Coleoptera: Curculionidae: Scolytinae) in avocado groves and estimation of lure sampling range. Agr. Forest Entomol., 21: 199–208. Paap, T., Z. W. de Beer, C. Migliorini, W. J. Nel, and M. J. Wingfield, 2018. The polyphagous shot hole borer (PSHB) and its fungal symbiont Fusarium euwallaceae: A new invasion in South Africa. Australasian Plant Pathol., 47: 231–237. Ploetz, R. C., J. M. Pérez-Martínez, J. A. Smith, M. Hughes, T. J. Dreaden, S. A. Inch, and Y. Fu, 2012. Responses of avocado to laurel wilt, caused by Raffaelea lauricola. Plant Pathol., 61: 801–808. Ploetz, R. C., M. A. Hughes, P. E. Kendra et al., 2017a. Recovery plan for laurel wilt of avocado, caused by Raffaelea lauricola. Plant Health Prog., 18: 51–77. Ploetz, R. C., P. E. Kendra, R. A. Choudhury, J. Rollins, A. Campbell, K. Garrett, M. Hughes, and T. Dreaden, 2017b. Laurel wilt in natural and agricultural ecosystems: Understanding the drivers and scales of complex pathosystems. Special Issue: Forest Pathology and Plant Health. Forests, 8: 48. Ploetz, R. C., J. L. Konkol, T. Narvaez, R. E. Duncan, R. J. Saucedo, A. Campbell, J. Mantilla, D. Carrillo, and P. E. Kendra, 2017c. Presence and prevalence of Raffaelea lauricola, cause of laurel wilt, in different species of ambrosia beetle in Florida USA. J. Econ. Entomol., 110: 347–354. Rabaglia, R. J., S. A. Dole, and A. I. Cognato, 2006. Review of American Xyleborina (Coleoptera: Curculionidae: Scolytinae) occurring north of Mexico, with an illustrated key. Ann. Entomol. Soc. Amer., 99: 1034–1056. Rabaglia, R., D. Duerr, R. Acciavatti, and I. Ragenovich, 2008. Early Detection and Rapid Response for NonNative Bark and Ambrosia Beetles. Washington, D. C.: US Department of Agriculture, Forest Service, Forest Health Protection, http://www.fs.fed.us/foresthealth/publications/EDRRProjectReport.pdf Ranger, C. M., M. E. Reding, A. B. Persad, and D. A. Herms, 2010. Ability of stress-related volatiles to attract and induce attacks by Xylosandrus germanus and other ambrosia beetles (Coleoptera: Curculionidae, Scolytinae). Agr. Forest Entomol., 12: 177–185. Stouthamer, R., P. Rugman-Jones, P. Q. Thu et al., 2017. Tracing the origin of a cryptic invader: Phylogeography of the Euwallacea fornicatus (Coleoptera: Curculionidae: Scolytinae) species complex. Agr. Forest Entomol., 19: 366–375. USDA-ARS, 2018. United States Department of Agriculture, Agricultural Research Service, National Plant Disease Recovery System. https://www.ars.usda.gov/crop-production-and-protection/plant-diseases/ docs/npdrs USDA-FS, 2018. United States Department of Agriculture, Forest Service, Southern Regional Extension Forestry. Distribution of Counties with Laurel Wilt Disease as of September 20, 2018. https://www. fs.usda.gov/Internet/FSE_DOCUMENTS/fseprd571973.pdf van den Dool, H, and P. D. Kratz, 1963. A generalization of the retention index system including linear temperature programmed gas-liquid partition chromatography. J. Chromatogr., A., 11: 463–471. Wiley, 2017. Wiley Registry of Mass Spectral Data, 11th edition. Ringoes, NJ, USA: Scientific Instrument Services, Inc..

19

Adverse Effects and Intoxication with Essential Oils Rosa Lemmens-Gruber

CONTENTS 19.1 Introduction......................................................................................................................... 518 19.1.1 General Side Effects.............................................................................................. 518 19.2 Camphor and Camphor-Containing Essential Oils............................................................. 519 19.2.1 Case Reports......................................................................................................... 519 19.2.1.1 Dose Range for Oral Intoxication......................................................... 519 19.2.1.2 Neurotoxic Effects................................................................................ 520 19.2.1.3 Effects Following Inhalative Application............................................. 520 19.3 Eucalyptus Oil..................................................................................................................... 521 19.3.1 Case Reports......................................................................................................... 521 19.3.1.1 Dose Range for Oral Intoxication......................................................... 521 19.3.1.2 Intoxication after Ingestion................................................................... 521 19.3.1.3 Intoxication after Topical Application and Inhalation......................... 522 19.3.2 Eucalyptol (1,8-Cineole)........................................................................................ 522 19.4 Thujone-Containing Essential Oils..................................................................................... 523 19.4.1 Case Reports......................................................................................................... 523 19.5 Peppermint Oil.................................................................................................................... 523 19.5.1 Adulterations......................................................................................................... 523 19.5.2 Case Reports......................................................................................................... 524 19.5.3 Menthol................................................................................................................. 525 19.5.3.1 Cooling vs. Irritating Effect................................................................. 525 19.5.3.2 Menthol-Induced Analgesia................................................................. 525 19.5.3.3 Menthol and Tobacco-Related Chemicals............................................ 525 19.5.3.4 Menthol and Dermal Penetration......................................................... 526 19.6 Pennyroyal Oil..................................................................................................................... 526 19.6.1 Case Reports......................................................................................................... 526 19.6.2 Pulegone and Menthofuran................................................................................... 527 19.6.3 Precautions............................................................................................................ 527 19.7 Wintergreen Oil................................................................................................................... 528 19.7.1 Methyl Salicylate................................................................................................... 528 19.8 Tea Tree Oil (Melaleuca Oil).............................................................................................. 528 19.8.1 Toxicity Following Oral Exposure........................................................................ 528 19.8.2 Toxicity Following Dermal Exposure................................................................... 529 19.8.3 Systemic Reactions................................................................................................ 529 19.8.4 Ototoxicity............................................................................................................. 529 19.8.5 Developmental Toxicity......................................................................................... 530 19.8.6 In Vitro Toxicity.................................................................................................... 530 19.9 Sassafras Oil........................................................................................................................ 530 19.9.1 Safrole................................................................................................................... 530

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19.10 Clove Oil (Oleum Caryophylli, Caryophylli floris aetheroleum)....................................... 531 19.10.1 Case Reports......................................................................................................... 531 19.10.2 Eugenol, Isoeugenol.............................................................................................. 531 19.10.2.1 Repeated Dose Toxicity........................................................................ 531 19.10.2.2 Developmental and Reproductive Toxicity........................................... 532 19.10.2.3 Skin Sensitization................................................................................. 532 19.10.3 Methyleugenol....................................................................................................... 533 19.11 Bergamot Oil....................................................................................................................... 533 19.12 Essential Oils of Nutmeg and Other Spices........................................................................ 534 19.13 Conclusion........................................................................................................................... 534 References....................................................................................................................................... 534

19.1 INTRODUCTION Studies on the toxicity of essential oils are comprehensive due to the fact that essential oils are composed of numerous compounds. In most cases, knowledge about intoxication with essential oils is based on case studies. In the absence of data for whole oil, insight may be derived from data for individual components. However, it is noteworthy to keep in mind that data for essential oils may not necessarily apply to individual components, and vice versa data for components may not apply to the whole oil. In most cases, essential oils are safe in use except in overdosage, wrong application route, and in hypersensitive persons. As essential oils are highly lipophilic, they are absorbed readily, and thus, also after topical application, systemic reactions may occur. Noteworthy, lipophilic constituents of essential oils may also pass the blood–brain and placental barriers, and therefore the application during pregnancy and lactation has to be done carefully. It is well documented that the toxicity of essential oils is concentration-dependent, and thus most unwanted side effects can be avoided by application of low doses. This advice, however, is not relevant for hypersensitive people, because allergic reactions can occur independently of the concentration. Adverse effects attributed to compounds of essential oils can be initiated by oxidative metabolism and conversion of components to reactive intermediates. Oxidation products formed by exposure to light and/or air may play an important role. Thus, formation of reactive compounds should be prevented by adequate storage. Further causes for intolerance and intoxication comprise impurity and contamination, adulteration, and wrong declaration of the essential oil.

19.1.1 General Side Effects Common side effects of essential oils include irritation of skin and mucosa. Besides these effects, for some furocoumarin-containing essential oils, photosensitivity has been reported. Noteworthy, in sensitive persons, signs of allergy on skin and in the respiratory tract have been reported in several publications, including following examples. In a five-year study with 3065 patients suffering from contact dermatitis, 16.6% patients were found to be allergic to a fragrance mix. In these patients, the most frequent allergens were identified as isoeugenol (57.9%), eugenol (55.4%), cinnamyl alcohol (34.4%), and oak moss (24.2%) (Turić et al., 2011). A 53-year-old patient who used volatile essential oils in aroma lamps over years suffered from relapsing eczema contact allergy to various essential oils. Sensitization was due to previous exposure to lavender, jasmine, and rosewood, and also, laurel, eucalyptus, and pomerance showed positive tests, although without known previous exposure. A diagnosis of allergic airborne contact dermatitis could be established (Schaller and Korting, 1995). Vicks VapoRub is a commonly used inhalant ointment that helps relieve symptoms of upper respiratory tract infections. It contains several plant substances, including turpentine oil, eucalyptus

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oil, and cedar leaf oil, which can potentially irritate or sensitize the skin, as well as camphor, menthol, nutmeg oil, and thymol. Many reports describe allergic contact dermatitis to the various constituents in Vicks VapoRub ointment (Noiles and Pratt, 2010).

19.2  CAMPHOR AND CAMPHOR-CONTAINING ESSENTIAL OILS The dietary exposure (herbs such as basil, coriander, marjoram, rosemary, and sage) to camphor was estimated to be 1.5 mg/person/day and thus is safe (Council of Europe, 2001). However, the essential oil from the so-called camphor tree (Cinnamomum camphora) contains up to 84% d-camphor and may be the source for intoxication. For the use of camphor as cough suppressant and decongestant, a limit of 11% allowable camphor in consumer products is set. The complex sensory properties of camphor can be explained by the modulation of several temperature-activated ion channels from the TRP family, such as TRPV3 (Moqrich et al., 2005), TRPV1 (Xu et al., 2005), TRPM8 (Selescu et al., 2013), and TRPA1 (Alpizar et al., 2013). Numerous reports have described the consequences of overdosage/poisoning with essential oils or their constituents like camphor (see inchem.org), eucalyptol, and thujone. Camphor is a well-known toxin responsible for thousands of poisonings per year. A review of the American Association of Poison Control Centers reports about almost 10,000 cases of intoxication with camphor-containing products in the period between 1990 and 2003 in the US (Manoguerra et al., 2006). Nonetheless, camphor still can be found in many over-the-counter remedies. Severe intoxication is mostly caused by accidental oral use or mistake with castor oil. After ingestion, symptoms of burning sensation in the mouth, nausea, and vomiting occur within 5–15 min (Committee on Drugs, 1978). But significant resorption of camphor may also occur after substantial cutaneous and inhalative application as the risk for intoxication depends on concentration, rate of absorption, exposition, and mixture of components. Locally, it can produce irritation of skin, eyes, and mucous membranes of the respiratory tract. Even usual topical application of camphor may cause contact eczema (Committee on Drugs, 1994). As camphor is easily absorbed in the gastrointestinal tract, the toxidrome manifests within minutes and includes gastrointestinal, hepato-, nephro-, and neurotoxic symptoms (Jimenez et al., 1983; Von Bruchhausen et al., 1993), as well as pulmonary and cardiac effects (Santos and Cabot, 2015). Severe ingestions may progress to seizures, apnea, and coma. Most individuals are no longer symptomatic after 24–48 h, but physiologic derangement may persist for far longer in some instances (Santos and Cabot, 2015).

19.2.1 Case Reports Especially children react sensitively when essential oils are ingested accidentally. Flaman et al. (2001) reported intoxications in 251 children after ingestion of essential oil or products. Out of these 251 children, 50 ingested eucalyptus oil, 18 camphorated oil, 93 a vaporizing liquid, and 90 Vicks VapoRub. The most common symptoms were cough, lethargy, and vomiting, whereas two children had seizures but recovered, and even one child died. 19.2.1.1  Dose Range for Oral Intoxication In humans admitted to the hospital in a state of acute intoxication after ingestion of 6–10 g camphor, camphor hydroxylated in the positions 3, 5, and 8 (or 9) were identified as major metabolites in the urine. Hydroxylated intermediates were subsequently oxidized to the corresponding ketones and carboxylic acids, the latter being conjugated with glucuronic acid (Köppel et al., 1982). Incubation of human liver microsomes with l-camphor resulted in the formation of 5-exo-hydroxycamphor as the only oxidation product (Gyoubu and Miyazawa, 2007). Among the 11n enzymes tested, only CYP2A6 was found to hydroxylate l-camphor. The important role of CYP2A6 was supported by the good correlation between contents of CYP2A6 and rates of formation of 5-exo-hydroxycamphor in liver microsomes from nine human samples (The EFSA Journal, 2008).

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In infants, a dosage of 0.5–1.0 g (Siegel and Wason, 1986), respectively a lowest lethal dose of 70 mg/kg body weight (inchem.org), may cause death or severe intoxication, probably due to immature hepatic metabolism (Uc et al., 2000). This dose range confirms previous reports. According to Smith and Margolis (1954), as little as 1 g camphor ingested in one teaspoonful of camphorated oil (20% camphor in cottonseed oil) was fatal in a 19-month-old child. Ragucci et al. (2007) presented the case of a 10-year-old boy who was admitted to the emergency room with symptoms of lethargy, nausea, vomiting, and rigors. He had chewed three over-the-counter cold remedy transdermal patches containing 4.7% (95.4 mg/patch) camphor and 2.6% menthol as active ingredients approximately 24 h before admission to the hospital. Assuming a body weight of 30 kg, this would correspond to an intake of camphor of approximately 10 mg/kg body weight. On the basis of these and similar data, a probable lethal dose was estimated to be in the range of 50–500 mg/kg body weight (Gleason et al., 1969; Phelan, 1976). In sensitive people, clinically insignificant signs of toxicity may be observed already at 5 mg/kg body weight, whereas clinically manifest toxicity may require doses higher than 30 mg/kg body weight, and indeed, the study of Geller et al. (1984) demonstrates a large variation in sensitivity of humans to the acute toxicity of camphor. 19.2.1.2  Neurotoxic Effects Neurotoxic effects include symptoms such as excitement, hallucinations, delirium, tremors, and convulsions (Opdyke, 1978). Camphor and camphor-containing essential oils have been included in the list of potential epileptogenics (Burkhard et al., 1999; Spinella, 2001). In a retrospective case series from 2007 to 2014, the patient-reported ingestion rate of camphor was 1.5–15 g. In 30 patients, nausea and vomiting occurred in 73.3% cases, and tonic-clonic seizure was seen in 40% of patients. Mean exposure time was significantly longer in patients who experienced seizure (Rahimi et al., 2017). Confirming the occurrence of seizures in animal toxicity tests, convulsions have been observed, especially in children following ingestion and also dermal or inhalation exposure of camphor- and thujone-containing products or eucalyptus oil (Craig, 1953; Millet et al., 1981; Melis et al., 1989; Committee on Drugs, 1994; Gouin and Patel, 1996; Burkhard et al., 1999; Woolf, 1999; Flaman et  al., 2001; Ruha et  al., 2003; Stafstrom, 2007; Khine et  al., 2009; Halicioglu et  al., 2011; see Bazzano et al., 2017). In lethal cases of camphor intoxication, postmortem necrosis of neurons was detected, similar to lesions observed in animal experiments (Smith and Margolis, 1954). The risk of serious neurological side effects in young children such as convulsions, accompanied by apnea and asystole, has led the Committee for Medicinal Products for Human Use (CHMP) to implement contraindication for the use of suppositories containing terpenic constituents of essential oils in children under 30 months old (EMA, 2012). 19.2.1.3  Effects Following Inhalative Application Camphor-containing products are almost exclusively used topically or for inhalation purpose. Nonetheless, topical and inhalative application is not free of risk due to the narrow therapeutic index of camphor. In infants, vapors of camphor and eucalyptus oil my cause laryngospasm with apnea and asystole (Kratschmer–Holmgren reflex). But, also, adults suffering from bronchial asthma or airway hypersensitivity may react with bronchoconstriction (inchem.org). No critical threshold could be determined for induction of laryngospasm. Vicks VapoRub (VVR) that contains 5.46% camphor and 1.35% eucalyptus oil is often used to relieve symptoms of chest congestion. A report about a toddler in whom severe respiratory distress developed after VVR was applied directly under her nose led Abanses et  al. (2009) to study the effect of VVR on mucociliary function in an animal model of airway inflammation. They could document a VVR-induced stimulation of mucin secretion (63% above control) and tracheal mucociliary transport velocity in the lipopolysaccharide-inflamed ferret airway. Findings were similar to the acute inflammatory stimulation observed with exposure to irritants and may lead to mucus obstruction of small airways and increased nasal resistance (Abanses et al., 2009).

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19.3  EUCALYPTUS OIL Human data are derived from accidental intoxications, which have been reported following ingestion of eucalyptus oil. However, it is difficult to draw general conclusions from reports on accidental intoxications with eucalyptus oil, because in many cases the amounts ingested could only be estimated roughly and the precise composition of the ingested product was not reported. Distilled eucalyptus oil contains at least 70% 1,8-cineole (eucalyptol) and a complex mixture of other compounds. Whitman and Ghazizadeh (1994) argue that the component hydrocyanic acid (prussic acid) may contribute to the toxicity of eucalyptus oil.

19.3.1 Case Reports 19.3.1.1  Dose Range for Oral Intoxication Reports about lethal doses of eucalyptus oil vary markedly (De Vincenzi et al., 2002). Although it is reported that in adults, less than 2.5 mL is usually asymptomatic, transient coma has followed ingestion of 1 mL, and adult fatalities have been recorded with as little as 3.5 mL (Hindle, 1994). In general, minor depression of consciousness is presumed if more than 3 mL of 100% oil is ingested and significant central nervous system depression with more than 5 mL (Darben et al., 1998). Death is commonly seen after ingestion of 30 mL, but also has occurred after 4–5 mL (Patel and Wiggins, 1980; Flaman et al., 2001; Karunakara and Jyotirmanju, 2012). In particular, infants are prone to severe intoxication. In 27 infant patients who ingested known doses of eucalyptus oil, 10 had no effects after a mean of 1.7 mL, 11 had minor poisoning after a mean of 2.0 mL, five had moderate poisoning after a mean of 2.5 mL and one had major poisoning after 7.5 mL (Tibballs, 1995). 19.3.1.2  Intoxication after Ingestion The toxic symptoms are rapid in onset, which include a burning sensation in the mouth and throat, abdominal pain, and spontaneous vomiting (Patel and Wiggins, 1980; Flaman et al., 2001). The initial central nervous system effects are giddiness, ataxia, and disorientation followed by loss of consciousness occurring in 10–15 min (Flaman et al., 2001; Karunakara and Jyotirmanju, 2012). Oil of eucalyptus was capable of producing severe cardiovascular, respiratory, and central nervous system manifestations after ingestion of as little as 4 mL by an adult (Gurr, 1965). In particular, ingestion of eucalyptus oil causes significant morbidity in infants and young children. There are several reports which document severe intoxication following oral application of eucalyptus oil. For example, accidental ingestion of eucalyptus oil by a 3-year-old boy caused profound central nervous system depression within 30 minutes and rapid recovery after gastric lavage (Patel and Wiggins, 1980). Out of 109 children (mean age 23.5 months) with eucalyptus oil ingestion, 59% of them were symptomatic; 28% had depression of conscious state, ranging from drowsiness to unconsciousness after ingesting 5–10 mL. Vomiting occurred in 37%, ataxia in 15%, and pulmonary disease in 11%. No treatment was given for 12% (Tibballs, 1995). Convulsions are rare in adults, but Kumar et al. (2015) reported two cases of 3- and 6-yearold boys, who presented with status epilepticus within 10 min of accidental ingestion of 10 mL of eucalyptus oil. They had four and eight episodes of tonic-clonic convulsions, respectively, without known previous history of seizures. The children recovered completely within 20 h (Kumar et al., 2015). Similarly, Karunakara and Jyotirmanju (2012) reported two cases, one of a 2-year-old female child with convulsions who accidentally ingested around 5–10 mL eucalyptus oil. Immediately after ingestion of the oil, the child had two episodes of vomiting, and within 20 minutes generalized tonicclonic convulsions started. In the second case, a 2-year, 7-month old male child, who was treated for two days with 5 mL of eucalyptus oil as a home remedy for cold, started having generalized tonicclonic convulsions within 10 minutes after oral application (Karunakara and Jyotirmanju, 2012).

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In contrast, Webb and Pitt (1993) identified 42 cases of oral eucalyptus oil poisoning in children under 14 years of age between 1984 and 1991 with less severe symptoms. Thirty-three children (80%) were asymptomatic, although four of these children ingested more than 30 mL of eucalyptus oil. Only two children had clear symptoms or clinical signs. Thus, the authors speculate that eucalyptus oil may be a less toxic compound than previously thought (Webb and Pitt, 1993). Due to lack of a specific antidote, the management of eucalyptus oil poisoning is mainly supportive and symptomatic. The main risk is aspiration following vomiting and depression of the central nervous system. Therefore, emesis is contraindicated (Patel and Wiggins, 1980; Flaman et al., 2001; Karunakara and Jyotirmanju, 2012). The role of activated charcoal is controversial (Flaman et al., 2001), because once ingested, eucalyptus oil is readily distributed throughout the body and efforts at elimination by using activated charcoal is unlikely to help significantly. In a case of severe intoxication with prolonged coma, successful management of intoxication was enabled by the use of mannitol, hemodialysis, and peritoneal dialysis (Gurr, 1965). Although reports about dosages causing toxic effects are divergent, already relatively small amounts of eucalyptus oil are also reported to be fatal, notably in children in whom severe toxicity may predispose to the development of status epilepticus. Thus, eucalyptus oil should never be given orally. 19.3.1.3  Intoxication after Topical Application and Inhalation Eucalyptus oil is well documented as being extremely toxic if ingested. But Darben et al. (1998) also present a case of systemic eucalyptus oil intoxication from topical application of a home remedy for urticaria, containing eucalyptus oil. The 6-year-old girl presented with slurred speech, ataxia, and muscle weakness progressing to unconsciousness following the widespread application. Six hours following removal of the topical preparation her symptoms had resolved (Darben et al., 1998).

19.3.2  Eucalyptol (1,8-Cineole) The case reports on acute toxicity in humans refer to the ingestion of eucalyptus oil and not to eucalyptol as such. Distilled eucalyptus oil typically contains at least 70% eucalyptol (1,8-cineole) and a complex mixture of other compounds. Thus, the contribution of eucalyptol and other constituents to the reported intoxications with eucalyptus oil do not provide information for adequate estimates of toxic dose levels for eucalyptol. Moreover, toxicological data available on eucalyptol are rather limited. However, similar to camphor essential oil, eucalyptol also produced electrocortical seizure activity in rat brain (Culic et al., 2009; Kolassa, 2013), which supports the context between eucalyptol and convulsions in children intoxicated with eucalyptus oil. Eucalyptol is rapidly absorbed from the gastrointestinal tract. In a subacute toxicity study, no significant differences in body weight and relative organ weight between the control group and eucalyptol treatment groups were found. The histopathological examinations showed dose-related granular degeneration and vacuolar degeneration in liver and kidney tissue after administration of high doses. The electron microscopy assays indicated that the influence of eucalyptol on the target organ at the subcellular level were mainly on the mitochondria, endoplasmic reticulum, and other membrane-type structures of liver and kidney (Hu et al., 2014; Xu et al., 2014). A limited long-term study with mice did not show treatment-related effects, including effects on tumor incidence. The study was performed with males only and the histopathological examination was limited to only a few organs. No evidence for genotoxicity has been found in bacterial tests. In Chinese hamster ovary cells, chromosomal aberrations were not induced, and sister chromatid exchanges were only observed at cytotoxic doses (Opinion of the Scientific Committee on Food on eucalyptol, EU, 2002). Acute and subacute oral toxicity studies of eucalyptol in Kunming mice revealed an LD50 value (95% CI) of 3849 mg/kg (Xu et al., 2014). Even if eucalyptol were responsible for the acute toxicity of eucalyptus oil, the estimated daily intake of eucalyptol from food would be much lower than the amount tentatively assumed to be present in the lowest lethal doses of eucalyptus oil reported (Opinion of the Scientific Committee on Food on eucalyptol, EU, 2002).

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19.4  THUJONE-CONTAINING ESSENTIAL OILS Thujones are renowned as the possible psychoactive and toxic principles of absinthe (Olsen, 2000). Thujone is a common name for two naturally occurring monoterpene diastereomeric ketones, (−)-α-thujone and (+)-β-thujone, that can be found in essential oils of sage (Salvia officinalis L.), absinthe wormwood (Artemisia absinthium L.), eastern arborvitae (Thuja occidentalis L.), and tansy (Tanacetum vulgare L.) in different quantities (Blagojevic et al., 2006). These plants are wildgrowing and ornamental plant species with ethnopharmacological usages. Their essential oils are important components of numerous flavoring, perfumery, cosmetic, and pharmaceutical products. However, their potential toxic effects, in particular the known neurotoxicity of thujones, are the main reasons why some countries impose restriction upon their utilization.

19.4.1 Case Reports Precautions should be taken into account because there are several case reports in which sage oil induced the development of vertical nystagmus, hyperreflexia, clonic spasms, and tonic-clonic convulsions, with muscle dystonia before seizures and postictal muscle weakness occurred (Burkhard et al., 1999; Halicioglu et al., 2011). The convulsive properties of a diluted essential oil (1:99 dilution, and repeating this dilution 30 times) containing thujones were observed in a 7-month-old baby (Stafstrom, 2007). In adults, the oil of A. absinthium caused convulsions and paralysis, probably due to its content of thujones, camphor, and 1,8-cineole (see Radulović et al., 2017). α-Thujone in absinthe and herbal medicines is a rapid-acting and readily detoxified modulator of the GABAgated chloride channel (Höld et al., 2000). Although all of the described neurotoxic effects could be attributed to the presence of thujones in these oils, numerous other oil constituents could also cause the observed neurotoxicity (Radulović et al., 2017).

19.5  PEPPERMINT OIL The main component of peppermint oil (Oleum Menthae piperitae) is menthol (Table 19.1).

19.5.1 Adulterations Dementholized essential oil of Mentha arvensis L. var. piperascens Malinv., which is mainly produced in Brazil and China, has to be labelled correctly as mint oil or Japanese mint oil. The composite of the oil is comparable to peppermint oil of Mentha x piperita L., but it is much cheaper. Synthetic

TABLE 19.1 Content of Components of Peppermint Oil Content (%)

Component

∼45

Menthol

∼24 1–9

Menthon

∼5

Menthofurane Eucalyptol (1,8-cineol)

∼4

Menthyl acetate

∼3

Isomenthon

∼1.5

Limonene

∼1

Pulegon

300 volunteers) trials at concentrations ranging from 1% to 100% in different formulations and time span. Patients did not demonstrate any or only weak irritant reactions (see Hammer et al., 2006). However, numerous case reports of contact allergy have been published (see Hammer et al., 2006). The reactions occurred in response to 100% pure tea tree oil as well as lower concentrations in various formulations. The frequency of allergic reaction reported in numerous trials ranged from 0.1% to 10.7% (Hammer et al., 2006). Patch testing of allergic patients with tea tree oil components showed that they reacted mostly to the sesquiterpenoid fractions but not the pure monoterpenes (Southwell et  al., 1997). Probably the oxidation products formed during prolonged storage are the main allergens (Hausen, 2004), while newly distilled tea tree oil seems to have a relatively low sensitizing capacity. The most important allergens formed could be terpinolene, α-terpinene, ascardiole, and 1,2,4-trihydroxymethane (Hausen et al., 1999; Hausen, 2004).

19.8.3 Systemic Reactions Toxicity following dermal application of inappropriately high doses of melaleuca oil to cats or dogs treated for fleas has been described. Animals had typical signs of depression, weakness, un-coordination and muscle tremors (Villar et al., 1994), and especially cats experienced severe symptoms of hypothermia, incoordination, dehydration, and trembling. One cat which had preexisting renal damage even died after three days (Bischoff and Guale, 1998). Only one report of a systemic reaction to tea tree oil in humans is published (Mozelsio et al., 2003). Following dermal application, a 38-year-old man suffered immediate flushing, pruritus, a constricted throat, and lightheadedness. His reaction, however, was not due to an IgG or IgE response. Henley et al. (2007) investigated possible causes of gynecomastia in three prepubertal boys who repeatedly applied lavender and tea tree oil topically and who were otherwise healthy and had normal serum concentrations of endogenous steroids. In all three boys, gynecomastia coincided with the topical application of products that contained lavender and tea tree oils. Studies in human cell lines confirmed the findings of estrogenic and antiandrogenic activities of both essential oils. Gynecomastia resolved in each patient shortly after the use of products containing these oils was discontinued (Henley et  al., 2007). However, this matter is discussed controversially (Carson et al., 2014).

19.8.4  Ototoxicity Tea tree oil has been suggested as an effective treatment for a number of microorganisms commonly associated with otitis externa and otitis media, but it is possible ototoxicity has only been evaluated in a single study by Zhang and Robertson (2000) in guinea pigs. The authors showed that concentrations up to 2% tea tree oil may be safe for use within the ear, whereas higher concentrations applied for

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a relatively short time were, to some extent, ototoxic to the high-frequency region of the cochlea (Zhang and Robertson, 2000).

19.8.5 Developmental Toxicity No studies of potential developmental toxicity following exposure to tea tree oil have been published yet. However, an embryo- and fetotoxicity study with α-terpinene, a constituent of tea tree oil with a content of approximately 9%, has demonstrated significant toxicity in a rat model (Araujo et al., 1996), with delayed ossification and skeletal malformations. These limited data suggest that tea tree oil might be embryo- and fetotoxic, although only if ingested at relatively high concentrations.

19.8.6  In Vitro Toxicity Several cytotoxicity studies have been performed in numerous cell lines and showed that tea tree oil and/or its components are cytotoxic at concentrations between 20 and 2700 µg/mL (Hammer et al., 2006). However, for topically applied products, data on percutaneous penetration and in vivo disposition including metabolism are needed for any relevant assessment of a potential in vivo risk. Furthermore, in bacterial reverse mutation assays, tea tree oil and its components were not mutagenic (Hammer et al., 2006) and not genotoxic (Pereira et al., 2014). To conclude, if oxidation products can be avoided, the available literature suggests that tea tree oil can be used topically in diluted form by the majority of individuals without adverse effects.

19.9  SASSAFRAS OIL The main constituent of sassafras oil is safrole, which is also present in small amounts in a number of spices. Sassafras oil is extracted from the bark and roots of the tree Sassafras albidum. It has had a traditional and widespread use as a natural diuretic, as well as a remedy against urinary tract disorders or kidney problems until safrole was discovered to be hepatotoxic and weakly carcinogenic (Fennell et al., 1984; Rietjens et al., 2005). Thus, the FDA banned the use of sassafras oil as a food and flavoring additive because of the high content of safrole and its proven carcinogenic effects. However, pure sassafras oil is still available online and also in some health-food stores.

19.9.1 Safrole Safrole has been demonstrated to be genotoxic and carcinogenic. It produces liver tumors in mice and rats by oral administration, and it produces also liver and lung tumors in male infant mice by subcutaneous application. It has been shown that safrole is able to induce chromosome aberrations, sister chromatid exchanges, and DNA adducts in hepatocytes of F344 rats exposed in vivo (Daimon et al., 1998). However, the carcinogenic potency appears to be relatively low and dependent on the metabolism. Safrole is metabolically activated through the formation of intermediates, and these are able to directly react with DNA. Two bioactivation pathways of safrole to potentially hepatotoxic intermediates have been reported. One involves P450-catalyzed hydroxylation, thereby producing 1′-hydroxysafrole, and on further conjugation with sulfate, generating a reactive sulfate ester. This ester creates a highly reactive carbocation, which alkylates DNA. The second pathway involves P450-catalyzed hydroxylation of the methylenedioxy ring and finally formation of reactive p-quinone methide. Both pathways could explain the genotoxic effects of safrole. DNA adducts have been identified in vitro and in vivo (see Dietz and Bolton, 2005). In vitro experiments confirm the genotoxic effects of safrole. Therefore, the existence of a threshold cannot be assumed and the EU Scientific Committee on Food could not establish a safe exposure limit. Consequently, reductions in the exposure and restrictions in the use levels by health authorities are justified (Opinion of the Scientific Committee on Food on the safety of the presence of safrole, 2002).

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19.10 CLOVE OIL (Oleum Caryophylli, Caryophylli floris aetheroleum) Clove oil is the common name for an extract from the flower buds of Syzygium aromaticum (L.) Merrill et L. M. Perry, also known as Caryophyllus aromaticus L., Eugenia aromatica (L.) Baill., Eugenia caryophyllus (Spreng.) Bullock & S. G. Harrison, and Eugenia caryophyllata Thunberg. It is a complex mixture of chemical substances, the main component being eugenol. The essential oil is widely used and well known for its medicinal properties. Traditional uses of clove oil include use in dental care, as an antiseptic and analgesic, where the undiluted oil may be rubbed on the gums to treat toothache. It is active against oral bacteria associated with dental caries and periodontal disease (Cai and Wu, 1996) and effective against a large number of other bacteria (Chaieb et al., 2007). The major component of clove oil is usually considered to be eugenol with a content of 88.6% (Chaieb et al., 2007).

19.10.1 Case Reports Recent growth in aromatherapy sales has been accompanied by an unfortunate increase in accidental poisoning from these products. Clove oil warrants special attention, because it has hepatotoxic effects at high concentrations. Eugenol is a material commonly used in dentistry with few reported side effects. It is not, however, a bio-friendly material when in contact with oral soft tissues. It can produce both local irritative and cytotoxic effects, as well as hypersensitivity reactions, as reported by Sarrami et al. (2002) in two cases and by Barkin et al. (1984), who reported one case with a serious allergic reaction. Also, an allergic reaction after use of a eucalyptol-containing anti-inflammatory cream was reported (Vilaplana and Romaguera, 2000). A case report about a 24-year-old woman who spilled a small amount of clove oil on her face in an attempt to relieve a toothache is described by Nelson et al. (2011). She experienced permanent infraorbital anesthesia and anhidrosis. Ingestion of 1–2 teaspoon of clove oil in a 2-year-old boy resulted in metabolic acidosis, coma, seizures, hypoglycemia, and liver failure (Ford et al., 2001). Also, Janes et al. (2005) described the development of fulminant hepatic failure in a 15-monthold boy after ingestion of 10 mL clove oil. The hepatic impairment resolved after intravenous administration of N-acetylcysteine, and recovery occurred over the next four days.

19.10.2  Eugenol, Isoeugenol For some data, isoeugenol was used as a read-across for eugenol, because both belong to the generic class of phenols with common structures and the only differ in the position of the double bond in the alkene chain. It is assumed that the structural differences do not essentially change the physicochemical properties nor raise any additional structural alerts. In addition, they are predicted to have similar metabolites. Therefore, the toxicity profiles are expected to be similar. 19.10.2.1  Repeated Dose Toxicity The margin of exposure for eugenol is adequate for the repeated dose toxicity endpoint at the current level of use. From a dietary 13-week subchronic toxicity study conducted in rats, the no-observed-adverseeffect level (NOAEL) for repeated dose toxicity was determined to be 300 mg/kg/day for eugenol and 37.5 mg/kg/day for isoeugenol (Api et al., 2016b), based on reduced body weights. The Research Institute for Fragrance Materials (RIFM) and the US National Toxicology Program (US NTP) concluded that hepatocellular tumors observed following eugenol administration were considered to be associated with the dietary administration of eugenol, but because of the lack of a dose-response effect in male mice and the marginal combined increases in female mice, there was equivocal evidence of carcinogenicity (Api et al., 2016c). In rats, it was not carcinogenic. Similarly,

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TABLE 19.2 Repeated Dose Toxicity Eugenol

Isoeugenol

Total systemic exposure (mg/kg/day)

0.019

0.0065 Dermal uptake 38.4%

Lowest dose level in mouse NTP study

>23,600 times higher 300

>11,500 times higher 37.5

300/0.019 = 15789

37.5/0.0004 = 93,000

NOAEL (mg/kg/day) Margin of Exposure (mg/kg/day)

the US NTP reported that isoeugenol is hepatocarcinogenic in male mice at 75 mg/kg/day and equivocally carcinogenic in female mice and male rats (thymoma, mammary gland carcinoma) at 300 mg/kg/day (see Api et al., 2016b). However, hepatotoxicity might have played a role in the development of the hepatic tumors in B6C3F1 mice, which are known to be sensitive to the development of liver tumors by nongenotoxic mechanisms. Thus, it is assumed that these data are not relevant to human risk, as the increase in the incidence of tumors in male B6C3F1 mice reflects the impact of high-dose liver damage to an organ already prone to spontaneous development of liver neoplasms (Haseman et al., 1986) (Table 19.2). A margin of exposure (MOE) of >100 is deemed acceptable. 19.10.2.2  Developmental and Reproductive Toxicity A gavage developmental toxicity study with isoeugenol, conducted in rats, revealed intrauterine growth retardation and delayed skeletal ossification at maternally toxic dosages (George et al., 2001) (Table 19.3). Reproductive toxicity of isoeugenol was confirmed in rats in a gavage multigenerational continuous breeding study based on a decreased number of male pups per litter during the F0 cohabitation and decreased male and female pup weights during the F1 cohabitation (Layton et al., 2001) (Table 19.3). Although estimated values for margin of exposure (MOE) suggest no developmental and reproductive toxicity of isoeugenol. The developmental toxicity data on eugenol are insufficient. There are no reproductive toxicity data on eugenol. Taking the NOAEL of 230 mg/kg/day from the read across analogue isoeugenol and considering 22.6% absorption from skin contact and 100% from inhalation, a MOE of 12,105 was estimated. As a MOE of >100 is deemed acceptable, the margin of exposure for eugenol is adequate for the developmental and reproductive toxicity endpoints at the current level of use. 19.10.2.3  Skin Sensitization The available data demonstrate that eugenol is a weak sensitizer with a weight-of-evidence no-expected-sensitization-induction level (WoE NESIL) of 5900 mg/cm2 (Gerberick et al., 2009).

TABLE 19.3 Developmental Toxicity Eugenol

Isoeugenol

Gavage developmental toxicity study

No data

Gavage multigenerational continuous breeding study

No data

NOAEL: 500 mg/kg/day MOE: 1,250,000 NOAEL: 230 mg/kg/day MOE: 575,000

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19.10.3 Methyleugenol Methyleugenol, the methyl ether of eugenol, is a natural constituent of numerous essential oils in very low amounts. However, it is not allowed to be added to cosmetic products and beverages as it has been listed by the National Toxicology Program’s Report on Carcinogens as reasonably anticipated to be a human carcinogen. This finding is based on the observation of increased incidence of malignant tumors at multiple tissue sites in experimental animals of different species. High doses of methyleugenol (at least 30 mg/kg body weight for 25 days) cause autoinduction of 1′-hydroxylation by P450 cytochromes, with formation of the proximate carcinogen 1′-hydroxymethyleugenol (Jeurissien et  al., 2006). Methyleugenol and its two metabolites 1-hydroxymethyleugenol and 2′,3′-epoxymethyleugenol induced unscheduled DNA synthesis (UDS) in vitro. In addition, methyleugenol formed DNA adducts in vitro and in vivo (Cartus et  al., 2012; Herrmann et  al., 2013). Sipe et al. (2014) showed that both methyleugenol and eugenol readily undergo peroxidative metabolism in vitro to form free radicals. Due to auto-oxidation of methyleugenol, commercial products contain 10–30 mg/L hydroperoxide. Spectroscopic studies show that the hydroperoxide is not a good substrate for catalase, which suggests that glutathione peroxidase may be important in the inhibition of this pathway in vivo. Thus, the authors suggest that peroxidase metabolism may contribute to the observed carcinogenicity of methyleugenol in extrahepatic tissues (Sipe et al., 2014). Since methyleugenol has been demonstrated to be genotoxic and carcinogenic, the existence of a threshold cannot be assumed and the EU Scientific Committee on Food could not establish a safe exposure limit. Consequently, reductions in exposure and restrictions in use levels are indicated (European Commission Opinion of the Scientific Committee on Food, 2001).

19.11  BERGAMOT OIL The main components of volatile fraction of bergamot oil, a cold-pressed essential oil from the peel of bergamot (Citrus bergamia Risso et Poiteau) fruit, are limonene (25%–53%), linalyl acetate (15%–40%), linalool (2%–20%), γ-terpinene, and β-pinene (Sawamura et al., 2006; see Navarra et al., 2015). In the non-volatile fraction, the main compounds are coumarins and furanocoumarins such as bergapten (5-methoxypsoralen) and bergamottin (5-geranyloxypsoralen) (Dugo et al., 2000). The bergamot essential oil is produced in relatively small quantities and thus it is rather expensive. This makes it particularly subject to adulteration (Schipilliti et al., 2011) either by adding distilled essences of poor quality and low cost, for example of bitter orange and bergamot mint, or by adding mixtures of natural or synthetic terpenes. An increase in limonene accompanied by a decrease in linalool hints at adulteration with cheaper citrus oils. In addition, bergamot oil should not contain more than 1.5% p-cymene, which is an indicator for degradation and may cause dry skin and redness of eyes and skin, as well as drowsiness and nausea. In several studies, application of bergamot oil directly to the skin was shown to have a concentration-dependent phototoxic effect after exposure to UV light, presumably due to bergapten and also bergamottin (Kaddu et al., 2001; Kejlova et al., 2007). Cutaneous lesions developed gradually within 48–72 h when aromatherapy with bergamot oil was applied 2–3 days prior to ultraviolet (UV) exposure (Kaddu et al., 2001). Thus, for applications on areas of skin exposed to sunshine, excluding bath preparations, soaps, and other products which are washed off the skin, bergamot oil should not be used such that the level in the consumer products exceeds 0.4% (International Fragrance Association, IFRA). Cases have become much more rare since the introduction of psoralen-free bergamot oil. Earl Grey tea is black tea flavored with bergamot essential oil. In one case study, a patient who consumed four liters of Earl Grey tea per day, suffered from paresthesias, fasciculations, and muscle cramps (Finsterer, 2002). The author explained the adverse effects in this patient by the effect of bergapten, which is reported to be an axolemmal potassium channel blocker (Bohuslavizki et al., 1994; Wulff et al., 1998; During et al., 2000). This may lead to hyperexcitability of the axonal membrane, causing fasciculations and muscle cramps.

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19.12  ESSENTIAL OILS OF NUTMEG AND OTHER SPICES Myristicin (methoxysafrole) and elemecin, along with safrole, are present in small amounts in the essential oil of nutmeg and, to an even lesser extent, in other spices such as parsley and dill. The essential oil Myristicae aetheroleum is used in the form of balsam, pastilles, drops, and massage oil for the treatment of common cold, digestive disorders, and rheumatism. The main components myristicin and elemecin are psychoactive compounds with anticholinergic activity causing dry mouth and skin, obstipation, urinary retention, and tachycardia. Intoxication with high concentrations may cause severe hypotension and liver injury, as well as central nervous symptoms such as memory disturbances, visual distortions, excitation, confusion, anxiety, and hallucinations. Although misuse is voluntary, intoxication is invariably accidental. Although the risks of nutmeg intoxication after voluntary use are known, some people still use it for a low-cost recreational-drug alternative (Demetriades et al., 2005). In case reports (e.g., Abernethy and Becker, 1992; Sangalli and Chiang, 2000; Stein et al., 2001; Kelly and Clarke, 2003; McKenna et al., 2004), intoxications with nutmeg had effects that varied from person to person. With a slow onset of symptoms, the effects persisted for several hours. Aftereffects even lasted up to several days.

19.13 CONCLUSION To conclude, in most cases, essential oils are safe in use except in overdosage, wrong application route, and in hypersensitive persons: • Toxicity of essential oils is concentration-dependent, and thus most unwanted side effects can be avoided by application of low doses and restriction to small skin areas. • Hypersensitive people should avoid essential oils because allergic reactions occur independently of the applied concentration and route of application. • Adequate storage of essential oils is mandatory, because oxidation products formed by exposure to light and/or air may play an important role in toxicity. • Most severe intoxications occur in children following accidental ingestion. Subjects of special concern are the • Epileptogenic effect of camphor-, eucalyptol-, and thujone-containing essential oils • Severe hepatotoxicity of pennyroyal oil due to pulegone and menthofuran, and at high concentrations of clove oil due to eucalyptol • Salicylate intoxication with wintergreen oil • Phototoxicity of coumarin- and furocoumarin-containing bergamot oil • Application in pregnant women, because components of essential oils easily cross the placental barrier and may harm the fetus and neonates • The use particularly in infants and young children as they are prone to severe intoxication

REFERENCES Abanses, J.C., S. Arima, and B.K. Rubin. 2009. Vicks VapoRub induces mucin secretion, decreases ciliary beat frequency, and increases tracheal mucus transport in the ferret trachea. Chest, 135: 143–148. Abernethy, M.K., and L.B. Becker. 1992. Acute nutmeg intoxication. Am. J. Emerg. Med., 10: 429–430. Alpizar, Y.A., M. Gees, A. Sanchez, A. Apetrei, T. Voets, B. Nilius, and K. Talavera. 2013. Bimodal effects of cinnamaldehyde and camphor on mouse TRPA1. Pflugers Arch., 465: 853–864. Anderson, I.B., W.H. Mullen, J.E. Meeker, S.C. Khojasteh Bakht, S. Oishi, S.D. Nelson, and P.D. Blanc. 1996. Pennyroyal toxicity: Measurement of toxic metabolite levels in two cases and review of the literature. Ann. Intern. Med., 124: 726–734.

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Andersson, M., and M. Hindsen. 2007. Rhinitis because of toothpaste and other menthol-containing products. Allergy, 62: 336–337. Api, A.M., D. Belsito, S. Bhatia et al. 2016a. RIFM fragrance ingredient safety assessment, 2-Hydroxy-α,α,4trimethylcyclohexanemethanol, CAS Registry Number 42822–86–6. Food Chem. Toxicol., 97S: S209–S215. Api, A.M., D. Belsito, S. Bhatia et al. 2016b. RIFM fragrance ingredient safety assessment, isoeugenol, CAS Registry Number 97–54–1. Food Chem. Toxicol., 97S: S49–S56. Api, A.M., D. Belsito, S. Bhatia et al. 2016c. RIFM fragrance ingredient safety assessment, eugenol, CAS Registry Number 97–53–0. Food Chem. Toxicol., 97S: S25–S37. Araujo, I.B., C.A.M. Souza, R.R. De-Carvalho, S.N. Kuriyama, R.P. Rodrigues, R.S. Vollmer, E.N. Alves, and F.J.R. Paumgartten. 1996. Study of the embryo foetotoxicity of a-terpinene in the rat. Food Chem. Toxicol., 34: 477–482. Arikan-Ayyildiz, Z., F. Akgül, Ş. Yılmaz, D. Özdemir, and N. Uzuner. 2012. Anaphylaxis in an infant caused by menthol-containing cologne. Allergol. Immunopathol. (Madr), 40: 198. Bakerink, J.A., S.M. Gospe Jr., R.J. Dimand, and M.W. Eldridge. 1996. Multiple organ failure after ingestion of pennyroyal oil from herbal tea in two infants. Pediatrics, 98: 944–947. Barkin, M.E., J.P. Boyd, and S. Cohen. 1984. Acute allergic reaction to eugenol. Oral Surg. Oral Med. Oral Pathol., 57: 441–442. Bautista, D.M., S.E. Jordt, T. Nikai, P.R. Tsuruda, A.J. Read, J. Poblete, E.N. Yamoah, A.I. Basbaum, and D. Julius. 2006. TRPA1 mediates the inflammatory actions of environmental irritants and proalgesic agents. Cell, 124: 1269–1282. Bazzano, A.N., C. Var, F. Grossman, and R.A. Oberhelman. 2017. Use of camphor and essential oil balms for infants in Cambodia. J. Trop. Pediatr., 63: 65–69. Bischoff, K., and F. Guale. 1998. Australian tea tree (Melaleuca alternifolia) oil poisoning in three purebred cats. J. Vet. Diagn. Invest., 10: 208–210. Blagojevic, P., N. Radulovic, R. Palic, and G. Stojanovic. 2006. Chemical composition of the essential oils of Serbian wild-growing Artemisia absinthium and Artemisia vulgaris. J. Agr. Food Chem., 54: 4780–4789. Blake, K.D., C.R. Fertleman, and M.A. Meates. 1993. Dangers of common cold treatments in children [letter] [published erratum appears in Lancet 1993; 341(8848):842]. Lancet, 341: 640. Bohuslavizki, K.H., W. Hänsel, A. Kneip, E. Koppenhöfer, E. Niemöller, and K. Sanmann. 1994. Mode of action of psoralens, benzofurans, acridinons, and coumarins on the ionic currents in intact myelinated nerve fibres and its significance in demyelinating diseases. Gen. Physiol. Biophys., 13: 309–328. Burkhard, P.R., K. Burkhardt, C.A. Haenggeli, and T. Landis. 1999. Plant induced seizures: Reappearance of an old problem. J. Neurol., 246: 667–670. Caceres, A.I., M. Brackmann, M.D. Elia et al. 2009. A sensory neuronal ion channel essential for airway inflammation and hyperreactivity in asthma. Proc. Natl. Acad. Sci. USA, 106: 9099–9104. Cai L., and C.D. Wu. 1996. Compounds from syzygium aromaticum possessing growth inhibitory activity against oral pathogens. J. Nat. Prod. 59(10), 987–990. Carson, C.F., R. Tisser, and T. Larkman. 2014. Lack of evidence that essential oils affect puberty. Reprod. Toxicol., 44: 50–51. Cartus, A.T., K. Herrmann, L.W. Weishaupt, K.-H. Merz, W. Engst, H. Glatt, and D. Schrenk. 2012. Metabolism of methyleugenol in liver microsomes and primary hepatocytes: Pattern of metabolites, cytotoxicity, and DNA-adduct formation. Toxicol. Sci., 129: 21–34. Casarin, M., J. Pazinatto, R.C.V. Santos, and F.B. Zanatta. 2018. Melaleuca alternifolia and its application against dental plaque and periodontal diseases: A systematic review. Phytother. Res., 32: 230–242. Chaieb, K., H. Hajlaoui, T. Zmantar, A.B. Kahla-Nakbi, M. Rouabhia, K. Mahdouani, and A. Bakhrouf. 2007. The chemical composition and biological activity of clove essential oil, Eugenia caryophyllata (Syzigium aromaticum L. Myrtaceae): A short review. Phytother. Res., 21: 501–506. Chen, L.J., E.H. Lebetkin, and L.T. Burka. 2003. Metabolism of (R)-(+)-menthofuran in Fischer-344 rats: Identification of sulfonic acid metabolites. Drug Metab. Dispos., 31: 1208–1213. Committee on Drugs. 1978. Camphor: Who needs it? American Academy of Pediatrics. Pediatrics, 62: 404–406. Committee on Drugs. 1994. Camphor revisited. Focus on toxicity. American Academy of Pediatrics. Pediatrics, 94: 127–128. Council of Europe. 2001. Committee of Experts on Flavouring Substances of the Council of Europe. 48th Meeting, Strasbourg, 2–6 April 2001. Craig, J.O. 1953. Poisoning by the volatile oils in childhood. Arch. Dis. Child., 28: 475–483. Culic, M., G. Kekovic, G. Grbic, L. Martac, M. Sokovic, J. Podgorac, and S. Sekulic. 2009. Wavelet and fractal analysis of rat brain activity in seizures evoked by camphor essential oil and 1,8-cineole. Gen. Physiol. Biophys., 28: 33–40.

536

Handbook of Essential Oils

Daimon, H., S. Sawada, S. Asakura, and F. Sagani. 1998. In vivo genotoxicity and DNA adduct levels in the liver of rats treated with safrole. Carcinogenesis, 19: 141–146. Darben, T., B. Cominos, and C.T. Lee. 1998. Topical eucalyptus oil poisoning. Australas. J. Dermatol., 39: 265–267. Davis, J.E. 2007. Are one or two dangerous? Methyl salicylate exposure in toddlers. J. Emerg. Med., 32: 63–69. Del Beccaro, M.A. 1995. Melaleuca oil poisoning in a 17-month-old. Vet. Hum Toxicol., 37: 557–558. Demetriades, A.K., P.D. Wallman, A. McGuiness, and M.C. Gavalas. 2005. Low cost, high risk: Accidental nutmeg intoxication. Emerg. Med. J., 22: 223–225. De Souza, M.E., D.J. Clerici, C.M. Verdi et  al. 2017. Antimicrobial activity of Melaleuca alternifolia nanoparticles in polymicrobial biofilm in situ. Microb. Pathog., 113: 432–437. De Vincenzi, M., M. Silano, A. De Vincenzi, F. Maialetti, and B. Scazzocchio. 2002. Constituents of aromatic plants: Eucalyptol. Fitoterapia, 73: 269–275. Dietz, B., and J.L. Bolton. 2007. Botanical dDietary sSupplements gGone bBad. Chem. Res. Toxicol., 20: 586–590. Dost, S., and B. Leiber. 1967. Menthol and Menthol containing External Remedies. Use, Mode of Effect and Tolerance in Children. Stuttgart: George Thieme Verlag. Dugo, P., L. Mondello, L. Dugo, R. Stancanelli, and G. Dugo. 2000. LC-MS for the identification of oxygen heterocyclic compounds in citrus essential oils. J. Pharm. Biomed. Anal., 24: 147–150. Düring, T., F. Gerst, W. Hansel, H. Wulff, and E. Koppenhofer. 2000. Effects of three alkoxypsoralens on voltage gated ion channels in Ranvier nodes. Gen. Physiol. Biophys., 19: 345–364. Eccles, R. 1994. Menthol and related cooling compounds. J. Pharm. Pharmacol., 46: 618–630. Ellenhorn, M.J., and D.G. Barceloux. 1988. Medical Toxicology - Diagnosis and Treatment of Human Poisoning. New York, NY: Elsevier Science Publishing Co., Inc., p. 562. Elliott, C. 1993. Tea tree oil poisoning. Med. J. Aust., 159: 830–831. EMA. 2012. EMA/67070/2012. Assessment report for suppositories containing terpenic derivatives. 20 January 2012. Procedure number: EMEA/H/A-1284 (http://www.ema.europa.eu/docs/en_GB/document_library/ Referrals_document/Terpenic_31/WC500122526.pdf) EMA. 2014. EMA/HMPC/138386/2005. Rev. 1 Committee on Herbal Medicinal Products (HMPC) Public statement on the use of herbal medicinal products containing pulegone and menthofuran. November 24, 2014. (http://www.ema.europa.eu/docs/en_GB/document_library/Public_statement/2014/12/ WC500179556.pdf) Engel, W. 2003. In vivo studies on the metabolism of the monoterpene pulegone in humans using the metabolism of ingestion-correlated amounts (MICA) approach: Explanation for the toxicity differences between (S)(−) and (R)-(+)-pulegone. J. Agric. Food Chem., 51: 6589–6597. European Commission Opinion of the Scientific Committee on Food on Methyleugenol (4-Allyl-1,2dimethoxybenzene). 2001. SCF/CS/FLAV/FLAVOUR/4 ADD1 FINAL (http://europa.eu.int/comm/ food/fs/sc/scf/index_en.html) European Commission Opinion of the Scientific Committee on Food on pulegone and menthofuran. 2002. SCF/CS/FLAV/FLAVOUR/3 ADD2 Final (https://ec.europa.eu/food/sites/food/files/safety/docs/ sci-com_scf_out133_en.pdf) Farco, J.A., and O. Grundmann. 2013. Menthol - Pharmacology of an important naturally medicinal “cool”. Mini Rev. Med. Chem., 13: 124–131. Fennell, T.R., J.A. Miller, and E.C. Miller. 1984. Characterization of the biliary and urinary glutathione and N-acetylcysteine metabolites of the hepatic carcinogen 1′-hydroxysafrole and its 1′-oxo metabolite in rats and mice. Cancer Res., 44: 3231–3240. Finsterer, J. 2002. Earl Grey tea intoxication. Lancet, 359(9316): 1484. Flaman, Z., S. Pellechia-Clark, B. Bailey, and M. McGuigan. 2001. Unintentional exposure of young children to camphor and eucalyptus oils. Paediatr. Child. Health, 6: 80–83. Ford, M.D., K.A. Delaney, L.J. Ling, and T. Erickson. 2001. Clinical Toxicology. Philadelphia, PA: W.B. Saunders Company, p. 346. Gardiner, P. 2000. Peppermint (Mentha piperita). The Longwood Herbal Task Force and The Center for Holistic Pediatric Education and Research (http://www.mcp.edu/herbal/default.htm) Gaudioso, C., J. Hao, M.F. Martin-Eauclaire, M. Gabriac, and P. Delmas. 2012. Menthol pain relief through cumulative inactivation of voltage-gated sodium channels. Pain, 153: 473–484. Geller, R.J., D.A. Spyker, L.K. Garretson, and A.D. Rogol. 1984. Camphor toxicity: Development of a triage strategy. Vet. Hum. Toxicol., 26(Suppl 2): 8–10. George, J.D., C.J. Price, M.C. Marr, C.B. Myers, and G.D. Jahnke. 2001. Evaluation of the developmental toxicity of isoeugenol in Sprague-Dawley (CD) rats. Toxicol. Sci. Former Fundam. Appl. Toxicol., 60: 112–120.

Adverse Effects and Intoxication with Essential Oils

537

Gerberick G.F., J.A. Troutman, L.M. Foertsch, J.D. Vassallo, M. Quijano, R.L.M. Dobson, C. Goebel, and J-P. Lepoittevin. 2009. Investigation of peptide reactivity of pro-hapten skin snsitizers using a peroxidaseperoxide oxidation system. Toxico.l Sci. 112(1), 164–174. Gleason, M.N., R.E. Gosselin, H.C. Hodge, and R.P. Smith. 1969. Clinical Toxicology of Commercial Products, 3rd ed. Baltimore: Williams & Williams Co, pp. 56–57. Gordon, W.P., A.C. Huitric, C.L. Seth, R.H. McClanahan, and S.D. Nelson. 1987. The metabolism of the abortifacient terpene, (R)-(+)-pulegone, to a proximate toxin, menthofuran. Drug Metab. Dispos., 15: 589–594. Gouin, S., and H. Patel. 1996. Unusual cause of seizure. Pediatr. Emerg. Care, 12: 298–300. Gurr, F.W. 1965. Scroggie JG: Eucalyptus oil poisoning treated by dialysis and mannitol infusion, with an appendix on the analysis of biological fluids for alcohol and eucalyptol. Australas. Ann. Med., 14: 238–249. Ha, M.A., G.J. Smith, J.A. Cichocki, L. Fan, Y.S. Liu, A.I. Caceres, S.E. Jordt, and J.B. Morris. 2015. Menthol attenuates respiratory irritation and elevates blood cotinine in cigarette smoke exposed mice. PLOS ONE, 10: e0117128. Halicioglu, O., G. Astarcioglu, I. Yaprak, and H. Aydinlioglu. 2011. Toxicity of Salvia officinalis in a newborn and a child: An alarming report. Pediatr. Neurol., 45: 259–260. Hammer, K.A., C.F. Carson, T.V. Riley, and J.B. Nielsen. 2006. A review of the toxicity of Melaleuca alternifolia (tea tree) oil. Food Chem. Toxicol., 44: 616–625. Hans, M., M. Wilhelm, and D. Swandulla. 2012. Menthol suppresses nicotinic acetylcholine receptor functioning in sensory neurons via allosteric modulation. Chem. Senses, 37: 463–469. Haseman, J.K., J.S. Winbush, and M.W. O’Donnell Jr. 1986. Use of dual control groups to estimate false positive rates in laboratory animal carcinogenicity studies. Fundam. Appl. Toxicol., 7: 573–584. Hausen, B.M. 2004. Evaluation of the main contact allergens in oxidized tea tree oil. Dermatitis, 15: 213–214. Hausen, B.M., J. Reichling, and M. Harkenthal. 1999. Degradation products of monoterpenes are the sensitizing agents in tea tree oil. Am. J. Contact Dermat., 10: 68–77. Hawthorn, M., J. Ferrante, E. Luchowski, A. Rutledge, X.Y. Wei, and D.J. Triggle. 1988. The actions of peppermint oil and menthol on calcium channel dependent processes in intestinal, neuronal and cardiac preparations. Aliment. Pharmacol. Ther., 2: 101–118. Heimes, K., F. Hauk, and E.J. Verspohl. 2011. Mode of action of peppermint oil and (−)-menthol with respect to 5-HT3 receptor subtypes: Binding studies, cation uptake by receptor channels and contraction of isolated rat ileum. Phytother. Res., 25: 702–708. Henley, D.V., N. Lipson, K.S. Korach, and C.A. Bloch. 2007. Prepubertal gynecomastia linked to lavender and tea tree oils. N. Engl. J. Med., 356: 479–485. Herrmann, K., F. Schumacher, W. Engst, K.E. Appel, K. Klein, U.M. Zanger, and H. Glatt. 2013. Abundance of DNA adducts of methyleugenol, a rodent hepatocarcinogen, in human liver samples. Carcinogenesis, 34: 1025–1030. Hindle, R.C. 1994. Eucalyptus oil ingestion. N. Z. Med. J., 107: 185–186. Hold, K.M., N.S. Sirisoma, T. Ikeda, T. Narahashi, and J.E. Casida. 2000. α-Thujone (the active component of absinthe): γ-Aminobutyric acid type A receptor modulation and metabolic detoxification. Proc. Natl. Acad. Sci. USA, 97: 3826–3831. Hu, Z., R. Feng, F. Xiang et al. 2014. Acute and subchronic toxicity as well as evaluation of safety pharmacology of eucalyptus oil-water emulsions. Int. J. Clin. Exp. Med., 7: 4835–4845. inchem.org (http://www.inchem.org/documents/pims/pharm/camphor.htm#SectionTitle) Jacobs, M.R., and C.S. Hornfeldt. 1994. Melaleuca oil poisoning. J. Toxicol. Clin. Toxicol., 32: 461–464. Janes, S.E., C.S. Price, and D. Thomas. 2005. Essential oil poisoning: N-acetylcysteine for eugenol-induced hepatic failure and analysis of a national database. Eur. J. Pediatr., 164: 520–522. Javorka, K., Z. Tomori, and L. Zavarská. 1980. Protective and defensive airway reflexes in premature infants. Physiol. Bohemoslov., 29: 29–35. Jeurissien, S.M.F., J.J.P. Bogaards, M.G. Boersma et al. 2006. Human cytochrome P450 enzymes of importance for the bioactivation of methyleugenol to the proximate carcinogen 1′-hydroxymethyleugenol. Chem. Res. Toxicol., 19: 111–116. Jimenez, J.F., A.L. Brown, W.C. Arnold, and W.J. Byrne. 1983. Chronic Camphor Ingestion Mimicking Reye’s Syndrome. Gastroenterology, 84: 394–398. Johnson, P.N. 1985. Methyl salicylate/aspirin equivalence. Vet. Hum. Toxicol., 26: 317–318. Kaddu, S., H. Kerl, and P. Wolf. 2001. Accidental bullous phototoxic reactions to bergamot aromatherapy oil. J. Am. Acad. Dermatol., 45: 458–461. Kamatou, G.P., I. Vermaak, A.M. Viljoen, and B.M. Lawrence. 2013. Menthol: A simple monoterpene with remarkable biological properties. Phytochemistry, 96: 15–25.

538

Handbook of Essential Oils

Karashima, Y., N. Damann, J. Prenen, K. Talavera, A. Segal, T. Voets, and B. Nilius. 2007. Bimodal action of menthol on the transient receptor potential channel TRPA1. J. Neurosci., 27: 9874–9884. Karunakara, B.P., and C.S. Jyotirmanju. 2012. Eucalyptus oil poisoning in children. J. Pediatr. Sci., 4: e132. Kawane, H. 1996. Menthol and aspirin induced asthma. Respir. Med., 90: 247. Kejlova, K., D. Jirova, H. Bendova, H. Kandarova, Z. Weidenhoffer, H. Kolarova, and M. Liebsch. 2007. Phototoxicity of bergamot oil assessed by in vitro techniques in combination with human patch tests. Toxicol. In Vitro, 21: 1298–1303. Kelly, B.D., and M.M. Clarke. 2003. Nutmeg and psychosis. Schizophr. Res., 60: 95–96. Khine, H., D. Weiss, N. Graber, R.S. Hoffman, N. Esteban-Cruciani, and J.R. Avner. 2009. A cluster of children with seizures caused by camphor poisoning. Pediatrics, 123: 1269–1272. Khojasteh-Bakht, S.C., L.L. Koenigs, R.M. Peter, W.F. Trager, and S.D. Nelson. 1998. (R)-(+)-Menthofuran is a potent, mechanism-based inactivator of human liver cytochrome P450 2A6. Drug Metab. Dispos., 26: 701–704. Khojasteh-Bakht, S.C., S.D. Nelson, and W.M. Atkins. 1999. Glutathione S-transferase catalyzes the isomerization of (R)-2-hydroxymenthofuran to mintlactones. Arch. Biochem. Biophys., 370: 59–65. Kogel, U., B. Titz, W.K. Schlage et al. 2016. Evaluation of the Tobacco Heating System 2.2. Part 7: Systems toxicological assessment of a mentholated version revealed reduced cellular and molecular exposure effects compared with mentholated and non-mentholated cigarette smoke. Regul. Toxicol. Pharmacol., 81(Suppl 2): S123–S138. Kolassa, N. 2013. Menthol differs from other terpenic essential oil constituents. Regul. Toxicol. Pharmacol., 65: 115–118. Köppel, C., J. Tenczer, Th. Schirop, and K. Ibe. 1982. Camphor poisoning. Abuse of camphor as a stimulant. Arch. Toxicol., 51: 101–106. Kumar, K.J., S. Sonnathi, C. Anitha, and M. Santhoshkumar. 2015. Eucalyptus oil poisoning. Toxicol. Int., 22(1): 170–171. Kumar, P., V.M. Caradonna-Graham, S. Gupta, X. Ci, P.N. Rao, and J. Thompson. 1995. Inhalation challenge effects of perfume scent strips in patients with asthma. Ann. Allergy Asthma Immunol., 75: 429–433. Layton, K.A., G.W. Wolfe, Y. Wang, J. Bishop, and R.E. Chaping. 2001. Reproductive effects of isoeugenol in Sprague-Dawley rats when assessed by the continuous breeding protocol. Toxicologist, 60(1): 384. Lessenger, J.E. 2001. Occupational acute anaphylactic reaction to assault by perfume spray in the face. J. Am. Board Fam. Pract., 14: 137–140. Liu, B., L. Fan, S. Balakrishna, A. Sui, J.B. Morris, and S.E. Jordt. 2013. TRPM8 is the principal mediator of menthol-induced analgesia of acute and inflammatory pain. Pain, 154(10): 2169–2177. Lucas, G.N. 2000. Acute drug poisoning in children. Sri Lanka J. Child Health, 29: 45–48. MacDougall, J.M., K. Fandrick, X. Zhang, S.V. Serafin, and J.R. Cashman. 2003. Inhibition of human liver microsomal (S)-nicotine oxidation by (–)-menthol and analogues. Chem. Res. Toxicol., 16: 988–993. Madyastha, K.M., and B. Moorthy. 1989. Pulegone mediated hepatotoxicity: Evidence for covalent binding of R(+)-[14C]pulegone to microsomal proteins in vitro. Chem. Biol. Interact., 72: 325–333. Madyastha, K.M., and C.P. Raj. 2002. Stereoselective hydroxylation of 4-methyl-2-cyclohexenone in rats: Its relevance to R-(+)-pulegone-mediated hepatotoxicity. Biochem. Biophys. Res. Commun., 297(2): 202–205. Manoguerra, A.S., A.R. Erdman, P.M. Wax et al. 2006. Camphor Poisoning: An evidence-based practice guideline for out-of-hospital management. Clin. Toxicol (Phila), 44(4): 357–370. Marlowe, K.F. 2003. Urticaria and asthma exacerbations after ingestion of menthol-containing lozanges. Am. J. Health Syst. Pharm., 60: 1657–1659. McClanahan, R.H., D. Thomassen, J.T. Slattery, and S.D. Nelson. 1989. Metabolic activation of (R)-(+)-pulegone to a reactive enonal that covalently binds to mouse liver proteins. Chem. Res. Toxicol., 2: 349–355. McKenna, A., S.P. Nordt, and J. Ryan. 2004. Acute nutmeg poisoning. Eur. J. Emerg. Med., 11(4): 240–241. Melis, K., A. Bochner, and G. Janssens. 1989. Accidental nasal eucalyptol and menthol instillation. Eur. J. Pediatr., 148: 786–787. Millet, Y., J. Jouglard, M.D. Steinmetz, P. Tognetti, P. Joanny, and J. Arditti. 1981. Toxicity of some essential plant oils. Clinical and experimental study. Clin. Toxicol., 18: 1485–1498. Moqrich, A., S.W. Hwang, T.J. Earley, M.J. Petrus, A.N. Murray, K.S. Spencer, M. Andahazy, G.M. Story, and A. Patapoutian. 2005. Impaired thermosensation in mice lacking TRPV3, a heat and camphor sensor in the skin. Science, 307(5714): 1468–1472. Morris, M.C., A. Donoghue, J.A. Markowitz, and K.C. Osterhoudt. 2003. Ingestion of tea tree oil (Melaleuca oil) by a 4-year-old boy. Pediatr. Emerg. Care, 19: 169–171. Morton, C.A., J. Garioch, P. Todd, P.J. Lamey, and A. Forsyth. 1995. Contact sensitivity to menthol and peppermint in patients with intra-oral symptoms. Contact Dermatitis, 32: 281–284.

Adverse Effects and Intoxication with Essential Oils

539

Mozelsio, N.B., K.E. Harris, K.G. McGrath, and L.C. Grammer. 2003. Immediate systemic hypersensitivity reaction associated with topical application of Australian tea tree oil. Allergy Asthma Proc., 24: 73–75. Nair, B. 2001. Final report on the safety assessment of Mentha piperita (peppermint) oil, Mentha piperita (peppermint) leaf extract, Mentha piperita (peppermint) leaf, and Mentha piperita (peppermint) leaf water. Int. J. Toxicol., 20(Suppl 3): 61–73. Najafi-Taher, R., B. Ghaemi, S. Kharazi, S. Rasoulikoohi, and A. Amani. 2018. Promising antibacterial effects of silver nanoparticle-loaded tea tree oil nanoemulsion: A synergistic combination against resistance threat. AAPS PharmSciTech, 19(3): 1133–1140. Nash, P., S.R. Gould, and D.E. Bernardo. 1986. Peppermint oil does not relieve the pain of irritable bowel syndrome. Br. J. Clin. Pract., 40: 292–293. Nath, S.S., C. Pandey, and D. Roy. 2012. A near fatal case of high dose peppermint oil ingestion - Lessons learnt. Indian J. Anaesth., 56(6): 582–584. National Cancer Institute. 1979. Carcinogenesis, Technical Report Series No. 98. Bioassay of dl-menthol for possible carcinogenicity (https://ntp.niehs.nih.gov/ntp/htdocs/lt_rpts/tr098.pdf) Navarra, M., C. Mannucci, M. Delbò, and G. Calapai. 2015. Citrus bergamia essential oil: From basic research to clinical application. Front. Pharmacol., 6: 36. Nelson, L., N. Lewin, M.A. Howland, R. Hoffman, L. Goldfrank, and N. Flomenbaum. 2011. Goldfrank’s Toxicologic Emergencies, 9th ed. New York, N.Y.: McGraw-Hill, p. 625. Noiles, K., and M. Pratt. 2010. Contact dermatitis to Vicks VapoRub. Dermatitis, 21(3): 167–169. Olowe, S.A., and O. Ransome-Kuti. 1980. The risk of jaundice in glucose-6-phosphate dehydrogenase deficient babies exposed to menthol. Acta Paediatr Scand, 69: 341–345. Olsen, R.W. 2000. Absinthe and g-aminobutyric acid receptors. P. Natl. Acad. Sci. USA, 97: 4417–4418. O’Mullane, N.M., P. Joyce, S.V. Kamath, M.K. Tham, and D. Knass. 1982. Adverse CNS effects of mentholcontaining olbas oil. Lancet, 1(8281): 1121. Opdyke, D.L.J. 1978. Camphor USP in monographs on fragrance raw materials. Food Cosmet .Toxicol., 16(Suppl 1): 665–1669. Opinion of the Scientific Committee on Food on eucalyptol. 2002. (https://ec.europa.eu/food/sites/food/files/ safety/docs/sci-com_scf_out126_en.pdf) Opinion of the Scientific Committee on Food on the safety of the presence of safrole. 2002. (https://ec.europa. eu/food/sites/food/files/safety/docs/sci-com_scf_out116_en.pdf) Oviedo, A., S. Lebrun, U. Kogel et al. 2016. Evaluation of the Tobacco Heating System 2.2. Part 6: 90–day OECD 413 rat inhalation study with systems toxicology endpoints demonstrates reduced exposure effects of a mentholated version compared with mentholated and non-mentholated cigarette smoke. Regul. Toxicol. Pharmacol., 81(Suppl 2): S93–S122. Paiva, M., S. Piedade, and A. Gaspar. 2010. Toothpaste-induced anaphylaxis caused by mint (Mentha) allergy. Allergy, 65: 1201–1202. Pan, R., Y. Tian, R. Gao, H. Li, X. Zhao, J.E. Barrett, and H. Hu. 2012. Central mechanisms of menthol-induced analgesia. J. Pharmacol. Exp. Ther., 343(3): 661–672. Parys, B.T. 1983. Chemical burns resulting from contact with peppermint oil mar: A case report. Burns Incl. Therm Inj., 9: 374–375. Patel, S., and J. Wiggins. 1980. Eucalyptus oil poisoning. Arch. Dis. Child., 55(5): 405–406. Peier, A.M., A. Moqrich, A.C. Hergarden et al. 2002. A TRP channel that senses cold stimuli and menthol. Cell, 108(5): 705–715. Pereira, T.S., J.R. de Sant’anna, E.L. Silva, A.L. Pinheiro, and M.A. de Castro-Prado. 2014. In vitro genotoxicity of Melaleuca alternifolia essential oil in human lymphocytes. J. Ethnopharmacol., 151(2): 852–857. Phelan, W.J. 1976. Camphor poisoning: Over-the-counter dangers. Pediatrics, 57, 428–431. Radulović, N.S., M.S. Genčić, N.M. Stojanović, P.J. Randjelović, Z.Z. Stojanović-Radić, and N.I. Stojiljković. 2017. Toxic essential oils. Part V: Behaviour modulating and toxic properties of thujones and thujonecontaining essential oils of Salvia officinalis L., Artemisia absinthium L., Thuja occidentalis L. and Tanacetum vulgare L. Food Chem. Toxicol., 105: 366–369. Ragucci, K.R., P.R. Trangmar, J.G. Bigby, and T.D. Detar. 2007. Camphor ingestion in a 10-year-old male. South. Med. J., 100: 204–207. Rahimi, M., F. Shokri, H. Hassanian-Moghaddam, N. Zamani, A. Pajoumand, and S. Shadnia. 2017. Severe camphor poisoning, a seven-year observational study. Environ. Toxicol. Pharmacol., 52: 8–13. Rees, W.D., B.K. Evans, and J. Rhodes. 1979. Treating irritable bowel syndrome with peppermint oil. Br. Med. J., 2: 835–836. Rietjens, I.M., M.J. Martena, M.G. Boersma, W. Spiegelenberg, and G.M. Alink. 2005. Molecular mechanisms of toxicity of important food-borne phytotoxins. Mol. Nutr. Food Res., 49: 131–158.

540

Handbook of Essential Oils

Ruha, A.M., K.A. Graeme, and A. Field. 2003. Late seizure following ingestion of Vicks VapoRub. Acad. Emerg. Med., 10: 691. Russell, M. 1999. Toxicology of tea tree oil. In: Southwell, I., and Lowe, R. (Eds.), Tea Tree: The Genus Melaleuca. Amsterdam: Harwood Academic Publishers, pp. 191–201. Sainio, E.L., and L. Kanerva. 1995. Contact allergens in toothpastes and a review of their hypersensitivity. Contact Dermatitis, 33: 100–105. Sangalli, B.C., and W. Chiang. 2000. Toxicology of nutmeg abuse. J. Toxicol. Clin. Toxicol., 38(6): 671–678. Santos, C.D., and J.C. Cabot. 2015. Persistent effects after camphor ingestion: A case report and literature review. J. Emerg. Med., 48(3): 298–304. Sarrami, N., M.N. Pemberton, M.H. Thornhill, and E.D. Theaker. 2002. Adverse reactions associated with the use of eugenol in dentistry. Br. Dent. J., 193(5): 257–259. Sawamura, M., Y. Onishi, J. Ikemoto, N.T.M. Tu, and N.T.L. Phi. 2006. Characteristic odour components of bergamot (Citrus bergamia Risso) essential oil. Flavour Fragr. J., 21(4): 609–615. Schaller, M., and H.C. Korting. 1995. Allergic airborne contact dermatitis from essential oils used in aromatherapy. Clin. Exp. Dermatol., 20(2): 143–145. Schipilliti, L., P. Dugo, L. Santi, G. Dugo, and L. Mondello. 2011. Authentication of bergamot essential oil by gaschromatography-combustion-isotope ratio mass spectrometer (GC-C-IRMS). J. Essent. Oil Res., 23: 60–671. Seawright, A. 1993. Tea tree oil poisoning - comment. Med. J. Aust., 159: 831. Selescu, T., A.C. Ciobanu, C. Dobre, G. Reid, and A. Babes. 2013. Camphor activates and sensitizes transient receptor potential melastatin 8 (TRPM8) to cooling and icilin. Chem. Senses, 38: 563–575. Seneviratne, M.P., S. Karunarathne, A.H. de Alwis, A.H.N. Fernando, and R. Fernando. 2015. Accidental methyl salicylate poisoning in two adults. Ceylon Med. J., 60: 65. Sidell, N., M.A. Verity, and E.P. Nord. 1990. Menthol blocks dihydropyridine-insensitive Ca 2+ channels and induces neurite outgrowth in human neuroblastoma cells. J. Cell. Physiol., 142: 410–419. Siegel, E., and S. Wason. 1986. Camphor toxicity. Pediatr. Clin. North Am., 33(2): 375–379. Sipe, H.J. Jr., O.M. Lardinois, and R.P. Mason. 2014. Free radical metabolism of methyleugenol and related compounds. Chem. Res. Toxicol., 27(4): 483–489. Smith A.G., and G. Margolis. 1954. Camphor poisoning; Anatomical and pharmacological study; report of a fatal case; experimental investigation of protective action of barbiturate. Am J Pathol, 30(5), 857–869. Southwell, I.A., S. Freeman, and D. Rubel. 1997. Skin irritancy of tea tree oil. J. Essent. Oil Res., 9: 47–852. Spindler, P., and C. Madsen. 1992. Subchronic toxicity study of peppermint oil in rats. Toxicol. Lett., 62(2–3): 215–220. Spinella, M. 2001. Herbal medicines and epilepsy: The potential for benefit and adverse effects. Epilepsy Behav., 2: 524–532. Squier, C.A., M.J. Mantz, and P.W. Wertz. 2010. Effect of menthol on the penetration of tobacco carcinogens and nicotine across porcine oral mucosa ex vivo. Nicotine Tob. Res., 12(7): 763–767. Stafstrom, C.E. 2007. Seizures in a 7-month-old child after exposure to the essential plant oil thuja. Pediatr. Neurol., 37: 446–448. Stein, U., H. Greyer, and H. Hentschel. 2001. Nutmeg (myristicin) poisoning - Report on a fatal case and a series of cases recorded by a poison information centre. Forensic. Sci. Int., 118(1): 87–90. Swandulla, D., E. Carbone, K. Schäfer, and H.D. Lux. 1987. Effect of menthol on two types of Ca currents in cultured sensory neurons of vertebrates. Pflugers Arch., 409: 52–59. Sztajnkrycer, M.D., E.J. Otten, G.R. Bond, C.J. Lindsell, and R.J. Goetz. 2003. Mitigation of pennyroyal oil hepatotoxicity in the mouse. Acad. Emerg. Med., 10: 1024–1028. The EFSA Journal. 2005. 298:1–32. Opinion of the Scientific Panel on Food Additives, Flavourings, Processing Aids and Materials in contact with Foods on a request from the Commission on Pulegone and Menthofuran in flavourings and other food ingredients with flavouring properties. Question number EFSA-Q-2003–119 (http://www.efsa.europa.eu/sites/default/files/scientific_output/files/main_documents/298.pdf) The EFSA Journal. 2008. 729:1–15. Camphor in flavourings and other food ingredients with flavouring properties Opinion of the Scientific Panel on Food Additives, Flavourings, Processing Aids and Materials in Contact with Food on a request from the Commission (http://www.efsa.europa.eu/sites/default/files/ scientific_output/files/main_documents/729.pdf) Thomassen, D., J.T. Slattery, and S.D. Nelson. 1990. Menthofuran dependent and independent aspects of pulegone hepatotoxicity: Roles of glutathione. J. Pharmacol. Exp. Ther., 253: 567–572. Thompson, M.F., G.L. Poirier, M.I. Dávila-García et al. 2017. Menthol enhances nicotine-induced locomotor sensitization and in vivo functional connectivity in adolescence. J. Psyhopharmacol., 32(3): 332–343. Tibballs, J. 1995. Clinical effects and management of eucalyptus oil ingestion in infants and young children. Med. J. Aust., 163(4): 177–180.

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Turić, P., J. Lipozencić, V. Milavec-Puretić, and S.M. Kulisić. 2011. Contact allergy caused by fragrance mix and Myroxylon pereirae (balsam of Peru)--A retrospective study. Coll. Antropol., 35(1): 83–87. Uc, A., W.P. Bishop, and K.D. Sanders. 2000. Camphor hepatotoxicity. South. Med. J., 93(6): 596–8. Vilaplana, J., and C. Romaguera. 2000. Allergic contact dermatitis due to eucalyptol in an anti-inflammatory cream. Contact Dermatitis, 43(2): 118. Villar, D., M.J. Knight, S.R. Hansen, and W.B. Buck. 1994. Toxicity of melaleuca oil and related essential oils applied topically on dogs and cats. Vet. Hum. Toxicol., 36: 139–142. Von Bruchhausen, F., G. Dannhardt, S. Ebel, and A. Frahm. 1993. Hagers Handbuch der Pharmazeutischen Praxis. Bd. 7. Stoffe A-D, 5. Auflage, Springer Verlag Berlin Heidelberg, 649–650. Webb, N.J.A., and W.R. Pitt. 1993. Eucalyptus oil poisoning in childhood: 41 cases in south-east Queensland. J. Paediatr. Child Health, 29(5): 368–371. Weston, C.F. 1987. Anal burning and peppermint oil [letter]. Postgrad. Med. J., 63: 717. Whitman, B.W., and H. Ghazizadeh. 1994. Eucalyptus oil: Therapeutic and toxic aspects of pharmacology in humans and animals. J. Paediatr. Child Health, 30(2): 190–191. Wilkinson, S.M., and M.H. Beck. 1994. Allergic contact dermatitis from menthol in peppermint. Contact Dermatitis, 30: 42–43. Woolf, A. 1999. Essential oil poisoning. Clin. Toxicol., 37: 721–727. Wulff, H., H. Rauer, T. Düring, C. Hanselmann, K. Ruff, A. Wrisch, S. Grissmer, and W. Hänsel. 1998. Alkoxypsoralens, novel nonpeptide blockers of Shaker-type K+ channels: Synthesis and photoreactivity. J. Med. Chem., 41(23): 4542–4549. Wyllie, J.P., and F.W. Alexander. 1994. Nasal instillation of “olbas oil” in an infant. Arch. Dis. Child., 70: 357–358. Xu, H., N.T. Blair, and D.E. Clapham. 2005. Camphor activates and strongly desensitizes the transient receptor potential vanilloid subtype 1 channel in a vanilloid-independent mechanism. J. Neurosci., 25(39): 8924–8937. Xu, J., Z.Q. Hu, C. Wang et al. 2014. Acute and subacute toxicity study of 1,8-cineole in mice. Int. J. Clin. Exp. Pathol., 7(4): 1495–1501. Yosipovitch, G., C. Szolar, X.Y. Hui, and H. Maibach. 1996. Effect of topically applied menthol on thermal, pain and itch sensations and biophysical properties of the skin. Arch. Dermatol. Res., 288(5–6): 245–248. Zhang, S.Y., and D. Robertson. 2000. A study of tea tree oil ototoxicity. Audiol. Neurootol., 5: 64–68. Zhang, X.B., P. Jiang, N. Gong, X.L. Hu, D. Fei, Z.Q. Xiong, L. Xu, and T.L. Xu. 2008. A-type GABA receptor as a central target of TRPM8 agonist menthol. PLOS ONE, 3: e3386.

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Adulteration of Essential Oils Erich Schmidt and Jürgen Wanner

CONTENTS 20.1 Introduction......................................................................................................................... 545 20.1.1 General Remarks.................................................................................................. 545 20.2 Definition and History.........................................................................................................546 20.3 Adulteration......................................................................................................................... 549 20.3.1 Unintended Adulteration...................................................................................... 549 20.3.2 Intentional Adulteration........................................................................................ 550 20.3.3 Prices.................................................................................................................... 551 20.3.4 Availability........................................................................................................... 551 20.3.5 Demand of Clients................................................................................................ 551 20.3.6 Regulations........................................................................................................... 551 20.3.7 Aging.................................................................................................................... 552 20.3.8 Cupidity................................................................................................................ 552 20.3.9 Simple Sports?...................................................................................................... 552 20.4 Possible Adulterations for Essential Oils............................................................................ 553 20.4.1 Water..................................................................................................................... 553 20.4.2 Ethanol.................................................................................................................. 553 20.4.3 Fatty Oils or Mineral Oils.................................................................................... 553 20.4.4 High Boiling Glycols............................................................................................ 553 20.4.5 Oils from Other Parts of the Same Species or Other Species with Similar Essential Oil Composition.................................................................................... 554 20.4.6 Related Botanical Species.................................................................................... 554 20.4.7 Fractions of Essential Oils.................................................................................... 554 20.4.8 Natural Isolates..................................................................................................... 555 20.4.9 Chemically Derived Synthetic Compounds, Which Are Proved to Appear in Nature................................................................................................................... 555 20.4.10 Steam Distilled Residues from Expression.......................................................... 556 20.4.11 Enzymatically Produced Chemicals (Natural by Law)........................................ 556 20.5 Methods to Detect Adulterations........................................................................................ 556 20.5.1 Organoleptic Methods.......................................................................................... 556 20.5.1.1 Appearance and Color........................................................................ 556 20.5.1.2 Odor.................................................................................................... 557 20.5.1.3 Physical–Chemical Methods.............................................................. 557 20.5.1.4 Calculation of Relationship Coefficient.............................................. 558 20.5.2 Analytical Methods.............................................................................................. 558 20.5.2.1 General Tests....................................................................................... 558 20.5.2.2 Thin-Layer Chromatography.............................................................. 559 20.5.2.3 Gas Chromatography (GC, GLC, HRGC, GC-FID, GC-MS)............ 559 20.5.2.4 Chiral Analysis................................................................................... 561 20.5.2.5 GC-GC and GC × GC (Two-Dimensional Gas Chromatography, 2D GC)................................................................... 561 20.5.2.6 13C NMR (Nuclear Magnetic Resonance).......................................... 562

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20.6 Important Essential Oils and Their Possible Adulteration.................................................. 562 20.6.1 Ambrette Seed Oil................................................................................................ 562 20.6.2 Amyris Oil............................................................................................................ 562 20.6.3 Angelica Oils........................................................................................................ 562 20.6.4 Anise Fruit Oil...................................................................................................... 562 20.6.5 Armoise Oil.......................................................................................................... 563 20.6.6 Basil Oils.............................................................................................................. 563 20.6.7 Bergamot Oil........................................................................................................ 563 20.6.8 Bitter Orange Oil.................................................................................................. 563 20.6.9 Bitter Orange Petitgrain Oil.................................................................................564 20.6.10 Cajeput Oil............................................................................................................564 20.6.11 Camphor Oil.........................................................................................................564 20.6.12 Cananga Oil..........................................................................................................564 20.6.13 Caraway Oil.......................................................................................................... 565 20.6.14 Cardamom Oil...................................................................................................... 565 20.6.15 Cassia Oil.............................................................................................................. 565 20.6.16 Cedar Leaf Oil...................................................................................................... 565 20.6.17 Cedarwood Oils.................................................................................................... 565 20.6.18 Celery Seed Oil..................................................................................................... 565 20.6.19 Chamomile Oil Blue............................................................................................. 566 20.6.20 Chamomile Oil Roman......................................................................................... 566 20.6.21 Cinnamon Bark Oil.............................................................................................. 566 20.6.22 Cinnamon Leaf Oil............................................................................................... 566 20.6.23 Citronella Oil........................................................................................................ 566 20.6.24 Clary Sage Oil...................................................................................................... 566 20.6.25 Clove Oils............................................................................................................. 566 20.6.26 Coriander Fruit Oil............................................................................................... 567 20.6.27 Corymbia Citriodora Oil....................................................................................... 567 20.6.28 Corn Mint Oil....................................................................................................... 567 20.6.29 Cumin Fruit Oil.................................................................................................... 567 20.6.30 Cypress Oil........................................................................................................... 567 20.6.31 Dill Oils................................................................................................................ 567 20.6.32 Dwarf Pine Oil..................................................................................................... 568 20.6.33 Elemi Oil.............................................................................................................. 568 20.6.34 Eucalyptus Oil...................................................................................................... 568 20.6.35 Fennel Oil Sweet................................................................................................... 568 20.6.36 Fennel Oil Bitter................................................................................................... 568 20.6.37 Geranium Oils...................................................................................................... 568 20.6.38 Grapefruit Oil....................................................................................................... 569 20.6.39 Juniper Berry Oil.................................................................................................. 569 20.6.40 Lavandin Oils....................................................................................................... 569 20.6.41 Lavender Oil......................................................................................................... 570 20.6.42 Lemon Oil............................................................................................................. 570 20.6.43 Lemongrass Oil.................................................................................................... 570 20.6.44 Lime Oil Distilled................................................................................................. 570 20.6.45 Lime Oil Expressed.............................................................................................. 571 20.6.46 Litsea cubeba Oil.................................................................................................. 571 20.6.47 Mandarin Oil........................................................................................................ 571 20.6.48 Melissa Oil (Lemon Balm)................................................................................... 571 20.6.49 Mentha citrata Oil................................................................................................ 572 20.6.50 Mountain Pine Oil................................................................................................ 572

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20.6.51 Neroli Oil.............................................................................................................. 572 20.6.52 Nutmeg Oil........................................................................................................... 572 20.6.53 Orange Oil Sweet.................................................................................................. 572 20.6.54 Origanum Oil........................................................................................................ 573 20.6.55 Palmarosa Oil....................................................................................................... 573 20.6.56 Parsley Oil............................................................................................................ 573 20.6.57 Pine Oil Siberian.................................................................................................. 573 20.6.58 Patchouli Oil......................................................................................................... 573 20.6.59 Pepper Oil............................................................................................................. 573 20.6.60 Peppermint Oil..................................................................................................... 573 20.6.61 Petitgrain Oil Paraguay Type............................................................................... 574 20.6.62 Pimento Oils......................................................................................................... 574 20.6.63 Rose Oil................................................................................................................ 574 20.6.64 Rosemary Oil........................................................................................................ 574 20.6.65 Rosewood Oil....................................................................................................... 574 20.6.66 Sage Oil (Salvia officinalis).................................................................................. 575 20.6.67 Sage Oil Spanish Type.......................................................................................... 575 20.6.68 Sandalwood Oil.................................................................................................... 575 20.6.69 Spearmint Oils...................................................................................................... 575 20.6.70 Spike Lavender Oil............................................................................................... 575 20.6.71 Star Anise Oil....................................................................................................... 576 20.6.72 Tarragon Oil.......................................................................................................... 576 20.6.73 Tea Tree Oil.......................................................................................................... 576 20.6.74 Thyme Oil............................................................................................................. 576 20.6.75 Turpentine Oil....................................................................................................... 576 20.6.76 Vetiver Oil............................................................................................................. 576 20.6.77 Ylang-Ylang Oils.................................................................................................. 577 References....................................................................................................................................... 577

20.1 INTRODUCTION 20.1.1 General Remarks Requirements of governmental bodies in the use of natural products increased tremendously in the last years. Not only the directions by the flavor regulation but also the demand of the consumer organizations presses the industry and producers to supply solely natural products, often desired in “bio” quality. The green responsibility for natural products brought synthetic aroma chemicals in a poor light. Regarding the mass market for cosmetic products, genuineness in fragrances is very limited because of the sources of raw material. On the other side, prices for raw materials increase since years for natural flavor and fragrance ingredients. The acceptance of fragrance compounds containing essential oils and natural aroma chemicals is a way to fulfill the consumer demand for safe, effective, and too affordable finished products. But even today with that claims and in spite of fabulous analytical equipment and highly sophisticated methods, many adulterated essential oils are found on the market. The question arises, what are the motivations for such a serious condition? This and above all, the manner of adulteration and the matching analysis methods shall be treated within this chapter. Essential oils are constituents of around 30,000 species of plants around the world. Only a few of them are used in today’s flavor, cosmetic, animal feeding, and pharmaceutical industry as well as in aromatherapy. Observing the product range of producers and dealers, about 250–300 essential oils are offered in varying quantities. Within those oils, 150 can be characterized as important oils in quantity and price.

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Consumption of these essential oils worldwide is tremendous. The data published by Lawrence (2009) show a total quantity of more than 120,000 metric tons. A monetary value can barely be calculated, as prices rise and fall over in a year; exchange rates in foreign currency and disposals within the dealer’s community falsify the result. Gambling with natural raw materials and the finished essential oil takes place with all the important ones. Comparison of production figures and those of export and import statistic will vary sometimes obviously; discrepancy appears in higher amounts. Reunion Island’s National Institute of Statistics and Economic Studies (INSEE, 2008–2009) reports for the year 2002 the low quantity of 0,4 to kg of “vetiver oil Bourbon” exported; the sold quantity in the market was more than ten times higher. From 2003 until today, no vetiver oil is produced on Reunion Island. The essential oils traded were proven pure and natural but were coming from other production areas.* However, “Bourbon” quality generated a much higher price. It must be assumed that the value of essential oils worldwide will be far above $10 billion and this dimension invites to make some extra money out of it. Screening essential oil qualities from the market with high-end analytical equipment leads to detection of many adulterated essential oils. Alarming is the fact that these adulterations appear not only within the consumer market but also in industry and trade. The questions arise again, what is the explanation for such a behavior and what are the reasons? Adulterations are subject of many publications with high impact; most of them came out in the beginning of the nineties, when new analytical methods were developed and applied. However, adulteration of essential oils began a long time ago. This chapter deals with adulteration starting in history up to the present. High criminal energy must be responsible for such sacrileges. Financial advantages, market shares, monopole status, and sometimes simply a sportive action are only some reasons.

20.2  DEFINITION AND HISTORY Observing the significant number of publications about essential oils using the Internet, the results show lack of knowledge, finding “extracted” essential oils, distilled ones by “supercritical fluids,” distilled Jasmine oil, therapeutic essential oil, and so on. It is remarkable that in scientific publications of serious publishers, there is the same confusion and ignorance about the term “essential oil.” Even in regulatory and institutional papers, this term is used falsely and unprofessionally. Essential oils are clearly defined by the International Standard Organization (ISO) in Draft International Standard (DIS) ISO, 9235, 2013 (former 9235.2)—Aromatic natural raw materials—Vocabulary. This DIS war, revised 2013, tightened the strict definition for an essential oil: Essential oil: Product obtained from a natural vegetable raw material of plant origin, by steam distillation, by mechanical process from the epicarp of Citrus fruits, or by dry distillation, after separation of the aqueous phase—if any—by physical process.

Further it is mentioned that steam distillation can be carried out with or without added water in a still. No change has been made with the note that “the essential oil can undergo physical treatments (e.g., filtrations, decantation, centrifugation) which do not result in any significant change in its composition.” This is a tightening of the old definition. Natural raw material also was updated in the definition: “Natural raw material of vegetal, animal or microbiologic origin, as such, obtained by physical, enzymatic or microbiological processes, or obtained by traditional preparation processes (e.g., extraction, distillation, heating, torrefaction, fermentation).” These definitions show clearly the importance of standards to avoid any adulteration during all stages of collection, production, and trade. Terms like “pure or natural essential oils” now are misleading as an essential oil has to fulfill this demand. Other regulatory elements are used for standardization like the pharmacopoeias around the world. Furthermore medicinal authorities and animal feeding product industry settled own regulatory standards. In the field of aromatherapy, no standards are settled although just in that field, the existence of ISO standards or pharmacopoeias as base should be implemented. The * Based on the authors’ private information and experiences.

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use of natural cosmetics is reported to result in a higher demand for oils but the quantity available on the market was and is insufficient to satisfy this developing market. Today many aromatic chemicals from “natural” source, produced by enzymatic reaction from sometimes not natural starters by microorganisms, help the producers to fill up that shortage. But are these chemicals really “natural”? Adulteration is defined as “making impure or inferior by adding foreign substances to something” or “is a legal term meaning that a product fails to meet international, federal or state standards” (The Free Dictionary n.d. [a]). In this context all kinds of adulterations are enclosed like poisonous, economical, and unethical ones. In the United States, FDA regulations for food and cosmetics are the basis of law; in Europe consumer protection and food regulations as well as the cosmetic regulation have to be applied. Adulterations must not always be deliberated with criminal or unethical background. Environmental pollution like herbicides or pesticides cannot be prevented; these are unavoidable “adulterations.” Negligently adulteration, basing on the lack of knowledge, still happens today. By selection of the botanical raw material, very often mistakes are done. Another instance is the missing of suitable production facilities and again, insufficient professionalism of the staff working there. The vocabulary for adulteration of essential oils is fanciful: extending, stretching, cutting, bouquetting, and rounding up to even “sophistication.” The history of adulteration reaches back to the turn of nineteenth to twentieth century. It is ongoing hand in hand with the establishment of chemistry and the development of synthetic aromatic chemicals. Already in 1834 it was possible to isolate cinnamaldehyde from cinnamon bark oil and as a result in 1856, it was synthesized. It is worth to know that Otto Wallach, a basic scientist for terpene chemistry at the University of Göttingen, already started in 1884 with the clarification of terpene structures. In 1885 he defined the contribution for the structure of pinenes, limonene, 1,8-cineole, dipentene, terpineol, pulegol, caryophyllene, and cadinene. The development of synthetic aroma chemicals found in essential oils and extracts started with the synthesis of vanillin in 1874 by W. Haarmann and F. Tiemann. Literature gives only a few information about adulteration and the authors” research is going back to publications from 1919. The company Schimmel & Co. in Leipzig, Germany, was founded in 1829 under the name Spahn & Büttner. In 1879 Schimmel was the first company to establish an industrial laboratory for the production of essential oils. Famous wellknown scientists like Wallach (Isolation of Muscon) and Eduard Gildemeister (the famous books of Gildemeister and Hofmann) were there at work. In 1909 Schimmel started publishing the famous “Schimmel-Berichte” (Schimmel reports). These reports about progress in research on essential oils, containing composition of essential oils, isolation of chemicals, cultivation of fragrance plants, and adulteration of essential oils, were discussed. In the report from Schimmel-Berichte (April/October, 1919), the following adulterations were reported: “Bergamot oil with phthalates and terpenes, bitter almond oil with raw benzaldehyde, cassia oil with phthalates, cinnamon oil Ceylon with camphor oil, lemon oil with water, lavender oil with phthalates and terpineol, peppermint oil with glycerol esters, menthol, phthalates and spirit, sandalwood oil with benzyl alcohol, star anise oil with fatty oils and rose oil with palmarosa oil and spermaceti.” That points out clearly that already at that time adulteration was omnipresent. Schimmel was the first company to create a synthetic neroli oil on the market. The cause of that was the establishment of a syndicate comprising of the South French growers and, what is more, frost and storm damages during orange flower blooming. In 1920, again in one of the “Schimmel-chronicles” (SchimmelBericht, April/May, 1920), the adulteration of lemon oil is reported: “Skilled mingled, with terpenes, sesquiterpenes and citral, the determination with specific gravity and optical rotation are not sufficient to detect adulteration in lemon oil.” As a solution for that problem, preparation of as much as possible fractions, tested for the solution in ethanol of 89% Vol., was proposed. Rochussen (1920) reported that several oils were adulterated, as other than the permitted parts of the plant were added. The utilization of turpentine oil was the most used component for adulteration of those essential oils, bearing large amounts of terpenes. Mineral oils (paraffin, petrolatum, benzene, and Vaseline [petrolatum]) were reported to adjust the change of physical data forced by other

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adulteration chemicals. For the same purpose cedarwood, gurjun balsam, and copaiba balsam oils were applied because of their weak odor. At that time, physical constants like optical rotation, density, refraction, boiling point, freezing point, residue on evaporation, ester value, acid value, carbonyl value, and water content could give satisfying data about the examined oils. In addition the so-called wet chemistry could give some more answers. Detection reaction, titration, photometry, and gravimetric analysis were used to achieve a more precise result about the genuineness of an essential oil (detailed information will follow). Bovill (2000) published in Perfumer & Flavorist an original text of E.W. Bovill, RC Treatt, London, about the essential oil market in 1934. One key sentence was as follows: “Essential oils, largely because they are liquids, are easy to adulterate in ways which are difficult to detect and – it must be admitted they very frequently adulterated.” At that time it was obvious that adulteration was a fight likely as between a thief and a policeman. Previously, adulteration was done by producers and brokers and customers had to believe the words about purity and nativeness. The only key of trustfulness could be the relationship between producers/traders and clients by permanent control in the harvest and production time locally. Ohloff (1990) confirmed that through hydrodistillation in industrial scale, progress was made in synthesizing compounds with effective methods. Camphor was one of the important chemical components (not only for the flavor and fragrance use but as starting substance for important “plastic” products), together with borneol. Starting with the synthesis of important aromatic chemicals at accessible prices like linalool, geraniol, phenylethanol, and the esters, the adulteration of essential oils started progressively. In a flash, essential oil components were available in sufficient quantities and at prices much cheaper than the essential oil itself. Another event was the problem of the two world wars. Especially in World War II, the acquisition of raw materials of natural or synthetic kind was no more possible at all. The military machine used nearly all raw material resources, chemists were forced to establish substitutes for essential oils, and that resulted in several new synthetic aroma chemicals, which now could be used to stretch or to substitute an essential oil. Thus, the ingenuity of corruption by essential oil producers increased rapidly, and even with general application of chromatographic analysis in industrial laboratories beginning in the 1980s, falsifications could hardly be detected. The authors noticed in the course of a visit to Provence area and producers of lavender and lavandin oil that starting with the analysis by gas chromatography using packed columns in 1979 led to the result that compositions of samples of essential oils were not adequate to literature of that time. By informing the producers about the establishment of chromatographic equipment, the quality was immediately rising. But not all was satisfying. So a “touristic” tour to the facilities of the producers was decided during harvest and production time, end of August 1981. Starting in Puimoisson and with the help of a little blue empty perfume bottle (“flacon montre,” used at that time for lavender perfumes), the staff was asked for a little bit of lavender oil. By giving them some Francs, he had the opportunity to watch around the distillation facility. Together with the oil sample, information about special observations was noticed. The same happened in facilities of Riez, Montguers, Simiane, Richerenches, Rosans, Remuzat, Banon, and Apt, and from all production units oil samples could be collected and these were analyzed in the laboratory by gas chromatography. As a result, only one from 10 samples was in accordance with the requirements. The others were adulterated mainly by synthetic linalool and linalyl acetate (recognized by dihydro- and dehydrolinalool as well as dihydro- and dehydrolinalyl acetate) but also with too high contents of limonene, borneol, and camphor (by adding lavandin oil), far too little amounts of lavandulol and lavandulyl acetate. Adulteration by adding those chemicals after distillation could not have happened, as the samples he received were filled directly from the Florentine flask. Finally he was in luck to observe that before closing the cover of the plant, linalyl acetate was spread by means of a portable pump with tank directly on the lavender. The cover was closed and the essential oil now running from the Florentine flask could be seriously called as natural. Interesting was the observation of the distillation facilities. In four cases drums with linalool and linalyl acetate from BASF (Badische Anilin- & Soda-Fabrik) could be detected. Concerning linalool and linalyl acetate, it is worth to mention that 1 year later other drums were occurring from

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a producer in Bulgaria produced by another synthesis pathway without de- and dihydro components.* At that time it was a customary practice, as reported by Touche et al. (1981), to use relationships of cis-β-ocimene to trans-β-ocimene (R1) and trans-β-ocimene to octanone-3 (R2) as well as linalool + linalyl acetate to lavandulol + lavandulyl acetate (R3). Statistically collected values over many years guaranteed the correct relations. Another observation was the use of lavender bushels together with some lavandin bushels. This could be transparently seen; they were larger in size and the leaves were characteristic of Lavandin Grosso. At least, there were other chemicals in a rack, such as camphor and geraniol, borneol, terpinen-4-ol, and β-caryophyllene. What should these chemicals be for? He was told that these are for special clients that are so-called comunelles, meaning a mixture of several charges from different farmers but from the same plant was established. The last production unit had constrained not only a laboratory but also a production unit with many chemicals. From that time on the author did no more trust any producer or trade companies.* Today such a way of working is no more possible. Ordering an essential oil means, with all possible parameters, the properties are scanned and tested and standards must be fulfilled. In spite of all loyalty of a producer/customer relationship, only analytical, physical, and sensory examination is inevitable.

20.3 ADULTERATION 20.3.1 Unintended Adulteration The term “adulteration” must be understood as a negative concept. It is in nearly all cases a criminal and unethical behavior. But there will be the possibility that it is an act of unwanted adulteration without any intent and has nothing to do with dishonesty. Reasons can be the lack of knowledge, bad equipment, wrong treatment of the plant material, not allowed methods for distillation and no good manufacturing practice. Of course, all these things should be avoided, but they happen. Selection of correct plant material is the basic for the production of essential oils. It is clearly confirmed in every ISO standard, from which botanical source the oil has to be produced. When a cultivation of the biomass is possible, it must be guaranteed that the correct species is used. By using wild collected plants, the possible risk of collecting similar species or chemotypes cannot be excluded. As an example, essential oil of Thymus serpyllum L. from Turkey origin was sent for analysis. As a result, the oil was absolutely different from experience and literature. Thymol and carvacrol were too low, and linalool was higher as well as geraniol and geranyl acetate: What could have happened? People collecting the wild thyme took everything that look like a thyme plant without looking for the chemotypes (which is hardly possible by visual selection). Higher quantities of linalool and geraniol chemotypes were present. The oil was natural at all, but it is not to be used in flavor application or aromatherapy. A further example is the petitgrain oil: The petitgrain from Paraguay is not a bitter orange plant as such. Citrus aurantium L. is the true bitter orange plant used for production of the essential oil from peel, but also from leaves to obtain petitgrain oil “bigarade.” The oil from Paraguayan plants is derived from Citrus sinensis L. Pers. × Citrus aurantium L. ssp. amara var. pumila. This is a hybrid from a sweet and bitter orange. The composition of that oil is different from the first because of its higher content of limonene, trans-β-ocimene, and mainly linalyl acetate (ISO/DIS, 8901 and ISO/DIS, 3064 2015). The Paraguayan oil was sold as bitter orange oil; however, curiously, the oil of petitgrain from Paraguay was much cheaper. A further mistake can be the inappropriate treatment of the biomass. Fresh distilled plant material results in a different final product than the oil from dried or fermented material. The question arises, is it a desired procedure or a mistake or an omission? Good manufacturing practice (GMP) is a standard application procedure in technologically developed countries, but what about emerging nations or those without any technical progress? Can such essential oils be traded or used? * Based on the authors’ private information and experiences.

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Hydrodistillation is nowadays a highly developed, automated and computerized method to produce essential oils. Applying computer programs for detection- and production parameters observing all conditions. Starting with the plant material (moisture content, degree of maturity, oil content) followed by temperature, steam quantity, steam pressure, cooling temperature, and subsequent treatment, all is handled electronically. In spite of such a technology, mistakes can happen as in the case of distilling Melissa officinalis L. from a biological culture. When analyzing that oil by gas chromatography– mass spectrometry (GC-MS), a content of more than 5% of thymol was detected and confirmed three times. After discussing the problem with the producer, he guaranteed that only balm, fresh cut, was used. Contamination in the analyzed sample could be excluded as the second sample showed identical results. After checking all distillation equipment, the reason was found. Before producing Melissa oil, thyme was distilled and he discovered a small chamber within his own constructed cooling unit. There remained about 50–60 mL of oil and it was mixed up with balm oil in the course of hydrodistillation. This is a great financial loss for the producer taking account of the price.* Adulteration can also be done by, for example, adding basic solution to the vessel while distilling. Sometimes it is used to avoid artifacts formation from the plant acids. By raw materials with high-grade acids, for example, barks like massoia, this is sometimes applied. The neutralization of the pH value inhibits the ester formation or degradation reactions in the course of the processing. The use of cohobation is another fact for discussion. This method uses the recovery of volatiles from distillation of water, remaining solved and without separation by Florentine flask. As long as this water is returned to the distillation vessel, there is no problem at all. For some oils like thyme oil, it is of importance to use cohobation, because of the high content of phenols, which dissolve not insignificantly in distilled water. Venskutonis (2002) reports that “cohobation minimizes the loss of oxygenated compounds but increases the risk of hydrolysis and degradation.” Similar is the situation with rose oil. Here the cohobation is an important step to receive a genuine reflection of rose odor. Treating the distillation water with salt (NaCl) or pH adjusting to remove plant acids seems to be discussed, as it will still remain an essential oil. By extracting the distillation water with any solvents, which then will be removed by distillation in vacuum, will lead to an extract. This extract, added to the distilled essential oil will result in the loss of the term “essential oil” by definition.! Codistillation is reported for several “essential oils.” Mainly sandalwood oils are mentioned to be extracted and redistilled. Valder (2003) reported that the production method of sandalwood oil is sometimes the hexane extraction followed by codistillation with propylene glycol and finally a rectification is done. Baldovini (2010) stated that this method is not used anymore as it does not longer comply with ISO rules. Rajeswara (2007) tested the effect of codistillation using crop weed with several plant materials like citronella, lemongrass, palmarosa, Corymbia citriodora, and basil. He observed the increase of yields and also changes in composition. The author was able to observe a similar effect, analyzing bio lavender oil. The oil differed in composition from standards with higher ester content, higher oxidation compounds, and less 3-octanone. The cultivation was bio, without removing any weed. The percentage of weed was esteemed to be more than 30%. The oil did not match ISO standard.* Contaminated containers for production and transport are another possibility of adulteration. Reused drums, not cleaned, are often the cause for worst damage of the essential oil. Plastic containers contain phthalates as plasticizers and the hydrocarbons solve these. Phthalates contaminate even in smallest traces the oil for the use in natural finished products.

20.3.2 Intentional Adulteration If anybody adulterates willfully and knowingly, it is a criminal act and never a wangle or minor offense. The target to cheat a customer by supplying an essential oil being not conform to any standard is a felony. There is no excuse for such an unethical behavior. Again, until now, adulteration can be found on essential oil market. Terms like cutting, stretching, blending, bouquetting, or sophistication try to moderate this act. The causes for adulteration are manifold and often related with economical and environmental reasons. * Based on the authors’ private information and experiences.

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20.3.3  Prices The cutting of essential oil happens in most cases because of the price. They are subject to falling and increasing prices, depending on crops and the demand but also speculation. The last is often the reason why essential oils come to market with adulteration in every dimension. All economical ambition is the increase of profit, the more the better to fulfill shareholders demand. Essential oil market makes no distinction as it is the same as the law of supply and demand. Speculation was always a reason to ameliorate the process. By hoarding raw materials, the profit can go up in high levels. Lavender oil is an example for that. Most of the farmers in France keep parts of the production in the cellars to wait as far as the new crop starts, until the price seems to be the best. Another example from the authors’ experience was the competitive battle with cornmint oil in the 1980s. China and Brazil were the two countries with the highest production at that time. Both increased the cultivation area with the result that in the next season prices fall down tremendously because of oversupply. So Brazil decided to decrease the cultivations and in the next year, prices were going up like a rocket, as China too had decreased production, and demand of the market could not be fulfilled. At that time, most menthol was produced from cornmint and consumption was rising. Stabilization was achieved by supplement of sufficient synthetic menthol and India coming to the market as third production continent. Nowadays speculation has another dimension. Likely nearly all foodstuffs and plant crops are merchandized at the stocks and of course this has influence on the oil market. Increased prices in essential oil market cannot be transferred simply onto the industry of finished products. Mass market and global players will not accept any price advance, so the only solution is cutting or stretching. This situation leads to a competitive market, where adulterated essential oils with certificate of naturalness are sold.

20.3.4 Availability The market changes from monopole situation for suppliers to buyers market within a few months. Shortage of biomass leads to a rapid increase of prices. The causes will be dryness in the production area, flood during harvest time, and ice and frost like in the case of citrus fruits in California or Florida. Tempests are crucial for destruction of stretches of land as well as deletion caused by insects or microbes. Annual rainfalls in tropic regions and lack of sunshine in Mediterranean region are reasons for partial failure. Unavailability or decreasing availability is a high risk for global players in the fine perfumery industry. The costs of launching a perfume worldwide exceed the amount of $100 million easily and one can imagine that the lack of an important natural raw material and therefore, unavailability of that perfume can lead to ruinous losses. In consequence of that, natural essential oils and also extracts are no more or only in small quantities added to the perfume compounds. It is easy to replace a natural product by a compound. Lemon oil, bergamot oil, rose oil, lavender oil, and many others can be substituted by basic synthetic compositions. Only some special oils like vetiver, sandalwood, patchouli, and orris are never perfect to compensate.

20.3.5 Demand of Clients The various fields of applications sometimes cause the client industries to require special essential oils. One of the reasons is of course the price. The end-consumer industry settles the prices for the market. Competitive prices have to be adjusted and the problem for the use of essential oils is that prices can change from crop to crop or even from week to week.

20.3.6 Regulations A huge number of regulations have been established for the protection of consumers worldwide. Cosmetic regulation, flavor regulation, and REACH (Registration, Evaluation, Authorisation and

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Restriction of Chemicals; the European chemical law) extend to the Globally Harmonized System of Classification and Labeling of Chemicals (GHS). All these provisions cover essential oils, but not per se. They are dealt with their contents. Most oils contain one or more chemicals, classified as toxic, allergic, sensitizing, or carcinogenic. Consequently these oils are no more or only in very limited quantities applicable. As an example, the methyl eugenol must be mentioned. Methyl eugenol is a native ingredient of distilled rose oil. Between 1.5% and 3.5% is found in this essential oil. Considering the European Cosmetic Regulation, the maximum dose in finished leave-on products might not exceed 0.001%, which results in the fact that rose oil is no more applicable in cosmetic products. The same pertains to pimento oil from leaves and berries, oil of myrtle, magnolia flower oil, laurel leaf oil, and bay oil. The demand of the industry is rose oil with nearly no or only traces of methyl eugenol. Applying various methods like fractioning and remixing and chemical reactions it was tempted to fulfill the regulation. The result was not as desired, as the odor was not a rose anymore and the finished product could not be named as “essential oil” or “natural.” To avoid an increase of cases of allergic contact dermatitis in the public, many essential oils nowadays have to labeled as “sensitizing—can cause allergic reaction”! Cosmetic producers as well as discounters with private labels avoid naming and labeling and thus, force the replacement of natural products. Essential oil producers offer natural oils with reduced values of sensitizers and this is not possible. Maybe the essential oils are treated in some way, but then are no more “natural,” according to ISO standard.

20.3.7 Aging It is impossible to characterize aging as adulteration without intent. Changes over storing, either correct or false, always result in quantitative modification of the composition. The formation of oxidative compounds like epoxides, peroxides, and hyperoxides is a dangerous effect. Terpenoids show the properties to be volatile but also to be thermolabile. The mentioned oxidative substances show the negative reaction activities coming in contact with the skin. Sensitizing and allergic reactions are consecutive. Countless components of essential oils are subject to oxidation reaction with atmospheric oxygen. Citrus oils implicate such chemicals. Dugo and Mondello (2011) confirm that the reaction of degradation of sabinene, limonene, γ-terpinene, neral, and geranial in lemon oil results in the enlargement of p-cymene, cis-8,9-limonene oxide, (E)-dihydro carvone, (Z)-carveol, 2,3-epoxy geraniol, and carvone. Particularly the increase of p-cymene in citrus essential oils is a distinct detection for heterodyning and aging. Sawamura et  al. (2004) demonstrated in a poster compositional changes in commercial mandarin essential oil and detected in a 12-month test the decrease of limonene (60%) and myrcene (nearly 100%) and the increase of (Z)-carveol (from 0.0% to 2.8%) and carvones (from 0.2% to 3.0%). Brophy et al. (1989) could observe degradation in terpinen-4-ol, γ-terpinene, and β-pinene but rise of p-cymene and α-terpineol. Examination of peroxide value seems not to be the suitable tool to recognize aging of essential oils, as some of the reactions are reversible. Eucalyptus oil, rosemary oil, and of course citrus oils have demonstrated that warming of these oils in a closed system at 40°C reduces obviously the peroxide value. The most suitable method of recognizing aging is analysis by GC-MS. It has to be the responsibility of producers and traders to take care for the safe quality of their essential oils.

20.3.8 Cupidity This of course is the main reason for adulteration. Cupidity is defined as “strong desire, especially for possessions or money; greed” (The Free Dictionary n.d. [b]). Although this is the main reason, here is not the place to discuss anything about cupidity as it is human fault.

20.3.9 Simple Sports? The knowledge about absence of qualified analytical equipment on the side of the client seduces producers as well as the trade to cut essential oils. With the development of new chromatographic

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methods and an increase in sensitivity as well as selectivity, the inventiveness of the producers shows high wealth of ideas. Again, the race between a thief and a police could have been stopped by the costs of investments in analytical systems, but there seems no end by observing bad qualities on the market.

20.4  POSSIBLE ADULTERATIONS FOR ESSENTIAL OILS 20.4.1 Water Adding of water is very simple and cheap. This is not possible for all essential oils but is likely for those possessing compounds with high affinity of binding to water. Conifer varieties and citrus oils are examples. Siberian pine (Abies sibirica Ledeb.) was supplied in August at 25°C–30°C. Visually no water could be detected. In January the oil came from stock (−5°C) and the quantity of water could be esteemed to 8%. Is that adulteration or natural behavior? Rajeswara Rao et al. (2002) mentioned water contents up to 20%, but that level seems too high by following GMP. Responsible for this effect are monoterpenes, and like conifer oils, citrus oils contain higher quantities of these compounds. Citrus expression techniques use a lot of water to spray away the oil/wax emulsion. Centrifugation will separate water and waxes from essential oil. Unfortunately also aldehydes and alcohols contained in the water/wax phase are removed. Cotroneo et al. (1987) observed this effect by comparing oils from manual sponge method without water and the industrial technical methods. An easy method to detect higher value of water in citrus oil is the visible cloudiness and the deposit of waxes when cooled down to 10°C. A validated method to detect the water content in essential oils is the Karl Fischer titration (ISO, 11021, 1999).

20.4.2  Ethanol Mostafa (1990) reports that ethanol is the main alcohol used in moderate quantities to dilute essential oils. Ethanol is a component of rose oil. The process of water/steam distillation forms alcohol in certain quantities. In the chromatographic profile in ISO, 9842, 2003, essential oil of rose, the value of ethanol is specified from 0% to 7%, depending on the origin (maxima: Bulgaria 2%, Turkey 7%, and Morocco 3%). The question arises whether the alcohol is coming directly from the process or if the distilled water is washed and extracted by alcohol and this extract is added to the essential oil. If it is the case it is no more an essential oil. Rose oil is a very expensive product and 1% or 2% of added ethanol increases the profit.

20.4.3 Fatty Oils or Mineral Oils As mentioned in the introduction, adulteration with fatty oils in history is well known. Rochussen (1920) confirms that and mentions paraffin, petrolatum, and castor oil. The easiest way to detect was the blotting paper test. Essential oils containing fatty oils leave behind a lasting grease spot. By testing the solubility in ethanol of 90%, a blur occurs, showing the presence of fatty oils. Only castor oil is soluble in ethanol but can be detected by evaluation of nonvolatile residue content method. Since more than three decades, such an adulteration became no more apparent.

20.4.4 High Boiling Glycols For a long time such adulterations remained undiscovered. High boiling materials like polyethylene glycols could not be detected with normal GC-MS method, even at 280°C column temperature. It takes many hours before these chemicals leave the column and give a long, small hill in chromatogram. Not only expensive essential oils like sandalwood, vetiver, and orris were cut with those chemicals but also extracts like rose absolute. By happenstance the author found the stretching

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of this rose absolute. Typically this product is of deep red color and its use in white cosmetic cream was impossible. A high vacuum distillation was performed to remove the color, which really seemed to be not a problem. The end temperature was 200°C in high vacuum but still about 20% remained in the retort. Never a component of rose absolute with that boiling point was detected or mentioned in the literature before.* At least in, 1974, Peyron et al. reported about the use of hexylene glycol up to 40% in vetiver and cananga oils (Peyron et al., 1974). By treatment with water, separation by rectification, and liquid gas chromatography, it could be detected. In 1991, John et al. reported the thin-layer chromatography method for detection. This method is simple and effective and can be used for routine analysis.

20.4.5 Oils from Other Parts of the Same Species or Other Species with Similar Essential Oil Composition This is a very simple method to stretch. Clove leaves are easier to harvest and bring similar oil as the stem or bud oil. Clove bud oil is nearly three times higher in price than clove leaf oil. Therewith the concocting of that oil makes sense. Pimento berry oil and leaf oil are closely related in composition. The price of the berry oil is double the leaf oil; consequently, adulteration is interesting. By GC-MS method and regarding minor components, this immixture will be detected. Eucalyptus oil from China is derived from Cinnamomum longepaniculatum (Gamble) N. Chao ex H.W. Li 1,8-cineole type. Either the whole oil sold as eucalyptus oil or Eucalyptus sp. added is clearly not allowed and must be seen as adulteration.

20.4.6 Related Botanical Species Something of the kind can only happen in the origin, where biomass is growing and distillation is proceeded. Some plants have relatives, which then might have comparable oil compositions. Ylang oil is produced from the flowers of Cananga odorata (Lam.) Hook. F. Thomson forma genuina. The other form of Cananga odorata is cananga oil with Cananga odorata (Lam.) Hook. F. Thomson forma macrophylla as plant source. On the market, cananga oil is cheaper than ylang oil. Instead of mixing the distilled oils of both, the flowers are distilled together. Furthermore flowers of climbing ylang-ylang Artabotrys uncinatus (Lam.) Merill are sometimes added before distillation. When producing patchouli oil, the vessel is filled with leaves, and to avoid that they stick together during processing, branches of the gurjun tree (Dipterocarpus alatus Roxb. Ex G. Don and Dipterocarpus turbinatus C.F. Gaertn.) are added to avoid that. As a result, the oil of patchouli is contaminated with α-Gurjunene (up to 3%),which is not part of a pure patchouli oil (private information to the author).

20.4.7 Fractions of Essential Oils This is one of the most applied adulteration methods. Fractions arise in all cases, when essential oils or extracts are concentrated, washed out, and rectified or are residues of removal from centrifugation and distillation as well as from recovery of water streams of expressed citrus peels. Heads and tails from rectification are added to similar essential oils, like peppermint terpenes to mint oil. Essential oils that are high in terpene content, cheap, and available on the market are citrus, eucalyptus, Litsea cubeba, cornmint and mint, petitgrain, spearmint, vetiver, lavender and lavandin, cedarwood, citronella, clove, and Corymbia citriodora terpenes. Especially citrus terpenes from various species are appropriate for stretching. Dugo (1993) recognized the contamination of bitter orange oil by the use of sweet orange and lemon oils and its terpenes. By applying liquid chromatography (LC), highresolution gas chromatography (HRGC), and GC-MS methods, they confirmed the addition of less than 3% of these components. Limonene is the main component of many citrus oils. On the other side, limonene is a component of nearly all essential oils in variable quantities. Limonene is a chiral * Based on the authors’ private information.

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terpene; chirality describes a basic property of nature. Some molecules show the property to have three-dimensional structures that possess a chiral center. These molecules show a basic property of nature, to look like image and mirror image, but both cannot be aligned. It is a matter of common knowledge that chemical compounds in essential oils tend to be chiral and the preference of one form can be observed. In contrary synthetic molecules are racemic, meaning both forms of the molecule are balanced. This is a real fingerprint for detection of adulterations with terpenes owing different chiral properties. On normal GC columns a separation of chiral molecules is impossible. With the use of columns coated with modified cyclodextrins, those mirror-imaged molecules will be separated and appear as two different signals. In 1995 bergamot essential oils, sold in 10 mL bottles to end users, were subject for investigation of the German journal ÖKO Test (ÖKO Test, 1995). This magazine, dealing with naturality and safety of consumer products, charged the University of Frankfurt, Prof. Mosandl, to check the authenticity of the oils by chiral separation of linalool and linalyl acetate. Thirty-seven of fifty-three samples could be confirmed not being natural and “pure.” In a private letter of Prof. König (University of Hamburg) to the author, he confirmed: “Linalool and linalyl acetate are present in bergamot oil only in pure R-enantiomers. That for bergamot oil— unnatural—(S)-linalyl acetate cannot be found even in traces.” The invention of this technique by König, Mosandl, Casabianca, and Dugo, improved by Mondello, Bicchi, and Rubiolo, was a huge step to diminish the cases of adulterations in essential oils. Limonene occurs in d-, l-, and dlform in nature. Limonene in sweet orange oil is always in d-form; most of the pine oils show pure l-forms. For the detection of stretching, other terpenes and terpene alcohols are used like α- and β-thujene, α- and β-pinene, α- and β-camphene, α- and β-sabinene, δ3-carene, linalool, β-borneol, myrtenol, linalyl acetate, menthol, cis- and trans-terpineol, 1-phenylethanol, nerolidol, terpinen4-ol, and many more. Whenever the chiral distribution differs from normal proportion, a manual intervention was made. However, the last sentence has to be relativized carefully. Chiral composition of camphor and borneol in natural essential oil of rosemary is very variable, depending on the origin! Another method of stretching is the use of distilled residues of expressed citrus peels. After citrus production process, some quantities of essential oil components still can be found in the “waste products.” In addition, Reeve et al. (2002) report of the use of distilled mandarin oil, produced from recovered water streams from spiking and pressing. Because of the contact with acidic juice, the composition of such an oil shows significant difference in composition from peel oil. This oil can be sold as “distilled” mandarin oil, but never be used to be blended with pressed oils. Such a handling must be seen as adulteration and is not covered by ISO definitions.

20.4.8 Natural Isolates Single chemicals can be derived from essential oils by methods of fractioning and rectification. As this procedure is from a natural source, of course, this chemical must be characterized as natural. Numerous components are offered on the market like citral from Litsea cubeba oil, geraniol from palmarosa oil, linalool from ho oil, coriander oil or lavandin, pinenes from different Pinus species, citronellal from Corymbia citriodora, cedrol from cedarwood, or even the santalols from different sandalwood species. All these are added to “finish” essential oils. As already mentioned before, synthetic chemicals can no more be applied as enantiomeric separation is a state of the art today and will convict the matter of fact of adulteration. By using synthesized, correct chiral compounds, the detection is hardly to be recognized but with NMR method, but this is an expensive analysis.

20.4.9 Chemically Derived Synthetic Compounds, Which Are Proved to Appear in Nature Compounds formerly named “nature identical” are to be defined by law since some years as synthetic products. These molecules have been found in nature by analytical proof and are published in an authorized scientific journal. The term “nature identical” is no more valid and allowed in Europe

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in relation to flavor and fragrance substances. Such molecules are identical with those appearing in nature but are produced by a synthetic process. These processes contain undesired by-products. The use of such synthetic compounds is easy to detect, as by-products from manufacturing can easily be detected by GC-MS systems. On the other hand, chiral separation will help to confirm adulteration.

20.4.10 Steam Distilled Residues from Expression After an expression process of citrus oils either in exhausted peels or in centrifugation residues, carryovers of the volatiles will remain. These can be removed by distillation with high-strung steam, and oils acquired are colorless and still have the smell of the starting material. Components are similar those of the expressed oils but contain for reason of the production method some oxidized chemicals. Nevertheless these oils are used to adulterate the expressed oils, and with that process the naturality per definition by ISO standard is lost. The main ingredient is limonene, followed by myrcene and γ-terpinene. Traces of aldehydes still can be found. Such adding can be detected by observing higher values of oxidation compounds.

20.4.11  Enzymatically Produced Chemicals (Natural by Law) Enzyme is defined as “Any of numerous proteins or conjugated proteins produced by living organisms and functioning as biochemical catalysts” (The Free Dictionary n.d. [c]). Enzymes of microorganisms in a medium with added nutrients produced such molecules. Isolation has to be done by any physical process to isolate the desired molecule designed by the biosystem. Although this manufacturing process is authorized within the applicable legal requirements for the use of such enzymes in the EU, it is somewhat uncertain. These processes will as a rule not happen in nature. Within a plant, production of hydrocarbons takes place without microorganisms by conversion of a starting material and by enzymatic reaction within a cell system. The question arises if that molecule from microorganisms can really be named as natural, particularly if it is generated from “any” starters. In the authors” mind, adding such a compound must also be named as adulteration within the definition of essential oil, which does not allow such a process and any additives. At least it must be mentioned that the mixture of essential oils from the same species but from different geographical sources cannot be called an adulteration. They fulfill the requirements of standards as long as the specific provenance is not laid down in the specification. Lavender oil from France can be mixed up with Chinese, Bulgarian, and Russian origin and is still lavender oil (Lavandula angustifolia Mill.), but may not be sold as “French lavender.” Furthermore if an essential oil from the same species but from another geographical area is recognizably different in composition, it must be mentioned in standards and certificates. One example is geranium oil from North Africa compared to other origins. 10-epi-γ-Eudesmol is only present in North African oil. The cause for this is adaptation to different conditions in a longer period.

20.5  METHODS TO DETECT ADULTERATIONS 20.5.1  Organoleptic Methods 20.5.1.1  Appearance and Color Appearance is the visual aspect of a thing or person. In this case it is the appearance of the “essential oil,” starting with color, going on with mobility, and finally with the odor itself. The color of essential oils is dependent on the starting material. Citrus oils are colored weak yellowish with lemon oil, light green to darker green with bergamot oil, and orange to brownish red with orange and mandarin oils. The color is dependent on the degree of ripeness of the fruits but they alter with storage and influence of light and warmth. Hereby the age of such an essential oil can be detected. Colors too will appear in hydrodistilled essential oils. The normal case for these oils is colorlessness accompanied by mobile

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fluidness. Weak yellow color can appear in oils like cardamom, rosewood, tarragon, or turpentine. Oils of lavender and lavandin are pale yellow and mobile; sandalwood oil is almost colorless to golden yellow but viscous. Lemongrass and citronella Java-type oils are pale yellow to yellowish brown; the latter is slightly viscous. Geranium oil has various shades, starting from amber yellow to greenish yellow. Yellow to light brown is the rectified oil of clove leaf, but the crude oil is black. Vetiver and patchouli oils are yellow to reddish brown; both are viscous or even highly viscous. The oils of thyme (Thymus vulgaris L. and Thymus zygis (Loefl.) L.) are red. The color results from a reaction between thymol and iron of the still. Divergent colors have to be observed critical as these can be a result of aging. Citrus oils can be clear at ambient temperature but becomes cloudy at lower temperatures. Sometimes waxes can undergo precipitation by storage under 4°C. 20.5.1.2 Odor The odor is the most important factor for the application of essential oils. The highest quantity of usage is in the flavor and fragrance industry. Hatt et al. (2011) confirmed that the human sense of scent is the most important, even if the visual is dominating today. Smelling influences the human being by affecting the limbic system and is responsible for feelings and instincts, to trigger hormonal effects and activate pheromone production. The fact, that the human species has 350 gene receptors for smelling but only 4 for seeing will demonstrate the vital necessity of this sense. In times of high sophisticated analytical technique with highest resolutions, the human sense of smell is not replaceable. Especially small traces of aromatic chemicals with very low odor threshold are quickly identifiable. Examples are vanillin from vanilla beans and maltol, found in malt and caramel. For perfumer beginners, a single chemical is easy to recognize again. Essential oils are multicomponent mixtures, varying sometimes in wider limits. In spite of electronic noses a well-trained perfumer can recognize more nuances. Common human nose might be duped by adulterations, but not those of professional perfumers. At least the sense of taste is used to recognize adulteration. A good example is rose oil. Part of a drop applied to the tongue and mixture with synthetic geraniol or citronellol reflects in soapy and bitter taste.* 20.5.1.3  Physical–Chemical Methods These are inevitable in spite of many analytical certificates according to regulations. In history these values have been the only reference points to confirm naturalness. Although the bandwidth is often too wide, the values can be used as simple and easy to establish proof-samples. Gross mistakes and adulteration can be detected as well as aging of the oil. A series of ISO standards have been established especially for the quality control of essential oils: Relative density (ISO 279): This is defined as the relation between the mass of a defined volume of substance at 20°C and the mass of a comparable volume of water at the same temperature. Refractive index (ISO 280): This is the ratio of the sine of the angle of refraction, when a ray of light is passing from one medium into another. Three decimals are obligatory for the result. Optical rotation (ISO 592): This is the property of defined substances, to turn the plane of polarized light. Residue on evaporation (ISO, 4715), 1978: This is the residue of the essential oil after vaporization on the water quench, using defined conditions and is expressed in percent (m/m). Determination of acid value (1242): This value shows the number of milligrams of potassium hydroxide required to neutralize free acid in one gram of essential oil. Miscibility with ethanol (ISO 875), 1999: For that key figure, a defined quantity of an essential oil is added to an also defined mixture of distilled water and ethanol at 20°C. As result a blur occures visual. The value before obscuration is the value for a clear solution. * Based on the authors’ private experience.

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Determination of ethanol content (ISO 17494): This is the method of detecting the quantity of ethanol contained in an essential oil by GC analysis on a suitable column. Determination of the carbonyl value—free hydroxylamine method (ISO 1271): The determination is done by converting the carbonyl compounds to oximes by reaction with free hydroxylamine liberated in a mixture of hydroxylammonium chloride and potassium hydroxide. Determination of water content—Karl Fischer method (ISO 11021): A reagent (Karl Fischer reagent without pyridine) is used for the reaction of the absorbed water from an essential oil. The reagent is produced by titration, using a Karl Fischer apparatus. The final result of the reaction is obtained by an electronic method. Determination of phenol content (ISO, 1272), 2000: The transformation of phenolic compounds into their alkaline phenol esters, then soluble in aqueous phase, in a defined volume of essential oil is proceeded. The volume of unabsorbed portion is measured. Determination of peroxide value (ISO 3960): The peroxide value is the quantity, expressed as oxygen, to oxidize potassium iodide under specific conditions, divided through the mass of testing substance. Determination of freezing point (ISO 1041): This is the highest temperature during freezing of an undercooled liquid. Iodine number: Kumar and Madaan (1979) report in their chapter the use of iodine number. This method is a measure of the unsaturation of a substance (as an oil or fat) expressed as the number of grams of iodine or equivalent halogen absorbed by 100 g of the substance (Merriam-Webster, n.d.). By using the iodine monobromide-mercuric acetate reagent for iodination, a much better result could be achieved, as reported by Kumar. With this method adulterations on essential oils can be successfully detected. Thin-layer chromatography: This method belongs to wet chemistry (see Section 20.18.5.2). All earlier-mentioned methods are still involved in the concerning standards and are useful tools to confirm naturalness and purity of essential oils. 20.5.1.4  Calculation of Relationship Coefficient This method might be derided by some people, but once a relationship is recognized, it will help to detect mixtures with natural fractions or isolates. Touche et al. (1981) presented reference factors for identification of adulteration in French lavender. The ratio between (Z)-β-ocimene and (E)β-ocimene (R1), (E)-β-ocimene and 3-Octanone (R2), and linalool + linalyl acetate to lavandulol + lavandulyl acetate had been determined using GC data of production values over more than 5 years. Schmidt (2003) showed in accordance to that the differentiation of the minor components of lavender oils as the possibility to detect even the provenance of the oil. Of course such a calculation of relationship can be done for many other essential oils. As long as typical minor substances are present, it is a valuable tool for genuineness control.

20.5.2 Analytical Methods 20.5.2.1  General Tests Before the availability of sophisticated analytical instrumentation, only rather simple tests could be carried out to detect adulterations in essential oils (EOs), and it can be assumed that at that time many falsifications went undetected and adulterations were widespread. To determine the identity and purity of EOs, several elementary methods are described in the European Pharmacopoeia (Ph. Eur.) and ISO Standards (ISO/TC 54). Physicochemical properties like relative density (Ph.Eur. 10th ed. 2.2.5, ISO 279), refractive index (Ph.Eur. 10th ed 2.2.6, ISO 280), optical rotation (Ph.Eur. 10th ed. 2.2.7, ISO 592), and flash point (ISO/DIS 3679:2015) should be within certain limits and numerical values can be found in

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the literature, on the Internet, and in ISO standards. Solubility tests can indicate the presence of contaminants (solubility in ethanol ISO 875), for instance, a turbidity of a solution of an EO in carbon disulfide is an indication of water (ISO 11021:1999). To detect fatty oil adulterations, a drop of the EO is placed on a filter paper and after 24 h no visible grease spot must be left behind (Ph. Eur. 7.0/2.08.07.00). Determination of the evaporation residue serves the same purpose and detects nonvolatile or low volatile compounds (ISO, 4715). An instrumentally more demanding method would be thermogravimetry. Basic chemical tests can also provide useful information on the properties of EOs. Ester and saponification value (ISO, 7660), content of free and total alcohol (ISO, 1241), carbonyl value (ISO, 1279, 1996), and phenol content (ISO, 1272, 2000) are helpful indicators of identity and purity of EOs. Last but not the least sensory evaluation is important especially in perfume houses and should be carried out by experts or a panel of fragrance professionals most effectively done with the help of reference samples of known status. But some parameters alone determined by the earlier-mentioned methods tell us nothing about the composition of the tested EO. Therefore, separation of the oil into their individual components is necessary. 20.5.2.2  Thin-Layer Chromatography One of the oldest and easiest separation methods is thin-layer chromatography (TLC), which nevertheless has the disadvantage of having a quite low separation efficiency. A drop of the EO is placed at one end of a plate (glass, aluminum foil) coated with a thin layer of an adsorbent (silica gel, diatomaceous earth, aluminum oxide, cellulose) as the stationary phase and placed in a glass chamber filled with a small amount of an appropriate solvent as a mobile phase. The solvent is drawn up the plate through capillary forces and separates the oil into several fractions. These fractions or spots can be made visible by derivatization with a chromophoric reagent or by inspection under UV light. A reference oil of known purity and composition must be analyzed simultaneously or under the same conditions for comparison (Ph. Eur. 7.0/2.02.26.00); (Jaspersen-Schib and Flueck, 1962; Phokas, 1965; Atal and Shah, 1966; Sen et al., 1974; Kubeczka and Bohn, 1985; Nova et al., 1986; John et al., 1991). 20.5.2.3  Gas Chromatography (GC, GLC, HRGC, GC-FID, GC-MS) The analysis of natural products can be demanding especially in the field of EO research since an EO can contain 300 or more components at a concentration ranging from more than 90% to a few ppm or less of the total oil amount. A separation technique of high efficiency is needed and gas liquid chromatography (GLC) or simply GC is one of the most importantly used analytical methods for volatile compounds and is especially suited in EO and fragrance analysis. With the introduction of capillary columns, the separation performance increased dramatically and theoretical plate numbers of more than 1 × 105 are commonplace and provide gas chromatograms of high resolution (HRGC). A small amount of the EO (nanograms to micrograms) is injected with a microliter syringe into a closed evaporation chamber (injector) held at elevated temperature (>200°C) where the sample evaporates and the vapor is transferred by an inert carrier gas (N2, H2, He, Ar) as the mobile phase to a quartz capillary column of an internal diameter ranging from 0.05 to 0.53 mm and a length between a few meters up to a hundred meters or more. The inner wall of the capillary is coated with a thin film of a liquid phase (polysiloxane, polyglycol) in which separation takes place through permanent absorption and desorption of the sample compounds between liquid and mobile phase. The column is mounted in an oven either held at constant temperature or more often programmed at a constant rising temperature rate and permanently flushed with the carrier gas. The substances pass through the column and leave at a reproducible time (retention time) and then need to be registered by an appropriate detector. The most common and versatile detector for organic compounds is the flame ionization detector (FID) in which the eluted substances are burnt in a tiny hot hydrogen/air flame

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producing charged carbon ions that are collected on an anode. The resulting current is registered after amplification by an electrometer. Amplitude and signal (peak) area are proportional to the amount of the corresponding compound. Retention times depend on many device parameters, and therefore, instrument-independent retention indices calculated according to Kovats (1958) or Van Den Dool and Kratz (1963) are used instead and can be compared with literature data for substance identification. To verify identification, a second analysis on a column of different polarity must be performed, and retention times or indices on both columns must match the data from the assumed compound or of those from a reference substance. This two-column approach is also useful to detect peak overlapping, which occurs quite often on single-column GC analysis of natural complex substances. Substance amounts are calculated from the individual peak areas as a percentage of the total peak areas of all substances multiplied by a substance-dependent correction factor, which accounts for the different sensitivities of each compound to FID detection. These correction factors should be determined at least for the major compound found in an EO relative to a standard substance (preferably a n-paraffin) whose factor is typically set to 1. Unfortunately many EO compounds are not available commercially in pure form or easily synthesized, so for convenience all correction factors are set to 1 to a first approximation. Chromatographic profiles and percentage composition of all important commercial essential oils and many oil-bearing plants can be found in the ISO standards and the literature of essential oil research (Formácek, 2002; Kubezka and Formácek, 2002; Adams, 2007), and genuine EOs should fit into this image within certain limits. Chemical profiling in this manner can detect adulterations if additional or missing peaks are found or percentage composition deviates substantially. Addition of synthetic compounds can in some cases be disclosed by the presence of by-products or impurities left behind from the synthesis pathway. Dihydro linalool, for example, is an intermediate in the industrial production of linalool and is always present in small amounts (≪1%), and if a trace of this substance is found in an EO, it is a clear indicator of an adulteration with synthetic linalool. However, fingerprinting of EOs by GC with FID detection alone cannot reveal the chemical identity of detected peaks if any deviations from the expected profiles occur. In that case further information is needed besides the chromatographic data, and mass spectrometry provides detailed information on the structure of the separated compounds. High-resolution gas chromatography coupled with a mass spectrometric detector (HRGC-MS or simply GC-MS), most commonly quadrupole ion trap detectors in EO analysis, together with sophisticated chromatographic software and special mass spectral libraries of essential oil components (Königet al., 2004; Adams, 2007; Mondello, 2011) separates and identifies most components of an EO. Since some classes of compounds show very similar mass spectra, like some groups of mono- and sesquiterpenes, retention indices must be taken into account as a second criterion for an unequivocal identification. Usually a GC-MS run is performed for identifying the EO components and a second GC-FID run for peak area, respectively, for percentage composition determination. Normally this is done on two different instruments. Identical capillary columns must be used in both GCs, and device parameters must be adjusted properly to obtain closely similar chromatographic profiles for both detectors to facilitate peak allocation between the two chromatograms. Peaks identified in the mass spectrometry (MS) chromatogram must be correctly assigned in the FID chromatogram for peak integration. Sometimes this is proving difficult since separation on two instruments is never exactly equal especially if one column ends up in a high vacuum (MS) and the other at atmospheric pressure (FID). Therefore, a series of closely eluting peaks may not be resolved in the same manner on both columns. To overcome this problem, an FID-MS splitter can be used since here the separation takes place on one GC column and the effluent is split to both detectors, MS and FID. To detect peak overlapping, the same procedure should be undertaken on another capillary column of different polarity. In EO analysis GC separations are preferably performed on 95% dimethylpolysiloxane/5% diphenylpolysiloxane and on Carbowax 20 M columns. In the end you come up with two FID and two MS chromatograms, each of the two carried on different capillary columns. These results in two analyses that should for the most part coincide and accept the overlapping peaks on the other GC column. EO analysis performed in this way confirms the chemical

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composition of an EO and detects adulterations with exogenic substances if they are amenable by GC, that is, volatile. Adulterations with gas chromatographic undetectable substances like very high boiling vegetable or mineral oils can be disclosed by a change in the percentage composition or a too low total peak area. Specific marker compounds or a diastereomeric isomer distribution can also be used for authentication of EO (Teisseire, 1987). In some unusual EOs not all detected peaks can be identified by MS library search because of missing entries but, with the necessary experience, it can often be estimated in which group of natural substances these peaks belong by looking at the mass spectrometric pattern. All in all an extensive experience in EO analysis and a thorough knowledge of the scientific literature is needed to evaluate the authenticity of an EO and detect adulterations. If this is the case the analyst should be in a position to disclose falsifications with cheaper or inappropriate EOs, synthetic natural substances, solvents, or other unnatural chemicals. 20.5.2.4  Chiral Analysis Many EOs contain substances with asymmetric carbon atoms, that is, there are two molecular forms of these molecules that behave like image und mirror image and cannot be brought into alignment (Busch and Busch, 2006). This is the reason why EOs rotate the oscillating plane of light and the resulting overall optical rotation is measured by a polarimeter. However, the enantiomeric distribution of single substances cannot be determined in this way because separation is needed in the first place. This will be enabled by GC, and additional separation of optically active compounds into their enantiomers can be achieved if a chiral selector is added to the stationary phase of the GC column. Cyclodextrin derivatives have proven to be very useful and chiral capillary columns with different optically active cyclodextrin selectors are commercially available. The enantiomeric separation should be done at rather low temperature since the interaction between the chiral molecules and the chiral selector is rather weak; therefore, the temperature gradient of the GC program should be low and the carrier gas flow is higher than the optimal flow. If the appropriate chiral GC column is used, the enantiomeric distribution of certain EO components can be determined, and since nature often prefers one enantiomer over the other, the determination of the enantiomeric excess is a valuable tool to detect adulterations by synthetic components that are racemic and change the enantiomeric ratio. The optical purity of authentic EOs can be found in the literature (Mosandl, 1998; Busch and Busch, 2006; Dugo and Mondello, 2011). 20.5.2.5  GC-GC and GC × GC (Two-Dimensional Gas Chromatography, 2D GC) Gas chromatography is one of the most widely deployed methods in analytical chemistry to investigate organic sample material due to its simple ease of use, the ready availability of sophisticated inexpensive instrumentation, and the large amount of qualitative and quantitative information that can be retrieved if the appropriate configuration is employed. Especially the high separation efficiency for volatiles makes GC very suitable to investigate complex mixtures and sample matrices. But for some applications, the separation performance is not sufficient when it comes to very complex mixtures like odors, flavors, crude oil products, and foodstuff. Co-elution with other analytes or sample matrix elements causes problems in detection and quantitation especially when the analytes differ greatly in their concentration. This problem can be solved by cutting out the co-eluting part of the chromatogram and a subsequent second chromatographic separation of the excised effluent preferably on a stationary phase of different polarity. This technique, called heart-cutting or twodimensional GC (2D GC or GC-GC), is done with the help of a diverting valve or a Deans switch. The sought-after substances are then resolved in the second GC column. An even more elaborate technique recently developed not only cuts out one or several parts of the first GC column but virtually cuts the whole 1D chromatogram into small equal pieces (each several seconds long), refocuses each effluent, and separates it on a short second GC column within a very short time (several seconds). Refocusing, which here means stopping the effluent of the first column for several seconds onto a very small area and then releasing the focused substances into the second column, is

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done with a cryogenic modulator, and this procedure is repeated until all substances are eluted from the first GC column. The overall separation efficiency is calculated by multiplying the theoretical plate numbers of the two columns, which results in rather large figures and a separation efficiency, which cannot be achieved by simple GC alone. Instruments and the appropriate data acquisition software for this comprehensive 2D GC (or GC × GC) are now commercially available. Instead of a 1D chromatogram, a 2D contour plot is generated, and since the “half-widths” of substances are very small, detection must proceed very fast, so in case a mass spectrometer is used as a detector, a fast scanning quadrupole or a time-of-flight MS (TOF-MS) must be chosen. Such an instrument arrangement like GC × GC-TOF-MS leaves almost nothing to be desired for EO analysis (Marsili, 2010). 20.5.2.6  13C NMR (Nuclear Magnetic Resonance) Another useful method to validate the identity of an EO offers the spectroscopy of the magnetic properties of 13C nuclei. 13C NMR spectroscopy is a very useful technique regularly used to elucidate the structure of individual substances and has been applied by Kubeczka and Formácek (2002) to a large number of essential oils and individual reference compounds. Even though it is not very common to apply this technique to substance mixtures, as it is with an EO, the spectrum of a genuine oil is very distinct and characterized by the chemical shifts, signal multiplicities, and intensities of all components. Functional groups can be identified by chemical shifts and provide information on chemical structures of individual substances within the oil. No separation of the oil into its components is necessary, and therefore, the approach is simply measuring a solution of the oil in an NMR tube and comparing the spectrum with literature data or reference oils and compounds. Unfortunately NMR instruments are rarely found in traditional analytical laboratories, and thus, this method has not gained the significance in the field of EO analysis it deserves.

20.6  IMPORTANT ESSENTIAL OILS AND THEIR POSSIBLE ADULTERATION 20.6.1 Ambrette Seed Oil This is a very expensive oil. The main ingredients are (E,E)-farnesyl acetate (60%) and ambrettolide (8.5%). Synthetic ambrettolide is used as a fixative in perfumery, is nearly odorless but has exalting properties. Compared to natural ambrette seed oil, the price of ambrettolide is only 10%. Detection of naturality can be done by isotope ratio mass spectrometry (IRMS).

20.6.2 Amyris Oil ISO standard 3525 shows character and data for this oil. Blending (professional term within dealers and perfumers for adulterating) is done by Virginia cedarwood oil, α-terpineol, and copaiba balm. Also, elemol distilled from elemi resin is used. Detection is done by GC-MS.

20.6.3 Angelica Oils The fruit oil contains up to 76% of β-phellandrene and α-pinene (13%) as main ingredients. As no chiral values for the β-phellandrene are described in literature, it must be assumed that adulteration is done by this compound. β-Phellandrene is naturally available by geranyl diphosphate cycling (BRENDA, BRaunschweig ENzyme DAtabase), 2007. For adulteration copaiva balm, gurjun balsam, lovage root oil, and amyris oil were used in the past. Cheap α-pinene can also be used to “improve” the composition.

20.6.4 Anise Fruit Oil ISO standard 3475 shows character and data for that oil. This oil often is produced not only from the fruits but also from the whole aerial part. Values of cis- and trans-anethole are then reduced in smaller amount. Adulteration with star anise oil can be easily detected. Anise fruit oil does not contain any

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foeniculin, but star anise oil contains up to 3% (ISO 11016). On the other hand pseudo-isoeugenol2-methylbutyrate is a component of anise fruit oil (0.3%–2%, ISO 3475) and does not appear in star anise oil. The midratio of cis-anethole to trans-anethole in anise seed oil in relation to star anise oil is 0.3%:0.6%. Synthetic anethole, fennel terpene limonene (80°), and terpineol were used for blending.

20.6.5 Armoise Oil This oil is often blended with α- and β-thujones from cheaper sources like Thuja orientalis. Furthermore camphor or white camphor oil and camphene are used. GC-MS method is used to detect these adulterations.

20.6.6 Basil Oils ISO standard 11043 shows character and data for the methyl chavicol–type oil. Synthetic methyl chavicol is used to adulterate that oil and can be detected by NMR. Basil oil from linalool type will be adulterated by synthetic linalool but can easily be detected by chiral separation. Casabianca (1996a) found the minimum value R-(−)-linalool with 99.8% and is congruent with the authors’ results.

20.6.7 Bergamot Oil ISO standard 3520 shows character and data for that oil. As mentioned in the text, this oil was adulterated with synthetic linalool and linalyl acetate but also with bergamot terpenes and distilled oil from bergamot peel residues. Blending was done with limonene (80°), nerol, geranyl acetate, petitgrain oil Paraguay, rosewood oil, and citral from Litsea cubeba oil. This oil was subject to many studies with chiral separation. König et al. (1992); Mosandl et al. (1991); Dugo et al. (1992) and Casabianca (1996a), reported similar values for linalool and linalyl acetate. (R)-(−)-linalyl acetate is present always in purity higher than 99.0% and (R)-(−)-linalool with a minimum of 99.0% too. (S-pinene value varies between α)-(−)- 68.0% and 71.1%, (S-pinene between 91.1% and 92.6%β)-(−)-, and (R)-(+)-limonene between 97.4% and 98.0% (Juchelka, 1996a). Blending is done by terpinyl acetate, citral synthetic, n-decanal, n-nonanal, nerol, limonene (80°), and sweet orange terpenes. Dugo and Mondello (2011) published the following data for chiral ratios: (S)-thujene (0.5%–1.3%):(α)-(+)-(R)-thujene (98.7%–99.5%); (α)-(−)-(R)-(+)-α-pinene (31.0%–36.1%):(S)-(−)-α-pinene (63.9–69.0); (1S,4R)-(−)-camphene (85.7%–92.7%):(1R,4S)(+)-camphene (7.3%–14.3%); (R)-(+)-β-pinene (7.6%–10.3%):(S)-(−)-β-pinene (89.7%–92.4%); (R)-(+)-sabinene (13.7%–19.8%):(S)-(−) (80.2%–86.3%); (R)-(−)-α-phellandrene (43.1%–54.7%):(S)(+)-α-phellandrene (45.3%–56.9%); (R)-(−)-β-phellandrene (24.1%–36.9%):(S)-(+)-β-phellandrene (63.1%–75.9%); (S)-(−)-limonene (1.2%–2.1%):(R)-(+)-limonene (97.9%–98.8%); (R)-(−)-linalool (97.8%–99.5%):(S)-(+)-linalool (0.5%–2.2%); (S)-(−)-citronellal (>98%):(R)-(+)-citronellal (99.0%):(1R,4S)-(−)-menthone (99.0%):(1R,4R)-(+)isomenthone (99%; (2S,4R)-cis-rose oxide as well as (2R,4R)-cis-rose oxide show a purity higher than 99.5%.

20.6.64 Rosemary Oil ISO standard 1342 shows character and data for oils from various sources. Turpentine oil, synthetic camphor, and limonene from orange terpenes are used to blend this oil. Adulteration is done by 1,8-cineole from eucalyptus or white camphor oil. Detection is usually done by GC-MS but also by multidimensional chiral separation. Enantiomeric ratio of linalool is reported by Casabianca (1996a) to be (R)-(−) 23 to (S)-(+) 77. Kreis (1991) published the following values of chiral separation: (−)-borneol (84.6%–97.8%):(+)-borneol (2.2%–15.4%) and (−)-isoborneol (29.1%–53.8%):(+)-isoborneol (46.2%–70.9%). König et al. (1992) reported the following chiral ratios: (+)-α-thujene 33.7%:(−)-α-thujene 66.3%; (−)-α-pinene 41.7%:(+)-α-pinene 58.3%; (−)-β-pinene 84.4%:(+)-β-pinene 15.6%; (−)-camphene 83.3%:(+)-camphene 16.7%; (−)-α-phellandrene 0%:(+)-α-phellandrene 100%; (−)-β-phellandrene 2.5%:(+)-β-phellandrene 97.5%; and (−)-limonene 64.8%:(+)-limonene 35.2%. The authors own results analyzing pure oils showed the following ratios: (+)-β-pinene 26.0%:(−)-β-pinene 74.0%; (−)-α-pinene 42.0%:(+)-α-pinene 58.0%; (S)-(−)-sabinene 66.5%:(R)-(+)-sabinene 33.5%; and (S)-(−)-camphene 31.4%:(R)-(+)-camphene 68.6%.

20.6.65 Rosewood Oil ISO standard 3761 shows character and data for this oil. Blending is done by synthetic linalool, α-terpineol, geraniol, and heads of rosewood oil. Eremophilane is a marker according to Lawrence (1999a,b) with values of 0.3%–0.9%. Detection is done by GC-MS on a chiral column. Casabianca (1996a) found chiral ratio for linalool with (S)-(+) 50.0%−51% and (R)-(−) 49.0%–50%. Lawrence

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(1999a,b) reported the following chiral ratios: (3R)-(−)-linalool 10.0%:(3S)-(+)-linalool 90.0%. (For information, rosewood leaf oil ratio is (3R)-(−)-linalool 22.2%:(3S)-(+)-linalool 77.8%).

20.6.66 Sage Oil (Salvia officinalis) ISO standard 9909 shows character and data for this oil. As the main components α-thujone β-thujone are known, adulteration is done with thuja oil or cedar leaf oil. β-Caryophyllene, 1,8-cineole, and borneol from other sources are used. α-Humulene is a marker with up to 12% total content. Detection is done by GC-MS analysis.

20.6.67 Sage Oil Spanish Type ISO standard 3526 shows character and data for this oil derived from Salvia lavandulifolia Vahl. Adulteration is done by eucalyptus oil, camphor oil chemotypes like 1,8-cineole and camphor, α-pinene, and limonene from different sources. Blending is done with synthetic linalyl acetate und terpinyl acetate. Detection can be achieved by using GC and GC-MS analytical equipment.

20.6.68 Sandalwood Oil ISO standard 3518 shows character and data for the oil derived from Santalum album L. This very expensive oil with source from India is no longer sold on the market these days. Today other varieties from various sources like New Caledonia (Santalum austrocaledonicum Vieill. and Santalum spicatum (R.Br.) A. DC) from Australia are used in flavor and fragrance industry. α- and β-Santalols are the main components and responsible for the fragrance. Adulteration will take place with cheaper varieties, coming from Australia. This is very simple to detect as Santalum spicatum contains cisnuciferol in high amounts. The same can be seen with S. austrocaledonicum, where the cis-lanceol can be found in high amounts compared to the other oils. Braun et al. (2005) found in analyzing S. album oil the following chiral substances and values: (1R,4R,5S)-α-acorenol (0.22%), (1R,4R,5R)-βacorenol (0.11%), (1R,4S,5S)-epi-α-acorenol (0.13%), and (1R,4S,5R)-epi-β-acorenol ( wheat > barnyard grass. Enantio- and diastereoselective biotransformation of trans- (81a and 81a′) and cis-carveols by E. gracilis Z. (Noma and Asakawa, 1992) and Chlorella pyrenoidosa IAM C-28 was studied (Noma et al., 1997). In the biotransformation of racemic trans-carveol (81a and 81a′), C. pyrenoidosa IAM C-28 showed high enantioselectivity for (−)-trans-carveol (81a′) to give (−)-carvone (93′), while (+)-­trans-carveol (81a) was not converted at all. The same C. pyrenoidosa IAM C-28 showed high enantioselectivity for (+)-cis-carveol (81b) to give (+)-carvone (93) in the biotransformation of racemic cis-carveol (81b and 81b′). (−)-cis-Carveol (81b′) was not converted at all. The same phenomenon was observed in the biotransformation of mixture of (−)-trans- and (−)-cis-carveol (81a′ and 81b′) and the mixture of (+)-trans- and (+)-cis-carveol (81a and 81b) as shown in Figure 22.90. The high enantioselectivity and the high diastereoselectivity for the dehydrogenation of (−)trans- and (+)-cis-carveols (81a and 81b′) were shown in E. gracilis Z. (Noma and Asakawa, 1992), C. ­pyrenoidosa IAM C-28 (Noma et al., 1997), N. tabacum, and other Chlorella spp. On the other hand, the high enantioselectivity for 81a′ was observed in the biotransformation of racemic (+)-trans-carveol (81a) and (−)-trans-carveol (81a′) by Chlorella sorokiniana SAG to give (−)-carvone (93′). It was considered that the formation of (−)-carvone (93′) from (−)-trans-carveol (81a′) by diastereo- and enantioselective dehydrogenation is a very interesting phenomenon in order to produce mosquito repellent (+)-p-menthane-2,8-diol (50a′) (Noma, 2007). (4R)-trans-Carveol (81a′) was converted by S. litura to give 1-p-menthene-6,8,9-triol (375) (Miyazawa et al., 1996b) (Figure 22.91).

OH

OH O

OH

81a

O

OH

81a΄

81b

OH

93

OH 93΄

50΄

81b΄

FIGURE 22.90  Enantio- and diastereoselective biotransformation of trans- (81a and a′) and cis-carveols (81b and b′) by Euglena gracilis Z and Chlorella pyrenoidosa IAM C-28. (Modified from Noma, Y. and Asakawa, Y., Phytochemistry, 31, 2009, 1992; Noma, Y. et al. Biotransformation of terpenoids and related compounds by Chlorella species, Proceedings of 41st TEAC, 1997, pp. 227–229.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

OH

669

OH

S. litura

OH OH

81a΄

375΄

FIGURE 22.91  Biotransformation of (4R)-trans-carveol (81a′) by Spodoptera litura. (Modified from Miyazawa, M. et al. Biotransformation of p-menthanes using common cutworm larvae, Spodoptera litura as a biocatalyst, Proceedings of 40th TEAC, 1996b, pp. 80–81.)

OH

102a (1S,2R,4R) (–)-Neo

OH

102a΄ (1R,2S,4S) (+)-Neo

OH

102b (1S,2S,4R) (+)-Dihydrocarveol

102d (1R,2R,4R) (+)-Neoiso

102c (1R,2S,4R) (+)-Iso

OH

OH

102b΄ (1R,2R,4S) (–)-Dihydrocarveol

OH

OH

102c΄ (1S,2R,4S) (–)-Iso

OH

102d΄ (1S,2S,4S) (–)-Neoiso

FIGURE 22.92  Chemical structure of eight kinds of dihydrocarveols.

22.3.3.10 Dihydrocarveol (+)-Neodihydrocarveol (102a′) was converted to p-menthane-2,8-diol (50a′), 8-p-menthene-2, 8-diol (107a′), and p-menthane-2,8,9-triols (104a′ and b′) by A. niger TBUYN-2 (Noma et al., 1985a,b; Noma and Asakawa, 1995) (Figures 22.92 and 22.93). In the case of E. gracilis Z., mosquito repellent (+)-p-menthane-2,8-diol (50a′) was formed stereospecifically from (−)-carvone (93′) via (+)-dihydrocarvone (101a′) and (+)-neodihydrocarveol (102a′) (Noma et al., 1993; Noma, 1988, 2007). (−)-Neodihydrocarveol (102a) was also easily and stereospecifically converted by E. gracilis Z. to give (−)-p-menthane-2,8-diol (50a) (Noma et al., 1993). On the other hand, A. glauca converted (−)-carvone (93′) stereospecifically to give (+)-8-p-menthene-2,8-diol (107a′) via (+)-dihydrocarvone (101a′) and (+)-neodihydrocarveol (102a′) (Demirci et al., 2004) (Figure 22.93). (+)- (102b) and (−)-Dihydrocarveol (102b′) were converted by 10 kinds of Aspergillus spp. to give mainly (+)- (107b′) and (−)-10-hydroxydihydrocarveol (107b, 8-p-menthene-2,10-diol) and (+)(50b′) and (−)-8-hydroxydihydrocarveol (50b, p-menthane-2,8-diol), respectively (Figure 22.94). The metabolic pattern of dihydrocarveols is shown in Table 22.4.

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OH

OH

OH 50a

102a OH

OH

OH

+

+

OH 102a΄

50a΄

OH

OH

+

OH

OH 104a΄

107a΄

OH

OH HO

104b΄

FIGURE 22.93  Biotransformation of (−)- and (+)-neodihydrocarveol (102a and a′) by Euglena gracilis Z, Aspergillus niger TBUYN-2, and Absidia glauca. (Modified from Noma, Y. et  al. Annual Meeting of Agricultural and Biological Chemistry, Sapporo, 1985a, p. 68; Noma, Y. et al. Biotransformation of carvone. 6. Biotransformation of (−)-carvone and (+)-carvone by a strain of Aspergillus niger, Proceedings of 29th TEAC, 1985b, pp. 235–237; Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25; Noma, Y., Aromatic Plants from Asia their Chemistry and Application in Food and Therapy, L. Jiarovetz, N.X. Dung, and V.K. Varshney, pp. 169–186, Har Krishan Bhalla & Sons, India, 2007; Noma, Y. and Asakawa, Y., Biotechnology in Agriculture and Forestry, Vol. 33. Medicinal and Aromatic Plants VIII, Y.P.S. Bajaj, ed., pp. 62–96, Springer, Berlin, Germany, 1995; Demirci, F. et al. Naturforsch., 59c, 389, 2004.)

OH

OH

OH Aspergillus sp. Aspergillus sp.

E.g. OH

OH

50b

107b

102b

OH

OH

OH Aspergillus sp. E.g.

Aspergillus sp.

OH 50b΄

OH 102b΄

107b΄

FIGURE 22.94  Biotransformation of (+)- (102b) and (−)-dihydrocarveol (102b′) by 10 kinds of Aspergillus spp. (Modified from Noma, Y., Formation of p-menthane-2,8-diols from (−)-dihydrocarveol and (+)-dihydrocarveol by Aspergillus spp. The Meeting of Kansai Division of The Agricultural and Chemical Society of Japan, Kagawa, 1988, p. 28) and Euglena gracilis Z (Modified from Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25.)

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Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

TABLE 22.4 Metabolic Pattern of Dihydrocarveols (102b and 102b′) by 10 Kinds of Aspergillus spp. Compounds Microorganisms

107bʹ

50bʹ

C.r. (%)

A. awamori, IFO 4033 A. fumigatus, IFO 4400

0 0

98 14

99 34

A. sojae, IFO 4389 A. usami, IFO 4338

0 0

47 32

59 52

A. cellulosae, M-77

0

27

52

A. cellulosae, IFO 4040 A. terreus, IFO 6123

0 0

30 79

55 92

A. niger, IFO 4034

0

29

49

A. niger, IFO 4049 A. niger, TBUYN-2

4 29

50 68

67 100

107b

50b

C.r. (%)

3

81 6

94 14

50 5

85 7

7

14

+

5 18

8 46

+ 9 30

8

12

34 53

59 100

+ 1 + + 1

Note: C.r.—conversion ratio.

OH

OH

O OH

O

O

O

105aa

HO

102b OH

OH

O 105aa΄

OH

OH

105ab

OH

O

O

O 102b΄

HO 105ab΄

FIGURE 22.95  Biotransformation of (+)- (102b) and (−)-dihydrocarveol (102b′) by Streptomyces bottropensis SY-2-1. (Modified from Noma, Y., Kagaku to Seibutsu, 22, 742, 1984.)

In the case of the biotransformation of S. bottropensis SY-2-1, (+)-dihydrocarveol (102b) was converted to (+)-dihydrobottrospicatol (105aa) and (+)-dihydroisobottrospicatol (105ab), whereas (−)-dihydrocarveol (102b′) was metabolized to (−)-dihydrobottrospicatol (105aa′) and (−)-dihydroisobottrospicatol (105ab′). (+)-Dihydroisobottrospicatol (105ab) and (−)-dihydrobottrospicatol (105aa′) are the major products (Noma, 1984) (Figure 22.95). E. gracilis Z. converted (−)-iso- (102c) and (+)-isodihydrocarveol (102c′) to give the corresponding 8-hydroxyisodihydrocarveols (50c and 50c′), respectively (Noma et al., 1993) (Figure 22.96). In the case of the biotransformation of S. bottropensis SY-2-1, (−)-neoisodihydrocarveol (102d) was converted to (+)-isodihydrobottrospicatol (105ba) and (+)-isodihydroisobottrospicatol (105bb), whereas (+)-neoisodihydrocarveol (102d′) was metabolized to (−)-isodihydrobottrospicatol (105ba′) and (−)-isodihydroisobottrospicatol (105bb′). (+)-Isodihydroisobottrospicatol (105bb) and (−)-isodihydrobottrospicatol (105ba′) are the major products (Noma, 1984) (Figure 22.97).

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OH

OH

OH

E. graclis

OH

E. graclis OH

102c

OH

50c

102c΄

50c΄

FIGURE 22.96  Biotransformation of (+)-iso- (102c) and (−)-dihydrocarveol (102c′) by Euglena gracilis Z. (Modified from Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25.)

OH

OH

O

O

OH

OH

O

105ba

O HO

102d

OH

OH

O

105bb

O

O

O

OH

OH

HO

105ba΄

102d΄

105bb΄

FIGURE 22.97  Formation of dihydroisobottrospicatols (105) from neoisodihydrocarveol (102d and d′) by Streptomyces bottropensis SY-2-1. (Modified from Noma, Y., Kagaku to Seibutsu, 22, 742, 1984.)

OH

OH

OH

E. graclis

OH

E. graclis

OH 102d

50d

OH 102d΄

50d΄

FIGURE 22.98  Biotransformation of (+)- (102c) and (−)-neoisodihydrocarveol (102c′) by Euglena gracilis Z. (Modified from Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

673

E. gracilis Z. converted (−)- (102d) and (+)-neoisodihydrocarveol (102d′) to give the corresponding 8-hydroxyneoisodihydrocarveols (50d and 50d′), respectively (Noma et al., 1993) (Figure 22.98). Eight kinds of 8-hydroxydihydrocarveols (50a–d and 50a′–d′; 8-p-menthane-2,8-diols) were obtained from carvone (93 and 93′), dihydrocarvones (101a–b and 101a′–b′), and dihydrocarveols (102a–d, 102a′–d′) by E. gracilis Z. as shown in Figure 22.99 (Noma et al., 1993). 22.3.3.11 Piperitenol

HO

458

Incubation of piperitenol (458) with A. niger gave a complex metabolites whose structures have not yet been determined (Noma, 2000). 22.3.3.12 Isopiperitenol

HO

110

Piperitenol (458) was metabolized by A. niger to give a complex alcohol mixtures whose structures have not yet been determined (Noma, 2000). 22.3.3.13  Perillyl Alcohol OH

74 R-(+)-

OH

74΄ S-(–)-

(−)-Perillyl alcohol (74′) was epoxidized by S. ikutamanensis Ya-2-1 to give 8,9-epoxy-(−)-perillyl alcohol (77′) (Noma et al., 1986) (Figure 22.100). (−)-Perillyl alcohol (74′) was glycosylated by E. perriniana suspension cells to (−)-perillyl alcohol monoglucoside (376′) and diglucoside (377′) (Hamada et al., 2002; Yonemoto et al., 2005) (Figure 22.101). Furthermore, 1-perillyl-β-glucopyranoside (376) was converted into the corresponding oligosaccharides (377–381) using a cyclodextrin glucanotransferase (Yonemoto et al., 2005) (Figure 22.102).

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Handbook of Essential Oils OH

OH OH

O

50a

102a

OH

OH

101a

O

OH 102b

50b

OH

93

OH OH

O 50c

102c

OH

OH

101b΄

OH 50d

102d

(a)

OH

OH

OH

O 50a΄

102a΄

OH

OH 101a΄

OH

O 50b΄

102b΄

OH

OH 93΄

OH

O 50c΄

102c΄

OH

OH 101b

OH (b)

102d΄

50d΄

FIGURE 22.99  Formation of eight kinds of 8-hydroxydihydrocarveols (50a–50d, 50a′–50d′), dihydrocarvones (101a–101b and 101a′–101b′), and dihydrocarveols (102a–102d and 102a′–102d′) from (+)- (93) and (−)-carvone (93′) by Euglena gracilis Z.: (a) denotes 8-hydroxydihydrocarveols and (b) denotes dihydrocarveols. (Modified from Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related ­compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

675

OH

OH S. ikutamanensis

O 74΄

77΄

FIGURE 22.100  Biotransformation of (−)-perillyl alcohol (74′) by Streptomyces ikutamanensis Ya-2-1. (Modified from Noma, Y. et al. Reduction of terpene aldehydes and epoxidation of terpene alcohols by S. ikutamanensis, Ya-2-1, Proceedings of 30th TEAC, 1986, pp. 204–206.) O-Glc-Glc

O-Glc

OH

+

E. perriniana

74΄

376΄

377΄

FIGURE 22.101  Biotransformation of (−)-perillyl alcohol (74′) by Eucalyptus perriniana suspension cell. (Modified from Hamada, H. et al. Glycosylation of monoterpenes by plant suspension cells, Proceedings of 46th TEAC, 2002, pp. 321–322; Yonemoto, N. et al. Preparation of (−)-perillyl alcohol oligosaccharides, Proceedings of 49th TEAC, 2005, pp. 108–110.)

22.3.3.14 Carvomenthol OH

OH

OH

OH

49a

49b

49c

49d

(1S,2R,4R)

(1S,2S,4R)

(1R,2R,4R)

(–)-Neo

(1R,2S,4R)

(+)-Carvomenthol

(–)-Iso

(–)-Neoiso

OH

OH

OH

OH

49a΄

49b΄

49c΄

49d΄

(1R,2S,4S)

(1R,2R,4S)

(1S,2R,4S)

(1S,2S,4S)

(+)-Neo

(–)-Carvomenthol

(+)-Iso

(+)-Neoiso

(+)-Iso- (49c) and (+)-neoisocarvomenthol (49d) were formed from (+)-carvotanacetone (47) via (−)-isocarvomenthone (48b) by P. ovalis strain 6-1, whereas (+)-neocarvomenthol (49a′) and (−)-carvomenthol (49b′) were formed from (−)-carvotanacetone (47′) via (+)-carvomenthone (48a′) by the same bacteria, of which 48b, 48a′, and 49d were the major products (Noma et al., 1974a) (Figure 22.103).

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Handbook of Essential Oils O-Glc

O-Glc-(Glc)n n=1–5 CGTase 55°C, pH 5.5

376

377–381

FIGURE 22.102  Biotransformation of (−)-perillyl alcohol monoglucoside (376) by CGTase. (Modified from Yonemoto, N. et al. Preparation of (−)-perillyl alcohol oligosaccharides, Proceedings of 49th TEAC, pp. 108–110, 2005.) OH

OH

O

+

P. ovalis 6-1

49c

47

49d

OH

O

OH

+

P. ovalis 6-1

47΄

49b΄

49a΄

FIGURE 22.103  Formation of (−)-iso- (49c), (−)-neoiso- (49d), (+)-neo- (49a′), and (−)-carvomenthol (49b′) from (+)- (47) and (−)-carvotanacetone (47′) by Pseudomonas ovalis strain 6-1. (Modified from Noma, Y. et al. Agric. Biol. Chem., 38, 1637, 1974a.)

Microbial resolution of carvomenthols was carried out by selected microorganisms such as Trichoderma S and B. subtilis var. niger (Oritani and Yamashita, 1973d). Racemic carvomenthyl acetate, racemic isocarvomenthyl acetate, and racemic neoisocarvomenthyl acetate were asymmetrically hydrolyzed to (−)-carvomenthol (49b′) with (+)-carvomenthyl acetate, (−)-isocarvomenthol (49c) with (+)-isocarvomenthyl acetate, and (+)-neoisocarvomenthol (49d′) with (−)-neoisocarvomenthyl acetate, respectively; racemic neocarvomenthyl acetate was not hydrolyzed (Oritani and Yamashita, 1973d) (Figure 22.104).

22.3.4 Monocyclic Monoterpene Ketone 22.3.4.1  α, β-Unsaturated Ketone 22.3.4.1.1 Carvone O

O S

93 (4S)-(+)-form

R 4 93΄ (4R)-(–)-form

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OAc

OAc

Trichoderma S B. subtilis 49a΄

677

Not hydrolyzed

49a΄ OAc

OH

OAc

OAc

Trichoderma S B. subtilis 49b΄ OAc

49b΄

49b

49b΄

OH

OAc

OAc

Trichoderma S B. subtilis 49c OAc

49c ΄

49c

49c

OH

OAc

OAc

Trichoderma S B. subtilis 49d 49d Racemic acetates

49d

49d΄

FIGURE 22.104  Microbial resolution of carvomenthols by Trichoderma S and Bacillus subtilis var. niger. (Modified from Oritani, T. and Yamashita, K., Agric. Biol. Chem., 37, 1691, 1973d.)

Carvone occurs as (+)-carvone (93), (−)-carvone (93′), or racemic carvone. (S)-(+)-Carvone (93) is the main component of caraway oil (ca. 60%) and dill oil and has a herbaceous odor reminiscent of caraway and dill seeds. (R)-(−)-Carvone (93′) occurs in spearmint oil at a concentration of 70%– 80% and has a herbaceous odor similar to spearmint (Bauer et al., 1990). The distribution of carvone convertible microorganisms is summarized in Table 22.5. When ethanol was used as a carbon source, 40% of bacteria converted (+)- (93) and (−)-carvone (93’). On the other hand, when glucose was used, 65% of bacteria converted carvone. In the case of yeasts, 75% converted (+)- (93) and (−)-carvone (93′). Of fungi, 90% and 85% of fungi converted 93 and 93′, respectively. In actinomycetes, 56% and 90% converted 93 and 93′, respectively. Many microorganisms except for some strains of actinomycetes were capable of hydrogenating the C = C double bond at C-1, 2 position of (+)- (93) and (−)-carvone (93′) to give mainly (−)-isodihydrocarvone (101b) and (+)-dihydrocarvone (101a′), respectively (Noma and Tatsumi, 1973; Noma et al., 1974b; Noma and Nonomura, 1974; Noma, 1976, 1977) (Figure 22.105) (Tables 22.6 and 22.7). Furthermore, it was found that (−)-carvone (93′) was converted via (+)-­isodihydrocarvone (101b′) to (+)-isodihydrocarveol (102c′) and (+)-neoisodihydrocarveol (102d′) by some strains of actinomycetes (Noma, 1979a,b). (−)-Isodihydrocarvone (101b) was epimerized to (−)-­dihydrocarvone (101a) after the formation of (−)-isodihydrocarvone (101b) from (+)-carvone (93) by the growing cells, the resting cells, and the cell-free extracts of Pseudomonas fragi IFO 3458 (Noma et al., 1975).

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TABLE 22.5 Distribution of (+)- (93) and (−)-Carvone (93′) Convertible Microorganisms Microorganisms

Number of Microorganisms Used

Numbers of Carvone Convertible Microorganisms

40

16 (ethanol, 93)

Bacteria

Ratio (%) 40 40

16 (ethanol, 93′) 26 (glucose, 93) Yeasts Fungi Actinomycetes

68

26 (glucose, 93′) 51 (93)

40

51 (93′) 34 (93)

48

36 (93′) 27 (93)

65 65 75 75 85 90 56 90

43 (93′)

Source: Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25.

OH

OH

O

O 102a΄

102a OH

OH

101a

101a΄ O

O

102b΄

102b OH

93΄

93

OH

O

O

102c΄

102c OH

101b΄

OH

101b

102d΄

102d

FIGURE 22.105  Biotransformation of (+)- (93) and (−)-carvone (93′) by various kinds of microorganisms. (Modified from Noma, Y. and Tatsumi, C., Nippon Nogeikagaku Kaishi, 47, 705, 1973; Noma, Y. et al. Agric. Biol. Chem., 38, 735, 1974b; Noma, Y. et al. Microbial transformation of carvone, Proceedings of 18th TEAC, pp. 20–23, 1974c; Noma, Y. and Nonomura, S., Agric. Biol. Chem., 38, 741, 1974; Noma, Y., Ann. Res. Stud. Osaka Joshigakuen Junior College, 20, 33, 1976; Noma, Y., Nippon Nogeikagaku Kaishi, 51, 463, 1977.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

679

TABLE 22.6 Ratio of Microorganisms That Carried Out the Hydrogenation of C = C Double Bond of Carvone by Si Plane Attack toward Microorganisms That Converted Carvone Microorganisms

Ratio (%)

Bacteria

100a 96b 74 80 39

Yeasts Fungi Actinomycetes a b

When ethanol was used. When glucose was used.

TABLE 22.7 Summary of Microbial and Chemical Hydrogenation of (−)-Carvone (93′) for the Formation of (+)-Dihydrocarvone (101a′) and (+)-Isodihydrocarvone (101b′) Compounds Microorganisms Amorphosporangium auranticolor Microbispora rosea IFO 3559 Bacillus subtilis var. niger Bacillus subtilis IFO 3007 Pseudomonas polycolor IFO 3918 Pseudomonas graveolens IFO 3460 Arthrobacter pascens IFO 121139 Pichia membranifaciens IFO 0128 Saccharomyces ludwigii IFO 1043 Alcaligenes faecalis IAM B-141-1 Zn-25% KOH–EtOH Raney-10% NaOH

101a′

101b′

100 86 85 67 75 74 73 70 69 70 73 71

0 0 13 11 15 17 12 16 18 13 27 19

Source: Noma, Y., Ann. Res. Stud. Osaka Joshigakuen Junior College, 20, 33, 1976.

Consequently, the metabolic pathways of carvone by microorganisms were summarized as the following eight groups (Figure 22.105): Group 1: (−)-Carvone (93′)- (+)-dihydrocarvone (101a′)-(+)-neodihydrocarveol (102a′) Group 2: 93′–101a′-(−)-Dihydrocarveol (102b′) Group 3: 93′–101a′-102a′ and 102b′ Group 4: 93′-(+)-Isodihydrocarvone (101b′)–102c′ and 102d′ Group 5: (+)-Carvone (93)-(−)-isodihydrocarvone (101b)-(−)-neoisodihydrocarveol (102d) Group 6: 93–101b–102c Group 7: 93–101b–102c and 102d Group 8: 93–101b–101a

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Handbook of Essential Oils

The result of the mode action of both the hydrogenation of carvone and the reduction for dihydrocarvone by microorganism is as follows. In bacteria, only two strains were able to convert (−)-carvone (93′) via (+)-dihydrocarvone (101a′) to (−)-dihydrocarveol (102b′) as the major product (group 3, when ethanol was used as a carbon source, 12.5% of (−)-carvone (93′) convertible microorganisms belonged to this group, and when glucose was used, 8% belonged to this group) (Noma and Tatsumi, 1973; Noma et al., 1975), whereas when (+)-carvone (93) was converted, one strain converted it to a mixture of (−)-isodihydrocarveol (102c) and (−)-neoisodihydrocarveol (102d) (group 7%, 6% and 4% of 93 convertible bacteria belonged to this group, when ethanol and glucose were used, respectively), and four strains converted it via (−)-isodihydrocarvone (101b) to (−)-dihydrocarvone (101a) (group 8%, 6% and 15% of (+)-carvone (93′) convertible bacteria belonged to this group, when ethanol and glucose were used, respectively.) (Noma et al., 1975). In yeasts, 43% of carvone convertible yeasts belong to group 1%, 14% to group 2%, and 33% to group 3 (of this group, three strains are close to group 1) and 12% to group 5%, 4% to group 6%, and 27% to group 7 (of this group, three strains are close to group 5 and one strain is close to group 6). In fungi, 51% of fungi metabolized (−)-­carvone (93′) by way of group 1% and 3% via group 3, but there was no strain capable of metabolizing (−)-carvone (93′) via group 2, whereas 20% of fungi metabolized (+)-carvone (93) via group 5% and 29% via group 7, but there was no strain capable of metabolizing (+)-carvone (93) via group 6. In actinomycetes, (−)-carvone (93′) was converted to dihydrocarveols via group 1 (49%), group 2 (0%), group 3 (9%), and group 4 (28%), whereas (+)-carvone (93) was converted to dihydrocarveols via group 5 (7%), group 6 (0%), group 7 (19%), and group 8 (0%). Furthermore, (+)-neodihydrocarveol (102a′) stereospecifically formed from (−)-carvone (93′) by A. niger TBUYN-2 was further biotransformed to mosquito repellent (1R,2S,4R)-(+)-p-menthane2,8-diol (50a′), (1R,2S,4R)-(+)-8-p-menthene-2,10-diol (107a′), and the mixture of (1R,2S,4R,8S/ R)-(+)-p-menthane-2,8,9-triols (104aa′ and 104ab′), while A. glauca ATCC 22752 gave 107a′ stereoselectively from 102a′ (Demirci et al., 2001) (Figure 22.106). On the other hand, (−)-carvone (93′) was biotransformed stereoselectively to (+)-neodihydrocarveol (102a′) via (+)-dihydrocarvone (101a′) by a strain of A. niger (Noma and Nonomura, 1974), E. gracilis Z. (Noma et al., 1993), and Chlorella miniata (Gondai et al., 1999). Furthermore, in E. gracilis Z., mosquito repellent (1R,2S,4R)-(+)-p-menthane-2,8-diol (50a′) was obtained stereospecifically from (−)-carvone (93′) via 101a′ and 102a′ (Figure 22.107). As the microbial method for the formation of mosquito repellent 50a′ was established, the production of (+)-dihydrocarvone (101a′) and (+)-neodihydrocarveol (102a′) as the precursor of

O

O

OH

OH

OH

+

S 93΄

101a΄

102a΄

OH OH

104aa΄

R

OH

104ab΄

OH

OH

OH

OH

107a΄

HO

50a΄

FIGURE 22.106  Metabolic pathways of (−)-carvone (93′) by Aspergillus niger TBUYN-2 and Absidia glauca ATCC 22752. (Modified from Demirci, F. et al. The biotransformation of thymol methyl ether by different fungi, Book of Abstracts of the XII Biotechnology Congr., p. 47, 2001.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

O

O

A.n.

OH E.g.

A.n.

681

OH

E.g. C.p.

E.g. C.p.

OH

93΄

101a΄

102a΄

50a΄

FIGURE 22.107  Metabolic pathway of (−)-carvone (93′) by Aspergillus niger, Euglena gracilis Z, and Chlorella miniata. (Modified from Noma, Y. and Nonomura, S., Agric. Biol. Chem., 38, 741, 1974; Noma, Y. et al. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII, Proceedings of 37th TEAC, 1993, pp. 23–25; Gondai, T. et al. Asymmetric reduction of enone compounds by Chlorella miniata, Proceedings of 43rd TEAC, 1999, pp. 217–219.)

mosquito repellent 50a′ was investigated by using 40 strains of bacteria belonging to Escherichia, Aerobacter, Serratia, Proteus, Alcaligenes, Bacillus, Agrobacterium, Micrococcus, Staphylococcus, Corynebacterium, Sarcina, Arthrobacter, Brevibacterium, Pseudomonas, and Xanthomonas spp.; 68 strains of yeasts belonging to Schizosaccharomyces, Endomycopsis, Saccharomyces, Schwanniomyces, Debaryomyces, Pichia, Hansenula, Lipomyces, Torulopsis, Saccharomycodes, Cryptococcus, Kloeckera, Trigonopsis, Rhodotorula, Candida, and Trichosporon spp.; 40 strains of fungi belonging to Mucor, Absidia, Penicillium, Rhizopus, Aspergillus, Monascus, Fusarium, Pullularia, Keratinomyces, Oospora, Neurospora, Ustilago, Sporotrichum, Trichoderma, Gliocladium, and Phytophthora spp.; and 48 strains of actinomycetes belonging to Streptomyces, Actinoplanes, Nocardia, Micromonospora, Microbispora, Micropolyspora, Amorphosporangium, Thermopolyspora, Planomonospora, and Streptosporangium spp. As a result, 65% of bacteria, 75% of yeasts, 90% of fungi, and 90% of actinomycetes converted (−)-carvone (93′) to (+)-dihydrocarvone (101a′) or (+)-neodihydrocarveol (102a′) (Figures 22.105 and 22.108). Many microorganisms are capable of converting (−)-carvone (93′) to (+)-neodihydrocarveol (102a′) stereospecifically. Some of the useful microorganisms are listed in TABLE 22.8 Summary of Microbial and Chemical Reduction of (−)-Carvone (93′) for the Formation of (+)-Neodihydrocarveol (102a′) Compounds Microorganisms Torulopsis xylinus IFO 454 Monascus anka var. rubellus IFO 5965 Fusarium anguioides Sherbakoff IFO 4467 Phytophthora infestans IFO 4872 Kloeckera magna IFO 0868 Kloeckera antillarum IFO 0669 Streptomyces rimosus Penicillium notatum Westling IFO 464 Candida pseudotropicalis IFO 0882 Candida parapsilosis IFO 0585 LiAlH4 Meerwein–Ponndorf–Verley reduction

101a′ 0 0 0 0 0 19 + 6 17 16 0 0

101b′

102a′

102b′

102c′

102d′

0 0 0 0 0 4 0

100 100 100 100 98 72 98

0 0 0 0 2 0 0

0 0 0 0 0 0 0

0 0 0 0 0 0 0

2 4 4 0 0

92 79 80 17 29

0 0 0 67 55

0 0 0 2 9

0 0 0 13 5

Source: Noma, Y., Ann. Res. Stud. Osaka Joshigakuen Junior College, 20, 33, 1976.

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OH

O 102d OH 101b O 102c OH 93 O 102b OH 101a

102a

FIGURE 22.108  Metabolic pathways of (+)-carvone (93) by Pseudomonas ovalis strain 6-1 and other many microorganisms. (Modified from Noma, Y. et al. Agric. Biol. Chem., 38, 735, 1974b.)

Tables 22.7 and 22.8. There is no good chemical method to obtain (+)-­neodihydrocarveol (102a′) in large quantity. It was considered that the method utilizing microorganisms is a very useful means and better than the chemical synthesis for the production of mosquito repellent precursor (+)-­neodihydrocarveol (102a′). (−)-Carvone (93) was biotransformed by A. niger TBUYN-2 to give mainly (+)-8-hydroxyneodihydrocarveol (50a′), (+)-8,9-epoxyneodihydrocarveol (103a′), and (+)-10-­hydroxyneodihydrocarveol (107a′) via (+)-dihydrocarvone (101a′) and (+)-neodihydrocarveol (102a′). A. niger TBUYN-2 dehydrogenated (+)-cis-carveol (81b) to give (+)-carvone (93), which was further converted to (−)-isodihydrocarvone (101b). Compound 101b was further metabolized by four pathways to give 10-hydroxy-(−)-isodihydrocarvone (106b), (1S,2S,4S)-p-menthane-1,2diol (71d) via 1α-hydroxy-(−)-isodihydrocarvone (72b) as intermediate, (−)-isodihydrocarveol (102c), and (−)-neoisodihydrocarveol (102d). Compound 102d was further converted to isodihydroisobottrospicatol (105bb) via 8,9-epoxy-(−)-neoisodihydrocarveol (103d); compound 105′ was a major product (Noma et al., 1985a) (Figure 22.109). In the case of the plant pathogenic fungus, A. glauca (−)-carvone (93′) was metabolized to give the diol 10-hydroxy-(+)-neodihydrocarveol (107a′) (Nishimura et al., 1983b). (+)-Carvone (93) was converted by five bacteria and one fungus (Verstegen-Haaksma et al., 1995) to give (−)-dihydrocarvone (101a), (−)-isodihydrocarvone (101b), and (−)-neoisodihydrocarveol (102d). Sensitivity of the microorganism to (+)-carvone (93) and some of the products prevented yields exceeding 0.35 g/L in batch cultures. The fungus Trichoderma pseudokoningii gave the highest yield of (−)-neoisodihydrocarveol (102d) (Figure 22.110). (+)-Carvone (93) is known to inhibit fungal growth of Fusarium sulphureum when it was administered via the gas phase (Oosterhaven et al., 1995a,b). Under the same conditions, the related fungus Fusarium solani var. coeruleum was not inhibited. In liquid medium, both fungi were found to convert (+)-carvone

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Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

O

O

OH

OH

OH

O 93΄

101a΄

102a΄

OH 103a΄

50a΄

OH

OH

OH

OH

HO

107a΄

104a΄

OH O

71d

OH 106b OH

OH

OH O

72b

O

OH

O 102c OH

OH 81b

93

101b O 102d

HO 103d

O 105bb

FIGURE 22.109  Possible main metabolic pathways of (−)-carvone (93′) and (+)-carvone (93) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et  al. Biotransformation of (−)-carvone and (+)-carvone by Aspergillus spp., Annual Meeting of Agricultural and Biological Chemistry, Sapporo, Japan, 1985a, p. 68.) O

93

OH

O

101b

102d

FIGURE 22.110  Biotransformation of (+)-carvone (93) by Trichoderma pseudokoningii. (Modified from Verstegen-Haaksma, A.A. et al. Ind. Crops Prod., 4, 15, 1995.)

(93), with the same rate, mainly to (−)-isodihydrocarvone (101b), (−)-isodihydrocarveol (102c), and (−)-­neoisodihydrocarveol (102d). 22.3.4.1.1.1   Biotransformation of Carvone to Carveols by Actinomycetes  The distribution of actinomycetes capable of reducing carbonyl group of carvone containing α, β-unsaturated ketone to (−)-trans- (81a′) and (−)-cis-carveol (81b′) was investigated. Of 93 strains of actinomycetes, 63 strains were capable of converting (−)-carvone (93′) to carveols. The percentage of microorganisms that

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produced carveols from (−)-carvone (93′) to total microorganisms was about 71%. Microorganisms that produced carveols were classified into three groups according to the formation of (−)-transcarveol (81a′) and (−)-cis-carveol (81b′): group 1, (−)-carvone-81b′ only; group 2, (−)-carvone-81a′ only; and group 3, (−)-carvone mixture of 81a′ and 81b′. Three strains belonged to group 1 (4.5%), 34 strains belonged to group 2 (51.1%), and 29 strains belonged to group 3 (44%; of this group two strains were close to group 1 and 14 strains were close to group 2). Streptomyces A-5-1 isolated from soil converted (−)-carvone (93′) to 101a′–102d′ and (−)-­transcarveol (81a′), whereas Nocardia, 1-3-11 converted (−)-carvone (93′) to (−)-cis-carveol (81b′) together with 101a′–81a′ (Noma, 1980). In the case of Nocardia, the reaction between 93′ and 81a′ was reversible and the direction from 81a′ to 93′ is predominantly (Noma, 1979a,b, 1980) (Figure 22.111). (−)-Carvone (93′) was metabolized by actinomycetes to give (−)-trans- (81a′) and (−)-ciscarveol (81b′) and (+)-dihydrocarvone (101a′) as reduced metabolites. Compound 81b′ was further metabolized to (+)-bottrospicatol (92a′). Furthermore, 93′ was hydroxylated at C-5 position and C-8, 9 position to give 5β-hydroxy-(−)-carvone (98a′) and (−)-carvone-8,9-epoxide (96′), respectively. Compound 98a′ was further metabolized to 5β-hydroxyneodihydrocarveol (100aa′) via 5β-hydroxydihydrocarvone (99a′) (Noma, 1979a,b, 1980) (Figure 22.111). Metabolic pattern of (+)-carvone (93) is similar to that of (−)-carvone (93′) in S. bottropensis. (+)-Carvone (93) was converted by S. bottropensis to give (+)-carvone-8,9-epoxide (96) and (+)-5α-hydroxycarvone (98a) (Figure 22.112). (+)-Carvone-8,9-epoxide (96) has light sweet aroma and has strong inhibitory activity for the germination of lettuce seeds (Noma and Nishimura, 1982). OH

OH

O 81a΄

O 92a΄

OH

OH

100aa΄

O

O

O

93΄ HO

HO

HO

101a΄

O

81b΄

O 99a΄

98a΄

96΄

FIGURE 22.111  Metabolic pathways of (−)-carvone (93′) by Streptomyces bottropensis SY-2-, Streptomyces ikutamanensis Ya-2-1, Streptomyces A-5-1, and Nocardia 1-3-11. (Modified from Noma, Y., Nippon Nogeikagaku Kaishi, 53, 35, 1979a; Noma, Y., Ann. Res. Stud. Osaka Joshigakuen Junior College, 23, 27, 1979b; Noma, Y., Agric. Biol. Chem., 44, 807, 1980; Noma, Y. and Nishimura, H., Biotransformation of (−)-carvone and (+)-carvone by S. ikutamanensis Ya-2-1, Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, p. 390, 1983a; Noma, Y. and Nishimura, H., Biotransformation of carvone. 5. Microbiological transformation of dihydrocarvones and dihydrocarveols. Proceedings of 27th TEAC, 1983b, pp. 302–305.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

685

O

OH

O HO 92b

O

81b

101b

O

O 93 HO

O 98a

96

FIGURE 22.112  Metabolic pathways of (+)-carvone (93′) by Streptomyces bottropensis SY-2-1 and Streptomyces ikutamanensis Ya-2-1. (Modified from Noma, Y. and Nishimura, H., Biotransformation of carvone. 4. Biotransformation of (+)-carvone by Streptomyces bottropensis, SY-2-1, Proceedings of 26th TEAC, 1982, pp. 156–159l; Noma, Y. and Nishimura, H., Biotransformation of (−)-carvone and (+)-carvone by S. ikutamanensis Ya-2-1, Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, 1983a, p. 390; Noma, Y. and Nishimura, H., Biotransformation of carvone. 5. Microbiological transformation of dihydrocarvones and dihydrocarveols, Proceedings of 27th TEAC, 1983b, pp. 302–305; Noma, Y., Kagaku to Seibutsu, 22, 742, 1984.)

The investigation of (−)-carvone (93′) and (+)-carvone (93) conversion pattern was carried out by using rare actinomycetes. The conversion pattern was classified as follows (Figure 22.113): Group 1: Carvone (93), dihydrocarvones (101), dihydrocarveol (102), dihydrocarveol-8,9epoxide (103), dihydrobottrospicatols (105), 5-hydroxydihydrocarveols (100) Group 2: Carvone (93), carveols (89), bottrospicatols (92), 5-hydroxy-cis-carveols (12) Group 3: Carvone (93), 5-hydroxycarvone (98), 5-hydroxyneodihydrocarveols (15) Group 4: Carvone (93), carvone-8,9-epoxides (96) Of 50 rare actinomycetes, 22 strains (44%) were capable of converting (−)-carvone (93′) to give (−)-carvone-8,9-epoxide (96′) via pathway 4 and (+)-5β-hydroxycarvone (98a′), (+)-5α-hydroxycarvone (98b′), and (+)-5β-hydroxyneodihydrocarveol (100aa′) via pathway 3 (Noma and Sakai, 1984). On the other hand, in the case of (+)-carvone (93) conversion, 44% of rare actinomycetes were capable of converting (+)-carvone (93) to give (+)-carvone-8,9-epoxide (96) via pathway 4 and (−)-5α-hydroxycarvone (98a), (−)-5β-hydroxycarvone (98b), and (−)-5α-hydroxyneodihydrocarveol (100aa) via pathway 3 (Noma and Sakai, 1984). 22.3.4.1.1.2  Biotransformation of Carvone by Citrus Pathogenic Fungi, A. niger Tiegh TBUYN  Citrus pathogenic A. niger Tiegh (CBAYN) and A. niger TBUYN-2 hydrogenated C = C double bond at C-1, 2 position of (+)-carvone (93) to give (−)-isodihydrocarvone (101b) as the major product together with a small amount of (−)-dihydrocarvone (101a), of which 101b was further metabolized through two kinds of pathways as follows: One is the pathway to give (+)-1α-hydroxyn eoisodihydrocarveol (71) via (+)-1α-hydroxyisodihydrocarvone (72) and the other one is the pathway to give (+)-4α-hydroxyisodihydrocarvone (378) (Noma and Asakawa, 2008) (Figure 22.114).

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Handbook of Essential Oils OH

OH O

O 102a

103a OH

101a O OH HO 105aa

O 102b OH

93

O 105ab

OH O

O 102c

103c OH

101b O

O

OH HO 102d

105ba

105bb OH

OH

O

O 102a΄

103a΄ OH

101a΄ O

O

OH HO

105ab΄

105aa΄

102b΄

O

OH

OH 93΄ O

O 102c΄

103c΄ OH

101b΄

O 102d'

105ba΄

OH

O HO

105bb΄

FIGURE 22.113  Metabolic pathways of (+)- (93) and (−)-carvone (93′) and dihydrocarveols (102a-d and 102a′-d′) by Streptomyces bottropensis SY-2-1 and Streptomyces ikutamanensis Ya-2-1. (Modified from Noma, Y., Kagaku to Seibutsu, 22, 742, 1984.)

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Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

O

93

OH

O

OH O

101b

72

OH

71

O

O

OH 101a

378

FIGURE 22.114  Metabolic pathways of (+)-carvone (93) by citrus pathogenic fungi, Aspergillus niger Tiegh CBAYN and A. niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of (+)-carvone by Citrus pathogenic Aspergillus niger Tiegh CBAYN and A. niger TBUYN-2, Proceedings of 52nd TEAC, 2008, pp. 206–208.) H From si face 2

H

re

re

H

O

si

H O

H

H OH

2H

2H

H

H

OH

2

H

H From re face

OH

93΄

101a΄

102a΄

50a΄

FIGURE 22.115  The stereospecific hydrogenation of the C = C double bond of α,β-unsaturated ketones, the reduction of saturated ketone, and the hydroxylation by Euglena gracilis Z. (Modified from Noma, Y. et  al. Biotransformation of [6-2H]-(−)-carvone by Aspergillus niger, Euglena gracilis Z and Dunaliella tertiolecta, Proceedings of 39th TEAC, 1995, pp. 367–368; Noma, Y. and Asakawa, Y., Euglena gracilis Z: Biotransformation of terpenoids and related compounds, in Bajaj, Y.P.S., (ed.), Biotechnology in Agriculture and Forestry, vol. 41, Medicinal and Aromatic Plants X, Springer, Berlin, Germany, 1998, pp. 194–237.)

The biotransformation of enones such as (−)-carvone (93′) by the cultured cells of C. miniata was examined. It was found that the cells reduced stereoselectively the enones from si face at α-position of the carbonyl group and then the carbonyl group from re face (Figure 22.115). Stereospecific hydrogenation occurs independent of the configuration and the kinds of the substituent at C-4 position, so that the methyl group at C-1 position is fixed mainly at R-configuration. [2-2H]-(−)-Carvone ([2-2H]-93′) was synthesized in order to clear up the hydrogenation mechanism at C-2 by microorganisms. Compound [2-2H]-93 was also easily biotransformed to [2-2H]-8-hydroxy(+)-neodihydrocarveol (50a′) via [2-2H]-(+)-neodihydrocarveol (102a′). On the basis of 1H-NMR spectral data of compounds 102a′ and 50a′, the hydrogen addition of the carbon–carbon double bond at the C1 and C2 position by A. niger TBUYN-2, E. gracilis Z., and D. tertiolecta occurs from the si face and re face, respectively, that is, anti addition (Noma et al., 1995) (Figure 22.115) (Table 22.9). 22.3.4.1.1.3   Hydrogenation Mechanisms of C = C Double Bond and Carbonyl Group  In order to understand the mechanism of the hydrogenation of α-, β unsaturated ketone of (−)-carvone (93′) and the reduction of carbonyl group of dihydrocarvone (101a′) (−)-carvone (93′), (+)-dihydrocarvone (101a′) and the analogues of (−)-carvone (93′) were chosen and the conversion of the analogues

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Handbook of Essential Oils

O

O

O

O

O O

O 93΄

93

47΄

O

96΄

47

O

O

OH

OH

44΄

96

44

O O

O

HO 98a΄

379

380

O

O

148

148

O

O

381

119

FIGURE 22.116  Substrates used for the hydrogenation of C = C double bond with Pseudomonas ovalis strain 6-1, Streptomyces bottropensis SY-2-1, Streptomyces ikutamanensis Ya-2-1, and Euglena gracilis Z.

was carried out by using P. ovalis strain 6-1. As the analogues of carvone (93 and 93′), (−)- (47′) and (+)-carvotanacetone (47), 2-methyl-2-cyclohexenone (379), the mixture of (−)-cis- (81b′) and (−)-trans-carveol (81a′), 2-cyclohexenone, racemic menthenone (148), (−)-piperitone (156), (+)-pulegone (119), and 3-methyl-2-cyclohexenone (381) were chosen. Of these analogues, (−)- (47′) and (+)-carvotanacetone (47) were reduced to give (+)-carvomenthone (48a′) and (−)-isocarvomenthone (48b′), respectively. 2-Methyl-2-cyclohexenone (379) was mainly reduced to (−)-2-methylcyclohexanone. But other compounds were not reduced. The efficient formation of (+)-dihydrocarvone (101a), (−)-isodihydrocarvone (101b′), (+)-carvomenthone (48a), (−)-isocarvomenthone (48b′), and (−)-2-methylcyclohexanone from (−)-carvone (93), (+)-carvone (93′), (−)-carvotanacetone (47), (+)-carvotanacetone (47′), and 2-methyl-2-cyclohexenone (379) suggested at least that C = C double bond conjugated with carbonyl group may be hydrogenated from behind (si plane) (Noma et al., 1974b; Noma, 1977) (Figure 22.116). 22.3.4.1.1.4   What Is Hydrogen Donor in the Hydrogenation of Carvone to Dihydrocarvone? What Is Hydrogen Donor in Carvone Reductase?  Carvone reductase prepared from E. gracilis Z., which catalyzes the NADH-dependent reduction of the C = C bond adjacent to the carbonyl group, was characterized with regard to the stereochemistry of the hydrogen transfer into the substrate. The reductase was isolated from E. gracilis Z. and was found to reduce stereospecifically the C = C double bond of carvone by anti addition of hydrogen from the si face at α-position to the carbonyl group and the re face at β-position (Tables 22.9 and 22.10). The hydrogen atoms participating in the TABLE 22.9 Summary for the Stereospecificity of the Reduction of the C = C Double Bond of [2-2H]-(−)-Carvone ([2-2H]-93) by Various Kinds of Microorganisms Stereochemistry at C-2H of Compounds Microorganisms

102a

Aspergillus niger TBUYN-2

β

Euglena gracilis Z

β

Dunaliella tertiolecta

β

The cultured cells of Nicotiana tabacum (Suga et al., 1986)

β

50a β

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Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

TABLE 22.10 Purification of the Reductase from Euglena gracilis Z. Total Activity Unit  ×  104

Total Protein (mg) Crude extract DEAE Toyopearl AF-Blue Toyopearl

125 7 0.1

Sp. Act. Units per Gram Protein

Fold

1.7 21 30

1 12 18

2.2 1.5 0.03

NADH Hs

N-R

HR

CONH2 re

O H O H

si

O

Anti-addition

101a΄

93΄

FIGURE 22.117  Stereochemistry in the reduction of (−)-carvone (93′) by the reductase from Euglena gracilis Z. (Modified from Shimoda, K. et al. Phytochemistry, 49, 49, 1998.)

enzymatic reduction at α- and β-position to the carbonyl group originate from the medium and the pro-4R hydrogen of NADH, respectively (Shimoda et al., 1998) (Figure 22.117). In the case of biotransformation by using cyanobacterium, (+)- (93) and (−)-carvone (93′) were converted with a different type of pattern to give (+)-isodihydrocarvone (101b′, 76.6%) and (−)-dihydrocarvone (101a, 62.2%), respectively (Kaji et al., 2002) (Figure 22.118). On the other hand, Catharanthus rosea–cultured cell biotransformed (−)-carvone (93′) to give 5β-hydroxy(+)-neodihydrocarveol (100aa′, 57.5%), 5α-hydroxy-(+)-neodihydrocarveol (100ab′, 18.4%), 5α-hydroxy-(−)-carvone (98b′), 4β-hydroxy-(−)-carvone (384′, 6.3%), 10-hydroxycarvone (390′), 5β-hydroxycarvone (98′), 5β-hydroxyneodihydrocarveol (100ab′), 5β-hydroxyneodihydrocarveol (100aa′), and 5α-hydroxydihydrocarvone (99b′) as the metabolites as shown in Figure 22.119, whereas (+)-carvone (93) gave 5α-hydroxy-(+)-carvone (98a, 65.4%) and 4α-hydroxy-(+)-carvone (384, 34.6%) (Hamada and Yasumune, 1995; Hamada et al., 1996; Kaji et al., 2002) (Figure 22.119) (Table 22.11). (−)-Carvone (93′) was incubated with cyanobacterium, enone reductase (43 kDa) isolated from the bacterium, and microsomal enzyme to afford (+)-isodihydrocarvone (101b′) and (+)-dihydrocarvone (101a′). Cyclohexenone derivatives (379 are 385) were treated in the same enone reductase with

O

O

O Cyanobacterium

93

O Cyanobacterium

101a

93΄

101b΄

FIGURE 22.118  Biotransformation of (−)- and (+)-carvone (93 and 93′) by cyanobacterium. (Modified from Kaji, M. et al. Glycosylation of monoterpenes by plant suspension cells, Proceedings of 46th TEAC, 2002, pp. 323–325.)

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O

O

O +

Catharanthus roseus 93΄

OH + HO

HO OH 384΄

98b΄

+ HO 100aa΄

O

O +

OH

100ab΄ O

+ HO

HO

OH 390΄ O

O

99ab΄ O

+

Catharanthus roseus 93

98a΄

OH 384

HO 98a

FIGURE 22.119  Biotransformation of (+)- and (−)-carvone (93 and 93′) by Catharanthus roseus. (Modified from Hamada, H. and Yasumune, H., The hydroxylation of monoterpenoids by plant cell biotransformation, Proceedings of 39th TEAC, pp. 375–377, 1995; Hamada, H. et al. The hydroxylation and glycosylation by plant catalysts, Proceedings of 40th TEAC, 1996, pp. 111–112; Kaji, M. et al. Glycosylation of monoterpenes by plant suspension cells, Proceedings of 46th TEAC, 2002, pp. 323–325.)

TABLE 22.11 Enantioselectivity in the Reduction of Enones (379 and 385) by Enone Reductase Microsomal Enzyme

Substrate

Product

ee

Configurationa

–

379

382a

>99

R

–

385

386a

379

382b

>99 85

R

+ +

385

386b

80

S

a

S

Preferred configuration at α-position to the carbonyl group of the products.

microsomal enzyme to give the dihydro derivative (382a, 386a) with R-configuration in excellent ee (over 99%) and the metabolites (382b, 386b) with S-configuration in relatively high ee (85% and 80%) (Shimoda et al., 2003) (Figure 22.120). In contrast, almost all the yeasts tested showed reduction of carvone, although the enzyme activity varied. The reduction of (−)-carvone (93′) was often much faster than the reduction of (+)-­carvone (93). Some yeasts only reduced the carbon–carbon double bond to yield the dihydrocarvone isomers (101a′ and b′ and 101a and b) with the stereochemistry at C-1 with R-configuration, while others also reduced the ketone to give the dihydrocarveols with the stereochemistry at C-2 always with S for

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

Enone O reductase

O

Enone reductase

691

O

Microsomal enzyme 93΄

101b΄ O

101a΄

O

382b

O

379

O

386b

382a

O

O

385

386a

FIGURE 22.120  Biotransformation of 2-methyl-2-cyclohexenone (379) and 2-ethyl-2-cyclohexenone (385) by enone reductase.

(−)-carvone (93′), but sometimes S and sometimes R for (+)-carvone (93). In the case of (−)-carvone (93′) yields increased up to 90% within 2 h (van Dyk et al., 1998). 22.3.4.1.2 Carvotanacetone O

O

47 S-(+)-

47΄ R-(–)

In the conversion of (+)- (47) and (−)-carvotanacetone (47′) by P. ovalis strain 6-1, (−)-­carvotanacetone (47′) is converted stereospecifically to (+)-carvomenthone (48a′) an the latter compound is further converted to (+)-neocarvomenthol (49a′) and (−)-carvomenthol (49b′) in small amounts, whereas (+)-carvotanacetone (47) is converted mainly to (−)-isocarvomenthone (48b) and (−)-­neoisocarvomenthol (49d), forming (−)-carvomenthone (48a) and (−)-isocarvomenthol (49c) in small amounts as shown in Figure 22.121 (Noma et al., 1974a). Biotransformation of (−)-carvotanacetone (47) and (+)-carvotanacetone (47′) by S. bottropensis SY-2-1 was carried out (Noma et al., 1985c). As shown in Figure 22.122, (+)-carvotanacetone (47) was converted by S. bottropensis SY-2-1 to give 5β-hydroxy-(+)-neoisocarvomenthol (139db), 5α-hydroxy-(+)-carvotanacetone (51a), 5β-hydroxy-(−)-carvomenthone (52ab), 8-hydroxy-(+)-carvotanacetone (44), and 8-hydroxy-(−)carvomenthone (45a), whereas (−)-carvotanacetone (47′) was converted to give 5β-hydroxy(−)carvotanacetone (51a′) and 8-hydroxy-(−)-carvotanacetone (44′). A. niger TBUYN-2 converted (−)-carvotanacetone (47′) to (+)-carvomenthone (48a′), (+)-­carvomenthone (49a′), diastereoisomeric p-menthane-2,9-diols [55aa′ (8R) and 55ab′ (8S) in the ratio of 3:1], and 8-hydroxy-(+)-neocarvomenthol (102a′). On the other hand, the same fungus converted (+)-carvotanacetone (47) to (−)-isocarvomenthone (48b), 1α-hydroxy-(+)-neoisocarvomenthol (54)

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Handbook of Essential Oils

O

47΄

O

OH

O

48a΄

47

49a΄

OH

O

48b

OH

49d OH

O

49b΄

48a

49c

FIGURE 22.121  Metabolic pathways of (−)-carvotanacetone (47′) and (+)-carvotanacetone (47) by Pseudomonas ovalis strain 6-1. (Modified from Noma, Y. et al. Agric. Biol. Chem., 38, 1637, 1974a.)

O HO

HO

HO

52bb

51b

HO

O

O

51a΄

HO

HO

47

O

139db

O

O

OH

O

O

51a

52ab

47΄ OH

O

O

OH

44

44΄

OH

45a

FIGURE 22.122  Proposed the metabolic pathways of (+)-carvotanacetone (47) and (−)-carvotanacetone (47′) by Streptomyces bottropensis SY-2-1. (Modified from Noma, Y. et al. Microbiological conversion of (−)-carvotanacetone and (+)-carvotanacetone by S. bottropensis SY-2-1, Proceedings of 29th TEAC, 1985c, pp. 238–240.)

via 1α-hydroxy-(+)-isocarvomenthone (53), and 8-hydroxy-(−)-isocarvomenthone (45b) as shown in Figure 22.123 (Noma et al., 1988a). 22.3.4.1.3 Piperitone

O 156 R-(–)

O 156΄ S-(+)

A large number of yeasts were screened for the biotransformation of (−)-piperitone (156). A relatively small number of yeasts gave hydroxylation products of (−)-piperitone (156). Products obtained from (−)-piperitone (156) were 7-hydroxypiperitone (161), cis-6-hydroxypiperitone (158b), trans-6-hydroxypiperitone (158a), and 2-isopropyl-5-methylhydroquinone (180). Yields for the

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals O

47΄

OH

O

48a΄

OH

3 1

49a΄

R

H

OH

55aa΄ OH

OH

55ab΄

102a΄

47

OH

OH O

O

48b

H

S

OH

O

693

53

OH

OH

54 O OH

45b

FIGURE 22.123  Proposed metabolic pathways of (−)-carvotanacetone (47) and (+)-carvotanacetone (47′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Microbiological conversion of (−)-carvotanacetone and (+)-carvotanacetone by a strain of Aspergillus niger, Proceedings of 32nd TEAC, 1988b, pp. 146–148.)

hydroxylation reactions varied between 8% and 60%, corresponding to the product concentrations of 0.04–0.3 g/L. Not one of the yeasts tested reduced (−)-piperitone (156) (van Dyk et al., 1998). During the initial screen with (−)-piperitone (156), only hydroxylation products were obtained. The hydroxylation products (161, 158a, and 158b) obtained with nonconventional yeasts from the genera Arxula, Candida, Yarrowia, and Trichosporon have recently been described (van Dyk et al., 1998) (Figure 22.124). OH HO

HO +

O

O 161

O 158a

156

OH 180

HO O 158b

FIGURE 22.124  Hydroxylation products of (R)-(−)-piperitone (156) by yeast. (Modified from van Dyk, M.S. et al. J. Mol. Catal. B: Enzym., 5, 149, 1998.)

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Handbook of Essential Oils

O

OH

149a

137a

OH H+ O

O

O 119

387

112

O

O

HO

388

OH–

OH 121

O

OH 389

FIGURE 22.125  Biotransformation of (+)-pulegone (119) by Aspergillus sp., Botrytis allii, and H. isolate (UOFS Y-0067). (Modified from Miyazawa, M. et al. Chem. Express, 6, 479, 1991a; Miyazawa, M. et al. Chem. Express, 6, 873, 1991b; Ismaili-Alaoui, M. et al. Tetrahedron Lett., 33, 2349, 1992; van Dyk, M.S. et al. J. Mol. Catal. B: Enzym., 5, 149, 1998.)

22.3.4.1.4 Pulegone (R)-(+)-Pulegone (119), with a mint-like odor monoterpene ketone, is the main component (up to 80%–90%) of Mentha pulegium essential oil (pennyroyal oil), which is sometimes used in beverages and food additive for human consumption and occasionally in herbal medicine as an abortifacient drug. The biotransformation of (+)-pulegone (119) by fungi was investigated (Ismaili-Alaoui et al., 1992). Most fungal strains grown in a usual liquid culture medium were able to metabolize (+)-pulegone (119) to some extent in a concentration range of 0.1–0.5 g/L; higher concentrations were generally toxic, except for a strain of Aspergillus sp. isolated from mint leaves infusion, which was able to survive to concentrations of up to 1.5 g/L. The predominant product was generally l-hydroxy-(+)-pulegone (384) (20%–30% yield). Other metabolites were present in lower amounts (5% or less) (see Figure 22.125). The formation of 1-hydroxy-(+)-pulegone (387) was explained by hydroxylation at a tertiary position. Its dehydration to piperitenone (112), even under the incubation conditions, during isolation or derivative reactions precluded any tentative determination of its optical purity and absolute configuration. Botrytis allii converted (+)-pulegone (119) to (−)-(1R)-8-hydroxy-4-p-menthen-3-one (121) and piperitenone (112) (Miyazawa et al., 1991b,c). Hormonema isolate (UOFS Y-0067) quantitatively reduced (+)-pulegone (119) and (−)-menthone (149a) to (+)-neomenthol (137a) (van Dyk et al., 1998) (Figure 22.125). Biotransformation by the recombinant reductase and the transformed Escherichia coli cells were examined with pulegone, carvone, and verbenone as substrates (Figure 22.126). The recombinant reductase catalyzed the hydrogenation of the exocyclic C = C double bond of pulegone (119) to give menthone derivatives (Watanabe et al., 2007) (Tables 22.12 and 22.13).

695

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

O O

O

O

O 24΄

24 119΄

119

O

O

149a

93΄

149b

O

93

O

O

149a΄

149b΄

FIGURE 22.126  Chemical structures of substrate reduced by the recombinant pulegone reductase and the transformed Escherichia coli cells.

TABLE 22.12 Substrate Specificity in the Reduction of Enones with the Recombinant Pulegone Reductase Entry No. (Reaction Time) 1 (3 h)

Substrates (R)-(+)-Pulegone (119) (R)-Pulegone (119) (S)-(–)-Pulegone (119ʹ) (S)-Pulegone (119ʹ) (R)-(–)-Carvone (93ʹ)

2 (12 h) 3 (3 h) 4 (12 h) 5 (12 h) 6 (12 h)

(S)-(+)-Carvone (93) (1S, 5S)-Verbenone (24) (1R, 5R)-Verbenone (24ʹ)

7 (12 h) 8 (12 h)

Products

Conversions (%)

(1R, 4R)-Isomenthone (149b)

4.4

(1S, 4R)-Menthone (149a) (1R, 4R)-Isomenthone (149b) (1S, 4R)-Menthone (149a) (1S, 4S)-Isomenthone (149bʹ) (1R, 4S)-Menthone (149aʹ)

6.8 14.3 15.7 0.3 0.5

(1S, 4S)-Isomenthone (149bʹ) (1R, 4S)-Menthone (149aʹ) — — — —

1.6 2.1 N.d. N.d. N.d. N.d.

Note: N.d., denotes not detected.

TABLE 22.13 Biotransformation of Pulegone (119 and 119′) with the Transformed Escherichia coli Cellsa Substrates (R)-(+)-Pulegone (119) (S)-(–)-Pulegone (119ʹ)

a

Products

Conversion (%)

(1R, 4R)-Isomenthone (149b)

26.8

(1S, 4R)-Menthone (149a) (1S, 4S)-Isomenthone (149bʹ) (1R, 4S)-Menthone (149aʹ)

30.0 32.3 7.1

Reaction times of the transformation reaction are 12 h.

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Handbook of Essential Oils

22.3.4.1.5  Piperitenone and Isopiperitenone Piperitenone (112) is metabolized to 5-hydroxypiperitenone (117), 7-hydroxypiperitenone (118), and 7,8-dihydroxypiperitenone (157). Isopiperitenol (110) is reduced to give isopiperitenone (111), which is further metabolized to piperitenone (112), 7-hydroxy- (113), 10-hydroxy- (115), 4-hydroxy(114), and 5-hydroxyisopiperitenone (116). Compounds 111 and 112 are isomerized to each other. Pulegone (119) was metabolized to 112, 8,9-dehydromenthenone (120) and 8-hydroxymenthenone (121) as shown in the biotransformation of the same substrate using B. allii (Miyazawa et al., 1991c) (Figure 22.127). H. isolate (UOFS Y-0067) reduced (4S)-isopiperitenone (111) to (3R,4S)-isopiperitenol (110), a precursor of (−)-menthol (137b) (van Dyk et al., 1998) (Figure 22.128). OH

OH

O

O

HO

O

OH 113

157

117

OH

O

OH 110

111

O

HO

112

118

O

HO

O

O

O

O 120

O

O OH

OH 116

114

115

119

121

FIGURE 22.127  Biotransformation of isopiperitenone (111) and piperitenone (112) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Biotransformation of isopiperitenone, 6-gingerol, 6-shogaol and neomenthol by a strain of Aspergillus niger, Proceedings of 37th TEAC, 1992c, pp. 26–28.)

O

111

OH

110a

OH

137b

FIGURE 22.128  Biotransformation of isopiperitenone (111) by H. isolate (UOFS Y-0067). (Modified from van Dyk, M.S. et al. J. Mol. Catal. B: Enzym., 5, 149, 1998.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

697

22.3.4.2  Saturated Ketone 22.3.4.2.1 Dihydrocarvone

101b΄ (1S,4S) (+)-Iso

101a΄ (1R,4S) (+)

O

O

O

O

101a (1S,4R) (–)

101b (1R,4R) (–)-Iso

In the reduction of saturated carbonyl group of dihydrocarvone by microorganism, (+)-­dihydrocarvone (101a′) is converted stereospecifically to either (+)-neodihydrocarveol (102a′) or (−)-­dihydrocarveol (102b′) or nonstereospecifically to the mixture of 102a′ and 102b′, whereas (−)-isodihydrocarvone (101b) is converted stereospecifically to either(−)-neoisodihydrocarveol (102d) or (−)-­isodihydrocarveol (102c) or nonstereospecifically to the mixture of 102c and 102d by various microorganisms (Noma and Tatsumi, 1973; Noma et al., 1974c; Noma and Nonomura, 1974; Noma, 1976, 1977). (+)-Dihydrocarvone (101a′) and (+)-isodihydrocarvone (101b′) are easily isomerized chemically to each other. In the microbial transformation of (−)-carvone (93′), the formation of (+)-dihydrocarvone (101a′) is predominant. (+)-Dihydrocarvone (101a′) was reduced to both/either (+)-neodihydrocarveol (102a′) and/or (−)-dihydrocarveol (102b), whereas in the biotransformation of (+)-carvone (93), (+)-isodihydrocarvone (101b) was formed predominantly. (+)-Isodihydrocarvone (101b) was reduced to both (+)-isodihydrocarveol (102c) and (+)-neoisodihydrocarveol (102d) (Figure 22.129). However, P. fragi IFO 3458, Pseudomonas fluorescens IFO 3081, and Aerobacter aerogenes IFO 3319 and IFO 12059 formed (−)-dihydrocarvone (101a) predominantly from (+)-carvone (93). OH O

102d OH

101b O Epimerization

102c OH

93 O

102b OH

101a

102a

FIGURE 22.129  Proposed metabolic pathways of (+)-carvone (93) and (−)-isodihydrocarvone (101b) by Pseudomonas fragi IFO 3458. (Modified from Noma, Y. et al. Agric. Biol. Chem., 39, 437, 1975.)

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Handbook of Essential Oils

In the time course study of the biotransformation of (+)-carvone (93), it appeared that predominant formation of (−)-dihydrocarvone is due to the epimerization of (−)-isodihydrocarvone (101b′) by epimerase of P. fragi IFO 3458 (Noma et al., 1975). 22.3.4.2.2  Isodihydrocarvone Epimerase 22.3.4.2.2.1   Preparation of Isodihydrocarvone Epimerase  The cells of P. fragi IFO 3458 were harvested by centrifugation and washed five times with 1/100 M KH2PO4 –Na2HPO4 buffer (pH 7.2). Bacterial extracts were prepared from the washed cells (20 g from 3 L medium) by sonic lysis (Kaijo Denki Co., Ltd., 20Kc., 15 min, at 5°C–7°C) in 100 mL of the same buffer. Sonic extracts were centrifuged at 25, 500 g for 30 min at −2°C. The opalescent yellow supernatant fluid had the ability to convert (−)-isodihydrocarvone (101b) to (−)-dihydrocarvone (101a). On the other hand, the broken cell preparation was incapable of converting (−)-isodihydrocarvone (101b) to (−)-­dihydrocarvone (101a). The enzyme was partially purified from this supernatant fluid about 56-fold with heat treatment (95°C–97°C for 10 min), ammonium sulfate precipitation (0.4–0.7 saturation), and DEAE-Sephadex A-50 column chromatography. The reaction mixture consisted of a mixture of (−)-isodihydrocarvone (101b) and (−)-­dihydrocarvone (101a) (60:40 or 90:10), 1/30 M KH2PO4 –Na2HPO4 buffer (pH 7.2), and the crude or partially purified enzyme solution. The reaction was started by the addition of the enzyme solution and stopped by the addition of ether. The ether extract was applied to analytical gas–liquid chromatography (GLC) (Shimadzu GC-4A 10% PEG-20M, 3 m × 3 mm, temperature 140°C–170°C at the rate of 1°C/min, N2 35 mL/min), and epimerization was assayed by measuring the peak areas of (−)-­isodihydrocarvone (101b) and (−)-dihydrocarvone (101a) in GLC before and after the reaction. The crude extract and the partially purified preparation were found to be very stable to heat treatment; 66% and 36% of the epimerase activity remained after treatment at 97°C for 60 and 120 min, respectively (Noma et al., 1975). A strain of A. niger TBUYN-2 hydroxylated at C-1 position of (−)-isodihydrocarvone (101b) to give 1α-hydroxyisodihydrocarvone (72b), which was easily and smoothly reduced to (1S, 2S, 4S)-(−)-8-p-menthene-1,2-trans-diol (71d), which was also obtained from the biotransformation of (−)-cis-limonene-1,2-epoxide (69) by microorganisms and decomposition by 20% HCl (Figure 22.127) (Noma et al., 1985a,b). Furthermore, A. niger TBUYN-2 and A. niger Tiegh (CBAYN) biotransformed (−)-isodihydrocarvone (101b) to give (−)-4α-hydroxyisodihydrocarvone (378b) and (−)-8-p-menthene-1,2-trans-diol (71d) as the major products together with a small amount of 1α-hydroxyisodihydrocarvone (72b) (Noma and Asakawa, 2008) (Figure 22.130). 22.3.4.2.3  Menthone and Isomenthone

O

149a (1R,4S) (–)-Menthone

O

O

149b (1S,4S) (–)-Isomenthone

149a΄ (1S,4R) (+)-Menthone

O

149b΄ (1R,4R) (+)-Isomenthone

The growing cells of P. fragi IFO 3458 epimerized 17% of racemic isomenthone (149b and b′) to menthone (149a and a′) (Noma et  al., 1975). (−)-Menthone (149a) was converted by P. fluorescens M-2 to (−)-3-oxo-4-isopropyl-1-cyclohexanecarboxylic acid (164a), (+)-3-oxo4-isopropyl-1-­ c yclohexanecarboxylic acid (164b), and (+)-3-hydroxy-4-isopropyl-1cyclohexanecarboxylic acid (165ab). On the other hand, (+)-menthone (149a′) was converted to

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH O

O

O

OH

699

O

OH

OH

378b

101b

72b

71d

69b

FIGURE 22.130  Biotransformation of (+)-carvone (93), (−)-isodihydrocarvone (101b), and (−)-cis-limonene1,2-epoxide (69b) by Aspergillus niger TBUYN-2 and A. niger Tiegh (CBAYN). (Modified from Noma, Y. et al. Biotransformation of (−)-carvone and (+)-carvone by Aspergillus spp., Annual Meeting of Agricultural and Biological Chemistry, Sapporo, Japan, 1985a, p. 68; Noma, Y. and Asakawa, Y., New metabolic pathways of (+)-carvone by Citrus pathogenic Aspergillus niger Tiegh CBAYN and A. niger TBUYN-2, Proceedings of 52nd TEAC, 2008, pp. 206–208.)

give (+)-3-oxo-4-isopropyl-1-cyclohexane carboxylic acid (164a′) and (−)-3-oxo-4-isopropyl1-cyclohexane carboxylic acid (164b′). Racemic isomenthone (149b and b′) was converted to give racemic 1-hydroxy-1-methyl-4-isopropylcyclohexane-3-one (150), racemic piperitone (156), racemic 3-oxo-4-isopropyl-1-­cyclohexene-1-carboxylic acid (162), 3-oxo-4-isopropyl-1-cyclohexane carboxylic acid (164b), 3-oxo-4-isopropyl-1-cyclohexane carboxylic acid (164a), and (+)-3-hydroxy4-isopropyl-1-­cyclohexane carboxylic acid (165ab) (Figure 22.131). Soil plant pathogenic fungi Rhizoctonia solani 189 converted (−)-menthone (149a) to 4β-hydroxy(−)menthone (392%, 29%) and 1 α, 4 β-dihydroxy-(−)-menthone (393%, 71%) (Nonoyama et al., 1999) (Figure 22.131). (−)-Menthone (149a) was transformed by S. litura to give 7-hydroxymenthone (151a), 7-hydroxyneomenthol (165c), and 7-hydroxy-9-carboxymenthone (394a) (Hagiwara et al., 2006) (Figure 22.132). (−)-Menthone (149a) gave 7-hydroxymenthone (151a) and (+)-­neomenthol (137c) by human liver microsome (CYP2B6). Of 11 recombinant human P450 enzymes (expressed in Trichoplusia ni cells) tested, CYP2B6 catalyzed oxidation of (−)-menthone (149a) to 7-hydroxymenthone (151a) (Nakanishi and Miyazawa, 2004; Miyazawa et al., 1992b,c) (Figure 22.132). 22.3.4.2.4 Thujone O

28a΄ O

4

O

5

4

O

4

1

28a 1S,4S,5R (+)-3-

28a΄ 1R,4R,5S (–)-3-

4

5

1

1

28b

28b΄ O

5

1

O

5

28b 1S,4R,5R (–)-3-iso

O

28b΄ 1R,4S,5S (+)-3-iso

β-Pinene (1) is metabolized to 3-thujone (28) via α-pinene (4) (Gibbon and Pirt, 1971). α-Pinene (4) is metabolized to give thujone (28). Thujone (28) was biotransformed to thujoyl alcohol (29) by A. niger TBUYN-2 (Noma, 2000). Furthermore, (−)-3-isothujone (28b) prepared from Armoise oil was biotransformed by plant pathogenic fungus B. allii IFO 9430 to give 4-hydroxythujone (30) and 4,6-dihydroxythujone (31) (Miyazawa et al., 1992a) (Figure 22.133).

700

Handbook of Essential Oils COOH

OH

COOH 165ab O

O

COOH

164a

149a

O 164b

CH2OH

O

O 151

149a΄

COOH

COOH

164a΄

164b΄ COOH

HO

149a and a΄

150

156

COOH

162

COOH

O 164b

O

O

O O

O

O

COOH

O 164a

OH 165

FIGURE 22.131  Biotransformation of (−)- (149a) and (+)-menthone (149a′) and racemic isomenthone (149b and 149b′) by Pseudomonas fluorescens M-2. (Modified from Sawamura, Y. et al. Microbiological oxidation of p-menthane 1. Formation of formation of p-cis-menthan-1-ol, Proceedings of 18th TEAC, 1974, pp. 27–29.)

22.3.4.3  Cyclic Monoterpene Epoxide 22.3.4.3.1 1,8-Cineole 1,8-Cineole (122) is a main component of the essential oil of Eucalyptus radiata var. australiana leaves, comprising ca. 75% in the oil, which corresponds to 31 mg/g fr.wt. leaves (Nishimura et al., 1980). The most effective utilization of 122 is very important in terms of renewable biomass production. It would be of interest, for example, to produce more valuable substances, such as plant growth regulators, by the microbial transformation of 122. The first reported utilization of 122 was presented by MacRae et al. (1979), who showed that it was a carbon source for Pseudomonas flava growing on Eucalyptus leaves. Growth of the bacterium in a mineral salt medium containing 122 resulted in the oxidation at the C-2 position of 122 to give the metabolites (1S,4R,6S)-(+)-2α-hydroxy-1,8cineole (225a), (1S,4R,6R)-(−)-2β-hydroxy-1,8-cineole (125a), (1S,4R)-(+)-2-oxo-1,8-cineole (126), and (−)-(R)-5,5-dimethyl-4-(3′-oxobutyl)-4,5-dihydrofuran-2(3H)-one (128) (Figure 22.134).

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

701

OH +

R. solani

OH

O

O OH

392 OH

393 OH

S. litura

+

+

O

O 149a

O

OH

151a OH

394a COOH

165c

+

CYP2B6

O

OH

151a

137c

FIGURE 22.132  Metabolic pathway of (−)-menthone (149a) by Rhizoctonia solani 189, Spodoptera litura, and human liver microsome (CYP2B6). (Modified from Nonoyama, H. et al. Biotransformation of (−)-menthone using plant parasitic fungi, Rhizoctonia solani as a biocatalyst, Proceedings of 43rd TEAC, 1999, pp. 387–388; Nakanishi, K. and Miyazawa, M., Biotransformation of (−)-menthone by human liver microsomes, Proceedings of 48th TEAC, 2004, pp. 401–402; Hagiwara, Y. et al. Biotransformation of (+)and (−)-menthone by the larvae of common cutworm (Spodoptera litura) as a biocatalyst, Proceedings of 50th TEAC, 2006, pp. 279–280.)

O

OH A.n.

28

29

OH O

OH

O

B. allii

O +

HO 28b

30b

31b

FIGURE 22.133  Biotransformation of (−)-3-isothujone (28b) by Aspergillus niger TBUYN-2 and plant pathogenic fungus Botrytis allii IFO 9430. (Modified from Gibbon, G.H. and Pirt, S.J., FEBS Lett., 18, 103, 1971; Miyazawa, M. et al. Biotransformation of thujone by plant pathogenic microorganism, Botrytis allii IFO 9430, Proceedings of 36th TEAC, 1992a, pp. 197–198.)

S. bottropensis SY-2-1 biotransformed 1,8-cineole (122) stereochemically to (+)-2α-hydroxy-1,8cineole (125b) as the major product and (+)-3α-hydroxy-1,8-cineole (123b) as the minor product. Recovery ratio of 1,8-cineole metabolites as ether extract was ca. 30% in S. bottropensis SY-2-1 (Noma and Nishimura, 1980, 1981) (Figure 22.135).

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Handbook of Essential Oils

HO

HO

O O

P. flava

+

+

O

O

+ O

O

H

O O

122

125a

125b

126

128

FIGURE 22.134  Biotransformation of 1,8-cineole (84) by Pseudomonas flava. (Modified from MacRae, I.C. et al. Aust. J. Chem., 32, 917, 1979.)

HO S. bottropensis

+ HO

O 122

O

O

125b Major

123b Minor

S. ikutamanensis + HO

O 123b 46%

HO

O 123a 29%

FIGURE 22.135  Biotransformation of 1,8-cineole (122) by Streptomyces bottropensis SY-2-1 and Streptomyces ikutamanensis Ya-2-1. (Modified from Noma, Y. and Nishimura, H., Microbiological transformation of 1,8-cineole. Oxidative products from 1,8-cineole by S. bottropensis, SY-2-1, Book of abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, 1980, p. 28; Noma, Y. and Nishimura, H., Microbiological transformation of 1,8-cineole. Production of 3β-hydroxy-1,8-cineole from 1,8-cineole by S. ikutamanensis, Ya-2-1, Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, 1981, p. 196.)

In the case of S. ikutamanensis Ya-2-1, 1,8-cineole (122) was biotransformed regioselectively to give (+)-3α-hydroxy-1,8-cineole (123b, 46%) and (+)-3β-hydroxy-1,8-cineole (123b, 29%) as the major product. Recovery ratio as ether extract was ca. 8.5% in S. ikutamanensis,Ya-2-1 (Noma and Nishimura, 1980, 1981) (Figure 22.135). When (+)-3α-hydroxy-1,8-cineole (123b) was used as substrate in the cultured medium of S. ikutamanensis Ya-2-1, (+)-3β-hydroxy-1,8-cineole (123a, 32%) was formed as the major product together with a small amount of (+)-3-oxo-1,8-cineole (126a, 1.6%). When (+)-3β-hydroxy-1,8 cineole (123a) was used, (+)-3-oxo-1,8-cineole (126a, 9.6%) and (+)-3α-hydroxy-1,8-cineole (123b, 2%) were formed. When (+)-3-oxo-1,8-cineole (126a) was used, (+)-3α-hydroxy- (123b, 19%) and (+)-3β-hydroxy-1,8-cineole (123a, 16%) were formed. Based on the aforementioned results, it is obvious that (+)-3β-hydroxy-1,8-cineole (123b) is formed mainly in the biotransformation of 1,8-cineole (122), (+)-3α-hydroxy-1,8-cineole (123b), and (+)-3-oxo-1,8-cineole (126a) by S. ikutamanensis Ya-2-1. The production of (+)-3β-hydroxy1,8-cineole (123b) is interesting, because it is a precursor of mosquito repellent, p-menthane-3,8-diol (142aa′) (Noma and Nishimura, 1981) (Figure 22.136).

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

HO

46%

O

123b O

O

703

O

29%

126a

122 HO

O

123a

HO HO

142aa΄

FIGURE 22.136  Biotransformation of 1,8-cineole (122), (+)-3α-hydroxy-1,8-cineole (123b), (+)-3β-hydroxy1,8-cineole(123a), and (+)-3-oxo-1,8-cineole (126a) by Streptomyces ikutamanensis Ya-2-1. (Modified from Noma, Y. and Nishimura, H., Microbiological transformation of 1,8-cineole. Production of 3β-hydroxy-1,8cineole from 1,8-cineole by S. ikutamanensis, Ya-2-1, Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, 1981, p. 196.)

When A. niger TBUYN-2 was cultured in the presence of 1,8-cineole (122) for 7 days, it was transformed to three alcohols [racemic 2α-hydroxy-1,8-cineoles (125b and b′), racemic 3α-hydroxy(123b and b′), and racemic 3β-hydroxy-1,8-cineoles (123a and 123a′)] and two ketones [racemic 2-oxo- (126 and 126′) and racemic 3-oxo-1,8-cineoles (124 and 124′)] (Figure 22.135). The formation of 3α-hydroxy- (123b and b′) and 3β-hydroxy-1,8-cineoles (123a and 123a′) is of great interest not only due to the possibility of the formation of p-menthane-3,8-diol (142 and 142′), the mosquito repellents, and plant growth regulators that are synthesized chemically from 3α-hydroxy- (123b and b′) and 3β-hydroxy-1,8-cineoles (123a and 123a′), respectively, but also from the viewpoint of the utilization of E. radiata var. australiana leaves oil as biomass. An Et2O extract of the culture broth (products and 122 as substrate) was recovered in 57% of substrate (w/w) (Nishimura et al., 1982; Noma et al., 1996) (Figure 22.137). Plant pathogenic fungus Botryosphaeria dothidea converted 1,8-cineole (122) to optical pure (+)-2α-hydroxy-1,8-cineole (125b) and racemic 3α-hydroxy-1,8-cineole (123b and b′), which were oxidized to optically active 2-oxo- (126) (100% ee) and racemic 3-oxo-1,8-cineole (124 and 124′), respectively (Table 22.14). The cytochrome P450 inhibitor 1-aminobenzotriazole inhibited the hydroxylation of the substrate (Noma et al., 1996) (Figure 22.138). S. litura also converted 1,8-­cineole (122) to give three secondary alcohols (123b, 125a, and b) and two primary alcohols (395 and 127) (Hagiwara and Miyazawa, 2007). Salmonella typhimurium OY1001/3A4 and NADPH-P450 reductase hydroxylated 1,8-cineole (122) to 2β-hydroxy-1,8-cineole (125a, [α]D + 9.3, 65.3% ee) and 3β-hydroxy-1 to 8-cineole (123a, [α]D−27.8, 24.7% ee) (Saito and Miyazawa, 2006). Extraction of the urinary metabolites from brushtail possums (Trichosurus vulpecula) maintained on a diet of fruit impregnated with 1,8-cineole (122) yielded p-cresol (129) and the novel C-9 oxidated products 9-hydroxy-1,8-cineole (127a) and 1,8-cineole-9-oic acid (462a) (Flynn and Southwell, 1979; Southwell and Flynn, 1980) (Figure 22.139).

704

Handbook of Essential Oils

OH

HO O

O O

O

O 125a΄

125a

HO

OH

126

O 126΄

O 125b

O O

125b΄

122

HO

O 123a

O 124

OH

123a΄

O

O

124΄

HO

O 123b

OH

123b΄

FIGURE 22.137  Biotransformation of 1,8-cineole (122) by Aspergillus niger TBUYN-2. (Modified from Nishimura, H. et al. Agric. Biol. Chem., 46, 2601, 1982; Noma, Y. et al. Biotransformation of 1,8-cineole. Why do the biotransformed 2α- and 3α-hydroxy-1,8-cineole by Aspergillus niger have no optical activity? Proceedings of 40th TEAC, 1996, pp. 89–91.)

TABLE 22.14 Stereoselectivity in the Biotransformation of 1,8-Cineole (122) by Aspergillus niger, Botryosphaeria dothidea, and Pseudomonas flava Products Microorganisms Aspergillus niger TBUYN-2 Botryosphaeria dothidea

125a and a′, 125b and b′, 123b and b′, 123a and a′ 2:43:49:6 50:50:41:59 4:59:34:3 100:0:53:47

Pseudomonas flava 29:71:0:0 100:0 Source: Noma, Y. et al. Biotransformation of 1,8-cineole. Why do the biotransformed 2α- and 3α-hydroxy-1,8-cineole by Aspergillus niger have no optical activity? Proceedings of 40th TEAC, 1996, pp. 89–91.

1,8-Cineole (122) gave 2β-hydroxy-1,8-cineole (125a) by CYP450 human and rat liver microsome. Cytochrome P450 molecular species responsible for metabolism of 1,8-cineole (122) was determined to be CYP3A4 and CYP3A1/2 in human and rat, respectively. Kinetic analysis showed that K m and Vmax values for the oxidation of 1,8-cineole (122) by human and rat treated with pregnenolone-16αcarbonitrile recombinant CYP3A4 were determined to be 50 µM and 90.9 nmol/min/nmol P450, 20 µM and 11.5 nmol/min/nmol P450, and 90 µM and 47.6 nmol/min/nmol P450, respectively (Shindo et al., 2000).

705

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

HO + B. dothidea

+ OH HO

O

O

123b

123b΄

125b

OH

HO

HO

+ HO

+

S. litura O

O

O

125a

O

O

O

125b

+

+

123b

O HO

395

127b

122 HO

S. typhimurium +

HO

O

O

123a

125a

FIGURE 22.138  Biotransformation of 1,8-cineole (122) by Botryosphaeria dothidea, Spodoptera litura, and Salmonella typhimurium. (Modified from Noma, Y. et al. Biotransformation of 1,8-cineole. Why do the biotransformed 2α- and 3α-hydroxy-1,8-cineole by Aspergillus niger have no optical activity? Proceedings of 40th TEAC, 1996, pp. 89–91; Saito, H. and Miyazawa, M., Biotransformation of 1,8-cineole by Salmonella typhimurium OY1001/3A4, Proceedings of 50th TEAC, pp. 275–276, 2006; Hagiwara, Y. and Miyazawa, M., Biotransformation of cineole by the larvae of common cutworm (Spodoptera litura) as a biocatalyst, Proceedings of 51st TEAC, 2007, pp. 304–305.)

T. vulpecula O

+ O

+ O

OH

122

127a

462a

COOH

OH

129

FIGURE 22.139  Metabolism of 1,8-cineole in Trichosurus vulpecula. (Modified from Southwell, I.A. and Flynn, T.M., Xenobiotica, 10, 17, 1980.)

Microbial resolution of racemic 2α-hydroxy-1,8-cineoles (125b and b′) was carried out by using G. cingulata. The mixture of 125b and b′ was added to a culture of G. cingulata and esterified to give after 24 h (1R,2R,4S)-2α-hydroxy-1,8-cineole-2-yl-malonate (130b′) in 45% yield (ee 100%). The recovered alcohol showed 100% ee of the (1S,2S,4R)-enantiomer (125b) (Miyazawa et  al., 1995c). On the other hand, optically active (+)-2α-hydroxy-1,8-cineole (125b) was also formed from (+)-limonene (68) by a strain of citrus pathogenic fungus P. digitatum (Saito and Miyazawa, 2006; Noma and Asakawa, 2007a) (Figure 22.140). Esters of racemic 2α-hydroxy-1,8-cineole (125b and b′) were prepared by a convenient method (Figure 22.141). Their odors were characteristic. Then products were tested against antimicrobial activity and their microbial resolution was studied (Hashimoto and Miyazawa, 2001) (Table 22.15).

706

Handbook of Essential Oils

O

OH 68

122 A. niger

A. niger B. dothidea P. flava

HO

68 G. cyanea

O HO

OH

OH

+ O

O

O

130b΄

125b

125b΄

125b

O

O

G. cingulata O

P. digitatum

Hydrolysis 125b΄

(1S, 2S, 4R) [α]D + 29.6

FIGURE 22.140  Formation of 2α-hydroxy-1,8-cineoles (125b and b′) from 1,8-cineole (122) and optical resolution by Glomerella cingulata and Aspergillus niger TBUYN-2 and 125b′ from (+)-limonene (68) by Penicillium digitatum. (Modified from Nishimura, H. et al. Agric. Biol. Chem., 46, 2601, 1982; Abraham, W.-R. et al. Appl. Microbiol. Biotechnol., 24, 24, 1986; Miyazawa, M. et al. Biotransformation of 2-endo-hydroxy-1,4cineole by plant pathogenic microorganism, Glomerella cingulata, Proceedings of 39th TEAC, 1995b, pp. 352– 353; Noma, Y. et al. Reduction of terpene aldehydes and epoxidation of terpene alcohols by S. ikutamanensis, Ya-2-1, Proceedings of 30th TEAC, 1986, pp. 204–206; Noma, Y. and Asakawa, Y., Biotransformation of limonene and related compounds by newly isolated low temperature grown citrus pathogenic fungi and red yeast, Book of Abstracts of the 38th ISEO, 2007a, p. 7.)

HO

OH

MCPBA in dichloromethane

O

125b΄

125b

Acid chloride, pyridine

OH

HO 34a΄

34a

O R

O

O O

O O

O

R=CH3(396) R

C2H5(397) C3H7(398) C4H9(399) CH(CH3)2(400) C(CH3)3(401)

125b΄s Ester

CH2CH(CH3)2(402) 125b Ester

CH2C(CH3)3(403)

FIGURE 22.141  Chemical synthesis of esters of racemic 2α-hydroxy-1,8-cineole (125b and b′). (Modified from Hashimoto Y. and Miyazawa, M., Microbial resolution of esters of racemic 2-endo-hydroxy-1,8-cineole by Glomerella cingulata, Proceedings of 45th TEAC, 2001, pp. 363–365.)

707

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

TABLE 22.15 Yield and Enantiomer Excess of Esters of Racemic 2α-Hydroxy-1,8-Cineole (125b and b′) on the Microbial Resolution by Glomerella cingulata 0 h

24 h

Compounds

% ee

% ee

Yield (%)

% ee

Yield (%)

396

(−)36.3

(+)85.0

24.0

(+)100

14.1

397

(−)36.9

(+)73.8

18.6

(+)100

8.6

398

(−)35.6

(+)33.2

13.7

(+)75.4

3.5

(+)45.4

14.4

(+)100

2.3

(−)21.4

25.2

(+)20.6

8.0

399

(−)36.8

400

(−)35.4

48 h

401

(−)36.7

(−)37.8

31.5

(−)40.6

15.2

402

(−)36.1

(−)29.8

46.8

(−)15.0

24.0

403

(−)36.3

(−)37.6

72.2

(−)39.0

36.9

Source: Hashimoto, Y. and Miyazawa, M., Microbial resolution of esters of racemic 2-endo-hydroxy-1,8-cineole by Glomerella cingulata, Proceedings of 45th TEAC, 2001, pp. 363–365.

Glu-O

Glu-Glu-O

O-Glc

+

E. perriniana O

O 405

E. perriniana

404

O 122

O 404΄

FIGURE 22.142  Biotransformation of 1,8-cineole (122) by Eucalyptus perriniana suspension cell. (Modified from Hamada, H. et al. Glycosylation of monoterpenes by plant suspension cells, Proceedings of 46th TEAC, 2002, pp. 321–322.)

1,8-Cineole (122) was glucosylated by E. perriniana suspension cells to 2α-hydroxy-1,8 cineole monoglucoside (404, 16.0% and 404′, 16.0%) and diglucosides (405, 1.4%) (Hamada et al., 2002) (Figure 22.142). 22.3.4.3.2 1,4-Cineole Regarding the biotransformation of 1,4-cineole (131), S. griseus transformed it to 8-hydroxy-1,4cineole (134), whereas Bacillus cereus transformed 1,4-cineole (131) to 2α-hydroxy-1,4 ­cineole (132b, 3.8%) and 2β-hydroxy-1,4-cineoles (132a, 21.3%) (Liu et  al., 1988) (Figures 22.143 and 22.144). On the other hand, a strain of A. niger biotransformed 1,4-cineole (131) regiospecifically to 2α-hydroxy-1,4-cineole (132b) (Miyazawa et al., 1991e) and (+)-3α-hydroxy-1,4-cineole (133b) (Miyazawa et al., 1992d) along with the formation of 8-hydroxy-1,4-cineole (134) and 9-hydroxy-1,4 cineole (135) (Miyazawa et al., 1992e) (Figure 22.144). Microbial optical resolution of racemic 2α-hydroxy-1,4-cineoles (132b and b′) was carried out by using G. cingulata (Liu et al., 1988). The mixture of 2α-hydroxy-1,4-cineoles (132b and b′) was added to a culture of G. cingulata and esterified to give after 24 h (1R,2R,4S)-2α-hydroxy-1,4cineole-2-yl-malonate (136′) in 45% yield (ee 100%). The recovered alcohol showed an ee of 100% of the (1S,2S,4R)-enantiomer (132b). On the other hand, optically active (+)-2α-hydroxy-1,4-cineole

708

Handbook of Essential Oils HO

OH O

O

132a (1S, 2R, 4R)

132a΄ (1R, 2S, 4S)

HO

OH O

O O

132b [α]D +19.6 (1S, 2S, 4R)

132b΄ (1R, 2R, 4S) [α]D –19.6

131 O

HO

O OH

133a (1S, 3S, 4R)

133a΄ (1R, 3R, 4S)

O

O

HO

OH

133b (1S, 3R, 4R)

133b΄ (1R, 3S, 4S)

FIGURE 22.143  Metabolic pathways of 1,4-cineole (131) by microorganisms. HO O

O

OH

B. cereus

+ 132a

OH

O

A.n. S. griseus

132a΄

HO

406

O

OH

A.n. B. cereus

131

OH O

O

+

A.n. 132b

132b΄

OH

134

O

A.n. HO

OH

+

OH

135

O

O

133b

133b΄

FIGURE 22.144  Metabolic pathways of 1,4-cineole (131) by Aspergillus niger TBUYN-2, Bacillus cereus, and Streptomyces griseus. (Modified from Liu, W. et al. J. Org. Chem., 53, 5700, 1988; Miyazawa, M. et al. Chem. Express, 6, 771, 1991c; Miyazawa, M. et al. Chem. Express, 7, 305, 1992b; Miyazawa, M. et al. Chem. Express, 7, 125, 1992c; Miyazawa, M. et  al. Biotransformation of 2-endo-hydroxy-1,4-cineole by plant pathogenic microorganism, Glomerella cingulata, Proceedings of 39th TEAC, 1995b, pp. 352–353.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

709

O OH 342

131 A. niger

A. niger G. cyanea

A. niger

HO

OH O

O

O

HO G. cingulata

O

O +

O

OH

O 132b΄

132b

132b΄

132b (1S, 2S, 4R)

rac-132b and b΄ [α]D +0.13

136΄

(1R, 2R, 4S) [α]D –19.6

[α]D +19.6

FIGURE 22.145  Formation of optically active 2α-hydroxycineole from 1,4-cineole (131) and terpinen-4-ol (342) by Aspergillus niger TBUYN-2, Gibberella cyanea, and Glomerella cingulata. (Modified from Abraham, W.-R. et al. Appl. Microbiol. Biotechnol., 24, 24, 1986; Miyazawa, M. et al. Chem. Express, 6, 771, 1991c; Miyazawa, M. et al. Biotransformation of 2-endo-hydroxy-1,4-cineole by plant pathogenic microorganism, Glomerella cingulata, Proceedings of 39th TEAC, 1995b, pp. 352–353; Noma, Y. and Asakawa, Y., Microbial transformation of limonene and related compounds, Proceedings of 51st TEAC, 2007b, pp. 299–301.)

(132b) was also formed from (−)-terpinen-4-ol (342) by G. cyanea DSM (Abraham et al., 1986) and A. niger TBUYN-2 (Noma and Asakawa, 2007b) (Figure 22.145).

22.4  METABOLIC PATHWAYS OF BICYCLIC MONOTERPENOIDS 22.4.1 Bicyclic Monoterpene 22.4.1.1  α-Pinene

4 (+)-α-Pinene

4΄ (–)-α-Pinene

α-Pinene (4 and 4′) is the most abundant terpene in nature and obtained industrially by fractional distillation of turpentine (Krasnobajew, 1984). (+)-α-Pinene (4) occurs in oil of Pinus palustris Mill. at concentrations of up to 65% and in oil of Pinus caribaea at concentrations of 70% (Bauer et al., 1990). On the other hand, P. caribaea contains (−)-α-pinene (4′) at the concentration of 70%–80% (Bauer et al., 1990). The biotransformation of (+)-α-pinene (4) was investigated by A. niger NCIM 612 (Bhattacharyya et al., 1960; Prema and Bhattachayya, 1962). A 24 h shake culture of this strain metabolized 0.5% (+)-α-pinene (4) in 4–8 h. After the fermentation of the culture broth contained (+)-verbenone (24) (2%–3%), (+)-cisverbenol (23b) (20%–25%), (+)-trans-sobrerol (43a) (2%–3%), and (+)-8-hydroxycarvotanacetone (44) (Bhattacharyya et al., 1960; Prema and Bhattachayya, 1962) (Figure 22.146). The degradation of (+)-α-pinene (4) by a soil Pseudomonas sp. (PL strain) was investigated by Hungund et al. (1970). A terminal oxidation pattern was proposed, leading to the formation of organic

710

Handbook of Essential Oils OH

+ 4

23b

+

+

O

OH

O

OH

24

OH

43a

44

FIGURE 22.146  Biotransformation of (+)-α-pinene (4) by Aspergillus niger NCIM 612. (Modified from Bhattacharyya, P.K. et al. Nature, 187, 689, 1960; Prema, B.R. and Bhattachayya, P.K., Appl. Microbiol., 10, 524, 1962.) COOH

COOH

OH OH + 4

+

36

+

5

7 65

FIGURE 22.147  Biotransformation of (+)-α-pinene (4) by Pseudomonas sp. (PL strain). (Modified from Shukla, O.P., and Bhattacharyya, P.K., Indian J. Biochem., 5, 92, 1968.)

CHO O

4

38

CHO

39

40

FIGURE 22.148  Biotransformation of (+)-α-pinene (4) by Pseudomonas fluorescens NCIMB 11671. (Modified from Best, D.J. et al. Biocatal. Biotransform., 1, 147, 1987.)

acids through ring cleavage. (+)-α-Pinene (4) was fermented in shake cultures by a soil Pseudomonas sp. (PL strain) that is able to grow on (+)-α-pinene (4) as the sole carbon source, and borneol (36), myrtenol (5), myrtenic acid (84), and α-phellandric acid (65) (Shukla and Bhattacharyya, 1968) (Figure 22.147) were obtained. The degradation of (+)-α-pinene (4) by P. fluorescens NCIMB11671 was studied, and a pathway for the microbial breakdown of (+)-α-pinene (4) was proposed as shown in Figure 22.148 (Best et al., 1987; Best and Davis, 1988). The attack of oxygen is initiated by enzymatic oxygenation of the 1,2-double bond to form α-pinene epoxide (38), which then undergoes rapid rearrangement to produce a unsaturated aldehyde, occurring as two isomeric forms. The primary product of the reaction (Z)-2-methyl-5-isopropylhexa-2,5-dien-1-al (39, isonovalal) can undergo chemical isomerization to the E-form (novalal, 40). Isonovalal (39), the native form of the aldehyde, possesses citrus, woody, and spicy notes, whereas novalal (40) has woody, aldehydic, and cyclone notes. The same biotransformation was also carried out by Nocardia sp. strain P18.3 (Griffiths et al., 1987a,b). Pseudomonas PL strain and PIN 18 degraded α-pinene (4) by the pathway proposed in Figure 22.149 to give two hydrocarbon, limonene (68) and terpinolene (346), and neutral metabolite, borneol (36). A probable pathway has been proposed for the terminal oxidation of β-isopropylpimelic acid (248) in the PL strain and PIN 18 (Shukla and Bhattacharyya, 1968).

711

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH

COOH

5

4

7

+

HO

+

32 COOH

1

COOH COOH

36 (–)-from 1 (+)-from 4

(–)-from 1 (+)-from 4 OH

+ 33

86

68

COOH

COOH

61

346 +

OH

OH 463

COOH

OH

?

464

35

83 COOH

H–

OH COOH COOH

59

62

65

248 (–)-from 1 and 4

FIGURE 22.149  Metabolic pathways of degradation of α- and β-pinene by a soil Pseudomonad (PL strain) and Pseudomonas PIN 18. (Modified from Shukla, O.P. and Bhattacharyya, P.K., Indian J. Biochem., 5, 92, 1968.)

Pseudomonas PX 1 biotransformed (+)-α-pinene (4) to give (+)-cis-thujone (29) and (+)-transcarveol (81a) as major compounds. Compounds 81a, 171, 173, and 178 have been identified as fermentation products (Gibbon and Pirt, 1971; Gibbon et al., 1972) (Figure 22.150). A. niger TBUYN-2 biotransformed (−)-α-pinene (4′) to give (−)-α-terpineol (34′) and (−)-transsobrerol (43a′) (Noma et al., 2001). The mosquitocidal (+)-(1R,2S,4R)-1-p-menthane-2,8-diol (50a′) was also obtained as a crystal in the biotransformation of (−)-α-pinene (4′) by A. niger TBUYN-2 (Noma et al., 2001; Noma, 2007) (Figure 22.151). (1R)-(+)-α-Pinene (4) and its enantiomer (4′) were fed to S. litura to give the corresponding (+)- and (−)-verbenones (24 and 24′) and (+)- and (−)-myrtenols (5 and 5′) (Miyazawa et al., 1996c) (Figure 22.152). (−)-α-Pinene (4′) was treated in human liver microsomes CYP2B6 to afford (−)-trans-verbenol (23′) and (−)-myrtenol (5′) (Sugie and Miyazawa, 2003) (Figure 22.153). In rabbit, (+)-α-pinene (4) was metabolized to (−)-trans-verbenols (23) as the main metabolites together with myrtenol (5) and myrtenic acid (7). The purities of (−)-verbenol (23) from (−)- (4′), (+)- (4), and (+/−)-α-pinene (4 and 4′) were 99%, 67%, and 68%, respectively. This means that the biotransformation of (−)-4′ in rabbit is remarkably efficient in the preparation of (−)-trans-verbenol (23a) (Ishida et al., 1981b) (Figure 22.154).

712

Handbook of Essential Oils

O

4 29 OH

OH

O

OH

+ 81a

42

COOH

COOH

173

HO

O

171

56

O

COOH

174

O

COOH

172

COOH

47

46a

175

COOH

COOH

176

178

FIGURE 22.150  Proposed metabolic pathways for (+)-α-pinene (4) degradation by Pseudomonas PX 1. (Modified from Gibbon, G.H. and Pirt, S.J., FEBS Lett., 18, 103, 1971; Gibbon, G.H. et al. Degradation of α-pinene by bacteria, Proceedings of IV IFS: Fermentation Technology Today, 1972, pp. 609–612.) HO

HO

A. niger

A. niger

A. niger

OH

OH

4΄ 34΄

43a΄

OH 50a΄

FIGURE 22.151  Biotransformation of (−)-α-pinene (4) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Microbiological transformation of β-pinene, Proceedings of 45th TEAC, 2001, pp. 88–90.)

(−)-α-Pinene (4′) was biotransformed by the plant pathogenic fungus B. cinerea to afford 3α-hydroxy-(−)-β-pinene (2a′, 10%), 8-hydroxy-(−)-α-pinene (434′, 12%), 4β-hydroxy-(−)-pinene6-one (468′, 16%), and (−)-verbenone (24′) (Farooq et al., 2002) (Figure 22.155). 22.4.1.2  β-Pinene

1 (+)-β-Pinene

1΄ (–)-β-Pinene

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

713

OH S. litura + O

4

5

24

OH S. litura + O

4΄

5΄

24΄

FIGURE 22.152  Biotransformation of (+)- (4) and (−)-α-pinene (4′) by Spodoptera litura. (Modified from Miyazawa, M. et al. Biotransformation of pinanes by common cutworm larvae, Spodoptera litura as a biocatalyst, Proceedings of 40th TEAC, 1996c, pp. 84–85.) OH

P450 2B6

+

HO

4΄

5΄

23΄

FIGURE 22.153  Biotransformation of (−)-α-pinene (4′) by human liver microsomes CYP2B6. (Modified from Sugie, A. and Miyazawa, M., Biotransformation of (−)-α-pinene by human liver microsomes, Proceedings of 47th TEAC, 2003, pp. 159–161.) OH

Rabbit

4

+

23a

OH

COOH

+

5

7

Rabbit

4΄

HO

23a΄

FIGURE 22.154  Biotransformation of α-pinene by rabbit. (Modified from Ishida, T. et al. J. Pharm. Sci., 70, 406, 1981b.)

(+)-β-Pinene (1) is found in many essential oils. Optically active and racemic β-pinenes are present in turpentine oils, although in smaller quantities than (+)-α-pinene (4) (Bauer et al., 1990). Shukla et  al. (1968) obtained a similarly complex mixture of transformation products from (−)-β-pinene (1′) through degradation by a Pseudomonas sp/(PL strain). On the other hand, Bhattacharyya and Ganapathy (1965) indicated that A. niger NCIM 612 acts differently and more specifically on the pinenes by preferably oxidizing (−)-β-pinene (1′) in the allylic position to form the interesting products pinocarveol (2′) and pinocarvone (3′), besides myrtenol (5′) (see Figure 22.156). Furthermore, the conversion of (−)-β-pinene (1′) by Pseudomonas putida arvilla (PL strain) gave borneol (36′) (Rama Devi and Bhattacharyya, 1978) (Figure 22.156).

714

Handbook of Essential Oils

HO B. cinerea +

O

4΄

OH

+

2a΄

24΄

+

O

HO

434΄

468΄

FIGURE 22.155  Microbial transformation of (−)-α-pinene (4′) by Botrytis cinerea. (Modified from Farooq, A. et al. Z. Naturforsch., 57c, 686, 2002.) OH HO

O

A. niger NCIM 612

2΄

3΄

5΄

OH

P. pudidaarvilla PL strain

1΄

+

+

36΄

FIGURE 22.156  Biotransformation of (−)-β-pinene (1′) by Aspergillus niger NCIM 612 and Pseudomonas putida arvilla (PL strain). (Modified from Bhattacharyya, P.K. and Ganapathy, K., Indian J. Biochem., 2, 137, 1965; Rama Devi, J. and Bhattacharyya, P.K., J. Indian Chem. Soc., 55, 1131, 1978.)

P. pseudomallei isolated from local sewage sludge by the enrichment culture technique utilized caryophyllene as the sole carbon source (Dhavalikar et al., 1974). Fermentation of (−)-β-pinene (1′) by P. pseudomallei in a mineral salt medium (Seubert’s medium) at 30°C with agitation and aeration for 4 days yielded camphor (37′), borneol (36a′), isoborneol (36b′), α-terpineol (34′), and β-isopropyl pimelic acid (248′) (see Figure 22.154). Using modified Czapek Dox medium and keeping the other conditions the same, the pattern of the metabolic products was dramatically changed. The metabolites were trans-pinocarveol (2′), myrtenol (5′), α-fenchol (11′), á-terpineol (34′), myrtenic acid (7′), and two unidentified products (see Figure 22.157). COOH

P.p.

OH

OH

+

+

Seubert

36b΄

36a΄

37΄

OH 1΄

P.p.

COOH

+

O +

OH 34΄

248΄

COOH

HO

HO +

+

+

Czapek-dox

2΄

5΄

7΄

+

11b΄

OH 34΄

FIGURE 22.157  Biotransformation of (−)-β-pinene (1′) by Pseudomonas pseudomallei. (Modified from Dhavalikar, R.S. et al. Dragoco Rep., 3, 47, 1974.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

715

OH HO +

B.c. 1΄

408΄

+ 409΄

OH

+

HO

HO

410΄ OH

O 411΄

FIGURE 22.158  Biotransformation of (−)-β-pinene (1′) by Botrytis cinerea. (Modified from Farooq, A. et al. Z. Naturforsch., 57c, 686, 2002.) OH A.n.

A.n.

1

OH

OH

34

204 OH

A.n.

A.n.

1΄

OH

OH 34΄

204΄

FIGURE 22.159  Biotransformation of (+)- (1) and (−)-β-pinene (1′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Microbiological transformation of β-pinene, Proceedings of 45th TEAC, 2001, pp. 88–90.)

(−)-β-Pinene (1′) was converted by plant pathogenic fungi, B. cinerea, to give four new compounds such as (−)-pinane-2α,3α-diol (408′), (−)-6β-hydroxypinene (409′), (−)-4α,5-dihydroxypinene (410′), and (−)-4α-hydroxypinene-6-one (411′) (Figure 22.158). This study progressed further biotransformation of (−)-pinane-2α,3α-diol (408′) and related compounds by microorganisms as shown in Figure 22.158. As shown in Figure 22.159, (+)- (1) and (−)-β-pinenes (1′) were biotransformed by A. niger TBUYN-2 to give (+)-α-terpineol (34) and (+)-oleuropeyl alcohol (204) and their antipodes (34′ and 204′), respectively. The hydroxylation process of α-terpineol (34) to oleuropeyl alcohol (204) was completely inhibited by 1-aminotriazole as a cytochrome P450 inhibitor. (−)-β-Pinene (1′) was at first biotransformed by A. niger TBUYN-2 to give (+)-trans-pinocarveol (2a′) (274). (+)-trans-Pinocarveol (2a′) was further transformed by three pathways: First, (+)-transpinocarveol (2a′) was metabolized to (+)-pinocarvone (3′), (−)-3-isopinanone (413′), (+)-2α-hydroxy3-pinanone (414′), and (+)-2α,5-dihydroxy-3-pinanone (415′). Second, (+)-trans-pinocarveol (2a′) was metabolized to (+)-6β-hydroxyfenchol (349ba′), and third, (+)-trans-pinocarveol (2a′) was metabolized to (−)-6β,7-dihydroxyfenchol (412ba′) via epoxide and diol as intermediates (Noma and Asakawa, 2005a) (Figure 22.160). (–)-β-Pinene (1′) was metabolized by A. niger TBUYN-2 with three pathways as shown in Figure 22.154 to give (–)-α-pinene (4′), (–)-α-terpineol (34′), and (+)-trans-pinocarveol (2a′). (–)-α-Pinene (4′) is further metabolized by three pathways. At first, (–)-α-pinene (4′) was metabolized via (–)-α-pinene epoxide (38′), trans-sobrerol (43a′), (–)-8-hydroxycarvotanacetone (44′), and (+)-8-­hydroxycarvomenthone (45a) to (+)-p-menthane-2,8-diol (50a′), which was also metabolized

716

Handbook of Essential Oils OH HO

O

O

1΄

2a΄

O

3΄

413΄

414΄

OH 415΄

OH

H+ HO

OH O

O

OH

OH OH

OH

HO

OH

349ba΄

412ba΄

FIGURE 22.160  The metabolism of (−)-β-pinene (1′) and (+)-trans-pinocarveol (2a′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

in (–)-carvone (93′) metabolism. Second, (–)-α-pinene (4′) is metabolized to myrtenol (83′), which is metabolized by rearrangement reaction to give (–)-oleuropeyl alcohol (204′). (–)-α-Terpineol (34′), which is formed from β-pinene (1′), was also metabolized to (–)-oleuropeyl alcohol (204′), and (+)-trans-pinocarveol (2a′), formed from (–)-β-pinene (1′), was metabolized to pinocarvone (3′), 3-pinanone (413′), 2α-hydroxy-3-pinanone (414′), 2α,5-dihydroxy-3-pinanone (415′), and 2α,9-dihydroxy-3-pinanone (416′). Furthermore, (+)-trans-pinocarveol (2a′) was metabolized by rearrangement reaction to give 6β-hydroxyfenchol (349ba′) and 6β,7-dihydroxyfenchol (412ba′) (Noma and Asakawa, 2005a) (Figure 22.161). OH

HO

o

O

OH

OH

38΄

83΄

OH

4΄

204΄

OH

HO

417΄

102a΄

50a΄ OH O

HO

412ba΄

OH

45a΄

OH HO

HO

OH

44΄

43a΄

OH

HO

O

418΄

O

93΄ OH

HO

OH 34΄

1΄

HO

3΄

2b΄

HO

11b΄

O

O

OH

349ba΄

101a΄ HO

OH

O

413΄

OH 419΄

414΄ OH O

OH O OH

416΄

OH 415΄

FIGURE 22.161  Biotransformation of (−)-β-pinene (1′), (−)-α-pinene (4′), and related compounds by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

717

OH HO

HO

2b΄

1΄

HO

HO

11b΄

412ba΄

OH

13b΄

OH

13a΄

OH

O

349ba΄

O

OH

O

12΄

FIGURE 22.162  Relationship of the metabolism of (−)-β-pinene (1′), (+)-fenchol (11′), and (−)-fenchone (12′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

(–)-β-Pinene (1′) was metabolized by A. niger TBUYN-2 to give (+)-trans-pinocarveol (2a′), which was further metabolized to 6β-hydroxyfenchol (349ba′) and 6β, 7-dihydroxyfenchol (412ba′) by rearrangement reaction (Noma and Asakawa, 2005a) (Figure 22.162). 6β-Hydroxyfenchol (349ba′) was also obtained from (–)-fenchol (11b′). (–)-Fenchone was hydroxylated by the same fungus to give 6β- (13a′) and 6α-hydroxy-(–)-fenchone (13b′). There is a close relationship between the metabolism of (–)-β-pinene (1′) and those of (–)-fenchol (11′) and (–)-fenchone (12′). (–)-β-Pinene (1′) and (–)-α-pinene (4′) were isomerized to each other. Both are metabolized via (–)-α-terpineol (34’) to (–)-oleuropeyl alcohol (204′) and (–)-oleuropeic acid (61′). (–)-Myrtenol (5′) formed from (–)-α-pinene (1′) was further metabolized via cation to (–)-oleuropeyl alcohol (204′) and (–)-oleuropeic acid (61′). (–)-α-Pinene (4′) is further metabolized by A. niger TBUYN-2 via (–)-α-pinene epoxide (38′) to trans-sobrerol (43a′), (–)-8-hydroxycarvotanacetone (44′), (+)-8-hydroxycarvomenthone (45a), and mosquitocidal (+)-p-menthane-2,8-diol (50a′) (Bhattacharyya et al., 1960; Noma et al., 2001, 2002, 2003) (Figure 22.163). The major metabolites of (–)-β-pinene (1′) were trans-10-pinanol (myrtanol) (8ba′) (39%) and (–)-1-p-menthene-7,8-diol (oleuropeyl alcohol) (204′) (30%). In addition, (+)-trans-pinocarveol (2a′) (11%) and (–)-α-terpineol (34′) (5%) and verbenol (23a and 23b) and pinocarveol (2a′) were oxidation products of α-(4) and β-pinene (1), respectively, in bark beetle, Dendroctonus frontalis. (–)-cis- (23b′) and (+)-trans-verbenols (23a′) have pheromonal activity in Ips paraconfusus and Dendroctonus brevicomis, respectively (Ishida et al., 1981b) (Figure 22.164). 22.4.1.3 (±)-Camphene Racemate camphene (437 and 437′) is a bicyclic monoterpene hydrocarbon found in Liquidambar species, Chrysanthemum, Zingiber officinale, Rosmarinus officinalis, and among other plants. It was administered into rabbits. Six metabolites, camphene-2,10-glycols (438a, 438b), which were the major metabolites, together with 10-hydroxytricyclene (438c), 7-hydroxycamphene (438d), 6-exohydroxycamphene (438e), and 3-hydroxytricyclene (438f), were obtained (Ishida et al., 1979). On the basis of the production of the glycols (438a and 438b) in good yield, these alcohols might be formed

718

Handbook of Essential Oils OH HO

+

HO

OH OH–

+ OH

+ OH+

HOH HO

HO OH

4΄

OH

50a΄

H+

OH

OH

5΄

OH

43a΄

O

204΄

O

1΄

205΄

34΄

OH

OH

44΄

45a΄

OH

COOH

OH

OH

CHO

61΄

FIGURE 22.163  Metabolic pathways of (−)-β-pinene (1′) and related compounds by Aspergillus niger TBUYN-2. (Modified from Bhattacharyya, P.K. et al. Nature, 187, 689, 1960; Noma, Y. et al. Microbiological transformation of β-pinene, Proceedings of 45th TEAC, 2001, pp. 88–90; Noma, Y. et al. Stereoselective formation of (1R, 2S, 4R)-(+)-p-menthane-2,8-diol from α-pinene, Book of Abstracts of the 33rd ISEO, 2002, p. 142; Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, 2003, pp. 91–93.)

through their epoxides as shown in Figure 22.165. The homoallyl camphene oxidation products (438c–f) apparently were formed through the nonclassical cation as the intermediate. 22.4.1.4  3-Carene and Carane

1

6

6

439 1S,6R (+)-3-carene

1

1 6

439b 1S,3S,6R (–)-cis-carane

6

439a 1S,3S,6R (+)-trans-carane

1

439΄ 1R,6S (–)-3-carene

6

1

439a΄ 1R,3R,6S (–)-trans-carane

6

1

439b΄ 1R,3R,6S (+)-cis-carane

(+)-3-Carene (439) was biotransformed by rabbits to give m-mentha-4,6-dien-8-ol (440) (71.6%) as the main metabolite together with its aromatized m-cymen-8-ol (441). The position of C-5 in the substrate is thought to be more easily hydroxylated than C-2 by enzymatic systems in the rabbit liver. In addition to the ring opening compound, 3-carene-9-ol (442), 3-carene-9-carboxylic acid

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

719

HO

8ba΄

1΄

HO

OH

23b΄

4΄

HO OH 2a΄

HO

OH

34΄

23a΄

204΄

FIGURE 22.164  Metabolism of β-pinene (1) by bark beetle, Dendroctonus frontalis. (Modified from Ishida, T. et al. J. Pharm. Sci., 70, 406, 1981b.)

437 and 437΄ O

O OH

HO

HO

HO OH

OH

OH 438a

438e

438d

OH

438c

438f

438b

FIGURE 22.165  Biotransformation of (±)-camphene (437 and 437′) by rabbits. (Modified from Ishida, T. et al. J. Pharm. Sci., 68, 928, 1979.)

(443), 3-carene-9,10-dicarboxylic acid (445), chamic acid, and 3-caren-10-ol-9-carboxylic acid (444) were formed. The formation of such compounds is explained by stereoselective hydroxylation and carboxylation of gem-dimethyl group (Ishida et al., 1981b) (Figure 22.166). In the case of (–)-ciscarane (446), two C-9 and C-10 methyl groups were oxidized to give dicarboxylic acid (447) (Ishida et al., 1981b) (Figure 22.166). 3-(+)-Carene (439) was converted by A. niger NC 1M612 to give either hydroxylated compounds of 3-carene-2-one or 3-carene-5-one, which was not fully identified (Noma et al., 2002) (Figure 22.167).

22.4.2 Bicyclic Monoterpene Aldehyde 22.4.2.1  Myrtenal and Myrtanal CHO

CHO

CHO

CHO

6 (+)-

6΄ (–)-

435b (+)-cisBinihiol

435a (–)-trans-

Myrtenal

Myrtanal

COH

435b΄ (–)-cis-

CHO

435a΄ (+)-trans-

720

Handbook of Essential Oils

4

6 1 OH 1

OH

440

441

6 1

439

1

6

1

6

1

6

HO 9

OH

COOH 9 443

442

6

HOOC

COOH 9 444

COOH 9 445

FIGURE 22.166  Metabolic pathways of (+)-3-carene (439) by rabbit (Modified from Ishida, T. et al. J. Pharm. Sci., 70, 406, 1981b). 3-(+)-Carene (439) was converted by Aspergillus niger NC 1M612 to give either hydroxylated compounds of 3-carene-2-one or 3-carene-5-one, which was not fully identified (Figure 22.167). (Modified from Noma, Y. et al. Stereoselective formation of (1R, 2S, 4R)-(+)-p-menthane-2,8-diol from α-pinene, Book of Abstracts of the 33rd ISEO, 2002, p. 142.) O

1

1 6

OH

6

HO

or

1

6

O

439

FIGURE 22.167  Metabolic pathways of (+)-3-Carene (439) by Aspergillus niger NC 1M612. (Modified from Noma, Y. et al. Stereoselective formation of (1R, 2S, 4R)-(+)-p-menthane-2,8-diol from α-pinene, Book of Abstracts of the 33rd ISEO, 2002, p. 142.)

E. gracilis Z. biotransformed (–)-myrtenal (6′) to give (–)-myrtenol (5′) as the major product and (–)-myrtenoic acid (7′) as the minor product. However, further hydrogenation of (–)-myrtenol (5′) to trans- and cis-myrtanol (8a and 8b) did not occur even at a concentration less than ca. 50 mg/L. (S)-trans- and (R)-cis-myrtanal (435a′ and 435b′) were also transformed to trans- and cis-myrtanol (8a′ and 8b′) as the major products and (S)-trans- and (R)-cis-myrtanic acid (436a′ and 436b′) as the minor products, respectively (Noma et al., 1991b) (Figure 22.168). In the case of A. niger TBUYN-2, Aspergillus sojae, and Aspergillus usami, (–)-myrtenol (5′) was further metabolized to 7-hydroxyverbenone (25′) as a minor product together with (–)-oleuropeyl alcohol (204′) as a major product (279, 280). (–)-Oleuropeyl alcohol (204′) is also formed from (–)-α-terpineol (34) by A. niger TBUYN-2 (Noma et al., 2001) (Figure 22.168). Rabbits metabolized myrtenal (6′) to myrtenic acid (7′) as the major metabolite and myrtanol (8a′ or 8b′) as the minor metabolite (Ishida et al., 1981b) (Figure 22.168).

22.4.3 Bicyclic Monoterpene Alcohol 22.4.3.1 Myrtenol OH

5

OH

5΄

721

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH

COOH

CHO

OH

CHO

8b΄

435b΄ OH

7΄

6΄

CHO

COOH

OH 8a΄

34΄

436b΄

5΄ OH

OH

COOH

435a΄

436a΄

O

OH

25΄

204΄

FIGURE 22.168  Biotransformation of (−)-myrtenal (6′) and (+)-trans- (435a′) and (−)-cis-myrtanal (435b′) by microorganisms. (Modified from Noma, Y. et al. Phytochemistry, 30, 1147, 1991a; Noma, Y. and Asakawa, Y., Microbial transformation of (−)-myrtenol and (−)-nopol, Proceedings of 49th TEAC, 2005b, pp. 78–80; Noma, Y. and Asakawa, Y., Biotransformation of β-pinene, myrtenol, nopol and nopol benzyl ether by Aspergillus niger TBUYN-2, Book of Abstracts of the 37th ISEO, 2006b, p. 144.)

(–)-Myrtenol (5′) was biotransformed mainly to (–)-oleuropeyl alcohol (204′), which was formed from (–)-α-terpineol (34′) as a major product by A. niger TBUYN-2. In the case of A. sojae IFO 4389 and A. usami IFO 4338, (–)-myrtenol (5′) was metabolized to 7-hydroxyverbenone (25′) as a minor product together with (–)-oleuropeyl alcohol (204′) as a major product (Noma and Asakawa, 2005b) (Figure 22.169). 22.4.3.2 Myrtanol S. litura converted (–)-trans-myrtanol (8a) and its enantiomer (8a′) to give the corresponding myrtanic acid (436 and 436′) (Miyazawa et al., 1997b) (Figure 22.170). 22.4.3.3 Pinocarveol OH

OH

2a 1S,3R,5S (–)-trans

HO

HO

2a΄ 1R,3S,5R (+)-trans

2b 1S,3S,5S (+)-cis

2b΄ 1R,3R,5R (–)-cis

Pinocarveol

(+)-trans-Pinocarveol (2a′) was biotransformed by A. niger TBUYN-2 to the following two pathways. That is, (+)-trans-pinocarveol (2a′) was metabolized via (+)-pinocarvone (3′), (–)-3-­isopinanone (413′), and (+)-2α-hydroxy-3-pinanone (414′) to (+)-2α,5-dihydroxy-3-pinanone (415′) (pathway 1). Furthermore, (+)-trans-pinocarveol (2a′) was metabolized to epoxide followed by rearrangement reaction to give 6β-hydroxyfenchol (349ba′) and 6β,7-dihydroxyfenchol (412ba′) (Noma and Asakawa, 2005a) (Figure 22.171). S. litura converted (+)-trans-pinocarveol (2a′) to (+)-pinocarvone (3′) as a major product (Miyazawa et al., 1995b) (Figure 22.171).

722

Handbook of Essential Oils OH

OH

OH

O 5΄

25΄

OH

OH 34΄

204΄

FIGURE 22.169  Biotransformation of (−)-myrtenol (5′) and (−)-α-terpineol (34′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., Microbial transformation of (−)-myrtenol and (−)nopol, Proceedings of 49th TEAC, 2005b, pp. 78–80.)

OH

COOH S. litura

8a

436 OH

COOH S. litura

8a΄

436΄

FIGURE 22.170  Biotransformation of (−)-trans-myrtanol (8a) and its enantiomer (8a′) by Spodoptera litura. (Modified from Miyazawa, M. et al. Biotransformation of (+)-trans myrtanol and (−)-trans-myrtanol by common cutworm Larvae, Spodoptera litura as a biocatalyst, Proceedings of 41st TEAC, 1997b, pp. 389–390.) OH HO

O

O

2a΄

O

O

A.n.

A.n.

A.n.

A.n.

S. litula.

OH

412΄

3΄

413΄

OH 414΄ OH

H+ O

HO

OH

HO

OH

A.n.

OH

A.n.

349ba΄

412ba΄

FIGURE 22.171  Biotransformation of (+)-trans-pinocarveol (2a′) by Aspergillus niger TBUYN-2 and Spodoptera litura. (Modified from Miyazawa, M. et  al. Biotransformation of trans-pinocarveol by plant pathogenic microorganism, Glomerella cingulata, and by the larvae of common cutworm, Spodoptera litura Fabricius, Proceedings of 39th TEAC, 1995c, pp. 360–361; Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

723

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

22.4.3.4 Pinane-2,3-Diol OH

OH

OH

OH

OH

OH HO

HO

418aa

418ab

418ab΄

418aa΄

()-Pinane-2,3-diol 1S,2S,3S,5S

(+)-Pinane-2,3-diol 1S,2S,3R,5S

(–)-Pinane-2,3-diol 1R,2R,3S,5R

()-Pinane-2,3-diol 1R,2R,3R,5R

This results led us to study the biotransformation of (–)-pinane-2,3-diol (418ab′) and (+)-pinane-2,3 diol (418ab) by A. niger TBUYN-2. (–)-Pinane-2,3-diol (418ab′) was easily biotransformed to give (–)-pinane-2,3,5-triol (419ab′) and (+)-2,5-dihydroxy-3-pinanone (415a′) as the major products and (+)-2-hydroxy-3-pinanone (414a′) as the minor product. On the other hand, (+)-pinane-2,3-diol (418ab) was also biotransformed easily to give (+)-pinane2,3,5-triol (419ab) and (–)-2,5-dihydroxy-3-pinanone (415a) as the major products and (–)-2-hydroxy3-pinanone (414a) as the minor product (Noma et al., 2003) (Figure 22.172). G. cingulata transformed (–)-pinane-2,3-diol (418ab′) to a small amount of (+)-2α-hydroxy-3-pinanone (414ab′, 5%) (Kamino and Miyazawa, 2005), whereas (+)-pinane-2,3-diol (418ab) was transformed to a small amount of (–)-2α-hydroxy-3-pinanone (414ab, 10%) and (–)-3-acetoxy-2α-pinanol (433ab-Ac, 30%) (Kamino et al., 2004) (Figure 22.172). OH OH

OH

OH OAc

A.n. OH

OH 419ab

G.c.

O

O

G.c.

418ab

418ab΄-Ac

OH

OH

A.n.

A.n.

414a

OH 415a

HO

HO

419ab΄

O

O

A.n.

OH

OH

OH

OH

OH

A.n.

A.n. G.c. 418ab΄

414a΄

OH 415a΄

FIGURE 22.172  Biotransformation of (+)-pinane-2,3-diol (418ab′) and (−)-pinane-2,3-diol (418ab′) by Aspergillus niger TBUYN-2(276)] and Glomerella cingulata. (Modified from Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, 2003, pp. 91–93; Kamino, F. et al. Biotransformation of (1S,2S,3R,5S)-(+)-pinane-2,3-diol using plant pathogenic fungus, Glomerella cingulata as a biocatalyst, Proceedings of 48th TEAC, 2004, pp. 383–384; Kamino, F. and Miyazawa, M., Biotransformation of (+)-and (−)-pinane-2,3-diol using plant pathogenic fungus, Glomerella cingulata as a biocatalyst, Proceedings of 49th TEAC, 2005, pp. 395–396.)

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Handbook of Essential Oils

22.4.3.5  Isopinocampheol (3-Pinanol) OH

420 ba (–)-isopino 1R,2R,3R,5S

HO

OH

OH

420bb (+)-neoiso 1R,2R,3S,5S

HO

420aa (–)-neo 1R,2S,3R,5S

HO

420ba΄ (+)1S,2S,3S,5R

420bb΄ (–)1S,2S,3R,5R

OH

420ab (+)-pinocampherol 1R,2S,3S,5S

HO

420aa΄ (+)-neo 1S,2R,3S,5R

420ab΄ 1S,2R,3R,5R

22.4.3.5.1 Chemical Structure of (–)-Isopinocampheol (420ba) and (+)-Isopinocampheol (420ba′) Biotransformation of isopinocampheol (3-pinanol) with 100 bacterial and fungal strains yielded 1-, 2-, 4-, 5-, 7-, 8-, and 9-hydroxyisopinocampheol besides three rearranged monoterpenes, one of them bearing the novel isocarene skeleton. A pronounced enantioselectivity between (–)- (420ba) and (+)-isopinocampheol (420ba′) was observed. The phylogenetic position of the individual strains could be seen in their ability to form the products from (+)-isopinocampheol (420ba′). The formation of 1,3-dihydroxypinane (421ba′) is a domain of bacteria, while 3,5- (415ba′) or 3,6-dihydroxypinane (428baa′) was mainly formed by fungi, especially those of the phylum Zygomycotina. The activity of Basidiomycotina toward oxidation of isopinocampheol was rather low. Such informations can be used in a more effective selection of strains for screening (Wolf-Rainer, 1994) (Figure 22.173). (+)-Isopinocampheol (420ba′) was metabolized to 4β-hydroxy-(+)-isopinocampheol (424′), 2β-hydroxy-(+)-isopinocampheol acetate (425ba′-Ac), and 2α-methyl,3-(2-methyl-2-hydroxypropyl)cyclopenta-1β-ol (432′) (Wolf-Rainer, 1994) (Figure 22.174). (–)-Isopinocampheol (420ba) was converted by S. litura to give (1R,2S,3R,5S)-pinane-2,3-diol (418ba) and (–)-pinane-3,9-diol (423ba), whereas (+)-isopinocampheol (420ba′) was converted to (+)-pinane-3,9-diol (423ba′) (Miyazawa et al., 1997c) (Figure 22.175). (–)-Isopinocampheol (420ba) was biotransformed by A. niger TBUYN-2 to give (+)-(1S,2S,3S,5R)pinane-3,5-diol (422ba, 6.6%), (–)-(1R,2R,3R,5S)-pinane-1,3-diol (421ba, 11.8%), and pinane-2,3-diol (418ba, 6.6%), whereas (+)-isopinocampheol (420ba′) was biotransformed by A. niger TBUYN-2 to give (+)-(1S,2S,3S,5R)-pinane-3,5-diol (422ba′, 6.3%) and (–)-(1R,2R,3R,5S)-pinane-1,3,-diol (421ba′, 8.6%) (Noma et al., 2009) (Figure 22.176). On the other hand, G. cingulata converted (–)(420ba) and (+)-isopinocampheol (420ba′) mainly to (1R,2R,3S,4S,5R)-3,4-pinanediol (484ba) and (1S,2S,3S,5R,6R)-3,6-pinanediol (485ba′), respectively, together with (418ba), (422ba), (422ba′), and (486ba′) as minor products (Miyazawa et al., 1997c) (Figure 22.176). Some similarities exist between the main metabolites with G. cingulata and R. solani (Miyazawa et al., 1997c) (Figure 22.176). 22.4.3.6  Borneol and Isoborneol

36a (1R,2S)-(+)borneol (36a)

OH

OH

HO

HO

36b΄ (1R,2R)-(–)isoborneol (36b΄)

36b΄ (1S,2R)-(–)borneol (36a΄)

36b (1S,2S)-(+)isoborneol (36b)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH

HO

RO OH

HO

HO HO 426ba΄

427ba΄,R:H 427ba΄-Ac,R:Ac

HO

725

OH

HO

429΄

RO

HO

OH

HO

420ba΄

421ba΄

OH

418ab΄

430΄

O

OH 428baa΄

OH 415ba΄,R:H 415ba΄-Ac,R:Ac

OH 431b΄

FIGURE 22.173  Metabolic pathways of (+)-isopinocampheol (420ba′) by microorganisms. (Modified from Wolf-Rainer, A., Z. Naturforsch., 49c, 553, 1994.)

OH

AcO

HO

OH

OH 432΄

420ba΄

425ba΄-Ac

HO

HO

424΄

FIGURE 22.174  Metabolic pathways of (+)-isopinocampheol (420ba′) by microorganisms. (Modified from Wolf-Rainer, A., Z. Naturforsch., 49c, 553, 1994.)

(–)-Borneol (36a′) was biotransformed by P. pseudomallei strain H to give (–)-camphor (37′), 6-hydroxycamphor (228′), and 2,6-diketocamphor (229′) (Hayashi et al., 1969) (Figure 22.177). E. gracilis Z. showed enantio- and diastereoselectivity in the biotransformation of (+)- (36a), (–)(36a′), and (±)-racemic borneols (equal mixture of 36a and 36a′) and (+)- (36b), (–)- (36b′), and (±)-isoborneols (equal mixture of 36b and 36b′). The enantio- and diastereoselective dehydrogenation for (–)-borneol (36a′) was carried out to give (–)-camphor (37′) at ca. 50% yield (Noma et al., 1992d; Noma and Asakawa, 1998). The conversion ratio was always ca. 50% even at different kinds of concentration of (–)-borneol (36a′). When (–)-camphor (37′) was used as a substrate, it was also converted to (–)-borneol (36a′) in 22% yield for 14 days. Furthermore, (+)-camphor (37) was also

726

Handbook of Essential Oils

OH

OH S. litura

HO 423ba

HO S. litura

HO

OH

420ba΄

420ba

423ba΄

S. litura HO OH

418ab

FIGURE 22.175  Biotransformation of (−)- (420ba) and (+)-isopinocampheol (420ba′) by Spodoptera litura. (Modified from Miyazawa, M. et al. Phytochemistry, 45, 945, 1997c.) OH OH

HO

HO

OH OH

418ba G.c. A.n.

OH 422ba

G.c.

A.n. G.c. OH

OH

HO

485ba΄ A.n.

G.c.

HO

HO

A.n. 421ba

OH 422ba΄

OH

A.n. 420ba

G.c.

420ba΄

OH

421ba΄

G.c.

HO OH

484ba

OH 486ba΄

FIGURE 22.176  Biotransformation of (−)- (420ba) and (+)-isopinocampheol (420ba′) by Aspergillus niger TBUYN-2 and Glomerella cingulata. (Modified from Miyazawa, M. et al. Phytochemistry, 45, 945, 1997c; Noma, Y. et al. Unpublished data, 2009.)

reduced to (+)-borneol (36a) in 4% and 18% yield for 7 and 14 days, respectively (Noma et al., 1992d; Noma and Asakawa, 1998) (Figure 22.178). (+)- (36a) and (–)-Borneols (36a′) were biotransformed by S. litura to (+)- (370a) and (–)-bornane2,8-diols (370a′), respectively (Miyamoto and Miyazawa, 2001) (Figure 22.179). 22.4.3.7  Fenchol and Fenchyl Acetate 10 1

HO

6

2 3

8 9

7

HO

OH

OH

5

4

(1R,2R,4S) (+)-endo (+)-α-fenchol 11a

(1R,2S,4S) (+)-exo (+)-β-fenchol 11b

(1S,2S,4R) (–)-endo (–)-α-fenchol 11a΄

(1S,2R,4R) (–)-exo (–)-β-fenchol 11b΄

727

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

O

HO

O

OH

O

37΄

P. pseudomonari HO

36b΄

OH

O

OH

O

229΄

228΄

FIGURE 22.177  Biotransformation of (−)-borneol (36a′) by Pseudomonas pseudomallei strain. (Modified from Hayashi, T. et al. J. Agric. Chem. Soc. Jpn., 43, 583, 1969.) HO

OH

E. g

rac

ilis

O

O

36b΄ HO

37 (+)

s

cili

ra E. g

37΄ (–)

ilis

rac

E. g

E.

gr

36b OH

ac

ilis

36a

36a΄

FIGURE 22.178  Enantio- and diastereoselectivity in the biotransformation of (+)- (36a) and (−)-borneols (36a′) by Euglena gracilis Z. (Modified from Noma, Y. et al. Biotransformation of terpenoids and related compounds, Proceedings of 36th TEAC, 1992b, pp. 199–201; Noma, Y. and Asakawa, Y., Euglena gracilis Z: Biotransformation of terpenoids and related compounds, in Bajaj, Y.P.S. (ed.), Biotechnology in Agriculture and Forestry, Vol. 41, Medicinal and Aromatic Plants X, Springer, Berlin, Germany, 1998, pp. 194–237.)

OH S. litura

HO

HO

S. litura

36a

OH 370

36a΄

OH HO 370΄

FIGURE 22.179  Biotransformation of (+)- (36a) and (−)-borneols (36a′) by Spodoptera litura. (Modified from Miyamoto, Y. and Miyazawa, M., Biotransformation of (+)- and (−)-borneol by the larvae of common cutworm (Spodoptera litura) as a biocatalyst, Proceedings of 45th TEAC, 2001, pp. 377–378.)

(1R,2R,4S)-(+)-Fenchol (11a) was converted by A. niger TBUYN-2 and A. cellulosae IFO 4040 to give (–)-fenchone (12), (+)-6β-hydroxyfenchol (349ab), (+)-5β-hydroxyfenchol (350ab), and 5α-hydroxyfenchol (350aa) (Noma and Asakawa, 2005a) (Figure 22.180). The larvae of common cutworm, S. litura, converted (+)-fenchol (11a) to (+)-10-hydroxyfenchol (467a), (+)-8-hydroxyfenchol (465a), (+)-6β-hydroxyfenchol (349ab), and (–)-9-hydroxyfenchol (466a) (Miyazawa and Miyamoto, 2004) (Figure 22.180). (+)-trans-Pinocarveol (2), which was formed from (–)-β-pinene (1), was metabolized by A. niger TBUYN-2 to 6β-hydroxy-(+)-fenchol (349ab) and 6β,7-dihydroxy-(+)-fenchol (412ba′). (–)-Fenchone (12) was also metabolized to 6α-hydroxy- (13b) and 6β-hydroxy-(–)-fenchone (13a).

728

Handbook of Essential Oils OH HO HO O

OH

467a S. litura 10

A.n.

12

1

HO

HO

S. litura 8

HO

6

2 3 9

465a

7

349ab A.n. S. litura HO A.n.

5 4 11a

350ab

A.n.

OH

S. litura HO

HO

466a

HO

350aa

OH

FIGURE 22.180  Biotransformation of (+)-fenchol (11a) by Aspergillus niger TBUYN-2, Aspergillus cellulosae IFO 4040, and the larvae of common cutworm, Spodoptera litura. (Modified from Miyazawa, M. and Miyamoto, Y., Tetrahadron, 60, 3091, 2004; Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

(+)-Fenchol (11) was metabolized to 6β-hydroxy-(+)-fenchol (349ab) by A. niger TBUYN-2. The relationship of the metabolisms of (+)-trans-pinocarveol (2), (–)-fenchone (12), and (+)-fenchol (11) by A. niger TBUYN-2 is shown in Figure 22.181 (Noma and Asakawa, 2005a). (+)-α-Fencyl acetate (11a-Ac) was metabolized by G. cingulata to give (+)-5-β-hydroxy-αfencyl acetate (350a-Ac, 50%) as the major metabolite and (+)-fenchol (11a, 20%) as the minor metabolite (Miyazato and Miyazawa, 1999). On the other hand, (–)-α-fencyl acetate (11a′-Ac) was metabolized to (–)-5-β-hydroxy-α-fencyl acetate (350a′-Ac, 70%) and (–)-fenchol (11a′, 10%) as the minor metabolite by G. cingulata (Miyazato and Miyazawa, 1999) (Figure 22.182). 22.4.3.8 Verbenol

HO

HO

23a΄ (–)-transverbenol

OH

23b΄ (–+)-cisverbenol

23a (+)-transverbenol

OH

23b (–)-cisverbenol

(–)-trans-Verbenol (23a′) was biotransformed by S. litura to give 10-hydroxyverbenol (451a′). Furthermore, (–)-verbenone (24′) was also biotransformed in the same manner to give 10-hydroxyverbenone (25′) (Yamanaka and Miyazawa, 1999) (Figure 22.183).

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

729

HO

1

2a OH

HO

HO

OH

11a

HO

OH

349ab OH

O

412ba΄ OH

O

O

13a

13b

12

FIGURE 22.181  Metabolism of (+)-trans-pinocarveol (2), (−)-fenchone (12), and (+)-fenchol (11) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.) 10 HO

2 3

8 9

11a 20%

AcO

1

AcO

6 7

5 4 11a-Ac

OH

350a-Ac 50%

OH

OAc

OAc 2 5 HO

11a΄ 10%

350a΄-Ac 70%

11a΄-Ac

FIGURE 22.182  Biotransformation of (+)- (11a-Ac) and (−)-α-fencyl acetate (11a′-Ac) by Glomerella ­cingulata. (Modified from Miyazato, Y. and Miyazawa, M., Biotransformation of (+)- and (−)-α-fenchyl acetated using plant parasitic fungus, Glomerella cingulata as a biocatalyst, Proceedings of 43rd TEAC, 1999, pp. 213–214.) OH

OH S. litura

S. litura

HO

O

HO 23a΄ (–)-transverbenol

451a΄

24΄ (–)-verbenone

O 25΄

FIGURE 22.183  Metabolism of (–)-trans-verbenol (23a′) and (–)-verbenone (24′) by Spodoptera litura. (Modified from Yamanaka, T. and Miyazawa, M., Biotransformation of (−)-trans-verbenol by common cutworm larvae, Spodoptera litura as a biocatalyst, Proceedings of 43rd TEAC, 1999, pp. 391–392.)

730

Handbook of Essential Oils OH

OH

OH

O

452΄

454΄

OH

453΄

FIGURE 22.184  Biotransformation of (–)-nopol (452′) by Aspergillus niger TBUYN-2, Aspergillus sojae IFO 4389, and Aspergillus usami IFO 4338. (Modified from Noma, Y. and Asakawa, Y., Microbial transformation of (−)-myrtenol and (−)-nopol, Proceedings of 49th TEAC, 2005b, pp. 78–80; Noma, Y. and Asakawa, Y., Microbial transformation of (−)-nopol benzyl ether, Proceedings of 50th TEAC, 2006c, pp. 434–436.)

22.4.3.9  Nopol and Nopol Benzyl Ether Biotransformation of (–)-nopol (452′) was carried out at 30°C for 7 days at the concentration of 100 mg/200 mL medium by A. niger TBUYN-2, A. sojae IFO 4389, and A. usami IFO 4338. (–)Nopol (452′) was incubated with A. niger TBUYN-2 to give 7-hydroxymethyl-1-p-menthen-8-ol (453′). In cases of A. sojae IFO 4389 and A. usami IFO 4338, (–)-nopol (452′) was metabolized to 3-oxonopol (454′) as a minor product together with 7-hydroxymethyl-1-p-menthen-8-ol (453′) as a major product (Noma and Asakawa, 2005b, 2006c) (Figure 22.184). Biotransformation of (–)-nopol benzyl ether (455′) was carried out at 30°C for 8–13 days at the concentration of 277 mg/200 mL medium by A. niger TBUYN-2, A. sojae IFO 4389, and A. usami IFO 4338. (–)-Nopol benzyl ether (455′) was biotransformed by A. niger TBUYN-2 to give 4-oxonopol-2′, 4′-dihydroxy benzyl ether (456′), and (–)-oxonopol (454′). 7-Hydroxymethyl-1-p-menthen-8-ol benzyl ether (457′) was not formed at all (Figure 22.185). 4-Oxonopol-2′,4′-dihydroxybenzyl ether (456′) shows strong antioxidative activity (IC50 30.23 µM). The antioxidative activity of 4-oxonopol-2′,4′-dihydroxybenzyl ether (456′) is the same as that of butyl hydroxyl anisole (BHA) (Noma and Asakawa, 2006b,c). Citrus pathogenic fungi A. niger Tiegh (CBAYN) also transformed (–)-nopol (452′) to (–)-­oxonopol (454′) and 4-oxonopol-2′,4′-dihydroxybenzyl ether (456′) (Noma and Asakawa, 2006b,c) (Figure 22.186).

22.4.4 Bicyclic Monoterpene Ketones 22.4.4.1  α-, β-Unsaturated Ketone 22.4.4.1.1 Verbenone

O

O

24 (+)-verbenone

24΄ (–)-verbenone

(–)-Verbenone (24′) was hydrogenated by reductase of N. tabacum to give (–)-isoverbanone (458b′) (Suga and Hirata, 1990; Shimoda et al., 1996, 1998, 2002; Hirata et al., 2000) (Figure 22.187). 22.4.4.1.2 Pinocarvone O

3 (–)-Pinocarvone

O

3΄ (+)-Pinocarvone

731

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH O

O

OH OH +

O

O 455΄

454΄

456΄

FIGURE 22.185  Biotransformation of (–)-nopol benzyl ether (455′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., Biotransformation of β-pinene, myrtenol, nopol and nopol benzyl ether by Aspergillus niger TBUYN-2, Book of Abstracts of the 37th ISEO, 2006b, p. 144; Noma, Y. and Asakawa, Y., Microbial transformation of (−)-nopol benzyl ether, Proceedings of 50th TEAC, 2006c, pp. 434–436.) 2΄ O

O 4΄

4 OH

455΄ 457 OH

OH

O

O

OH

O OH

OH O

HO

HO

O

454΄ 456΄

FIGURE 22.186  Proposed metabolic pathways of (–)-nopol benzyl ether (455′) by microorganisms. (Modified from Noma, Y. and Asakawa, Y., Biotransformation of β-pinene, myrtenol, nopol and nopol benzyl ether by Aspergillus niger TBUYN-2, Book of Abstracts of the 37th ISEO, 2006b, p. 144; Noma, Y. and Asakawa, Y., Microbial transformation of (−)-nopol benzyl ether, Proceedings of 50th TEAC, 2006c, pp. 434–436.)

A. niger TBUYN-2 transformed (+)-pinocarvone (3′) to give (–)-isopinocamphone (413b′), 2α-hydroxy-3-pinanone (414b′), and 2α, 5-dihydroxy-3-pinanone (415b′) together with small amounts of 2α, 10-dihydroxy-3-pinanone (416b′) (Noma and Asakawa, 2005a) (Figure 22.188). 22.4.4.2  Saturated Ketone 22.4.4.2.1 Camphor O

O

37 (1R)-(+)-camphor (37)

37΄ (1S)-(–)-camphor (37΄)

732

Handbook of Essential Oils

N. tabacum Verbenone reductase O

O 24΄

458b΄

FIGURE 22.187  Hydrogenation of (–)-verbenone (24′) to (–)-isoverbanone (458b′) by verbenone reductase of Nicotiana tabacum. (Modified from Suga, T. and Hirata, T., Phytochemistry, 29, 2393, 1990; Shimoda, K. et al. J. Chem. Soc., Perkin Trans., 1, 355, 1996; Shimoda, K. et al. Phytochemistry, 49, 49, 1998; Shimoda, K. et al. Bull. Chem. Soc. Jpn., 75, 813, 2002; Hirata, T. et al. Chem. Lett., 29, 850, 2000.) OH O OH O

O

O OH 415b΄

3΄

413b΄

414b΄

OH O OH 416b΄

FIGURE 22.188  Biotransformation of (+)-pinocarvone (3′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. and Asakawa, Y., New metabolic pathways of β-pinene and related compounds by Aspergillus niger, Book of Abstracts of the 36th ISEO, 2005a, p. 32.)

(+)- (37) and (–)-Camphor (37′) are found widely in nature, of which (+)-camphor (37) is more abundant. It is the main component of oils obtained from the camphor tree C. camphora (Bauer et al., 1990). The hydroxylation of (+)-camphor (37) by P. putida C1 was described (Abraham et al., 1988). The substrate was hydroxylated exclusively in its 5-exo- (235b) and 6-exo- (228b) positions. Although only limited success was achieved in understanding the catabolic pathways of (+)-camphor (37), key roles for methylene group hydroxylation and biological Baeyer–Villiger monooxygenases in ring cleavage strategies were established (Trudgill, 1990). A degradation pathway of (+)-camphor (37) by P. putida ATCC 17453 and Mycobacterium rhodochrous T1 was proposed (Trudgill, 1990). The metabolic pathway of (+)-camphor (37) by microorganisms is shown in Figure 22.189. (+)-Camphor (37) is metabolized to 3-hydroxy- (243), 5-hydroxy- (235), 6-hydroxy- (228), and 9-hydroxycamphor (225) and 1,2-campholide (237). 6-Hydroxycamphor (228) is degradatively metabolized to 6-oxocamphor (229) and 4-carboxymethyl-2,3,3-trimethylcyclopentanone (230), 4-carboxymethyl-3,5,5-trimethyltetrahydro-2-pyrone (231), isohydroxycamphoric acid (232), isoketocamphoric acid (233), and 3,4,4-trimethyl-5-oxo-trans-2-hexenoic acid (234), whereas 1,2-campholide (237) is also degradatively metabolized to 6-hydroxy-1,2-campholide (238), 6-oxo-1,2-campholide (239), and 5-carboxymethyl-3,4,4-trimethyl-2-cyclopentenone (240), 6-carboxymethyl-4,5,5-trimethyl-5,6-dihydro-2-pyrone (241), and 5-carboxymethyl-3,4,4-trimethyl2-heptene-1,7-dioic acid (242). 5-Hydroxycamphor (235) is metabolized to 6-hydroxy-1,2-campholide (238), 5-oxocamphor (236), and 6-oxo-1,2-campholide (239). 3-Hydroxycamphor (243) is also metabolized to camphorquinone (244) and 2-hydroxyepicamphor (245) (Bradshaw et  al., 1959;

733

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

AcO

HO

O HO 225a

36a and b O

O

226a and b

O

O

HO

OH HO

37

225b

O

O

O

244

COOH

OH 235

237

O

O

O

COOH

228

o

391

O

O OH

O

236

245

OH

O

o

O

243

238

229

O

230

239

O

O

O

o

COOH

231

COOH

HO

COOH

240

O

O

O COOH

241

COOH

COOH COOH 232 O

COOH COOH 233

COOH 242

OH

O H COOH 234

FIGURE 22.189  Metabolic pathways of (+)-camphor (37) by Pseudomonas putida and Corynebacterium diphtheroides. (Modified from Bradshaw, W.H. et al. J. Am. Chem. Soc., 81, 5507, 1959; Conrad, H.E. et al. Biochem. Biophys. Res. Commun., 6, 293, 1961; Conrad, H.E. et al. J. Biol. Chem., 240, 495, 1965a; Conrad, H.E. et al. J. Biol. Chem., 240, 4029, 1965b; Gunsalus, I.C. et al. Biochem. Biophys. Res. Commun., 18, 924, 1965; Chapman, P.J. et al. J. Am. Chem. Soc., 88, 618, 1966; Hartline, R.A. and Gunsalus, I.C., J. Bacteriol., 106, 468, 1971; Oritani, T. and Yamashita, K., Agric. Biol. Chem., 38, 1961, 1974.)

734

Handbook of Essential Oils

O

O

O

O

+

Rat CYP2B1

OH

OH

225

228b

235b

OH

O

O

OH

+

+

235a

37

O

OH

O

O

Human CYP2A6

Human CYP2A6 HO 37

235a΄

37΄

228b

FIGURE 22.190  Biotransformation of (+)-camphor (37) by rat P450 enzyme (above) and (+)- (37) and (–)-camphor (37′) by human P450 enzymes.

OH

O

O

+

E. perriniana

E. perriniana 37

O-Glc

O

Another camphor glycoside

3 new

228

FIGURE 22.191  Biotransformation of (+)-camphor (37) by Eucalyptus perriniana suspension cell.

Conrad et al., 1961, 1965a,b; Gunsalus et al., 1965; Chapman et al., 1966; Hartline and Gunsalus, 1971; Oritani and Yamashita, 1974) (Figure 22.189). Human CYP2A6 converted (+)-camphor (37) and (–)-camphor (37′) to 6-endo-hydroxycamphor (228a) and 5-exo-hydroxycamphor (235b), while rat CYP2B1 did 5-endo- (235a), 5-exo- (235b), and 6-endo-hydroxycamphor (228a) and 8-hydroxycamphor (225) (Gyoubu and Miyazawa, 2006) (Figure 22.190). (+)-Camphor (37) was glycosylated by E. perriniana suspension cells to (+)-camphor monoglycoside (3 new, 11.7%) (Hamada et al., 2002) (Figure 22.191). 22.4.4.2.2 Fenchone O

12 (+)-

O

12΄ (–)-

(+)-Fenchone (12) was incubated with Corynebacterium sp. (Chapman et al., 1965) and Absidia orchidis (Pfrunder and Tamm, 1969a) give 6β-hydroxy- (13a) and 5β-hydroxyfenchones (14a) (Figure 22.191). On the other hand, A. niger biotransformed (+)-fenchone (12) to (+)-6α- (13b) and (+)-5α-hydroxyfenchones (14b) (Miyazawa et al., 1990a,b) and 5-oxofenchone (15), 9-formylfenchone (17b), and 9-carboxyfenchone (18b) (Miyazawa et al., 1990a,b) (Figure 22.192). Furthermore, A. niger biotransformed (–)-fenchone (12′) to 5α-hydroxy- (14b′) and 6α-hydroxyfenchones (13b′) (Yamamoto et al., 1984) (Figure 22.193). (+)- and (–)-Fenchone (12 and 12′) were converted to 6β-hydroxy- (13a, 13a′), 6α-hydroxyfenchone (13b, 13b′), and 10 hydroxyfenchone (4, 4′) by P450. Of the 11 recombinant human P450 enzymes

735

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

HO

O

O

O

6

A.n.

5

HO 13b

HO

O

6

A.n.

A.n.

O

15

14b

O

A. orchidis

O

Corynebacterium sp. A. orchidis

Corynebacterium sp.

5

HO

12

13a

5

14a

A.n. O

O

CHO

COOH

A.n.

17b

18b

FIGURE 22.192  Metabolic pathways of (+)-fenchone (12) by Corynebacterium sp., A. orchidis, and Aspergillus niger TBUYN-2. (Modified from Chapman, P.J. et al. Biochem. Biophys. Res. Commun., 20, 104, 1965; Pfrunder, B. and Tamm, Ch., Helv. Chim. Acta., 52, 1643, 1969a; Miyazawa, M. et al. Chem. Express, 5, 237, 1990a; Miyazawa, M. et al. Chem. Express, 5, 407, 1990b.) 10

O

O 8 14b΄

OH

9

1 6

2 3

7

OH

O

5

4 12΄

13b΄

FIGURE 22.193  Metabolic pathways of (−)-fenchone (12′) by Aspergillus niger TBUYN-2. (Modified from Yamamoto, K. et al. Biotransformation of d- and l-fenchone by a strain of Aspergillus niger, Proceedings of 28th TEAC, 1984, pp. 168–170.)

tested, CYP2A6 and CYP2B6 catalyzed oxidation of (+)- (12) and (–)-fenchone (12′) (Gyoubu and Miyazawa, 2005) (Figure 22.194). 22.4.4.2.3  3-Pinanone (Pinocamphone and Isopinocamphone) O

413a 1R2S5S (+)-pinocamphone

O

413b 1R,2R,5S (+)-isopinocamphone

O

413a΄ 1S2R5R (–)-pinocamphone

O

413b΄ 1S2S5R (–)-isopinocamphone

(+)- (413) and (−)-Isopinocamphone (413′) were biotransformed by A. niger to give (−)- (414) and (+)-2-hydroxy-3-pinanone (414′) as the main products, respectively, which inhibit strongly

736

Handbook of Essential Oils OH O

HO 6

P450

O

HO 6

O

O

+

+ 13a

12

4

13b

OH O

OH

O

P450

OH

O

+

+ 13a΄

12΄

4΄

13b΄

FIGURE 22.194  Biotransformation of (+)-fenchone (12) and (−)-fenchone (12′) by P450 enzymes. (Modified from Gyoubu, K. and Miyazawa, M., Biotransformation of (+)- and (−)-fenchone by liver microsomes, Proceedings of 49th TEAC, 2005, pp. 420–422.) OH

OH O

O

413

O

414

OH 415 OH

OH O

O

O

413΄

OH 415΄

414΄

FIGURE 22.195  Biotransformation of (+)-isopinocamphone (413b) and (−)-isopinocamphone (413b′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, pp. 91–93, 2003; Noma, Y. et al. Biotransformation of (+)- and (−)-3-pinanone by Aspergillus niger, Proceedings of 48th TEAC, 2004, pp. 390–392.)

germination of lettuce seeds, and (−)- (415) and (+)-2,5-dihydroxy-3-pinanone (415′) as the minor components, respectively (Noma et al., 2003, 2004) (Figure 22.195). 22.4.4.2.4 2-Hydroxy-3-Pinanone OH

OH

O

O

(1S,2R,5S) (–)-2-OH-3pinanone 414a

(1S,2R,5S) (+)-2-OH-3pinanone 414b

OH

OH O

O

(1R,2R,5R) (+)-2-OH-3pinanone 414a΄

(1R,2S,5R) (+)-2-OH-3pinanone 414b΄

(−)-2α-Hydroxy-3-pinanone (414) was incubated with A. niger TBUYN-2 to give (−)-2α, 5-dihydroxy3-pinanone (415) predominantly, whereas the fungus converted (+)-2α-hydroxy-3-pinanone (414′)

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

737

OH

OH O

O

A. niger

414

OH 415 OH

OH HO

O OH

A. niger

A. niger

O

OH 419΄

OH 415΄

A. niger

OH O

414΄

OH

416΄

FIGURE 22.196  Biotransformation of (−)- (414) and (+)-2-hydroxy-3-pinanone (414′) by Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, pp. 91–93, 2003; Noma, Y. et al. Biotransformation of (+)- and (−)-3-pinanone by Aspergillus niger, Proceedings of 48th TEAC, 2004, pp. 390–392.)

mainly to 2α, 5-dihydroxy-3-pinanone (415′), 2α,9-dihydroxy-3-pinanone (416′), and (−)-pinane2α,3α,5-triol (419ba′) (Noma et al., 2003, 2004) (Figure 22.196). A. niger TBUYN-2 metabolized β-pinene (1), isopinocamphone (414b), 2α-hydroxy-3-pinanone (414a), and pinane-2,3-diol (419ab) as shown in Figure 22.197. On the other hand, A. niger TBUYN-2 and B. cinerea metabolized β-pinene (1′), isopinocamphone (414b′), 2α-hydroxy-3-pinanone (414a′), OH

OH OH

OH

OH O

A.n. 418ab

1

A.n. A.n.

OH 415ab OH

OH

O

O

A.n. 420ba

OH 415

A.n.

414b

OH

A.n.

O

HO

414a 416b

FIGURE 22.197  Relationship of the metabolism of β-pinene (1), isopinocamphone (414b), 2α-hydroxy-3pinanone (414a), and pinane-2,3-diol (419ab) in Aspergillus niger TBUYN-2. (Modified from Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, 2003, pp. 91–93; Noma, Y. et al. Biotransformation of (+)- and (−)-3-pinanone by Aspergillus niger, Proceedings of 48th TEAC, 2004, pp. 390–392.)

738

Handbook of Essential Oils OH

OH

OH

O

HO

HO

A.n. B.c.

A.n. 418ab΄

1΄

OH 419ab΄

A.n.

OH 415a΄

A.n.

A.n. OH

OH HO

O

O

420ba΄

O

A.n.

414b΄

OH

A.n. 414a΄

416b΄

FIGURE 22.198  Relationship of the metabolism of β-pinene (1′), isopinocamphone (414b′), 2α-hydroxy3-pinanone (414a′), and pinane-2,3-diol (419ab′) in Aspergillus niger TBUYN-2 and Botrytis cinerea. (Modified from Noma, Y. et al. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger, Proceedings of 47th TEAC, 2003, pp. 91–93; Noma, Y. et al. Biotransformation of (+)- and (−)-3-pinanone by Aspergillus niger, Proceedings of 48th TEAC, 2004, pp. 390–392.)

and pinane-2,3-diol (419ab′) as shown in Figure 22.198. The relationship of the metabolism of β-pinene (1, 1′), isopinocamphone (414b, 414b′), 2α-hydroxy-3-pinanone (414a, 414a′), and pinane2,3-diol (419ab, 419ab′) in A. niger TBUYN-2 and B. cinerea is shown in Figures 22.197 and 22.198. 22.4.4.2.4.1  Mosquitocidal and Knockdown Activity  Knockdown and mortality activity toward mosquito, Culex quinquefasciatus, was carried out for the metabolites of (+)- (418ab) and (−)-pinane-2,3-diols (418ab′) and (+)- and (−)-2-hydroxy-3-pinanones (414 and 414′) by Dr. Radhika Samarasekera, Industrial Technology Institute, Sri Lanka. (−)-2-Hydroxy-3-pinanone (414′) showed the mosquito knockdown activity and the mosquitocidal activity at the concentration of 1% and 2% (Table 22.16). 22.4.4.2.4.2  Antimicrobial Activity  The microorganisms were refreshed in Mueller–Hinton broth (Merck) at 35°C–37°C and inoculated on Mueller–Hinton agar (Mast Diagnostics, Merseyside, United Kingdom) media for preparation of inoculum. E. coli (NRRL B-3008), P. aeruginosa (ATCC 27853), TABLE 22.16 Knockdown and Mortality Activity toward Mosquitoa Compounds (+)-2,5-Dihydroxy-3-pinanone (415%, 2%)

Knockdown (%) 27

Mortality (%) 20

(−)-2,5-Dihydroxy-3-pinanone (415′, 2%)

NT

7

(+)-2-Hydroxy-3-pinanone (414%, 2%)

40

33

(−)-2-Hydroxy-3-pinanone (414′, 2%)

100

40

(−)-2-Hydroxy-3-pinanone (414′, 1%)

53

7

(+)-Pinane-2,3,5-triol (419%, 2%)

NT

NT

(−)-Pinane-2,3,5-triol (419%, 2%)

13

NT

(+)-Pinane-2,3-diol (418%, 2%)

NT

NT

(−)-Pinane-2,3-diol (418′, 2%)

NT

NT

a

The results are against Culex quinquefasciatus.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

739

TABLE 22.17 Biological Activity of Pinane-2,3,5-Triol (419 and 419′), 2,5-Dihydroxy-3-Pinanone (415 and 415′), and 7-Hydroxymethyl-1-p-Menthene-8-ol (453′) toward MRSA MIC (mg/mL) Compounds

Control

Microorganisms

419

415′

415

419′

453′

ST1

ST2

ST3

Escherichia coli Pseudomonas aeruginosa Enterobacter aerogenes Salmonella typhimurium Candida albicans Staphylococcus epidermidis MRSA

0.5 0.5 0.5 0.25 0.5 0.5 0.25

0.5 0.125 0.5 0.125 0.125 0.5 0.125

0.25 0.125 0.25 0.125 0.125 0.25 0.125

0.5 0.25 0.5 0.25 0.25 0.5 0.25

0.25 0.25 1.00 0.25 1.00 1.00 0.125

0.007 0.002 0.007 0.01 NT 0.002 0.5

0.0039 0.0078 0.0019 0.0019 NT 0.0009 0.031

Nt Nt Nt Nt 0.0625 NT NT

Source: Iscan (2005, unpublished data). Note: MRSA, methicillin-resistant Staphylococcus aureus; NT, not tested; ST1, ampicillin Na (sigma); ST2, chloramphenicol (sigma); ST3, ketoconazole (sigma).

Enterobacter aerogenes (NRRL 3567), S. typhimurium (NRRL B-4420), Staphylococcus epidermidis (ATCC 12228), methicillin-resistant Staphylococcus aureus (MRSA, clinical isolate, Osmangazi University, Faculty of Medicine, Eskisehir, Turkey), and Candida albicans (clinical isolate, Osmangazi University, Faculty of Medicine, Eskisehir, Turkey) were used as pathogen test microorganisms. Microdilution broth susceptibility assay (R1, R2) was used for the antimicrobial evaluation of the samples. Stock solutions were prepared in DMSO (Carlo-Erba). Dilution series were prepared from 2 mg/mL in sterile distilled water in microtest tubes from where they were transferred to 96-well microtiter plates. Overnight grown bacterial and candial suspensions in double strength Mueller–Hilton broth (Merck) was standardized to approximately 108 CFU/mL using McFarland No. 0.5 (106 CFU/mL for C. albicans). A volume of 100 µL of each bacterial suspension was then added to each well. The last row containing only the serial dilutions of samples without microorganism was used as negative control. Sterile distilled water, medium, and microorganisms served as a positive growth control. After incubation at 37°C for 24 h, the first well without turbidity was determined as the minimal inhibition concentration (MIC), and chloramphenicol (sigma), ampicillin (sigma), and ketoconazole (sigma) were used as standard antimicrobial agents (Amsterdam, 1997; Koneman et al., 1997) (Table 22.17).

22.5 SUMMARY 22.5.1 Metabolic Pathways of Monoterpenoids by Microorganisms About 50 years is over since the hydroxylation of α-pinene (4) was reported by A. niger in 1960 (Bhattacharyya et al., 1960). During these years, many investigators have studied the biotransformation of a number of monoterpenoids by using various kinds of microorganisms. Now we summarize the microbiological transformation of monoterpenoids according to the literatures listed in the references including the metabolic pathways (Figures 22.199 through 22.206) for the further development of the investigation on microbiological transformation of terpenoids. Metabolic pathways of β-pinene (1), α-pinene (4), fenchol (11), fenchone (12), thujone (28), carvotanacetone (47), and sobrerol (43) are summarized in Figure 22.199. In general, β-pinene (1) is metabolized by six pathways. At first, β-pinene (1) is metabolized via α-pinene (4) to many metabolites such as myrtenol (5) (Shukla et al., 1968; Shukla and Bhattacharyya, 1968), verbenol (23) (Bhattacharyya et al., 1960; Prema and Bhattachayya, 1962), and thujone (28) (Gibbon and

740

Handbook of Essential Oils OH

OH

OH

O

OH

OH

O

O

O

OH 416

3

OH 415 +

OH

O

7

413

OH

O

26

204

CHO

6

CHO

OH

O

414

418

OH 419

OH

COOH

OH OH

O

27

OH

OH

O

24

O

25

OH

OH O

OH O HO 31

30 37

36

5

2

35

OH

O

OH

23

OH

+ OH

OH

453

+ 33

34

28

4

1

32

+

OH HO

HO

CHO

OH

466

O

476 OH

11

HO

40 O

O

12

19

OH O

OH

46

47

OH

O

16

COOH O

OH 34

OH

68

81

43

O

O

18 COOH

172

Figure 19.201

COOH

COOH

22 O

173

177

OH

HO COOH

O

o

O

COOH

45

17

O

334

93

O o

OH

O

O

OH

OH

44

OH CHO

15

O

171

OH

Figures 19.201 and 19.203

20

O 14

56

OH

OH 412

O

O

349

OH

+ 42

OH O

39 OH

Figures 19.202 and 19.205

O

+ 33

12

HO

OH

67 COOH

11 OH

38

O

13 HO

CHO

O

OH

32 OH

29

COOH

COOH

OH 21

50 176

175

174

FIGURE 22.199  Metabolic pathways of β-pinene (1), α-pinene (4), fenchone (9), thujone (28), and carvotanacetone (44) by microorganisms.

741

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

OH O

O

OH 121

OH

OH

CHO

COOH

64 OH

65

120 54

63

O

HO

O

O

O

119

OH O

OH

114

OH

116

OH

62

115

66

OH

O

O

O 112

118

+ 33

HO 111

OH

378

68 OH

O

110

71

69

OH

OH

OH

O

O

101

72

OH

OH OH O

O

O OH 157

117 OH

HO HO

113

O

O

OH

57 OH

81

34 HO HO

OH

OH

81

Figure 19.205 125 OH

OH

OH 93

101

O

OH OH

76

COOH HO

COOH

O

75 COOH

CHO

105

82

COOH COOH

COOH 88

86

87

O OH 102

OH

81

O

O OH 44

97

84

COOH

OH

O

78 COOH

O 109

OH

OH

OAc

COOH

Figure 19.205

O

334

85

OH

106 96

O

378

O

77

OH

OH

O

O

74

O O

122 OH

99

OH

HO O

HO

O

OH

90

O

98

94

OH

204

61

HO

70 COOH

100

O

OH

HO

O

58 COOH

92

91

68

68

OH

HO O OH

O

O 103

OH

OH

HO 104

O

OH 50

OH 45

COOH 89 83

OH OH

FIGURE 22.200  Metabolic pathways of limonene (68), perillyl alcohol (74), carvone (93), isopiperitenone (111), and piperitenone (112) by microorganisms.

742

Handbook of Essential Oils OH

HO O

O

158 COOH

159 OH

157

47

HOOC

CHO

O

OH

O

162

O

161

160

OH

144

166

O

151 CHO

COOH

COOH O 199

OH

OH COOH O

182 OH O

O OH 179

COOH

COOH

COOH O

184

186

195

COOH

OH COOH O

OH

180

OH

198

OH HO

OH

358

137-Ac

145 HO

O

COOH

COOH

OAc

OH OH

181 HO

O

OH

O

O 163

164

185

O 183

COOH 89 CH3COOH 187

185

OH

CHO

O COOH O

OH OH

OH OH

147

HO

O

OH

141

142

143

148

OH OH

OH OH

OH

O

O

167

COOH

OH

140

137 OH

OH

COOH 200

89

OH

OH

149

O OH

170

52

139

O

HO

OH

OH

O

168

OH

OH

138

O

COOH

OH

O

O

O

O

150

156 O

169

51

OH HO OH

O

O

OH

OH

146

155

154 O

O

O

Figure 19.200 OH OH

O

O OH

OH

OH OH

OH OH

197

196

194

193

COOH OH

O

477

OH

178

188

346

351

OH

OH

201

192

OH

OH

191

475

OH

OH

471 OH OH

OH

OH

OH

OH 189

352 OH OMe

OH

OMe

OH OH 470

474 OH

477

OH

472

COOH 190

COOH 475-Me OH OMe

473

OH 478

202 OMe

OH 471=Me

FIGURE 22.201  Metabolic pathways of menthol (137), menthone (149), p-cymene (178), thymol (179), carvacrol methyl ether (201), and carvotanacetone (47) by microorganisms and rabbit.

743

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals COOH

COOH

COOH

O

COOH

86

85

84

OH O

COOH

COOH

82

61

CHO

OH

CHO

HOOC

OH

OH

OH

COOH

205 OH

O OH

O 216

74

215

212

78

OH

+ 213

204

HO

O

O

43

O

O

218

O

217

HOOC

221

COOH HOOC 233

COOH 232

O

O

O

HO

O

COOH 231

COOH

211

34 Figure 19.200 and 19.203

+

O

O

COOH 241

O

COOH 240

207

OAc 227

228 O O

238

37

226

+ 33

+

+

35

32

OH

O

O

208

O

229

O

239

206

O HO

COOH 230

O HO

OH

COOH 219

O

O O

210

HO

+ 222

220

OH COOH 242

209

214

HOOC

HOOC HOOC 223 224

234 COOH HOOC

OH

HO

HO

O

125

HO

O

36

237 O

O O

O HO

236

243

OH

OH 34

235 OH

245

+

1

33

O

O

244

O +

OH . H2O 60

59

COOH 89 +

COOH O

251

COOH COOH

COOH

COOH

COOH CO2 + H2O

OH

CHO

COOH

OH

O

COOH

46

O 250

249

248

HO

COOH

257

247

HO

246

64

63

253

255

62

OH HO HO

OH HO

CHO HO

256

65

252

66

54

HO

OH 254

FIGURE 22.202  Metabolic pathway of borneol (36), camphor (37), phellandral (64), linalool (206), and p-menthane (252) by microorganisms.

744

Handbook of Essential Oils

CH2OH Figure 19.202

CH2OH

OH OH

263

260

OH

CH2OAc

OH

266

CH2OH

OAc 294 Figure 19.200 and 19. 201

CH2OH CH2OH

137 O 259

269

268

OH

OH

265

267

68

OH

OAc 137

294

CH2OAc CHO

COOH

CH2OH

261

262

258

O

CH2OH OH

264

270 HO

O

291 CH2OH

CH2OH

O 273

274

299

CH2OH CH2OH

CH2OH

CH2OH

272

OH

271

OH

CH2OH

OH

300

CH2OH

OH

OH

CHO 292

CHO

OH 206

275

276

CH O

O

CH2COOH

277

OH

278

288 COOH

O

H2O

COOH

O

O

CH2COOH OH

CH2OH COOH

COOH 283

282

OH

293

COOH

COOH

266

CH2OH

COOH

CO2

281

CH2OH

287 CH2OH

279 COOH

298 294

O

289

COOH

O 295

O

OH O

284

CO2 + H2O

COOH

HOOC 286

280

O

290

296

285

FIGURE 22.203  Metabolic pathways of citronellal (258), geraniol (271), nerol (272), and citral (275 and 276) by microorganisms.

745

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals OH

OH

HO

HO OH

373 HO 371

HO 372

HO OH

OAc

O

342

HO

O

HO

O

O

O

O HO

OH 50

49-Ac

O

45

49

133

102

OH OH

55

132

O

OH

OH

136

OH

O

O

O

O

131 OH

OH

OH O

OH

O

O

OH 135

134 101

93

OH 54

44

48

53

O

HO O O

O

OH

O 130

O

HO 47

51 1

HO

O 124

123

125

HO

O

O

O OH

OH

O

+

68

34

HO

126

4

32

O

O

OH 52

43

46

O

OH OH + + 33

COOH

OH 129

122 O

59

62

54 OH

CHO

O OH

Figure 19.203

O

O

127

OH

128

65

64

63

66

FIGURE 22.204  Metabolic pathways of 1,8-cineole (122), 1,4-cineole (131), phellandrene (62), and carvotanacetone (47) by microorganisms.

746

Handbook of Essential Oils OH OH

O

308

307

OH 303

OH 305

OH

302

OH 304 OH

306

OH OH 309

310

FIGURE 22.205  Metabolic pathways of myrcene (302) and citronellene (309) by rat and microorganisms.

OH

HO

OH

O O

454

OH

O

OH

452

OH 453

456

O

455

OH

OH

5

OH 204

FIGURE 22.206  Metabolic pathways of nopol (452) and nopol benzyl ether (455) by microorganisms.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

747

Pirt, 1971). Myrtenol (5) is further metabolized to myrtenal (6) and myrtenic acid (7). Verbenol (23) is further metabolized to verbenone (24), 7-hydroxyverbenone (25), 7-hydroxyverbenone (26), and 7-formyl verbanone (27). Thujone (28) is further metabolized to thujoyl alcohol (29), 1-hydroxythujone (30), and 1,3-dihydroxythujone (31). Second, β-pinene (1) is metabolized to pinocarveol (2) and pinocarvone (3) (Ganapathy and Bhattacharyya, unpublished data). Pinocarvone (3) is further metabolized to isopinocamphone (413), which is further hydroxylated to give 2-hydroxy-3-pinanone (414). Compound 414 is further metabolized to give pinane-2,3-diol (419), 2,5-dihydroxy- (415), and 2,9-dihydroxy-3-pinanone (416). Third, β-pinene (1) is metabolized to α-fenchol (11) and fenchone (12) (Dhavalikar et al., 1974), which are further metabolized to 6-hydroxy- (13) and 5-hydroxyfenchone (14), 5-oxofenchone (15), fenchone-9-al (17), fenchone9-oic acid (18) via 9-hydroxyfenchone (16), 2,3-fencholide (21), and 1,2-fencholide (22) (Pfrunder and Tamm, 1969a,b; Yamamoto et  al., 1984; Christensen and Tuthill, 1985; Miyazawa et  al., 1990a,b). Fenchol (12) is also metabolized to 9-hydroxyfenchol (466) and 7-hydroxyfenchol (467), 6-hydroxyfenchol (349), and 6,7-dihydroxyfenchol (412). Fourth, β-pinene (1) is metabolized via fenchoquinone (19) to 2-hydroxyfenchone (20) (Pfrunder and Tamm, 1969b; Gibbon et al., 1972). Fifth, β-pinene (1) is metabolized to α-terpineol (34) via pinyl cation (32) and 1-p-menthene-8cation (33) (Saeki and Hashimoto, 1968, 1971; Hosler, 1969; Hayashi et al., 1972). α-Terpineol (34) is metabolized to 8,9-epoxy-1-p-methanol (58) via diepoxide (57), terpin hydrate (60), and oleuropeic acid (204) (Saeki and Hashimoto, 1968, 1971; Shukla et al., 1968; Shukla and Bhattacharyya, 1968; Hosler, 1969; Hungund et al., 1970; Hayashi et al., 1972). As shown in Figure 22.202, oleuropeic acid (204) is formed from linalool (206) and α-terpineol (34) via 204, 205, and 213 as intermediates (Shukla et  al., 1968; Shukla and Bhattacharyya, 1968; Hungund et  al., 1970) and degradatively metabolized to perillic acid (82), 2-hydroxy-8-p-menthen-7-oic acid (84), 2-oxo-8-p-menthen-7-oic acid (84), 2-oxo-8-p-menthen-1-oic acid (85), and β-isopropyl pimelic acid (86) (Shukla et al., 1968; Shukla and Bhattacharyya, 1968; Hungund et al., 1970). Oleuropeic acid (204) is also formed from β-pinene (1) via α-terpineol (34) as the intermediate (Noma et al., 2001). Oleuropeic acid (204) is also formed from myrtenol (5) by rearrangement reaction by A. niger TBUYN-2 (Noma and Asakawa, 2005b). Finally, β-pinene (1) is metabolized to borneol (36) and camphor (37) via two cations (32 and 35) and to 1-p-menthene (62) via two cations (33 and 59) (Shukla and Bhattacharyya, 1968). 1-p-Menthene (62) is metabolized to phellandric acid (65) via phellandrol (63) and phellandral (64), which is further degradatively metabolized through 246–251 and 89 to water and carbon dioxide as shown in Figure 22.204 (Shukla et al., 1968). Phellandral (64) is easily reduced to give phellandrol (63) by Euglena sp. and Dunaliella sp. (Noma et al., 1984, 1986, 1991a,b, 1992b). Furthermore, 1-p-menthene (62) is metabolized to 1-p-menthen-2-ol (46) and p-menthane-1,2-diol (54) as shown in Figure 22.204. Perillic acid (82) is easily formed from perillaldehyde (78) and perillyl alcohol (74) (Figure 22.19) (Swamy et al., 1965; Dhavalikar and Bhattacharyya, 1966; Dhavalikar et al., 1966; Ballal et al., 1967; Shima et al., 1972; Kayahara et al., 1973). α-Terpineol (34) is also formed from linalool (206). α-Pinene (4) is metabolized by five pathways as follows: First, α-pinene (4) is metabolized to myrtenol (5), myrtenal (6), and myrtenoic acid (7) (Shukla et al., 1968; Shukla and Bhattacharyya, 1968; Hungund et al., 1970; Ganapathy and Bhattacharyya, unpublished results). Myrtenal (6) is easily reduced to myrtenol (5) by Euglena and Dunaliella spp., (Noma et al., 1991a,b, 1992b). Myrtanol (8) is metabolized to 3-hydroxy- (9) and 4-hydroxymyrtanol (10) (Miyazawa et al., 1994a). Second, α-pinene (4) is metabolized to verbenol (23), verbenone (24), 7-hydroxyverbenone (25), and verbanone-7-al (27) (Bhattacharyya et al., 1960; Prema and Bhattachayya, 1962; Miyazawa et al., 1991a). Third, α-pinene (4) is metabolized to thujone (28), thujoyl alcohol (29), 1-hydroxy(30), and 1,3-dihydroxythujone (31) (Gibbon and Pirt, 1971; Miyazawa et al., 1992a; Noma, 2000). Fourth, α-pinene (4) is metabolized to sobrerol (43) and carvotanacetol (46, 1-p-menthen-2-ol) via α-pinene epoxide (38) and two cations (41 and 42). Sobrerol (43) is further metabolized to 8-hydroxycarvotanacetone (44, carvonhydrate), 8-hydrocarvomenthone (45), and p-menthane2,8-diol (50) (Prema and Bhattachayya, 1962; Noma, 2007). In the metabolism of sobrerol (43), 8-hydroxycarvotanacetone (44), and 8-hydroxycarvomenthone (45) by A. niger TBUYN-2, the

748

Handbook of Essential Oils

formation of p-menthane-2,8-diol (50) is very highly enantio- and diastereoselective in the reduction of 8-hydroxycarvomenthone (Noma, 2007). 8-Hydroxycarvotanacetone (44) is a common metabolite from sobrerol (43) and carvotanacetone (47). That is, carvotanacetone (47) is metabolized to carvomenthone (48), carvomenthol (49), 8-hydroxycarvomenthol (50), 5-hydroxycarvotanacetone (51), 8-hydroxycarvotanacetone (44), 5-hydroxycarvomenthone (52), and 2,3-lactone (56) (Gibbon and Pirt, 1971; Gibbon et  al., 1972; Noma et  al., 1974a, 1985c, 1988a). Carvomenthone (48) is metabolized to 45, 8-hydroxycarvomenthol (50), 1-hydroxycarvomenthone (53), and p-menthane1,2-diol (54) (Noma et  al., 1985b, 1988a). Compound 52 is metabolized to 6-hydroxymenthol (139), which is the common metabolite of menthol (137) (see Figure 22.201). Carvomenthol (49) is metabolized to 8-hydroxycarvomenthol (50) and p-menthane-2, 9-diol (55). Finally, α-pinene (4) is metabolized to borneol (36) and camphor (37) via 32 and 35 and to phellandrene (62) via 32 and two cations (33 and 59) as mentioned in the metabolism of β-pinene (1). Carvotanacetone (47) is also metabolized degradatively to 3,4-dimethylvaleric acid (177) via 56 and 158–163 as shown in Figure 22.201 (Gibbon and Pirt, 1971; Gibbon et al., 1972). α-Pinene (4) is also metabolized to 2-(4-methyl3-cyclohexenylidene)-propionic acid (67) (Figure 22.199). Metabolic pathways of limonene (68), perillyl alcohol (74), carvone (93), isopiperitenone (111), and piperitenone (112) are summarized in Figure 22.199. Limonene (68) is metabolized by eight pathways. That is, limonene (68) is converted into α-terpineol (34) (Savithiry et al., 1997), limonene1,2-epoxide (69), 1-p-menthene-9-oic acid (70), perillyl alcohol (74), 1-p-menthene-8,9-diol (79), isopiperitenol (110), p-mentha-1,8-diene-4-ol (80, 4-terpineol), and carveol (81) (Dhavalikar and Bhattacharyya, 1966; Dhavalikar et al., 1966; Bowen, 1975; Noma et al., 1982, 1992b; Miyazawa et al., 1983; Savithiry et al., 1997; Van der Werf et al., 1997, 1998b; Van der Werf and de Bont, 1998a). Dihydrocarvone (101), limonene-1,2-diol (71), 1-hydroxy-8-p-menthene-2-one (72), and p-mentha-2,8-diene-1-ol (73) are formed from limonene (68) via limonene epoxide (69) as intermediate. Limonene (68) is also metabolized via carveol (78), limonene-1,2-diol (71), carvone (93), 1-p-menthene-6,9-diol (95), 8,9-dihydroxy-1-p-menthene (90), α-terpineol (34), 2α-hydroxy1,8-cineole (125), and p-menthane-1,2,8-triol (334). Bottrospicatol (92) and 5-hydroxycarveol (94) are formed from cis-carveol by S. bottropensis SY-2-1 (Noma et al., 1982; Nishimura et al., 1983a; Noma and Asakawa, 1992; Noma et al., 1982). Carvyl acetate and carvyl propionate (both are shown as 106) are hydrolyzed enantio- and diastereoselectively to carveol (78) (Oritani and Yamashita, 1980; Noma, 2000). Carvone (93) is metabolized through four pathways as follows: First, carvone (93) is reduced to carveol (81) (Noma, 1980). Second, it is epoxidized to carvone-8,9-epoxide (96), which is further metabolized to dihydrocarvone-8,9-epoxide (97), dihydrocarveol-8,9-epoxide (103), and menthane-2,8,9-triol (104) (Noma et  al., 1980; Noma and Nishimura, 1982; Noma, 2000). Third, 93 is hydroxylated to 5-hydroxycarvone (98), 5-hydroxydihydrocarvone (99), and 5-hydroxydihydrocarveol (100) (Noma and Nishimura, 1982). Dihydrocarvone (101) is metabolized to 8-p-menthene-1,2-diol (71) via l-hydroxydihydrocarvone (72), 10-hydroxydihydrocarvone (106), and dihydrocarveol (102), which is metabolized to 10-hydroxydihydrocarveol (107), p-menthane-2,8diol (50), dihydrocarveol-8,9-epoxide (100), p-menthane-2,8,9-triol (104), and dihydrobottrospicatol (105) (Noma et al., 1985a,b). In the biotransformation of (+)-carvone by plant pathogenic fungi, A. niger Tiegh, isodihydrocarvone (101) was metabolized to 4-hydroxyisodihydrocarvone (378) and 1-hydroxyisodihydrocarvone (72) (Noma and Asakawa, 2008). 8,9-Epoxydihydrocarvyl acetate (109) is hydrolyzed to 8,9-epoxydihydrocarveol (103). Perillyl alcohol (74) is metabolized through three pathways to shisool (75), shisool-8,9-epoxide (76), perillyl alcohol-8,9-epoxide (77), perillaldehyde (78), perillic acid (82), and 4,9-dihydroxy-1-p-­menthen-7-oic acid (83). Perillic acid (82) is metabolized degradatively to 84–89 as shown in Figure 22.200 (Swamy et al., 1965; Dhavalikar and Bhattacharyya, 1966; Dhavalikar et al., 1966; Ballal et al., 1967; Shukla et al., 1968; Shukla and Bhattacharyya, 1968; Shima et al., 1972; Kayahara et al., 1973; Hungund et al., 1970). Isopiperitenol (110) is reduced to isopiperitenone (111), which is metabolized to 3-hydroxy- (115), 4-hydroxy- (116), 7-hydroxy- (113), and 10-hydroxyisopiperitenone (114) and piperitenone (112). Compounds isopiperitenone (111) and piperitenone (112) are isomerized to each other (Noma et al.,

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

749

1992c). Furthermore, piperitenone (112) is metabolized to 8-hydroxypiperitone (157) and 5-hydroxy(117) and 7-hydroxypiperitenone (118). Pulegone (119) is metabolized to 112, 8-hydroxymenthenone (121), and 8,9-dehydromenthenone (120). Metabolic pathways of menthol (137), menthone (149), thymol (179), and carvacrol methyl ether (202) are summarized in Figure 22.201. Menthol (137) is generally hydroxylated to give 1-hydroxy(138), 2-hydroxy- (140), 4-hydroxy- (141), 6-hydroxy- (139), 7-hydroxy- (143), 8-hydroxy- (142), and 9-hydroxymenthol (144) and 1,8-dihydroxy- (146) and 7,8-dihydroxymenthol (148) (Asakawa et al., 1991; Takahashi et al., 1994; Van der Werf et al., 1997). Racemic menthyl acetate and menthyl chloroacetate are hydrolyzed asymmetrically by an esterase of microorganisms (Brit Patent, 1970; Moroe et al., 1971; Watanabe and Inagaki, 1977a,b). Menthone (149) is reductively metabolized to 137 and oxidatively metabolized to 3,7-dimethyl-6-hydroxyoctanoic acid (152), 3,7-dimethyl6-oxooctanoic acid (153), 2-methyl-2,5-oxidoheptenoic acid (154), 1-hydroxymenthone (150), piperitone (156), 7-hydroxymenthone (151), menthone-7-al (163), menthone-7-oic acid (164), and 7-carboxylmenthol (165) (Sawamura et al., 1974). Compound 156 is metabolized to menthone-1,2diol (155) (Miyazawa et al., 1991e, 1992d, e). Compound 148 is metabolized to 6-hydroxy- (158), 8-hydroxy- (157), and 9-hydroxypiperitone (159), piperitone-7-al (160), 7-hydroxypiperitone (161), and piperitone-7-oic acid (162) (Lassak et al., 1973). Compound 149 is also formed from menthenone (148) by hydrogenation (Mukherjee et al., 1973), which is metabolized to 6-hydroxymenthenone (181, 6-hydroxy-4-p-menthen-3-one). 6-Hydroxymenthenone (181) is also formed from thymol (179) via 6-hydroxythymol (180). 6-Hydroxythymol (180) is degradatively metabolized through 182–185 to 186, 187, and 89 (Mukherjee et al., 1974). Piperitone oxide (166) is metabolized to 1-hydroxymenthone (150) and 4-hydroxypiperitone (167) (Lassak et al., 1973; Miyazawa et al., 1991d,e). Piperitenone oxide (168) is metabolized to 1-hydroxymenthone (150), 1-hydroxypulegone (169), and 2,3-secop-menthalacetone-3-en-1-ol (170) (Lassak et al., 1973; Miyazawa et al., 1991e). p-Cymene (178) is metabolized to 8-hydroxy- (188) and 9-hydroxy-p-cymene (189), 2-(4-methylphenyl)-propanoic acid (190), thymol (179), and cumin alcohol (192), which is further converted degradatively to p-cumin aldehyde (193), cumic acid (194), cis-2,3-dihydroxy-2,3-dihydro-p-cumic acid (195), 2,3-dihydroxy-p-cumic acid (197), 198–200, and 89 as shown in Figure 22.3 (Chamberlain and Dagley, 1968; DeFrank and Ribbons, 1977a,b; Hudlicky et al., 1999; Noma, 2000). Compound 197 is also metabolized to 4-methyl-2-oxopentanoic acid (201) (DeFrank and Ribbons, 1977a). Compound 193 is easily metabolized to 192 and 194 (Noma et al., 1991b. Carvacrol methyl ether (202) is easily metabolized to 7-hydroxycarvacrol methyl ether (203) (Noma, 2000). Metabolic pathways of borneol (36), camphor (37), phellandral (64), linalool (206), and p-menthane (252) are summarized in Figure 22.202. Borneol (36) is formed from β-pinene (1), α-pinene (4), 34, bornyl acetate (226), and camphene (229), and it is metabolized to 36; 3-hydroxy- (243), 5-hydroxy(235), 6-hydroxy- (228), and 9-hydroxycamphor (225); and 1,2-campholide (23). Compound 228 is degradatively metabolized to 6-oxocamphor (229) and 230–234, whereas 237 is also degradatively metabolized to 6-hydroxy-1,2-campholide (238), 6-oxo-1,2-campholide (239), and 240–242. 5-Hydroxycamphor (235) is metabolized to 238, 5-oxocamphor (236), and 6-oxo-1,2-campholide (239). Compound 243 is also metabolized to camphorquinone (244) and 2-hydroxyepicamphor (245) (Bradshaw et al., 1959; Conrad et al., 1961, 1965a,b; Gunsalus et al., 1965; Chapman et al., 1966; Hartline and Gunsalus, 1971; Oritani and Yamashita, 1974). 1-p-Menthene (62) is formed 1 and 4 via three cations (32, 33, and 59) and metabolized to phellandrol (63) (Noma et al., 1991b) and p-menthane-1,2-diol (54). Compound 63 is metabolized to phellandral (64) and 7-hydroxyp-menthane (66). Compound 64 is furthermore metabolized degradatively to CO2 and water via phellandric acid (65), 246–251, and 89 (Dhavalikar and Bhattacharyya, 1966; Dhavalikar et al., 1966; Ballal et al., 1967; Shukla et al., 1968; Shukla and Bhattacharyya, 1968; Hungund et al., 1970). Compound 64 is also easily reduced to phellandrol (63) (Noma et al., 1991b, 1992a). p-Menthane (252) is metabolized via 1-hydroxy-p-menthane (253) to p-menthane-1,9-diol (254) and p-menthane1,7-diol (255) (Tsukamoto et al., 1974, 1975; Noma et al., 1990). Compound 255 is degradatively metabolized via 256–248 to CO2 and water through the degradation pathway of phellandric acid (65,

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Handbook of Essential Oils

246–251, and 89) as aforementioned. Linalool (206) is metabolized to α-terpineol (34), camphor (37), oleuropeic acid (61), 2-methyl-6-hydroxy-6-carboxy-2,7-octadiene (211), 2-methyl-6-hydroxy6-carboxy-7-octene (199), 5-methyl-5-vinyltetrahydro-2-furanol (215), 5-methyl-5-vinyltetrahydro2-furanone (216), and malonyl ester (218). 1-Hydroxylinalool (219) is metabolized degradatively to 2,6-dimethyl-6-hydroxy-trans-2,7-octadienoic acid (220), 4-methyl-trans-3, 5-hexadienoic acid (221), 4-methyl-trans-3,5-hexadienoic acid (222), 4-methyl-trans-2-hexenoic acid (223), and isobutyric acid (224). Compound 206 is furthermore metabolized via 213 to 61, 82, and 84–86 as shown in Figure 22.2 (Mizutani et al., 1971; Murakami et al., 1973; Madyastha et al., 1977; Rama Devi and Bhattacharyya, 1977a,b; Rama Devi et al., 1977; David and Veschambre, 1985; Miyazawa et al., 1994a,b). Metabolic pathways of citronellol (258), citronellal (261), geraniol (271), nerol (272), citral [neral (275) and geranial (276)], and myrcene (302) are summarized in Figure 22.203 (Seubert and Fass, 1964a,b; Hayashi et al., 1968; Rama Devi and Bhattacharyya, 1977a,b). Geraniol (271) is formed from citronellol (258), nerol (272), linalool (206), and geranyl acetate (270) and metabolized through 10 pathways. That is, compound 271 is hydrogenated to give citronellol (258), which is metabolized to 2,8-dihydroxy-2,6-dimethyl octane (260) via 6,7-epoxycitronellol (268), isopulegol (267), limonene (68), 3,7-dimethyloctane-1,8-diol (266) via 3,7-dimethyl-6-octene-1,8-diol (265), 267, citronellal (261), dihydrocitronellol (259), and nerol (272). Citronellyl acetate (269) and isopulegyl acetate (301) are hydrolyzed to citronellol (258) and isopulegol (267), respectively. Compound 261 is metabolized via pulegol (263) and isopulegol (267) to menthol (137). Compound 271 and 272 are isomerized to each other. Compound 272 is metabolized to 271, 258, citronellic acid (262), nerol-6-,7-epoxide (273), and neral (275). Compound 272 is metabolized to neric acid (277). Compounds 275 and 276 are isomerized to each other. Compound 276 is completely decomposed to CO2 and water via geranic acid (278), 2,6-dimethyl-8-hydroxy-7-oxo-2-octene (279), 6-methyl-5-heptenoic acid (280), 7-methyl-3-oxo-6-octenoic acid (283), 6-methyl-5-heptenoic acid (284), 4-methyl-3-heptenoic acid (284), 4-methyl-3-pentenoic acid (285), and 3-methyl-2-butenoic acid (286). Furthermore, compound 271 is metabolized via 3-hydroxymethyl-2,6-octadiene-1-ol (287), 3-formyl-2,6-octadiene-1-ol (288), and 3-carboxy-2,6-octadiene-1-ol (289) to 3-(4-methyl-3-pentenyl)-3-butenolide (290). Geraniol (271) is also metabolized to 3,7-dimethyl-2,3-dihydroxy-6-octen-1-ol (292), 3,7-dimethyl2-oxo-3-hydroxy-6-octen-1-ol (293), 2-methyl-6-oxo-2-heptene (294), 6-methyl-5-hepten-2-ol (298), 2-methyl-2-heptene-6-one-1-ol (295), and 2-methyl-γ-butyrolactone (296). Furthermore, 271 is metabolized to 7-methyl-3-oxo-6-octanoic acid (299), 7-hydroxymethyl-3-methyl-2,6-octadien-1-ol (291), 6,7-epoxygeraniol (274), 3,7-dimethyl-2,6-octadiene-1,8-diol (300), and 3,7-dimethyloctane1,8-diol (266). Metabolic pathways of 1,8-cineole (122), 1,4-cineole (131), phellandrene (62), carvotanacetone (47), and carvone (93) by microorganisms are summarized in Figure 22.204. 1,8-Cineole (112) is biotransformed to 2-hydroxy- (125), 3-hydroxy- (123), and 9-hydroxy-1,8cineole (127), 2-oxo- (126) and 3-oxo-1,8-cineole (124), lactone [128, (R)-5,5-dimethyl-4-(3′oxobutyl)-4,5-dihydrofuran-2-(3H)-one], and p-hydroxytoluene (129) (MacRae et al., 1979; Nishimura et al., 1982; Noma and Sakai, 1984). 2-Hydroxy-1,8-cineole (125) is further converted into 2-oxo1,8-cineole (126), 1,8-cineole-2-malonyl ester (130), sobrerol (43), and 8-hydroxycarvotanacetone (44) (Miyazawa et al., 1995b). 2-Hydroxy-1,8-cineole (125) and 2-oxo-1,8-cineole (126) are also biodegraded to sobrerol (43) and 8-hydroxycarvotanacetone (44), respectively. 2-Hydroxy-1,8-cineole (125) was esterified to give malonyl ester (130). 2-Hydroxy-1,8-cineole (125) was formed from limonene (68) by citrus pathogenic fungi P. digitatum (Noma and Asakawa, 2007b). 1,4-Cineole (131) is metabolized to 2-hydroxy- (132), 3-hydroxy- (133), 8-hydroxy- (134), and 9-hydroxy-1,4cineole (135). Compound 132 is also esterified to malonyl ester (136) as well as 125 (Miyazawa et  al., 1995b). Terpinen-4-ol (342) is metabolized to 2-hydroxy-1,4-cineole (132), 2-hydroxy(372) and 7-hydroxyterpinene-4-ol (342), and p-menthane-1,2,4-triol (371) (Abraham et al., 1986; Kumagae and Miyazawa, 1999; Noma and Asakawa, 2007a). Phellandrene (62) is metabolized to carvotanacetol (46) and phellandrol (63). Carvotanacetol (46) is further metabolized through the

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751

metabolism of carvotanacetone (47). Phelandrol (63) is also metabolized to give phellandral (64), phellandric acid (65), and 7-hydroxy-p-menthane (66). Phellandric acid (65) is completely degraded to carbon dioxide and water as shown in Figure 22.202. Metabolic pathways of myrcene (302) and citronellene (309) by microorganisms and insects are summarized in Figure 22.205. β-Myrcene (302) was metabolized with D. gossypina ATCC 10936 (Abraham et al., 1985) to the diol (303) and a side product (304). β-Myrcene (302) was metabolized with G. applanatum, P. flabellatus, and P. sajor-caju to myrcenol (305) (2-methyl-6-methylene-7octen-2-ol) and 306 (Busmann and Berger, 1994). β-Myrcene (302) was converted by common cutworm larvae, S. litura, to give myrcene-3, (10)-glycol (308) via myrcene-3,(10)-epoxide (307) (Miyazawa et  al., 1998). Citronellene (309) was metabolized by cutworm S. litura to give 3,7-dimethyl-6-octene-1,2-diol (310) (Takeuchi and Miyazawa, 2005). Myrcene (302) is metabolized to two kinds of diols (303 and 304), myrcenol (305), and ocimene (306) (Seubert and Fass, 1964a,b; Abraham et al., 1985). Citronellene (309) was metabolized to (310) by S. litura (Takeuchi and Miyazawa, 2005). Metabolic pathways of nopol (452) and nopol benzyl ether (455) by microorganisms are summarized in Figure 22.206. Nopol (452) is metabolized mainly to 7-hydroxyethyl-α-terpineol (453) by rearrangement reaction and 3-oxoverbenone (454) as minor metabolite by Aspergillus spp. including A. niger TBUYN-2 (Noma and Asakawa, 2006b,c). Myrtenol (5) is also metabolized to oleuropeic alcohol (204) by rearrangement reaction. However, nopol benzyl ether (455) was easily metabolized to 3-oxoverbenone (454) and 3-oxonopol-2′,4′-dihydroxybenzylether (456) as main metabolites without rearrangement reaction (Noma and Asakawa, 2006c).

22.5.2 Microbial Transformation of Terpenoids as Unit Reaction Microbiological oxidation and reduction patterns of terpenoids and related compounds by fungi belonging to Aspergillus spp. containing A. niger TBUYN-2 and E. gracilis Z. are summarized in Tables 22.18 and 22.19, respectively. Dehydrogenation of secondary alcohols to ketones, hydroxylation of both nonallylic and allylic carbons, oxidation of olefins to form diols and triols via epoxides, reduction of both saturated and α,β-unsaturated ketones, and hydrogenation of olefin conjugated with the carbonyl group were the characteristic features in the biotransformation of terpenoids and related compounds by Aspergillus spp. Compound names: 1, β-pinene; 2, pinocarveol; 3, pinocarvone; 4, α-pinene; 5, myrtenol; 6, myrtenal; 7, myrtenoic acid; 8, myrtanol; 9, 3-hydroxymyrtenol; 10, 4-hydroxymyrtanol; 11, α-fenchol; 12, fenchone; 13, 6-hydroxyfenchone; 14, 5-hydroxyfenchone; 15, 5-oxofenchone; 16, 9-hydroxyfenchone; 17, fenchone-9-al; 18, fenchone-9-oic acid; 19, fenchoquinone; 20, 2-hydroxyfenchone; 21, 2,3-fencholide; 22, 1,2-fencholide; 23, verbenol; 24, verbenone; 25, 7-hydroxyverbenone; 26, 7-hydroxyverbenone; 27, verbanone-4-al; 28, thujone; 29, thujoyl alcohol; 30, 1-hydroxythujone; 31, 1,3-dihydroxythujone; 32, pinyl cation; 33, 1-p-menthene-8-cation; 34, α-terpineol; 35, bornyl cation; 36, borneol; 37, camphor; 38, α-pinene epoxide; 39, isonovalal; 40, novalal; 41, 2-hydroxypinyl cation; 42, 6-hydroxy-1-p-menthene-8-cation; 43, trans-sobrerol; 44, 8-hydroxycarvotanacetone; (carvonehydrate); 45, 8-hydrocarvomenthone; 46, 1-p-menthen-2-ol; 47, carvotanacetone; 48, carvomenthone; 49, carvomenthol; 50, 8-hydroxycarvomenthol; 51, 5-hydroxycarvotanacetone; 52, 5-hydroxycarvomenthone; 53, 1-hydroxycarvomenthone; 54, p-menthane-1,2-diol; 55, p-menthane2,9-diol; 56, 2,3-lactone; 57, diepoxide; 58, 8,9-epoxy-1-p-methanol; 59, 1-p-menthene-4-cation; 60, terpin hydrate; 61, oleuropeic acid (8-hydroxyperillic acid); 62, 1-p-menthene; 63, phellandrol; 64, phellandral; 65, phellandric acid; 66, 7-hydroxy-p-menthane; 67, 2-(4-methyl-3-cyclohexenylidene)propionic acid; 68, limonene; 69, limonene-1,2-epoxide; 70, 1-p-menthene-9-oic acid; 71, limonene1,2-diol; 72, 1-hydroxy-8-p-menthene-2-one; 73, 1-hydroxy-p-menth-2,8-diene; 74, perillyl alcohol; 75, shisool; 76, shisool-8,9-epoxide; 77, perillyl alcohol-8,9-epoxide; 78, perillaldehyde; 79, 1-p-­ menthene-8,9-diol; 80, 4-hydroxy-p-menth-1,8-diene (4-terpineol); 81, carveol; 82, perillic acid; 83,

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TABLE 22.18 Microbiological Oxidation and Reduction Patterns of Monoterpenoids by Aspergillus niger TBUYN-2 Microbiological Oxidation Oxidation of Alcohols

Oxidation of Primary Alcohols to Aldehydes and Acids Oxidation of secondary alcohols to ketones

(−)-trans-Carveol (81a′), (+)-transcarveol (81a), (−)-cis-carveol (81b′), (+)-cis-carveol (81b), 2α-hydroxy-1,8 cineole (125b), 3α-hydroxy-1,8cineole (123b), 3β-hydroxy-1,8cineole (123a)

Oxidation of aldehydes to acids

Oxidation of olefins

Hydroxylation of nonallylic carbon

(−)-Isodihydrocarvone (101c′), (−)-carvotanacetone (47′), (+)-carvotanacetone (47), cis-pmenthane (252), 1α-hydroxy-pmenthane (253), 1,8-cineole (122), 1,4-cineole (131), (+)-fenchone (12), (−)-fenchone (12′), (−)-menthol (137b′), (+)-menthol (137b), (−)-neomenthol (137a), (+)-neomenthol (137a), (+)-isomenthol (137c)

Hydroxylation of allylic carbon

(−)-Isodihydrocarvone (101b), (+)-neodihydrocarveol (102a′), (−)-dihydrocarveol (102b′), (+)-dihydrocarveol (102b), (+)-limonene (68), (−)-limonene (68′)

Formation of epoxides and oxides Formation of diols

Formation of triols

(+)-Neodihydrocarveol (102a′), (+)-dihydrocarveol (102b), (−)-dihydrocarveol (102b′), (+)-limonene (68), (−)-limonene (68′) (+)-Neodihydrocarveol (102a′)

Lactonization Microbiological Reduction Reduction of aldehydes to alcohols Reduction of ketones to alcohols

Hydrogenation of olefins

Reduction of saturated ketones

Reduction of α,β-unsaturated ketones Hydrogenation of olefin conjugated with carbonyl group Hydrogenation of olefin not conjugated with a carbonyl group

(+)-Dihydrocarvone (101a′), (−)-isodihydrocarvone (101b), (+)-carvomenthone (48a′), (−)-isocarvomenthone (48b)

(−)-Carvone (93′), (+)-carvone (93), (−)-carvotanacetone (47′), (+)-carvotanacetone (47)

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TABLE 22.19 Microbiological Oxidation, Reduction, and Other Reaction Patterns of Monoterpenoids by Euglena gracilis Z Microbiological Oxidation Oxidation of Alcohols

Oxidation of Primary Alcohols to Aldehydes and Acids Oxidation of secondary alcohols to ketones

Oxidation of aldehydes to acids

Hydroxylation Oxidation of olefins

(−)-trans-Carveol (81a′), (+)-cis-carveol (81b), (+)-isoborneol (36b).a Myrtenal (6), myrtanal, (−)-perillaldehyde (78), trans- and cis-1,2-dihydroperillaldehydes (261a and 261b), (−)-phellandral (64), trans- and cis-tetrahydroperillaldehydes, cuminaldehyde (193), (+)- and (−)-citronellal (261 and 261′).b

Hydroxylation of nonallylic carbon Hydroxylation of allylic carbon Formation of epoxides and oxides Formation of diols Formation of triols

(+)-Limonene (68), (−)-limonene (68′).

(+)- and (−)-Neodihydrocarveol (102a′ and a), (−)- and(+)-dihydrocarveol (102b′ and b), (+)- and (−)-isodihydrocarveol (102c′ and c), (+)- and (−)-neoisodihydrocarveol (102d′ and d).

Lactonization Microbiological Reduction Reduction of aldehydes to alcohols

Reduction of ketones to alcohols

Reduction of terpene aldehydes to terpene alcohols

Reduction of aromatic and related aldehydes to alcohols Reduction of aliphatic aldehydes to alcohols Reduction of saturated ketones

Reduction of α,β-unsaturated ketones Hydrogenation of olefins Hydrogenation of olefin conjugated with carbonyl group

Myrtenal (6), myrtanal, (−)-perillaldehyde (78), trans- and cis-1,2-dihydroperillaldehydes (261a and 261b), phellandral (64), trans- and cis-1,2-dihydroperillaldehydes (261a and 261b), trans- and cis-tetrahydroperillaldehydes, cuminaldehyde (193), citral (275 and 276), (+)- (261) and (−)-citronellal (261′).

(+)-Dihydrocarvone (101a′), (−)-isodihydrocarvone (101b), (+)-carvomenthone (48a′), (−)-isocarvomenthone (48b), (+)-dihydrocarvone-8,9-epoxides (97a′), (+)-isodihydrocarvone-8,9-epoxides (97b′), (−)-dihydrocarvone-8,9-epoxides (97a). (−)-Carvone (93′), (+)-carvone (93), (−)-carvotanacetone (47′), (+)-carvotanacetone (47), (−)-carvone-8,9epoxides (96′), (+)-carvone-8,9-epoxides (96).

Hydrogenation of olefin not conjugated with a carbonyl group (Continued)

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TABLE 22.19 (Continued) Microbiological Oxidation, Reduction, and Other Reaction Patterns of Monoterpenoids by Euglena gracilis Z Hydrolysis Hydrolysis

Hydrolysis of ester

Hydration

Hydration of C  =  C bond in isopropenyl group to tertiary alcohol

(+)-trans- and cis-Carvyl acetates (108a and b), (−)-cis-carvyl acetate (108b′), (−)-cis-carvyl propionate, geranyl acetate (270). Hydration (+)-Neodihydrocarveol (102a′), (−)-dihydrocarveol (102b′), (+)-isodihydrocarveol (102c′), (+)-neoisodihydrocarveol (102d′), (−)-neodihydrocarveol (102a), (+)-dihydrocarveol (102b), (−)-isodihydrocarveol (102c), (−)-neoisodihydrocarveol (102d), trans- and cis-shisools (75a and 75b).

Isomerization Isomerization a b

Geraniol (271), nerol (272).

Diastereo- and enantioselective dehydrogenation is observed in carveol, borneol, and isoborneol. Acids were obtained as minor products.

4,9-dihydroxy-1-p-menthene-7-oic acid; 84, 2-hydroxy-8-p-menthen-7-oic acid; 85, 2-oxo-8-pmenthen-7-oic acid; 86, β-isopropyl pimelic acid; 87, isopropenylglutaric acid; 88, isobutenoic acid; 89, isobutyric acid; 90, 1-p-menthene-8,9-diol; 91, carveol-8,9-epoxide; 92, bottrospicatol; 93, carvone; 94, 5-hydroxycarveol; 95, 1-p-menthene-6,9-diol; 96, carvone-8,9-epoxide; 97, dihydrocarvone-8,9-epoxide; 98, 5-hydroxycarvone; 99, 5-hydroxydihydrocarvone; 100, 5-hydroxydihydrocarveol; 101, dihydrocarvone; 102, dihydrocarveol; 103, dihydrocarveol-8,9epoxide; 104, p-menthane-2,8,9-triol; 105, dihydrobottrospicatol; 106, 10-hydroxydihydrocarvone; 107, 10-hydroxydihydrocarveol; 108, carvyl acetate and carvyl propionate; 109, 8,9-epoxydihydrocarvyl acetate; 110, isopiperitenol; 111, isopiperitenone; 112, piperitenone; 113, 7-hydroxyisopiperitenone; 114, 10-hydroxyisopiperitenone; 115, 4-hydroxyisopiperitenone; 116, 5-hydroxyisopiperitenone; 117, 5-hydroxypiperitenone; 118, 7-hydroxypiperitenone; 119, pulegone; 120, 8,9-dehydromenthenone; 121, 8-hydroxymenthenone; 122, 1,8-cineole; 123, 3-hydroxy-1,8-cineole; 124, 3-oxo-1,8-cineole; 125, 2-hydroxy-1,8-cineole; 126, 2-oxo-1,8-cineole; 127, 9-hydroxy-1,8-cineole; 128, lactone (R)-5,5dimethyl-4-(3′-oxobutyl)-4,5-dihydrofuran-2-(3H)-one; 129, p-hydroxytoluene; 130, 1,8-cineole-2malonyl ester; 131, 1,4-cineole; 132, 2-hydroxy-1,4-cineole; 133, 3-hydroxy-1,4-cineole; 134, 8-hydroxy-1,4-cineole; 135, 9-hydroxy-1, 4-cineole; 136, 1,4-cineole-2-malonyl ester; 137, menthol; 138, 1-hydroxymenthol; 139, 6-hydroxymenthol; 140, 2-hydroxymenthol; 141, 4-hydroxymenthol; 142, 8-hydroxymenthol; 143, 7-hydroxymenthol; 144, 9-hydroxymenthol; 145, 7,8-dihydroxymenthol; 146, 1,8-dihydroxymenthol; 147, 3-p-menthene; 148, menthenone; 149, menthone; 150, 1-hydroxymenthone; 151, 7-hydroxymenthone; 152, 3,7-dimethyl-6-hydroxyoctanoic acid; 153, 3,7-dimethyl-6-oxooctanoic acid; 154, 2-methyl-2,5-oxidoheptenoic acid; 155, menthone-1,2-diol; 156, piperitone; 157, 8-hydroxypiperitone; 158, 6-hydroxypiperitone; 159, 9-hydroxypiperitone; 160, piperitone-7-al; 161, 7-hydroxypiperitone; 162, piperitone-7-oic acid; 163, menthone-7-al; 164, menthone-7-oic acid; 165, 7-carboxylmenthol; 166, piperitone oxide; 167, 4-hydroxypiperitone; 168, piperitenone oxide; 169, 1-hydroxypulegone; 170, 2,3-seco-p-methylacetone-3-en-1-ol; 171, 2-methyl5-­i sopropyl-2,5-hexadienoic acid; 172, 2,5,6-trimethyl-2,4-heptadienoic acid; 173,

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2,5,6-trimethyl-3-heptenoic acid; 174, 2,5,6-trimethyl-2-heptenoic acid; 175, 3-hydroxy-2,5,6trimethyl-3-heptanoic acid; 176, 3-oxo-2,5,6-trimethyl-3-heptanoic acid; 177, 3,4-dimethylvaleric acid; 178, p-cymene; 179, thymol; 180, 6-hydroxythymol 181, 6-hydroxymenthenone, 6-hydroxy-4p-menthen-3-one; 182, 3-hydroxythymo-1,4-quinol; 183, 2-hydroxythymoquionone; 184, 2,4-dimethyl-6-oxo-3,7-dimethyl-2,4-octadienoic acid; 185, 2,4,6-trioxo-3,7-dimethyl octanoic acid; 186, 2-oxobutanoic acid; 187, acetic acid; 188, 8-hydroxy-p-cymene; 189, 9-hydroxy-p-cymene; 190, 2-(4-methylphenyl)-propanoic acid; 191, carvacrol; 192, cumin alcohol; 193, p-cumin aldehyde; 194, cumic acid; 195, cis-2,3-dihydroxy-2, 3-dihydro-p-cumic acid; 196, 3-hydroxycumic acid; 197, 2,3-dihydroxy-p-cumic acid; 198, 2-hydroxy-6-oxo-7-methyl-2,4-octadien-1,3-dioic acid; 199, 2-methyl-6-hydroxy-6-carboxy-7-octene; 201, 4-methyl-2-oxopentanoic acid; 202, carvacrol methyl ether; 203, 7-hydroxycarvacrol methyl ether; 204, 8-hydroxyperillyl alcohol; 205, 8-hydroxyperillaldehyde; 206, linalool; 207, linalyl-6-cation; 208, linalyl-8-cation; 209, 6-hydroxymethyl linalool; 210, linalool-6-al; 211, 2-methyl-6-hydroxy-6-carboxy-2,7-octadiene; 212, 2-methyl-6-hydroxy-7-octen-6-oic acid; 213, phellandric acid-8-cation; 214, 2,3-epoxylinalool; 215, 5-methyl-5-vinyltetrahydro-2-furanol; 216, 5-methyl-5-vinyltetrahydro-2-­f uranone; 217, 2,2,6-trimethyl-3-hydroxy-6-vinyltetrahydropyrane; 218, malonyl ester; 219, 1-hydroxylinalool (3,7-dimethyl-1,6-octadiene-8-ol); 220, 2,6-dimethyl-6-hydroxy-trans-2,7-­octadienoic acid; 221, 4-methyl-trans-3,5-hexadienoic acid; 222, 4-methyl-trans-3,5-hexadienoic acid; 223, 4-methyl-trans2-hexenoic acid; 224, isobutyric acid; 225, 9-hydroxycamphor; 226, bornyl acetate; 228, 6-hydroxycamphor; 229, 6-oxocamphor; 230, 4-carboxymethyl-2,3,3-­trimethylcyclopentanone; 231, 4-carboxymethyl-3,5,5-trimethyltetrahydro-2-pyrone; 232, isohydroxycamphoric acid; 233, isoketocamphoric acid; 234, 3,4,4-trimethyl-5-oxo-trans-2-hexenoic acid; 235, 5-hydroxycamphor; 236, 5-oxocamphor; 237, 238, 6-hydroxy-1, 2-campholide; 239, 6-oxo-1,2-campholide; 240, 5-carboxymethyl-3,4,4-trimethyl-2-cyclopentenone; 241, 6-carboxymethyl-4,5,5-trimethyl-5,6dihydro-2-pyrone; 242, 5-hydroxy-3,4,4-trimethyl-2-heptene-1,7-dioic acid; 243, 3-hydroxycamphor; 244, camphorquinone; 245, 2-hydroxyepicamphor; 246, 2-hydroxy-p-menthan-7-oic acid; 247, 2-oxop-menthan-7-oic acid; 248, 3-isopropylheptane-1,7-dioic acid; 249, 3-isopropylpentane-1,5-dioic acid; 250, 4-methyl-3-oxopentanoic acid; 251, methylisopropyl ketone; 252, p-menthane; 253, 1-hydroxyp-menthane; 254, p-menthane-1,9-diol; 255, p-menthane-1,7-diol; 256, 1-hydroxy-p-menthene-7-al; 257, 1-hydroxy-p-menthene-7-oic acid; 258, citronellol; 259, dihydrocitronellol; 260, 2,8-dihydroxy2,6-dimethyl octane; 261, citronellal; 262, citronellic acid; 263, pulegol; 264, 7-hydroxymethyl-6octene-3-ol; 265, 3,7-dimethyl-6-octane-1,8-diol; 266, 3,7-dimethyloctane-1,8-diol; 267, isopulegol; 268, 6,7-epoxycitronellol; 269, citronellyl acetate; 270, geranyl acetate; 271, geraniol; 272, nerol 273, nerol-6,7-epoxide; 274, 6,7-epoxygeraniol; 275, neral; 276, geranial; 277, neric acid; 278, geranic acid; 279, 2,6-dimethyl-8-hydroxy-7-oxo-2-octene; 280, 6-methyl-5-heptenoic acid; 281, 7-methyl-3carboxymethyl-2,6-octadiene-1-oic acid; 282, 7-methyl-3-hydroxy-3-carboxymethyl-6-octen-1-oic acid; 283, 7-methyl-3-oxo-6-octenoic acid; 284, 6-methyl-5-heptenoic acid; 284, 4-methyl-3-heptenoic acid; 285, 4-methyl-3-pentenoic acid; 286, 3-methyl-2-butenoic acid; 287, 3-hydroxymethyl-2,6octadiene-1-ol; 288, 3-formyl-2,6-octadiene-1-ol; 289, 3-carboxy-2,6-octadiene-1-ol; 290, 3-(4-methyl3-pentenyl)-3-butenolide; 291, 7-hydroxymethyl-3-methyl-2,6-octadien-1-ol; 292, 3,7-dimethyl-2,3-dihydroxy-6-octen-1-ol; 293, 3,7-dimethyl-2-​oxo-6-octene-1,3-diol; 294, 6-methyl5-hepten-2-one; 295, 6-methyl-7-hydroxy-5-heptene-2-one; 296, 2-methyl-γ-butyrolactone; 297, 6-methyl-5-heptenoic acid; 298, 6-methyl-5-hepten-2-ol; 299, 7-methyl-3-oxo-6-octanoic acid; 300, 3,7-dimethyl-2,6-octadiene-1,8-diol; 301, isopulegyl acetate; 302, myrcene; 303, 2-methyl-6methylene-7-octene-2,3-diol; 304, 6-methylene-7-octene-2,3-diol; 305, myrcenol; 306, ocimene; 307, myrcene-3,(10)-epoxide; 308, myrcene-3,(10)-glycol; 309, (−)-citronellene; 309′, (+)-citronellene; 310, (3R)-3,7-dimethyl-6-octene-1,2-diol; 310′, (3S)-3,7-dimethyl-6-octene-1,2-diol; 311, (E)-3,7-dimethyl5-octene-1,7-diol; 312, trans-rose oxide; 313, cis-rose oxide; 314, (2Z,5E)-3,7-dimethyl-2,5-octadiene1 ,7 - d i o l ; 315, ( Z ) -3 ,7 - d i m e t h y l - 2 ,7 - o c t a d i e n e -1 , 6 - d i o l ; 316 , (2E,6Z)-2,6-dimethyl-2,6-octadiene-1,8-diol; 317, a cyclization product; 318, (2E,5E)-3,7-dimethyl2,5-octadiene-1,7-diol; 319, (E)-3,7-dimethyl-2,7-octadiene-1,6-diol; 320, 8-hydroxynerol; 321,

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10-hydroxynerol; 322, 1-hydroxy-3,7-dimethyl-2E,6E-octadienal; 323, 1-hydroxy-3-,7-dimethyl2E,6E-octadienoic acid; 324, 3,9-epoxy-p-menth-1-ene; 325, tetrahydrolinalool; 326, 3,7-dimethyloctane-3,5-diol; 327, 3,7-dimethyloctane-3,7-diol; 328, 3,7-dimethyloctane-3,8-diol; 329, dihydromyrcenol; 330, 1,2-epoxydihydromyrcenol; 331, 3β-hydroxydihydromyrcenol; 332, dihydromyrcenyl acetate; 333, 1,2-dihydroxydihydromyrcenyl acetate; 334, (+)-p-menthane-1-β,2α,8triol; 335, α-pinene-1,2-epoxide; 336, 3-carene; 337, 3-carene-1,2-epoxide; 338, (1R)-transisolimonene; 338, (1R,4R)-p-menth-2-ene-8,9-diol; 339, (1R,4R)-p-menth-2-ene-8,9-diol; 340, α-terpinene; 341, α-terpinene-7-oic acid; 342, (−)-terpinen-4-ol; 343, p-menthane-1,2,4-triol; 344, γ-terpinene; 345, γ-terpinene-7-oic acid; 346, terpinolene; 347, (1R)-8-hydroxy-3-p-menthen-2-one; 348, (1R)-1,8-dihydroxy-3-p-menthen-2-one; 349, 6β-hydroxyfenchol; 350, 5β-hydroxyfenchol (a,5β,b,5α); 351, terpinolene-1,2-trans-diol; 352, terpinolene-4,8-diol; 353, terpinolene-9-ol; 354, terpinolene-10-ol; 355, α-phellandrene; 356, α-phellandrene-7-oic acid; 357, terpinolene-7-oic acid; 358, thymoquinone; 359, 1,2-dihydroperillaldehyde; 360, 1,2-dihydroperillic acid; 361, 8-hydroxy1,2-dihydroperillyl alcohol; 362, tetrahydroperillaldehyde (a trans, b cis); 363, tetrahydroperillic acid (a trans, b cis); 364, (−)-menthol monoglucoside; 365, (+)-menthol diglucoside; 366, (+)-isopulegol; 367, 7-hydroxy-(+)-isopulegol; 368, 10-hydroxy-(+)-isopulegol; 369, 1,2-epoxy-α-terpineol; 370, bornane-2,8-diol; 371, p-menthane-1α,2β,4β-triol; 372, 1-p-menthene-4β,6-diol; 373, 1-p-menthene4a,7-diol; 374, (+)-bottrospicatal; 375, 1-p-menthene-2β,8,9-triol; 376, (−)-perillyl alcohol monoglucoside; 377, (−)-perillyl alcohol diglucoside; 378, 4α-hydroxy-(−)-isodihydrocarvone; 379, 2-methyl-2-cyclohexenone; 380, 2-cyclohexenone; 381, 3-methyl-2-cyclohexenone; 382, 2-methylcyclohexanone; 383, 2-methylcyclohexanol (a, trans; b, cis); 384, 4-hydroxycarvone; 385, 2-ethyl-2-cyclohexenone; 386, 2-ethylcyclohexenone (a1R) (b1S); 387, 1-hydroxypulegone; 388, 5-hydroxypulegone; 389, 8-hydroxymenthone; 390, 10-hydroxy-(−)-carvone; 391, 1,5,5-trimethylcyclopentane-1,4-dicarboxylic acid; 392, 4β-hydroxy-(−)-menthone; 393, 1α,4βdihydroxy-(−)-menthone; 394, 7-hydroxy-9-carboxymenthone; 395, 7-hydroxy-1,8-cineole; 396, methyl ester of 2α-hydroxy-1,8-cineole; 397, ethyl ester of 2α-hydroxy-1,8-cineole; 398, n-propyl ester of 2α-hydroxy-1,8-cineole; 399, n-butyl ester of 2α-hydroxy-1,8-cineole; 400, isopropyl ester of 2α-hydroxy-1,8-cineole; 401, tertiary butyl ester of 2α-hydroxy-1,8-cineole; 402, methylisopropyl ester of 2α-hydroxy-1,8-cineole; 403, methyl tertiary butyl ester of 2α-hydroxy-1,8-cineole; 404, 2α-hydroxy-1,8-cineole monoglucoside (404 and 404’); 405, 2α-hydroxy-1,8-cineole diglucoside; 406, p-menthane-1,4-diol; 407, 1-p-menthene-4β,6-diol; 408, (−)-pinane-2α,3α-diol; 409, (−)-6β-hydroxypinene; 410, (−)-4α,5-dihydroxypinene; 411, (−)-4α-hydroxypinene-6-one; 412, 6β,7-dihydroxyfenchol; 413, 3-oxo-pinane; 414, 2α-hydroxy-3-pinanone; 415, 2α, 5-dihydroxy-3pinanone; 416, 2α,10-dihydroxy-3-pinanone; 417, trans-3-pinanol; 418, pinane-2α,3α-diol; 419, pinane-2α, 3α, 5-triol; 420, isopinocampheol (3-pinanol); 421, pinane-1,3α-diol; 422, pinane-3α,5diol; 423, pinane-3β,9-diol; 424, pinane-3β,4β,-diol; 425, 426, pinane-3α,4β-diol; 427, pinane-3α,9diol; 428, pinane-3α,6-diol; 429, p-menthane-2α,9-diol; 430, 2-methyl-3α-hydroxy-1-hydroxyisopropyl cyclohexane propane; 431, 5-hydroxy-3-pinanone; 432, 2α-methyl,3-(2-methyl-2-hydroxypropyl)cyclopenta-1β-ol; 433, 3-acetoxy-2α-pinanol; 434, 8-hydroxy-α-pinene; 435, 436, myrtanic acid; 437, camphene; 438, camphene glycol; 439, (+)-3-carene; 440, m-mentha-4,6-dien-8-ol; 441, m-cymen8-ol; 442, 3-carene-9-ol; 443, 3-carene-9-carboxylic acid; 444, 3-caren-10-ol-9-carboxylic acid; 445, 3-carene-9,10-dicarboxylic acid; 446, (−)-cis-carane; 447, dicarboxylic acid of (−)-cis-carane; 448, (−)-6β-hydroxypinene; 449, (−)-4α,5-dihydroxypinene; 450, (−)-4α-hydroxypinene-6-one; 451, 10-hydroxyverbenol; 452, (−)-nopol; 453, 7-hydroxymethyl-1-p-menthen-8-ol; 454, 3-oxonopol; 455, nopol benzyl ether; 456, 4-oxonopol-2′,4′-dihydroxybenzyl ether; 457, 7-hydroxymethyl-1-p-menthen8-ol benzyl ether; 458, piperitenol; 459, thymol methyl ether; 460, 7-hydroxythymol methyl ether; 461, 9-hydroxythymol methyl ether; 462, 1,8-cineol-9-oic acid; 463, 4-hydroxyphellandric acid; 464, 4-hydroxydihydrophellandric acid; 465, (+)-8-hydroxyfenchol; 466, (−)-9-hydroxyfenchol; 467, (+)-10-hydroxyfenchol, 468, 4α-hydroxy-6-oxo-α-pinene; 469, dihydrolinalyl acetate; 470, 3-hydroxycarvacrol; 471, 9-hydroxycarvacrol; 472, carvacrol-9-oic acid; 473, 8,9 dehydrocarvacrol; 474, 8-hydroxycarvacrol; 475, 7-hydroxycarvacrol; 476, carvacrol-7-oic acid; 477,

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8,9-dihydroxycarvacrol; 478, 7,9-dihydroxycarvacrol methyl ether; 479, 7-hydroxythymol; 480, 9-hydroxythymol; 481, thymol-7-oic acid; 482, 7,9-dihydroxythymol; 483, thymol-9-oic acid; 484, (1R,2R,3S,4S,5R)-3,4-pinanediol.

REFERENCES Abraham, W.-R., H.-A. Arfmann, B. Stumpf, P. Washausen, and K. Kieslich, 1988. Microbial transformations of some terpenoids and natural compounds. In: Bioflavour’87. Analysis—Biochemistry—Biotechnology, P. Schreier, ed., pp. 399–414. Berlin, Germany: Walter de Gruyter and Co. Abraham, W.-R., H.M.R. Hoffmann, K. Kieslich, G. Reng, and B. Stumpf, 1985. Microbial transformation of some monoterpenes and sesquiterpenoids. In: Enzymes in Organic Synthesis, R. Porter and S. Clark, eds., Ciba Foundation Symposium 111, pp. 146–160. London, U.K.: Pitman Press. Abraham, W.-R., K. Kieslich, H. Reng, and B. Stumpf, 1984. Formation and production of 1,2-trans-glycols from various monoterpenes with 1-menthene skeleton by microbial transformations with Diplodia gossypina. In: Third European Congress on Biotechnology, Vol. 1, pp. 245–248. Verlag Chemie, Weinheim, Germany. Abraham, W.-R., B. Stumpf, and H.-A. Arfmann, 1990. Chiral intermediates by microbial epoxidations. J. Essent. Oil Res., 2: 251–257. Abraham, W.-R., B. Stumpf, and K. Kieslich, 1986. Microbial transformation of terpenoids with 1-p-menthene skeleton. Appl. Microbiol. Biotechnol., 24: 24–30. Amsterdam, D. 1997. Susceptibility testing of antimicrobials in liquid media. In: Antibiotics in Laboratory Medicine, V. Lorian, ed., 4th ed. Baltimore, MD: Williams & Wilkins, Maple Press. Asakawa, Y., H. Takahashi, M. Toyota, Y. and Noma, 1991. Biotransformation of monoterpenoids, (−)- and (+)-menthols, terpinolene and carvotanacetone by Aspergillus species. Phytochemistry, 30: 3981–3987. Asakawa, Y., M. Toyota, T. Ishida, T. Takemoto, 1983. Metabolites in rabbit urine after terpenoid administration. Proceedings of the 27th TEAC, pp. 254–256. Atta-ur-Rahman, M. Yaqoob, A. Farooq, S. Anjum, F. Asif, and M.I. Choudhary, 1998. Fungal transformation of (1R,2S,5R)-(−)-menthol by Cephalosporium aphidicola. J. Nat. Prod., 61: 1340–1342. Ausgulen, L.T., E. Solheim, and R.R. Scheline, 1987. Metabolism in rats of p-cymene derivatives: Carvacrol and thymol. Pharmacol. Toxicol., 61: 98–102. Babcka, J., J. Volf, J. Czchec, and P. Lebeda, 1956. Patent 56-9686b. Ballal, N.R., P.K. Bhattacharyya, and P.N. Rangachari, 1967. Microbiological transformation of terpenes. Part XIV. Purification and properties of perillyl alcohol dehydrogenase. Indian J. Biochem., 5: 1–6. Bauer, K., D. Garbe, and H. Surburg (eds.), 1990. Common Fragrance and Flavor Materials: Preparation, Properties and Uses. 2nd revised ed., 218pp. New York: VCH Publishers. Best, D.J. and K.J. Davis, 1988. Soap, Perfumery Cosmetics 4: 47. Best, D.J., N.C. Floyd, A. Magalhaes, A. Burfield, and P.M. Rhodes, 1987. Initial enzymatic steps in the degradation of alpha-pinene by Pseudomonas fluorescens Ncimb 11671. Biocatal. Biotransform., 1: 147–159. Bhattacharyya, P.K. and K. Ganapathy, 1965. Microbial transformation of terpenes. VI. Studies on the mechanism of some fungal hydroxylation reactions with the aid of model systems. Indian J. Biochem., 2: 137–145. Bhattacharyya, P.K., B.R. Prema, B.D. Kulkarni, and S.K. Pradhan, 1960. Microbiological transformation of terpenes: Hydroxylation of α-pinene. Nature, 187: 689–690. Bock, G., I. Benda, and P. Schreier, 1986. Biotransformation of linalool by Botrytis cinerea. J. Food Sci., 51: 659–662. Bock, G., I. Benda, and P. Schreier, 1988. Microbial transformation of geraniol and nerol by Botrytis cinerea. Appl. Microbiol. Biotechnol., 27: 351–357. Bouwmester, H.J., J.A.R. Davies, and H. Toxopeus, 1995. Enantiomeric composition of carvone, limonene, and carveols in seeds of dill and annual and biennial caraway varieties. J. Agric. Food Chem., 43: 3057–3064. Bowen, E.R., 1975. Potential by-products from microbial transformation of d-limonene. Proc. Fla. State Hort. Soc., 88: 304–308. Bradshaw, W.H., H.E. Conrad, E.J. Corey, I.C. Gunsalus, and D. Lednicer, 1959. Microbiological degradation of (+)-camphor. J. Am. Chem. Soc., 81: 5507. Brit Patent, 1970. No. 1,187,320. Brunerie, P., I. Benda, G. Bock, and P. Schreier, 1987a. Bioconversion of citronellol by Botrytis cinerea. Appl. Microbiol. Biotechnol., 27: 6–10. Brunerie, P., I. Benda, G. Bock, and P. Schreier, 1987b. Biotransformation of citral by Botrytis cinerea. Z. Naturforsch., 42C: 1097–1100.

758

Handbook of Essential Oils

Brunerie, P., I. Benda, G. Bock, and P. Schreier, 1988. Bioconversion of monoterpene alcohols and citral by Botrytis cinerea. In: Bioflavour’87. Analysis—Biochemistry—Biotechnology, P. Schreier, ed., pp. 435– 444. Berlin, Germany: Walter de Gruyter and Co. Busmann, D. and R.G. Berger, 1994. Conversion of myrcene by submerged cultured basidiomycetes. J. Biotechnol., 37: 39–43. Cadwallander, K.R. and R.J. Braddock, 1992. Enzymatic hydration of (4R)-(+)-limonene to (4R)-(+)-alphaterpineol. Dev. Food Sci., 29: 571–584. Cadwallander, K.R., R.J. Braddock, M.E. Parish, and D.P. Higgins, 1989. Bioconversion of (+)-limonene by Pseudomonas gladioli. J. Food Sci., 54: 1241–1245. Cantwell, S.G., E.P. Lau, D.S. Watt, and R.R. Fall, 1978. Biodegradation of acyclic isoprenoids by Pseudomonas species. J. Bacteriol., 135: 324–333. Chamberlain, E.M. and S. Dagley, 1968. The metabolism of thymol by a Pseudomonas. Biochem. J., 110: 755–763. Chapman, P.J., G. Meerman, and I.C. Gunsalus, 1965. The microbiological transformation of fenchone. Biochem. Biophys. Res. Commun., 20: 104–108. Chapman, P.J., G. Meerman, I.C. Gunsalus, R. Srinivasan, and K.L. Rinehart Jr., 1966. A new acyclic acid metabolite in camphor oxidation. J. Am. Chem. Soc., 88: 618–619. Christensen, M. and D.E. Tuthill, 1985. Aspergillus: An overview. In: Advances in Penicillium and Aspergillus Systematics, R.A. Samson and J.I. Pitt, eds., pp. 195–209. New York: Plenum Press. Conrad, H.E., R. DuBus, and I.C. Gunsalus, 1961. An enzyme system for cyclic ketone lactonization. Biochem. Biophys. Res. Commun., 6: 293–297. Conrad, H.E., R. DuBus, M.J. Mamtredt, and I.C. Gunsalus, 1965a. Mixed function oxidation II. Separation and properties of the enzymes catalyzing camphor ketolactonization. J. Biol. Chem., 240: 495–503. Conrad, H.E., K. Lieb, and I.C. Gunsalus, 1965b. Mixed function oxidation III. An electron transport complex in camphor ketolactonization. J. Biol. Chem., 240: 4029–4037. David, L. and H. Veschambre, 1984. Preparation d’oxydes de linalol par bioconversion. Tetrahadron Lett., 25: 543–546. David, L. and H. Veschambre, 1985. Oxidative cyclization of linalol by various microorganisms. Agric. Biol. Chem., 49: 1487–1489. DeFrank, J.J. and D.W. Ribbons, 1977a. p-Cymene pathway in Pseudomonas putida: Initial reactions. J. Bacteriol., 129: 1356–1364. DeFrank, J.J. and D.W. Ribbons, 1977b. p-Cymene pathway in Pseudomonas putida: Ring cleavage of 2,3-dihydroxy-p-cumate and subsequent reactions. J. Bacteriol., 129: 1365–1374. Demirci, F., 2000. Microbial transformation of bioactive monoterpenes. PhD thesis, pp. 1–137. Anadolu University, Eskisehir, Turkey. Demirci, F., H. Berber, and K.H.C. Baser, 2007. Biotransformation of p-cymene to thymoquinone. Book of Abstracts of the 38th ISEO, SL-1, p. 6. Demirci, F., N. Kirimer, B. Demirci, Y. Noma, and K.H.C. Baser, 2001. The biotransformation of thymol methyl ether by different fungi. Book of Abstracts of the XII Biotechnology Congress, p. 47. Demirci, F., Y. Noma, N. Kirimer, and K.H.C. Baser, 2004. Microbial transformation of (–)-carvone. Z.  Naturforsch., 59c: 389–392. Demyttenaere, J.C.R. and H.L. De Pooter, 1996. Biotransformation of geraniol and nerol by spores of Penicillium italicum. Phytochemistry, 41: 1079–1082. Demyttenaere, J.C.R. and H.L. De Pooter, 1998. Biotransformation of citral and nerol by spores of Penicillium digitatum. Flav. Fragr. J., 13: 173–176. Demyttenaere, J.C.R., M. del Carmen Herrera, and N. De Kimpe, 2000. Biotransformation of geraniol, nerol and citral by sporulated surface cultures of Aspergillus niger and Penicillium sp. Phytochemistry, 55: 363–373. Demyttenaere, J.C.R., I.E.I. Koninckx, and A. Meersman, 1996. Microbial production of bioflavours by fungal spores. In: Flavour Science. Recent Developments, A.J. Taylor and D.S. Mottram, eds., pp. 105–110. Cambridge, U.K.: The Royal Society of Chemistry. Demyttenaere, J.C.R. and H.M. Willemen, 1998. Biotransformation of linalool to furanoid and pyranoid linalool oxides by Aspergillus niger. Phytochemistry, 47: 1029–1036. Dhavalikar, R.S. and P.K. Bhattacharyya, 1966. Microbial transformation of terpenes. Part VIII. Fermentation of limonene by a soil Pseudomonad. Indian J. Biochem., 3: 144–157. Dhavalikar, R.S., A. Ehbrecht, and G. Albroscheit, 1974. Microbial transformations of terpenoids: β-pinene. Dragoco Rep., 3: 47–49. Dhavalikar, R.S., P.N. Rangachari, and P.K. Bhattacharyya, 1966. Microbial transformation of terpenes. Part IX. Pathways of degradation of limonene in a soil Pseudomonad. Indian J. Biochem., 3: 158–164.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

759

Farooq, A., M.I. Choudhary, S. Tahara, T.-U. Rahman, K.H.C. Baser, and F. Demirci, 2002. The microbial oxidation of (−)-p-pinene by Botrytis cinerea. Z. Naturforsch., 57c: 686–690. Fenaroli, G., 1975. Synthetic flavors. In: Fenaroli’s Handbook of Flavor Ingredients, T.E. Furia and N. Bellanca, eds., Vol. 2, pp. 6–563. Cleveland, OH: CRC Press. Flynn, T.M. and I.A. Southwell, 1979. 1,3–Dimethyl-2-oxabicyclo [2,2,2]-octane-3-methanol and 1,3-dimethyl2-oxabicyclo[2,2,2]-octane-3-carboxylic acid, urinary metabolites of 1,8-cineole. Aust. J. Chem., 32: 2093–2095. Ganapathy, K. and P.K. Bhattacharyya, unpublished data. Gibbon, G.H., N.F. Millis, and S.J. Pirt, 1972. Degradation of α-pinene by bacteria. Proc. IV IFS, Ferment. Technol. Today, pp. 609–612. Gibbon, G.H. and S.J. Pirt, 1971. The degradation of a-pinene by Pseudomonas PX 1. FEBS Lett., 18: 103–105. Gondai, T., M. Shimoda, and T. Hirata, 1999. Asymmetric reduction of enone compounds by Chlorella miniata. Proceedings of the 43rd TEAC, pp. 217–219. Griffiths, E.T., S.M. Bociek, P.C. Harries, R. Jeffcoat, D.J. Sissons, and P.W. Trudgill, 1987a. Bacterial metabolism of alpha-pinene: Pathway from alpha-pinene oxide to acyclic metabolites in Nocardia sp. strain P18.3. J. Bacteriol., 169: 4972–4979. Griffiths, E.T., P.C. Harries, R. Jeffcoat, and P.W. Trudgill, 1987b. Purification and properties of alpha-pinene oxide lyase from Nocardia sp. strain P18.3. J. Bacteriol., 169: 4980–4983. Gunsalus, I.C., P.J. Chapman, and J.-F. Kuo, 1965. Control of catabolic specificity and metabolism. Biochem. Biophys. Res. Commun., 18: 924–931. Gyoubu, K. and M. Miyazawa, 2005. Biotransformation of (+)- and (−)-fenchone by liver microsomes. Proceedings of the 49th TEAC, pp. 420–422. Gyoubu, K. and M. Miyazawa, 2006. Biotransformation of (+)- and (−)-camphor by liver microsome. Proceedings of the 50th TEAC, pp. 253–255. Hagiwara, Y. and M. Miyazawa, 2007. Biotransformation of cineole by the larvae of common cutworm (Spodoptera litura) as a biocatalyst. Proceedings of the 51st TEAC, pp. 304–305. Hagiwara, Y., H. Takeuchi, and M. Miyazawa, 2006. Biotransformation of (+)-and (−)-menthone by the larvae of common cutworm (Spodoptera litura) as a biocatalyst. Proceedings of the 50th TEAC, pp. 279–280. Hamada, H. and T. Furuya, 2000. Hydroxylation of monoterpenes by plant suspension cells. Proceedings of the 44th TEAC, pp. 167–168. Hamada, H., T. Furuya, and N. Nakajima, 1996. The hydroxylation and glycosylation by plant catalysts. Proceedings of the 40th TEAC, pp. 111–112. Hamada, H., T. Harada, and T. Furuya, 2001. Hydroxylation of monoterpenes by algae and plant suspension cells. Proceedings of the 45th TEAC, pp. 366–368. Hamada, H., M. Kaji, T. Hirata, T. Furuya, 2003. Enantioselective biotransformation of monoterpenes by Cyanobacterium. Proceedings of the 47th TEAC, pp. 162–163. Hamada, H., Y. Kondo, M. Kaji, and T. Furuta, 2002. Glycosylation of monoterpenes by plant suspension cells. Proceedings of the 46th TEAC, pp. 321–322. Hamada, H., A. Matsumoto, and J. Takimura, 2004. Biotransformation of acyclic monoterpenes by biocatalysts of plant cultured cells and Cyanobacterium. Proceedings of the 48th TEAC, pp. 393–395. Hamada, H. and H. Yasumune, 1995. The hydroxylation of monoterpenoids by plant cell biotransformation. Proceedings of the 39th TEAC, pp. 375–377. Hartline, R.A. and I.C. Gunsalus, 1971. Induction specificity and catabolite repression of the early enzymes in camphor degradation by Pseudomonas putida. J. Bacteriol., 106: 468–478. Hashimoto Y. and M. Miyazawa, 2001. Microbial resolution of esters of racemic 2-endo-hydroxy-1,8-cineole by Glomerella cingulata. Proceedings of the 45th TEAC, pp. 363–365. Hayashi, T., T. Kakimoto, H. Ueda, and C. Tatsumi, 1969. Microbiological conversion of terpenes. Part VI. Conversion of borneol. J. Agric. Chem. Soc. Jpn., 43: 583–587. Hayashi, T., H. Takashiba, S. Ogura, H. Ueda, and C. Tsutsumi, 1968. Nippon Nogei-Kagaku Kaishi, 42: 190–196. Hayashi, T., H. Takashiba, H. Ueda, and C. Tsutsumi, 1967. Nippon Nogei Kagaku Kaishi, 41(254): 79878g. Hayashi, T., S. Uedono, and C. Tatsumi, 1972. Conversion of a-terpineol to 8,9-epoxy-p-menthan-1-ol. Agric. Biol. Chem., 36: 690–691. Hirata, T., K. Shimoda, and T. Gondai, 2000. Asymmetric hydrogenation of the C–C double bond of enones with the reductases from Nicotiana tabacum. Chem. Lett., 29: 850–851. Hosler, P., 1969. U.S. Patent 3,458,399. Hudlicky, T., D. Gonzales, and D.T. Gibson, 1999. Enzymatic dihydroxylation of aromatics in enantioselective synthesis: Expanding asymmetric methodology. Aldrichim. Acta, 32: 35–61.

760

Handbook of Essential Oils

Hungund, B.L., P.K. Bhattachayya, and P.N. Rangachari, 1970. Methylisopropyl ketone from a terpene fermentation by the soil Pseudomonad, PL-strain. Indian J. Biochem., 7: 80–81. Ishida, T., Y. Asakawa, and T. Takemoto, 1981a. Metabolism of myrtenal, perillaldehyde and dehydroabietic acid in rabbits. Res. Bull. Hiroshima Inst. Technol., 15: 79–91. Ishida, T., Y. Asakawa, T. Takemoto, and T. Aratani, 1979. Terpenoid biotransformation in mammals. II. Biotransformation of dl-camphene. J. Pharm. Sci., 68: 928–930. Ishida, T., Y. Asakawa, T. Takemoto, and T. Aratani, 1981b. Terpenoids biotransformation in mammals. III. Biotransformation of α-pinene, (3–pinene, pinane, 3-carene, carane, myrcene, and p-cymene in rabbits. J. Pharm. Sci., 70: 406–415. Iscan, G., 2005. Unpublished data. Ismaili-Alaoui, M., B. Benjulali, D. Buisson, and R. Azerad, 1992. Biotransformation of terpenic compounds by fungi I. Metabolism of R-(+)-pulegone. Tetrahedron Lett., 33: 2349–2352. Janssens, L., H.L. De Pooter, N.M. Schamp, and E.J. Vandamme, 1992. Production of flavours by microorganisms. Process Biochem., 27: 195–215. Joglekar, S.S. and R.S. Dhavlikar, 1969. Microbial transformation of terpenoids. I. Identification of metabolites produced by a Pseudomonad from citronellal and citral. Appl. Microbiol., 18: 1084–1087. Kaji, M., H. Hamada, and T. Furuya, 2002. Biotransformation of monoterpenes by Cyanobacterium and plant suspension cells. Proceedings of the 46th TEAC, pp. 323–325. Kamino, F. and M. Miyazawa, 2005. Biotransformation of (+)-and (−)-pinane-2,3-diol using plant pathogenic fungus, Glomerella cingulata as a biocatalyst. Proceedings of the 49th TEAC, pp. 395–396. Kamino, F., Y. Noma, Y. Asakawa, and M. Miyazawa, 2004. Biotransformation of (1S,2S,3R,5S)-(+)-pinane2,3-diol using plant pathogenic fungus, Glomerella cingulata as a biocatalyst. Proceedings of the 48th TEAC, pp. 383–384. Kayahara, H., T. Hayashi, C. and Tatsumi, 1973. Microbiological conversion of (−)-perillaldehyde and p-mentha-1,3-dien-7-al. J. Ferment. Technol., 51: 254–259. Kieslich, K., W.-R. Abraham, and P. Washausen, 1985. Microbial transformations of terpenoids. In: Topics in Flavor Research, R.G. Berger, S. Nitz, and P. Schreier, eds., pp. 405–427. Marzling Hangenham, Germany: Eichborn. Koneman, E.W., S.D. Allen, W.M. Janda, P.C. Schreckenberger, and W.C. Winn, 1997. Color Atlas and Textbook of Diagnostic Microbiology, Philadelphia, PA: Lippincott-Raven Publishers. Kraidman, G., B.B. Mukherjee, and I.D. Hill, 1969. Conversion of limonene into an optically active isomer of α-terpineol by a Cladosporium species. Bacteriol. Proc., 69: 63. Krasnobajew, V., 1984. Terpenoids. In: Biotechnology, K. Kieslich, ed., Vol. 6a, pp. 97–125. Weinheim, Germany: Verlag Chemie. Kumagae, S. and M. Miyazawa, 1999. Biotransformation of p-menthanes using common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 43rd TEAC, pp. 389–390. Lassak, E.V., J.T. Pinkey, B.J. Ralph, T. Sheldon, and J.J.H. Simes, 1973. Extractives of fungi. V. Microbial transformation products of piperitone. Aust. J. Chem., 26: 845–854. Liu, W., A. Goswami, R.P. Steffek, R.L. Chemman, F.S. Sariaslani, J.J. Steffens, and J.P.N. Rosazza, 1988. Stereochemistry of microbiological hydroxylations of 1,4-cineole. J. Org. Chem., 53: 5700–5704. MacRae, I.C., V. Alberts, R.M. Carman, and I.M. Shaw, 1979. Products of 1,8-cineole oxidation by a Pseudomonad. Aust. J. Chem., 32: 917–922. Madyastha, K.M. 1984. Microbial transformations of acyclic monoterpenes. Proc. Indian Acad. Sci. (Chem. Sci.), 93: 677–686. Madyastha, K.M. and P.K. Bhattacharyya, 1968. Microbiological transformation of terpenes. Part XIII. Pathways for degradation of p-cymene in a soil pseudomonad (PL-strain). Indian J. Biochem., 5: 161–167. Madyastha, K.M., P.K. Bhattacharyya, and C.S. Vaidyanathan, 1977. Metabolism of a monoterpene alcohol, linalool, by a soil pseudomonad. Can. J. Microbiol., 23: 230–239. Madyastha, K.M. and N.S.R. Krishna Murthy, 1988a. Regiospecific hydroxylation of acyclic monoterpene alcohols by Aspergillus niger. Tetrahedron Lett., 29: 579–580. Madyastha, K.M. and N.S.R. Krishna Murthy, 1988b. Transformations of acetates of citronellol, geraniol, and linalool by Aspergillus niger: Regiospecific hydroxylation of citronellol by a cell-free system. Appl. Microbiol. Biotechnol., 28: 324–329. Madyastha, K.M. and V. Renganathan, 1983. Bio-degradation of acetates of geraniol, nerol and citronellol by P. incognita: Isolation and identification of metabolites. Indian J. Biochem. Biophys., 20: 136–140. Mattison, J.E., L.L. McDowell, and R.H. Baum, 1971. Cometabolism of selected monoterpenoids by fungi associated with monoterpenoid-containing plants. Bacteriol. Proc., 1971: 141.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

761

Miyamoto, Y. and M. Miyazawa, 2001. Biotransformation of (+)- and (−)-borneol by the larvae of common cutworm (Spodoptera litura) as a biocatalyst. Proceedings of the 45th TEAC, pp. 377–378. Miyazato, Y. and M. Miyazawa, 1999. Biotransformation of (+)- and (−)-a-fenchyl acetated using plant parasitic fungus, Glomerella cingulata as a biocatalyst. Proceedings of the 43rd TEAC, pp. 213–214. Miyazawa, M., H. Furuno, and H. Kameoka, 1992a. Biotransformation of thujone by plant pathogenic microorganism, Botrytis allii IFO 9430. Proceedings of the 36th TEAC, pp. 197–198. Miyazawa, M., H. Furuno, K. Nankai, and H. Kameoka, 1991a. Biotransformation of verbenone by plant pathogenic microorganism, Rhizoctonia solani. Proceedings of the 35th TEAC, pp. 274–275. Miyazawa, M., H. Huruno, and H. Kameoka, 1991b. Biotransformation of (+)-pulegone to (−)-1R-8-hydroxy4-p-menthen-3-one by Botrytis allii. Chem. Express, 6: 479–482. Miyazawa, M., H. Huruno, and H. Kameoka, 1991c. Chem. Express, 6: 873. Miyazawa, M., H. Kakita, M. Hyakumachi, and H. Kameoka, 1992b. Biotransformation of monoterpenoids having p-menthan-3-one skeleton by Rhizoctonia solani. Proceedings of the 36th TEAC, pp. 191–192. Miyazawa, M., H. Kakita, M. Hyakumachi, K. Umemoto, and H. Kameoka, 1991d. Microbiological conversion of piperitone oxide by plant pathogenic fungi Rhizoctonia solani. Proceedings of the 35th TEAC, pp. 276–277. Miyazawa, M., H. Kakita, M. Hyakumachi, K. Umemoto, and H. Kameoka, 1992c. Microbiological conversion of monoterpenoids containing p-menthan-3-one skeleton by plant pathogenic fungi Rhizoctonia solani. Proceedings of the 36th TEAC, pp. 193–194. Miyazawa, M., S. Kumagae, H. Kameoka, 1997a. Biotransformation of (−)-menthol and (+)-menthol by common cutworm Larvae, Spodoptera litura as a biocatalyst. Proceedings of the 41st TEAC, pp. 391–392. Miyazawa, M., S. Kumagae, H. Kameoka, 1997b. Biotransformation of (+)-trans myrtanol and (−)-transmyrtanol by common cutworm Larvae, Spodoptera litura as a biocatalyst. Proceedings of the 41st TEAC, pp. 389–390. Miyazawa, M. and Y. Miyamoto, 2004. Biotransformation of (+)-(1R, 2S)-fenchol by the larvae of common cutworm (Spodoptera litura). Tetrahadron, 60: 3091–3096. Miyazawa, M., T. Murata, and H. Kameoka, 1998. Biotransformation of β-myrcene by common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 42nd TEAC, pp. 123–125. Miyazawa, M., H. Nankai, and H. Kameoka, 1996a. Microbial oxidation of citronellol by Glomerella cingulata. Nat. Prod. Lett., 8: 303–305. Miyazawa, M., Y. Noma, K. Yamamoto, and H. Kameoka, 1983. Microbiological conversion of d- and l-limonene. Proceedings of the 27th TEAC, pp. 147–149. Miyazawa, M., Y. Noma, K. Yamamoto, and H. Kameoka, 1991e. Biotransformation of 1,4-cineole to 2-endohydroxy-1,4-cineole by Aspergillus niger. Chem. Express, 6: 771–774. Miyazawa, M., Y. Noma, K. Yamamoto, and H. Kameoka, 1992d. Biohydroxylation of 1,4-cineole to 9-hydroxy1,4-cineole by Aspergillus niger. Chem. Express, 7: 305–308. Miyazawa, M., Y. Noma, K. Yamamoto, and H. Kameoka, 1992e. Biotransformation of 1,4-cineole to 3-endohydroxy-1,4-cineole by Aspergillus niger. Chem. Express, 7: 125–128. Miyazawa, M., Y. Suzuki, and H. Kameoka, 1994a. Biotransformation of myrtanol by plant pathogenic microorganism, Glomerella cingulata. Proceedings of the 38th TEAC, pp. 96–97. Miyazawa, M., Y. Suzuki, and H. Kameoka, 1997c. Biotransformation of (−)- and (+)-isopinocampheol by three fungi. Phytochemistry, 45: 945–950. Miyazawa, M., T. Wada, and H. Kameoka, 1995a. Biotransformation of terpinene, limonene and α-phellandrene in common cutworm larvae, Spodoptera litura Fabricius. Proceedings of the 39th TEAC, pp. 362–363. Miyazawa, M., T. Wada, and H. Kameoka, 1996b. Biotransformation of p-menthanes using common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 40th TEAC, pp. 80–81. Miyazawa, M., K. Yamamoto, Y. Noma, and H. Kameoka, 1990a. Bioconversion of (+)-fenchone to (+)-6-endohydroxyfenchone by Aspergillus niger. Chem. Express, 5: 237–240. Miyazawa, M., K. Yamamoto, Y. Noma, and H. Kameoka, 1990b. Bioconversion of (+)-fenchone to 5-endohydroxyfenchone by Aspergillus niger. Chem. Express, 5: 407–410. Miyazawa, M., H. Yanagihara, and H. Kameoka, 1996c. Biotransformation of pinanes by common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 40th TEAC, pp. 84–85. Miyazawa, M., H. Yanahara, and H. Kameoka, 1995b. Biotransformation of trans-pinocarveol by plant pathogenic microorganism, Glomerella cingulata, and by the larvae of common cutworm, Spodoptera litura Fabricius. Proceedings of the 39th TEAC, pp. 360–361. Miyazawa, M., K. Yokote, and H. Kameoka, 1994b. Biotransformation of linalool oxide by plant pathogenic microorganisms, Glomerella cingulata. Proceedings of the 38th TEAC, pp. 101–102.

762

Handbook of Essential Oils

Miyazawa, M., K. Yokote, and H. Kameoka, 1995c. Biotransformation of 2-endo-hydroxy-1,4-cineole by plant pathogenic microorganism, Glomerella cingulata. Proceedings of the 39th TEAC, pp. 352–353. Mizutani, S., T. Hayashi, H. Ueda, and C. Tstsumom, 1971. Microbiological conversion of terpenes. Part IX. Conversion of linalool. Nippon Nogei Kagaku Kaishi, 45: 368–373. Moroe, T., S. Hattori, A. Komatsu, and Y. Yamaguchi, 1971. Japanese Patent, 2,036,875. No. 98195t. Mukherjee, B.B., G. Kraidman, and I.D. Hill, 1973. Synthesis of glycols by microbial transformation of some monocyclic terpenes. Appl. Microbiol., 25: 447–453. Mukherjee, B.B., G. Kraidman, I.D. Hill, 1974. Transformation of 1-menthene by a Cladosporium: Accumulation of β-isopropyl glutaric acid in the growth medium. Appl. Microbiol., 27: 1070–1074. Murakami, T., I. Ichimoto, and C. Tstsumom, 1973. Microbiological conversion of linalool. Nippon Nogei Kagaku Kaishi, 47: 699–703. Murata, T. and M. Miyazawa, 1999. Biotransformation of dihydromyrcenol by common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 43rd TEAC, pp. 393–394. Nakanishi, K. and M. Miyazawa, 2004. Biotransformation of (−)-menthone by human liver microsomes. Proceedings of the 48th TEAC, pp. 401–402. Nakanishi, K. and M. Miyazawa, 2005. Biotransformation of (+)- and (−)- menthol by liver microsomal humans and rats. Proceedings of the 49th TEAC, pp. 423–425. Nishimura, H., S. Hiramoto, and J. Mizutani, 1983a. Biological activity of bottrospicatol and related compounds produced by microbial transformation of (−)-cis-carveol towards plants. Proceedings of the 27th TEAC, pp. 107–109. Nishimura, H., S. Hiramoto, J. Mizutani, Y. Noma, A. Furusaki, and T. Matsumoto, 1983b. Structure and biological activity of bottrospicatol, a novel monoterpene produced by microbial transformation of (−)-cis-carveol. Agric. Biol. Chem., 47: 2697–2699. Nishimura, H. and Y. Noma, 1996. Microbial transformation of monoterpenes: Flavor and biological activity. In: Biotechnology for Improved Foods and Flavors, G.R. Takeoka, R. Teranishi, P.J. Williams, and A. Kobayashi, eds., ACS Symposium Series 637, pp. 173–187. Washington, DC: American Chemical Society. Nishimura, H., Y. Noma, and J. Mizutani, 1982. Eucalyptus as biomass. Novel compounds from microbial conversion of 1,8–cineole. Agric. Biol. Chem., 46: 2601–2604. Nishimura, H., D.M. Paton, and M. Calvin, 1980. Eucalyptus radiata oil as a renewable biomass. Agric. Biol. Chem., 44: 2495–2496. Noma, Y., 1976. Microbiological conversion of carvone. Biochemical reduction of terpenes, part VI. Ann. Res. Stud. Osaka Joshigakuen Junior College, 20: 33–47. Noma, Y., 1977. Conversion of the analogues of carvone and dihydrocarvone by Pseudomonas ovalis, strain 6-1, Biochemical reduction of terpenes, part VII. Nippon Nogeikagaku Kaishi, 51: 463–470. Noma, Y., 1979a. Conversion of (−)-carvone by Nocardia lurida A-0141 and Streptosporangium roseum IFO3776. Biochemical reduction of terpenes, part VIII. Nippon Nogeikagaku Kaishi, 53: 35–39. Noma, Y., 1979b. On the pattern of reaction mechanism of (+)-carvone conversion by actinomycetes. Biochemical reduction of terpenes, part X, Ann. Res. Stud. Osaka Joshigakuen Junior College, 23: 27–31. Noma, Y., 1980. Conversion of (−)-carvone by strains of Streptomyces, A-5–1 and Nocardia, 1-3-11. Agric. Biol. Chem., 44: 807–812. Noma, Y., 1984. Microbiological conversion of carvone, Kagaku to Seibutsu, 22: 742–746. Noma, Y., 1988. Formation of p-menthane-2,8-diols from (−)-dihydrocarveol and (+)-dihydrocarveol by Aspergillus spp. The Meeting of Kansai Division of The Agricultural and Chemical Society of Japan, Kagawa, Japan, p. 28. Noma, Y., 2000. unpublished data. Noma, Y., 2007. Microbial production of mosquitocidal (1R,2S,4R)-(+)-menthane- 2,8-diol. In: Aromatic Plants from Asia their Chemistry and Application in Food and Therapy, L. Jiarovetz, N.X. Dung, and V.K. Varshney, eds., pp. 169–186. Dehradun, Uttarakhand: Har Krishan Bhalla & Sons. Noma, Y., E. Akehi, N. Miki, and Y. Asakawa, 1992a. Biotransformation of terpene aldehyde, aromatic aldehydes and related compounds by Dunaliella tertiolecta. Phytochemistry, 31: 515–517. Noma, Y. and Y. Asakawa, 1992. Enantio- and diastereoselectivity in the biotransformation of carveols by Euglena gracilis Z. Phytochem., 31: 2009–2011. Noma, Y. and Y. Asakawa, 1995. Aspergillus spp.: Biotransformation of terpenoids and related compounds. In: Biotechnology in Agriculture and Forestry, Vol. 33. Medicinal and Aromatic Plants VIII, Y.P.S. Bajaj, ed., pp. 62–96. Berlin, Germany: Springer. Noma, Y. and Y. Asakawa, 1998. Euglena gracilis Z: Biotransformation of terpenoids and related compounds. In: Biotechnology in Agriculture and Forestry, Vol. 41. Medicinal and Aromatic Plants X, Y.P.S. Bajaj, ed., pp. 194–237. Berlin Heidelberg, Germany: Springer.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

763

Noma, Y. and Y. Asakawa, 2005a. New metabolic pathways of β-pinene and related compounds by Aspergillus niger. Book of Abstracts of the 36th ISEO, p. 32. Noma, Y. and Y. Asakawa, 2005b. Microbial transformation of (−)-myrtenol and (−)-nopol. Proceedings of the 49th TEAC, pp. 78–80. Noma, Y. and Y. Asakawa, 2006a. Biotransformation of (+)-limonene and related compounds by Citrus pathogenic fungi. Proceedings of the 50th TEAC, pp. 431–433. Noma, Y. and Y. Asakawa, 2006b. Biotransformation of β-pinene, myrtenol, nopol and nopol benzyl ether by Aspergillus niger TBUYN-2. Book of Abstracts of the 37th ISEO, p. 144. Noma, Y. and Y. Asakawa, 2006c. Microbial transformation of (−)-nopol benzyl ether. Proceedings of the 50th TEAC, pp. 434–436. Noma, Y. and Y. Asakawa, 2007a. Biotransformation of limonene and related compounds by newly isolated low temperature grown citrus pathogenic fungi and red yeast. Book of Abstracts of the 38th ISEO, p. 7. Noma, Y. and Y. Asakawa, 2007b. Microbial transformation of limonene and related compounds. Proceedings of the 51st TEAC, pp. 299–301. Noma, Y. and Y. Asakawa, 2008. New metabolic pathways of (+)-carvone by Citrus pathogenic Aspergillus niger Tiegh CBAYN and A. niger TBUYN-2. Proceedings of the 52nd TEAC, pp. 206–208. Noma, Y., M. Furusawa, T. Hashimoto, and Y. Asakawa, 2002. Stereoselective formation of (1R, 2S, 4R)-(+)p-menthane-2,8-diol from α-pinene. Book of Abstracts of the 33rd ISEO, p. 142. Noma, Y., M. Furusawa, T. Hashimoto, and Y. Asakawa, 2004. Biotransformation of (+)- and (−)-3-pinanone by Aspergillus niger. Proceedings of the 48th TEAC, pp. 390–392. Noma, Y., T. Hashimoto, S. Uehara, and Y. Asakawa, 2009. unpublished data. Noma, Y., T. Higata, T. Hirata, Y. Tanaka, T. Hashimoto, and Y. Asakawa, 1995. Biotransformation of [6-2H]-(−)-carvone by Aspergillus niger, Euglena gracilis Z and Dunaliella tertiolecta. Proceedings of the 39th TEAC, pp. 367–368. Noma, Y., K. Hirata, and Y. Asakawa, 1996. Biotransformation of 1,8-cineole. Why do the biotransformed 2α- and 3α-hydroxy-1,8-cineole by Aspergillus niger have no optical activity? Proceedings of the 40th TEAC, pp. 89–91. Noma, Y. and M. Iwami, 1994. Separation and identification of terpene convertible actinomycetes: S. bottropensis SY-2-1, S. ikutamanensis Ya-2–1 and S. humidus Tu-1. Bull. Tokushima Bunri Univ., 47: 99–110. Noma, Y., F. Kamino, T. Hashimoto, and Y. Asakawa, 2003. Biotransformation of (+)- and (−)-pinane-2,3-diol and related compounds by Aspergillus niger. Proceedings of the 47th TEAC, pp. 91–93. Noma, Y., K. Matsueda, I. Maruyama, and Y. Asakawa, 1997. Biotransformation of terpenoids and related compounds by Chlorella species. Proceedings of the 41st TEAC, pp. 227–229. Noma, Y., N. Miki, E. Akehi, E. Manabe, and Y. Asakawa, 1991a. Biotransformation of monoterpenes by photosynthetic marine algae, Dunaliella tertiolecta. Proceedings of the 35th TEAC, pp. 112–114. Noma, Y., M. Miyazawa, K. Yamamoto, H. Kameoka, T. Inagaki, and H. Sakai, 1984. Microbiological conversion of perillaldehyde. Biotransformation of l- and dl-perillaldehyde by Streptomyces ikutamanensis, Ya-2-1. Proceedings of the 28th TEAC, pp. 174–176. Noma, Y. and H. Nishimura, 1980. Microbiological transformation of 1,8-cineole. Oxidative products from 1,8-cineole by S. bottropensis, SY-2-1. Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, p. 28. Noma, Y. and H. Nishimura, 1981. Microbiological transformation of 1,8-cineole. Production of 3β-hydroxy1,8-cineole from 1,8-cineole by S. ikutamanensis, Ya-2-1. Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, p. 196. Noma, Y. and H. Nishimura, 1982. Biotransformation of carvone. 4. Biotransformation of (+)-carvone by Streptomyces bottropensis, SY-2-1. Proceedings of the 26th TEAC, pp. 156–159. Noma, Y. and H. Nishimura, 1983a. Biotransformation of (−)-carvone and (+)-carvone by S. ikutamanensis Ya-2-1. Book of Abstracts of the Annual Meeting of Agricultural and Biological Chemical Society, p. 390. Noma, Y. and H. Nishimura, 1983b. Biotransformation of carvone. 5. Microbiological transformation of dihydrocarvones and dihydrocarveols. Proceedings of the 27th TEAC, pp. 302–305. Noma, Y. and H. Nishimura, 1984. Microbiological conversion of carveol. Biotransformation of (−)-cis-carveol and (+)-cis-carveol by S. bottropensis, Sy-2-1. Proceedings of the 28th TEAC, pp. 171–173. Noma, Y. and H. Nishimura, 1987. Bottrospicatols, novel monoterpenes produced on conversion of (−)- and (+)-cis-carveol by Streptomyces. Agric. Biol. Chem., 51: 1845–1849. Noma, Y., H. Nishimura, S. Hiramoto, M. Iwami, and C. Tstsumi, 1982. A new compound, (4R, 6R)-(+)-6,8oxidomenth-1-en-9-ol produced by microbial conversion of (−)-cis-carveol. Agric. Biol. Chem., 46: 2871–2872.

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Noma, Y., H. Nishimura, and C. Tatsumi, 1980. Biotransformation of carveol by Actinomycetes. 1. Biotransformation of (−)-cis-carveol and (−)-trans-carveol by Streptomyces bottropensis, SY-2-1. Proceedings of the 24th TEAC, pp. 67–70. Noma, Y. and S. Nonomura, 1974. Conversion of (−)-carvone and (+)-carvone by a strain of Aspergillus niger. Agric. Biol. Chem., 38: 741–744. Noma, Y., S. Nonomura, and H. Sakai, 1974a. Conversion of (−)-carvotanacetone and (+)-carvotanacetone by Pseudomonas ovalis, strain 6-1. Agric. Biol. Chem., 38: 1637–1642. Noma, Y., S. Nonomura, and H. Sakai, 1975. Epimerization of (−)-isodihydrocarvone to (−)-dihydrocarvone by Pseudomonas fragi IFO 3458. Agric. Biol. Chem., 39: 437–441. Noma, Y., S. Nonomura, H. Ueda, H. Sakai, and C. Tstusmi, 1974b. Microbial transformation of carvone. Proceedings of the 18th TEAC, pp. 20–23. Noma, Y., S. Nonomura, H. Ueda, and C. Tatsumi, 1974c. Conversion of (+)-carvone by Pseudomonas ovalis, strain 6-1(1). Agric. Biol. Chem., 38: 735–740. Noma, Y. and H. Sakai, 1984. Investigation of the conversion of (−)-perillyl alcohol, 1,8–cineole, (+)-carvone and (−)-carvone by rare actinomycetes. Ann. Res. Stud. Osaka Joshigakuen Junior College, 28: 7–18. Noma, Y., A. Sogo, S. Miki, N. Fujii, T. Hashimoto, and Y. Asakwawa, 1992b. Biotransformation of terpenoids and related compounds. Proceedings of the 36th TEAC, pp. 199–201. Noma, Y., H. Takahashi, and Y. Asakawa, 1989. Microbiological conversion of menthol. Biotransformation of (+)-menthol by a strain of Aspergillus niger. Proceedings of the 33rd TEAC, pp. 124–126. Noma, Y., H. Takahashi, and Y. Asakawa, 1990. Microbiological conversion of p-menthane 1. Formation of p-menthane-1,9-diol from p-menthane by a strain of Aspergillus niger. Proceedings of the 34th TEAC, pp. 253–255. Noma, Y., H. Takahashi, and Y. Asakawa, 1991b. Biotransformation of terpene aldehyde by Euglena gracilis. Z. Phytochem., 30: 1147–1151. Noma, Y., H. Takahashi, and Y. Asakawa, 1993. Formation of 8 kinds of p-menthane-2,8-diols from carvone and related compounds by Euglena gracilis Z. Biotransformation of monoterpenes by photosynthetic microorganisms. Part VIII. Proceedings of the 37th TEAC, pp. 23–25. Noma, Y., H. Takahashi, M. Toyota, and Y. Asakawa, 1988a. Microbiological conversion of (−)-carvotanacetone and (+)-carvotanacetone by a strain of Aspergillus niger. Proceedings of the 32nd TEAC, pp. 146–148. Noma, Y., H. Takahashi, T. Hashimoto, and Y. Asakawa, 1992c. Biotransformation of isopiperitenone, 6-gingerol, 6-shogaol and neomenthol by a strain of Aspergillus niger. Proceedings of the 37th TEAC, pp. 26–28. Noma, Y. and C. Tatsumi, 1973. Conversion of (−)-carvone by Pseudomonas ovalis, strain 6-1(1), Microbial conversion of terpenes part XIII. Nippon Nogeikagaku Kaishi, 47: 705–711. Noma, Y., M. Toyota, and Y. Asakawa, 1985a. Biotransformation of (−)-carvone and (+)-carvone by Aspergillus spp. Annual Meeting of Agricultural and Biological Chemistry, Sapporo, Japan, p. 68. Noma, Y., M. Toyota, and Y. Asakawa, 1985b. Biotransformation of carvone. 6. Biotransformation of  (−)-carvone and (+)-carvone by a strain of Aspergillus niger. Proceedings of the 29th TEAC, pp. 235–237. Noma, Y., M. Toyota, Y. and Asakawa, 1985c. Microbiological conversion of (−)-carvotanacetone and (+)-carvotanacetone by S. bottropensis SY-2-1. Proceedings of the 29th TEAC, pp. 238–240. Noma, Y., M. Toyota, and Y. Asakawa, 1986. Reduction of terpene aldehydes and epoxidation of terpene alcohols by S. ikutamanensis, Ya-2-1. Proceedings of the 30th TEAC, pp. 204–206. Noma, Y., M. Toyota, and Y. Asakawa, 1988b. Microbial transformation of thymol formation of 2-hydroxy-3p-menthen-5-one by Streptomyces humidus, Tu-1. Proceedings of the 28th TEAC, pp. 177–179. Noma, Y., J. Watanabe, T. Hashimoto, and Y. Asakawa, 2001. Microbiological transformation of β-pinene. Proceedings of the 45th TEAC, pp. 88–90. Noma, Y., S. Yamasaki, and Asakawa Y. 1992d. Biotransformation of limonene and related compounds by Aspergillus cellulosae. Phytochemistry, 31: 2725–2727. Nonoyama, H., H. Matsui, M. Hyakumachi, and M. Miyazawa, 1999. Biotransformation of (−)-menthone using plant parasitic fungi, Rhizoctonia solani as a biocatalyst. Proceedings of the 43rd TEAC, pp. 387–388. Ohsawa, M. and M. Miyazawa, 2001. Biotransformation of (+)- and (−)-isopulegol by the larvae of common cutworm (Spodoptera litura) as a biocatalyst. Proceedings of the 45th TEAC, pp. 375–376. Omata, T., N. Iwamoto, T. Kimura, A. Tanaka, S. Fukui, 1981. Stereoselective hydrolysis of dl-menthyl succinate by gel-entrapped Rhodotorula minuta var. texensis cells in organic solvent. Appl. Microbiol. Biotechnol., 11: 119–204. Oosterhaven, K., K.J. Hartmans, and J.J.C. Scheffer, 1995a. Inhibition of potato sprouts growth by carvone enantiomers and their bioconversion in sprouts. Potato Res., 38: 219–230.

Biotransformation of Monoterpenoids by Microorganisms, Insects, and Mammals

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Oosterhaven, K., B. Poolman, and E.J. Smid, 1995b. S-Carvone as a natural potato sprouts inhibiting, fungistatic and bacteriostatic compound. Indian Crops Prod., 4: 23–31. Oritani, T. and K. Yamashita, 1973a. Microbial dl-acyclic alcohols. Agric. Biol. Chem., 37: 1923–1928. Oritani, T. and K. Yamashita, 1973b. Microbial resolution of racemic 2- and 3-alkylcyclohexanols. Agric. Biol. Chem., 37: 1695–1700. Oritani, T. and K. Yamashita, 1973c. Microbial resolution of dl-isopulegol. Agric. Biol. Chem., 37: 1687–1689. Oritani, T. and K. Yamashita, 1973d. Microbial resolution of racemic carvomenthols. Agric. Biol. Chem., 37: 1691–1694. Oritani, T. and K. Yamashita, 1974. Microbial resolution of (±)-borneols. Agric. Biol. Chem., 38: 1961–1964. Oritani, T. and K. Yamashita, 1980. Optical resolution of dl-(β, γ-unsaturated terpene alcohols by biocatalyst of microorganism. Proceedings of the 24th TEAC, pp. 166–169. Pfrunder, B. and Ch. Tamm, 1969a. Mikrobiologische Umwandlung von bicyclischen monoterpenen durch Absidia orchidis (Vuill.) Hagem. 2. Teil: Hydroxylierung von Fenchon und Isofenchon. Helv. Chim. Acta., 52: 1643–1654. Pfrunder, B. and Ch. Tamm, 1969b. Mikrobiologische Umwandlung von bicyclischen monoterpenen durch Absidia orchidis (Vuill.) Hagem. 1. Teil: Reduktion von Campherchinon und Isofenchonchinon. Helv. Chim. Acta., 52: 1630–1642. Prema, B.R. and P.K. Bhattachayya, 1962. Microbiological transformation of terpenes. II. Transformation of a-pinene. Appl. Microbiol., 10: 524–528. Rama Devi, J., S.G. Bhat, and P.K. Bhattacharyya, 1977. Microbiological transformations of terpenes. Part XXV. Enzymes involved in the degradation of linalool in the Pseudomonas incognita, linalool strain. Indian J. Biochem. Biophys., 15: 323–327. Rama Devi, J. and P.K. Bhattacharyya, 1977a. Microbiological transformations of terpenes. Part XXIV. Pathways of degradation of linalool, geraniol, nerol and limonene by Pseudomonas incognita, linalool strain. Indian J. Biochem. Biophys., 14: 359–363. Rama Devi, J. and P.K. Bhattacharyya, 1977b. Microbiological transformation of terpenes. Part XXIII. Fermentation of geraniol, nerol and limonene by soil Pseudomonad, Pseudomonas incognita (linalool strain). Indian J. Biochem. Biophys., 14: 288–291. Rama Devi, J. and P.K. Bhattacharyya, 1978. Molecular rearrangements in the microbiological transformations of terpenes and the chemical logic of microbial processes. J. Indian Chem. Soc., 55: 1131–1137. Rapp, A. and H. Mandery, 1988. Influence of Botrytis cinerea on the monoterpene fraction wine aroma. In: Bioflavour’87. Analysis—Biochemistry—Biotechnology, P. Schreier, ed., pp. 445–452. Berlin, Germany: Walter de Gruyter and Co. Saeki, M. and N. Hashimoto, 1968. Microbial transformation of terpene hydrocarbons. Part I. Oxidation products of d-limonene and d-pentene. Proceedings of the 12th TEAC, pp. 102–104. Saeki, M. and N. Hashimoto, 1971. Microorganism biotransformation of terpenoids. Part II. Production of cisterpin hydrate and terpineol from d-limonene. Proceedings of the 15th TEAC, pp. 54–56. Saito, H. and M. Miyazawa, 2006. Biotransformation of 1,8-cineole by Salmonella typhimurium OY1001/3A4. Proceedings of the 50th TEAC, pp. 275–276. Savithiry, N., T.K. Cheong, and P. Oriel, 1997. Production of alpha-terpineol from Escherichia coli cells expressing thermostable limonene hydratase. Appl. Biochem. Biotechnol., 63–65: 213–220. Sawamura, Y., S. Shima, H. Sakai, and C. Tatsumi, 1974. Microbiological conversion of menthone. Proceedings of the 18th TEAC, pp. 27–29. Schwammle, B., E. Winkelhausen, S. Kuzmanova, and W. Steiner, 2001. Isolation of carvacrol assimilating microorganisms. Food Technol. Biotechnol., 39: 341–345. Seubert, W. and E. Fass, 1964a. Studies on the bacterial degradation of isoprenoids. V. The mechanism of isoprenoid degradation. Biochem. Z., 341: 35–44. Seubert, W. and E. Fass, 1964b. Studies on the bacterial degradation of isoprenoids. IV. The purification and properties of beta-isohexenylglutaconyl-COA-hydratase and beta-hydroxy-beta-isohexenylglutarylCOA-lyase. Biochem. Z., 341: 23–34. Seubert, W., E. Fass, and U. Remberger, 1963. Studies on the bacterial degradation of isoprenoid compounds. III. Purification and properties of geranyl carboxylase. Biochem. Z., 338: 265–275. Seubert, W. and U. Remberger, 1963. Studies on the bacterial degradation of isoprenoid compounds. II. The role of carbon dioxide. Biochem. Z., 338: 245–246. Shima, S., Y. Yoshida, Y. Sawamura, and C. Tstsumi, 1972. Microbiological conversion of perillyl alcohol. Proceedings of the 16th TEAC, pp. 82–84. Shimoda, K., T. Hirata, and Y. Noma, 1998. Stereochemistry in the reduction of enones by the reductase from Euglena gracilis. Z. Phytochem., 49: 49–53.

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Shimoda, K., D.I. Ito, S. Izumi, and T. Hirata, 1996. Novel reductase participation in the syn-addition of hydrogen to the C = C bond of enones in the cultured cells of Nicotiana tabacum. J. Chem. Soc. Perkin Trans., 1: 355–358. Shimoda, K., S. Izumi, and T. Hirata, 2002. A novel reductase participating in the hydrogenation of an exocyclic C–C double bond of enones from Nicotiana tabacum. Bull. Chem. Soc. Jpn., 75: 813–816. Shimoda, K., N. Kubota, H. Hamada, and M. Kaji, 2003. Cyanobacterium catalyzed asymmetric reduction of enones. Proceedings of the 47th TEAC, pp. 164–166. Shindo, M., T. Shimada, and M. Miyazawa, 2000. Metabolism of 1,8-cineole by cytochrome P450 enzymes in human and rat liver microsomes. Proceedings of the 44th TEAC, pp. 141–143. Shukla, O.P., R.C. Bartholomeus, and I.C. Gunsalus, 1987. Microbial transformation of menthol and menthane3,4-diol. Can. J. Microbiol., 33: 489–497. Shukla, O.P., and P.K. Bhattacharyya, 1968. Microbiological transformations of terpenes: Part XI—Pathways of degradation of α- & β-pinenes in a soil Pseudomonad (PL-strain). Indian J. Biochem., 5: 92–101. Shukla, O.P., M.N. Moholay, and P.K. Bhattacharyya, 1968. Microbiological transformation of terpenes: Part X—Fermentation of α- & β-pinenes by a soil Pseudomonad (PL-strain). Indian J. Biochem., 5: 79–91. Southwell, I.A. and T.M. Flynn, 1980. Metabolism of α- and β-pinene, p-cymene and 1,8–cineole in the brush tail possum. Xenobiotica, 10: 17–23. Suga, T. and T. Hirata, 1990. Biotransformation of exogenous substrates by plant cell cultures. Phytochemistry, 29: 2393–2406. Suga, T., T. Hirata, and H. Hamada, 1986. The stereochemistry of the reduction of carbon–carbon double bond with the cultured cells of Nicotiana tabacum. Bull. Chem. Soc. Jpn., 59: 2865–2867. Sugie, A. and M. Miyazawa, 2003. Biotransformation of (−)-a-pinene by human liver microsomes. Proceedings of the 47th TEAC, pp. 159–161. Swamy, G.K., K.L. Khanchandani, and P.K. Bhattacharyya, 1965. Symposium on Recent Advances in the Chemistry of Terpenoids, Natural Institute of Sciences of India, New Delhi, India, p. 10. Takagi, K., Y. Mikami, Y. Minato, I. Yajima, and K. Hayashi, 1972. Manufacturing method of carvone by microorganisms, Japanese Patent 7,238,998. Takahashi, H., Y. Noma, M. Toyota, and Y. Asakawa, 1994. The biotransformation of (−)- and (+)-neomenthols and isomenthols by Aspergillus niger. Phytochemistry, 35: 1465–1467. Takeuchi, H. and M. Miyazawa, 2004. Biotransformation of nerol by the larvae of common cutworm (Spodoptera litura) as a biocatalyst. Proceedings of the 48th TEAC, pp. 399–400. Takeuchi, H. and M. Miyazawa, 2005. Biotransformation of (−)- and (+)-citronellene by the larvae of common cutworm (Spodoptera litura) as biocatalyst. Proceedings of the 49th TEAC, pp. 426–427. Trudgill, P.W., 1990. Microbial metabolism of terpenes—Recent developments. Biodegradation 1: 93–105. Tsukamoto, Y., S. Nonomura, and H. Sakai, 1975. Formation of p-cis-menthan-1-ol from p-menthane by Pseudomonas mendocina SF. Agric. Biol. Chem., 39: 617–620. Tsukamoto, Y., S. Nonomura, H. Sakai, and C. Tatsumi, 1974. Microbiological oxidation of p-menthane 1. Formation of formation of p-cis-menthan-1-ol. Proceedings of the 18th TEAC, pp. 24–26. Van der Werf, M.J. and J.A.M. de Bont, 1998a. Screening for microorganisms converting limonene into carvone. In: New Frontiers in Screening for Microbial Biocatalysts, Proceedings of the International Symposium, Ede, the Netherlands, K. Kieslich, C.P. Beek, J.A.M. van der Bont, and W.J.J. van den Tweel, eds., Vol. 53, pp. 231–234. Amsterdam, the Netherlands: Studies in Organic Chemistry. Van der Werf, M.J., J.A.M. de Bont, and D.J. Leak, 1997. Opportunities in microbial biotransformation of monoterpenes. Adv. Biochem. Eng. Biotechnol., 55: 147–177. Van der Werf, M.J., K.M. Overkamp, and J.A.M. de Bont, 1998b. Limonene-1,2-epoxide hydrolase from Rhodococcus erythropolis DCL14 belongs to a novel class of epoxide hydrolases. J. Bacteriol., 180: 5052–5057. van Dyk, M.S., E. van Rensburg, I.P.B. Rensburg, and N. Moleleki, 1998. Biotransformation of monoterpenoid ketones by yeasts and yeast-like fungi. J. Mol. Catal. B: Enzym., 5: 149–154. Verstegen-Haaksma, A.A., H.J. Swarts, B.J.M. Jansen, A. de Groot, N. Bottema-MacGillavry, and B. Witholt, 1995. Application of S-(+)-carvone in the synthesis of biologically active natural products using chemical transformations and bioconversions. Indian Crops Prod., 4: 15–21. Watanabe, Y. and T. Inagaki, 1977a. Japanese Patent 77,12,989. No. 187696x. Watanabe, Y. and T. Inagaki, 1977b. Japanese Patent 77, 122,690. No. 87656g. Watanabe, T., H. Nomura, T. Iwasaki, A. Matsushima, and T. Hirata, 2007. Cloning of pulegone reductase and reduction of enones with the recombinant reductase. Proceedings of the 51st TEAC, pp. 323–325. Wolf-Rainer, A., 1994. Phylogeny and biotransformation. Part 5. Biotransformation of isopinocampheol. Z. Naturforsch., 49c: 553–560.

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Yamada, K., S. Horiguchi, and J. Tatahashi, 1965. Studies on the utilization of hydrocarbons by microorganisms. Part VI. Screening of aromatic hydrocarbon-assimilating microorganisms and cumic acid formation from p-cymene. Agric. Biol. Chem., 29: 943–948. Yamaguchi, Y., A. Komatsu, and T. Moroe, 1977. Asymmetric hydrolysis of dl-menthyl acetate by Rhodotorula mucilaginosa. J. Agric. Chem. Soc. Jpn., 51: 411–416. Yamamoto, K., M. Miyazawa, H. Kameoka, and Y. Noma, 1984. Biotransformation of d- and l-fenchone by a strain of Aspergillus niger. Proceedings of the 28th TEAC, pp. 168–170. Yamanaka, T. and M. Miyazawa, 1999. Biotransformation of (−)-trans-verbenol by common cutworm larvae, Spodoptera litura as a biocatalyst. Proceedings of the 43rd TEAC, pp. 391–392. Yawata, T., M, Ogura, K. Shimoda, S. Izumi, and T. Hirata, 1998. Epoxidation of monoterpenes by the peroxidase from the cultured cells of Nicotiana tabacum, Proceedings of the 42nd TEAC, pp. 142–144. Yonemoto, N., S. Sakamoto, T. Furuya, and H. Hamada, 2005. Preparation of (−)-perillyl alcohol oligosaccharides. Proceedings of the 49th TEAC, pp. 108–110.

23

Biotransformation of Sesquiterpenoids, Ionones, Damascones, Adamantanes, and Aromatic Compounds by Green Algae, Fungi, and Mammals Yoshinori Asakawa and Yoshiaki Noma

CONTENTS 23.1 Introduction........................................................................................................................... 769 23.2 Biotransformation of Sesquiterpenoids by Microorganisms................................................. 770 23.2.1 Highly Efficient Production of Nootkatone (2) from Valencene (1).......................... 770 23.2.2 Biotransformation of Valencene (1) by Aspergillus niger and Aspergillus wentii..... 772 23.2.3 Biotransformation of Nootkatone (2) by Aspergillus niger....................................... 773 23.2.4 Biotransformation of Nootkatone (2) by Fusarium culmorum and Botryosphaeria dothidea........................................................................................... 775 23.2.5 Biotransformation of (+)-1(10)-Aristolene (36) from the Crude Drug Nardostachys chinensis by Chlorella fusca, Mucor species, and Aspergillus niger....781 23.2.6 Biotransformation of Various Sesquiterpenoids by Microorganisms....................... 784 23.3 Biotransformation of Sesquiterpenoids by Mammals, Insects, and Cytochrome P-450....... 850 23.3.1 Animals (Rabbits) and Dosing.................................................................................. 850 23.3.2 Sesquiterpenoids........................................................................................................ 850 23.4 Biotransformation of Ionones, Damascones, and Adamantanes........................................... 854 23.5 Biotransformation of Aromatic Compounds......................................................................... 858 References....................................................................................................................................... 865

23.1 INTRODUCTION Recently, environment-friendly green or clean chemistry is emphasized in the field of organic and natural product chemistry. Noyori’s highly efficient production of (−)-menthol using (S)-BINAP-Rh catalyst is one of the most important green chemistries (Tani et al., 1982; Otsuka and Tani, 1991), and 1000 ton of (−)-menthol has been produced by this method in 1 year. On the other hand, enzymes of microorganisms and mammals are able to transform a huge variety of organic compounds, such as mono-, sesqui-, and diterpenoids, alkaloids, steroids, antibiotics, and amino acids from crude drugs and spore-forming green plants to produce pharmacologically and medicinally valuable substances. Since Meyer and Neuberg (1915) studied the microbial transformation of citronellal, there are a great number of reports concerning biotransformation of essential oils, terpenoids, steroids, alkaloids, and acetogenins using bacteria, fungi, and mammals. In 1988, Mikami (1988) reported the review article of biotransformation of terpenoids entitled Microbial Conversion of Terpenoids. Lamare and Furstoss (1990) reviewed biotransformation of more than 25 sesquiterpenoids by microorganisms. 769

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In this chapter, the recent advances in the biotransformation of natural and synthetic compounds; sesquiterpenoids, ionones, α-damascone, and adamantanes; and aromatic compounds, using microorganisms including algae and mammals, are described.

23.2  BIOTRANSFORMATION OF SESQUITERPENOIDS BY MICROORGANISMS 23.2.1 Highly Efficient Production of Nootkatone (2) from Valencene (1) The most important and expensive grapefruit aroma, nootkatone (2), decreases the somatic fat ratio (Haze et al., 2002), and therefore, its highly efficient production has been requested by the cosmetic and fiber industrial sectors. Previously, valencene (1) from the essential oil of Valencia orange was converted into nootkatone (2) by biotransformation using Enterobacter species only in 12% yield (Dhavlikar and Albroscheit, 1973), Rhodococcus KSM-5706 in 0.5% yield with a complex mixture (Okuda et al., 1994), and cytochrome P450 (CYP450) in 20% yield with other complex products (Sowden et al., 2005). Nootkatone (2) was chemically synthesized from valencene (1) with AcOOCMe3 in three steps and chromic acid in low yield (Wilson and Saw, 1978) and using surface-functionalized silica supported by metal catalysts such as Co2+ and Mn2+ with tert-butyl hydroperoxide in 75% yield (Salvador and Clark, 2002). However, these synthetic methods are not safe because they involve toxic heavy metals. An environment-friendly method for the synthesis of nootkatone that does not use any heavy metals such as chromium and manganese must be designed. The commercially available and cheap sesquiterpene hydrocarbon (+)-valencene (1) ([α]D + 84.6°, c  = 1.0) obtained from Valencia orange oil was very efficiently converted into nootkatone (2) by biotransformations using Chlorella (Hashimoto et al., 2003b), Mucor species (Hashimoto et al., 2003a), Botryosphaeria dothidea, and Botryodiplodia theobromae (Noma et al., 2001b; Furusawa et al., 2005a,b). Chlorella fusca var. vacuolata IAMC-28 (Figure 23.1) was inoculated and cultivated while stationary under illumination in Noro medium MgCl2·6H2O (1.5 g), MgSO4·7H2O (0.5 g), KCl (0.2 g), CaCl2·2H2O (0.2 g), KNO3 (1.0 g), NaHCO3 (0.43 g), TRIS (2.45 g), K2HPO4 (0.045 g), Fe-EDTA (3.64 mg), EDTA-2Na (1.89 mg), ZnSO4·7H2O (1.5 g), H3BO2 (0.61 mg), CoCl2·6H2O (0.015 mg), CuSO4·5H2O (0.06 mg), MnCl2·4H2O (0.23 mg), and (NH4)6Mo7O24·4H2O (0.38 mg), in distilled

FIGURE 23.1  Chlorella fusca var. vacuolata.

771

Biotransformation of Sesquiterpenoids and Their Related Compounds 26.58

O

900,000 800,000

Nootkatone (2)

Abundance

700,000 600,000 500,000 400,000 300,000 200,000

Valencene (1)

100,000 0

5.00

25.33 22.70 25.88 29.89 19.39

10.00 15.00 20.00 25.00 30.00 35.00 40.00 45.00 50.00 55.00 Time

FIGURE 23.2  Total ion chromatogram of metabolites of valencene (1) by Chlorella fusca var. vacuolata.

H2O 1 L (pH 8.0). Czapek-peptone medium (1.5% sucrose, 1.5% glucose, 0.5% polypeptone, 0.1% K2HPO4, 0.05% MgSO4·7H2O, 0.05% KCl, and 0.001% FeSO4·7H2O, in distilled water [pH 7.0]) was used for the biotransformation of substrate by microorganism. Aspergillus niger was isolated in our laboratories from soil in Osaka prefecture and was identified according to its physiological and morphological characters. (+)-Valencene (1) (20 mg/50 mL) isolated from the essential oil of Valencia orange was added to the medium and biotransformed by Chlorella fusca for a further 18 days to afford nootkatone (2) (gas chromatography–mass spectrometry [GC-MS] peak area, 89%; isolated yield, 63%) (Figure 23.2) (Noma et al., 2001b; Furusawa et al., 2005a,b). The reduction of 2 with NaBH4 and CeCl3 gave 2α-hydroxyvalencene (3) in 87% yield, followed by Mitsunobu reaction with p-nitrobenzoic acid, triphenylphosphine, and diethyl azodicarboxylate to give nootkatol (2β-hydroxyvalencene) (4)—possessing calcium antagonistic activity—isolated from Alpinia oxyphylla (Shoji et al., 1984) in 42% yield. Compounds 3 and 4 thus obtained were easily biotransformed by C. fusca and Chlorella pyrenoidosa for only 1 day to give nootkatone (2) in good yield (80%–90%), respectively. The biotransformation of compound 1 was further carried out by C. pyrenoidosa and Chlorella vulgaris (Furusawa et al., 2005a,b) and soil bacteria (Noma et al., 2001a) to give nootkatone in good yield (Table 23.1). In the time course of the biotransformation of 1 by C. pyrenoidosa, the yield of nootkatone (2) and nootkatol (4) without 2α-hydroxyvalencene (3) increased with the decrease in that of 1,

TABLE 23.1 Conversion of Valencene (1) to Nootkatone (2) by Chlorella sp. for 14 Days Metabolites (% of the Total in GC-MS) Chlorella sp.

Valencene (1)

2α-Nootkatol (3)

2β-Nootkatol (4)

Nootkatone (2)

Conversion Ratio (%)

C. fusca C. pyrenoidosa C. vulgaris

11 7 0

0 0 0

0 0 0

89 93 100

89 93 100

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Handbook of Essential Oils 21 4

10 5

7 11

12 13

Valencene (1) Chlorella sp. slow

Chlorella sp. slow

HO

HO

3

4

Fast

Fast O

Nootkatone (2)

FIGURE 23.3  Biotransformation of valencene (1) by Chlorella species.

and subsequently, the yield of 2 increased with decrease in that of 3. In the metabolic pathway of valencene (1), 1 was slowly converted into nootkatol (4), and subsequently, 4 was rapidly converted into 2, as shown in Figure 23.3. A fungus strain from the soil adhering to the thalloid liverwort Pallavicinia subciliata, Mucor species, was inoculated and cultivated statically in Czapek-peptone medium (pH 7.0) at 30°C for 7 days. Compound 1 (20 mg/50 mL) was added to the medium and incubated for a further 7 days. Nootkatone (2) was then obtained in very high yield (82%) (Noma et al., 2001b; Furusawa et al., 2005a). The biotransformation from 1 to 2 was also examined using the plant pathogenic fungi B. dothidea and B. theobromae (a total of 31 strains) separated from fungi infecting various types of fruit, and so on. B. dothidea and B. theobromae were both inoculated and cultivated while stationary in Czapekpeptone medium (pH 7.0) at 30°C for 7 days. The same size of the substrate 1 was added to each medium and incubated for a further 7 days to obtain nootkatone (42%–84%) (Furusawa et al., 2005a). The expensive grapefruit aromatic nootkatone (2) used by cosmetic and fiber manufacturers was obtained in high yield by biotransformation of (+)-valencene (1), which can be cheaply obtained from Valencia orange, by Chlorella species, fungi such as Mucor species, B. dothidea, and B. theobromae. This is a very inexpensive and clean oxidation reaction, which does not use any heavy metals, and thus, this method is expected to find applications in the industrial production of nootkatone.

23.2.2 Biotransformation of Valencene (1) by Aspergillus niger and Aspergillus wentii Valencene (1) from Valencia orange oil was cultivated by A. niger in Czapek-peptone medium, for 5 days to afford six metabolites: 5 (1.0%), 6 and 7 (13.5%), 8 (1.1%), 9 (1.5%), 10 (2.0%), and 11 (0.7%), respectively. Ratio of compounds 6 (11S) and 7 (11R) was determined as 1:3 by high-performance liquid chromatography (HPLC) analysis of their thiocarbonates (12 and 13) (Noma et al., 2001b) (Figure 23.4).

773

Biotransformation of Sesquiterpenoids and Their Related Compounds 21 4

10 5

12

7 11

13

1

A. niger

O

O

O

OH 5 (1.0%)

S 6

OH

OH

R

OH

OH

7

1:3 (13.3%)

OH

OH O

8 (1.1%)

R

OH

OH HO

9 (1.5%)

O

OH

OH

10 (1.6%)

OH

OH

11 (0.7%)

FIGURE 23.4  Biotransformation of valencene (1) by Aspergillus niger.

Compounds 8–11 could be biosynthesized by elimination of a hydroxy group of 2-hydroxyvalencenes (3, 4). Compound 3 was biotransformed for 5 days by A. niger to give three metabolites: 6 and 7 (6.4%), 8 (34.6%), and 9 (5.5%), respectively. Compound 4 was biotransformed for 5 days by A. niger to give three metabolites: 6 and 7 (21.8%), 9 (5.5%), and 10 (10.4%), respectively (Figure 23.5). Both ratios of 6 (11S) and 7 (11R) obtained from 3 and 4 were 1:3, respectively. From the aforementioned results, plausible metabolic pathways of valencene (1) and 2-hydroxyvalencene (3, 4) by A. niger are shown in Figure 23.6 (Noma et al., 2001b). Aspergillus wentii and Epicoccum purpurascens converted valencene (1) to 11,12-epoxide (14a) and the same diol (6, 7) (Takahashi and Miyazawa, 2005) as well as nootkatone (2) and 2α-hydroxyvalencene (3) (Takahashi and Miyazawa, 2006). Kaspera et al. (2005) reported that valencene (1) was incubated in submerged cultures of the ascomycete Chaetomium globosum, to give nootkatone (2), 2α-hydroxyvalencene (3), and valencene 11,12-epoxide (14a), together with a valencene ketodiol, valencene diols, a valencene triol, or valencene epoxydiol that were detected by liquid chromatography–MS (LC-MS) spectra and GC-MS of trimethylsilyl derivatives. These metabolites are accumulated preferably inside the fungal cells (Figure 23.7). The metabolites of valencene, nootkatone (2), (3), and (14a), indicated grapefruit with sour and citrus with bitter odor, respectively. Nootkatone 11,12-epoxide (14) showed no volatile fragrant properties.

23.2.3 Biotransformation of Nootkatone (2) by Aspergillus niger A. niger was inoculated and cultivated rotatory (100 rpm) in Czapek-peptone medium at 30°C for 7 days. (+)-Nootkatone (2) ([α]D + 193.5°, c  = 1.0) (80 mg/200 mL), which was isolated from the

774

Handbook of Essential Oils HO

21

10 5

4

12

7 11

13

3

A. niger

O

O

S 6

OH

OH

R

OH

OH

OH

8 (34.6%)

7

1:3

OH

(6.4%)

O

R

OH

OH

9 (5.5%) HO

21 4

10 5

7

12

11 13

4 A. niger

O

O

S 6

OH

OH

1:3

R

OH

OH O

7

R

OH

R

OH

OH

9 (5.5%)

(21.8%)

HO

OH

10 (10.4%)

FIGURE 23.5  Biotransformation of 2α-hydroxyvalencene (3) and 2β-hydroxyvalencene (4) by Aspergillus niger.

essential oil of grapefruit, was added to the medium and further cultivated for 7 days to obtain two metabolites, 12-hydroxy-11,12-dihydronootkatone (5) (10.6%) and C11 stereo mixtures (51.5%), of nootkatone-11S,12-diol (6) and its 11R isomer (7) (11R:11S = 1:1) (Hashimoto et al., 2000b; Noma et al., 2001b; Furusawa et al., 2003) (Figure 23.8). 11,12-Epoxide (14) obtained by epoxidation of nootkatone (2) with meta-chloroperbenzoic acid (mCPBA) was biotransformed by A. niger for 1 day to afford 6 and 7 (11R:11S = 1:1) in good yield

775

Biotransformation of Sesquiterpenoids and Their Related Compounds

1 [O] H+

H

H+

HO

H

HO

4

3 –H2O

[O]

[O] O

OH 9

OH

[O] HO

OH

OH

10

OH

OH

8

FIGURE 23.6  Possible pathway of biotransformation of valencene (1) by Aspergillus niger.

(81.4%). 1-Aminobenzotriazole, an inhibitor of CYP450, inhibited the oxidation process of 1 into compounds 5–7 (Noma et al., 2001b). From the aforementioned results, possible metabolic pathways of nootkatone (2) by A. niger might be considered as shown in Figure 23.9. The same substrate was incubated with A. wentii to produce diol (6, 7) and 11,12-epoxide (14) (Takahashi and Miyazawa, 2005).

23.2.4 Biotransformation of Nootkatone (2) by Fusarium culmorum and Botryosphaeria dothidea (+)-Nootkatone (2) was added to the same medium as mentioned earlier including Fusarium culmorum to afford nootkatone-11R,12-diol (7) (47.2%) and 9β-hydroxynootkatone (15) (14.9%) (Noma et al., 2001b). Compound 7 was stereospecifically obtained at C11 by biotransformation of 1. Purity of compound 7 was determined as ca. 95% by HPLC analysis of the thiocarbonate (13). The biotransformation of nootkatone (2) was examined by the plant pathogenic fungus B. dothidea separated from the fungus that infected the peach. (+)-Nootkatone (2) was cultivated with B. dothidea (Peach PP8402) for 14 days to afford nootkatone diols (6 and 7) (54.2%) and 7α-hydroxynootkatone (16) (20.9%). Ratio of compounds 6 and 7 was determined as 3:2 by HPLC analysis of the thiocarbonates (12, 13) (Noma et al., 2001b). Nootkatone (2) was administered into rabbits to give the same diols (6, 7) (Asakawa et al., 1986; Ishida, 2005). E. purpurascens also biotransformed nootkatone (2) to 5–7, 14, and 15a (Takahashi and Miyazawa, 2006).

776

Handbook of Essential Oils HO c

3 HO c

1

4

a,c a,b,c

O 14a

O

O

O

OH

b 5

2

OH

OH

6,7 OH

b,c

O

O

O OH O 14

15a

15b

a: Aspergillus wentii b: Epicoccum purpurasens c: Chaetomium globosum

FIGURE 23.7  Biotransformation of valencene (1) and nootkatone (2) by Aspergillus wentii, Epicoccum purpurascens, and Chaetomium globosum.

The biotransformation of 2 by A. niger and B. dothidea resembled to that of the oral administration to rabbit since the ratio of the major metabolites 11S- (6) and 11R-nootkatone-11,12-diol (7) was similar. It is noteworthy that the biotransformation of 2 by F. culmorum affords stereospecifically nootkatone-11R,12-diol (7) (Noma et al., 2001b) (Figure 23.10). Metabolites 3–5, 12, and 13 from (+)-nootkatone (2) and 14–17 from (+)-valencene (1) did not show an effective odor. Dihydronootkatone (17), which shows that citrus odor possesses antitermite activity, was also treated in A. niger to obtain 11S-mono- (18) and 11R-dihydroxylated products (19) (the ratio 11S and 11R = 3:2). On the other hand, Aspergillus cellulosae reduced ketone group at C2 of 17 to give 2α- (20) (75.7%) and 2β-hydroxynootkatone (21) (0.7%) (Furusawa et al., 2003) (Figure 23.11). Tetrahydronootkatone (22) also shows antitermite and mosquito-repellant activity. It was incubated with A. niger to give two similar hydroxylated compounds (23, 13.6% and 24, 9.9%) to those obtained from 17 (Furusawa, 2006) (Figure 23.12).

777

Biotransformation of Sesquiterpenoids and Their Related Compounds OH O

O

R

OH

OH

7 (47.2%)

15a (14.9%)

Fusarium culmorum (7 days) O

21

10 5 7

4

12 11 13

2

Aspergillus niger (5 days)

O

O

O OH

OH OH

S

5 (10.6%)

R

6

OH

OH

7 (51.5%)

O

21

10 5 7

4

12 11

2

13

Botryosphaeria dothidea (14 days)

O

O

O OH

S 6

OH

OH

3:2

R 7

OH

OH 16(20.9%)

( (54.2%)

FIGURE 23.8  Biotransformation of nootkatone (2) by Fusarium culmorum, Aspergillus niger, and Botryosphaeria dothidea.

8,9-Dehydronootkatone (25) was incubated with A. niger to give four metabolites, a unique acetonide (26, 15.6%), monohydroxylated (27, 0.2%), dihydroxylated (28%, 69%), and a carboxyl derivative (29, 0.8%) (Figure 23.13). When the same substrate was treated in Aspergillus sojae IFO 4389, compound 25 was converted to the different monohydroxylated products (30, 15.8%) from that mentioned earlier. A. cellulosae is an interesting fungus since it did not give any same products as mentioned earlier; in place, it produced trinorsesquitepene ketone (31%, 6%) and nitrogen-containing aromatic compound (32) (Furusawa et al., 2003) (Figure 23.14).

778

Handbook of Essential Oils O 11

12

2 Cytochrome P-450

O

O

15b

O

14

OH

[H]

[H2O]

[H] O

O

11 12

12 11

OH

OH

OH

6: 11S 7: 11R

5

FIGURE 23.9  Possible pathway of biotransformation of valencene (1) by cytochrome P-450. O

O

O

OH

OH

OH

OH

6

5

OH O OH

OH

OH HO

9

8

OH

OH

O 11

OH

S 12

11

OH 10

O O

OH

7

O

11

O

R O

O

O

13

O

OH O

O

O

OH

11 12 14

O

15

16

OH O

14a

O

O

15a

15b

OH

FIGURE 23.10  Metabolites (5–11, 14–15b) from valencene (1) and nootkatone (2) by various microorganisms.

779

Biotransformation of Sesquiterpenoids and Their Related Compounds O 11 O

R

H

H

OH

18 (4%)

A. niger

O

17

11 OH

OH

19 (62%) 11S:11 R = 2:3 H

HO 2

O

H 20 (75.7%)

A. cellulosae

17

H

HO 2

21 (0.7%)

FIGURE 23.11  Biotransformation of dihydronootkatone (17) by Aspergillus niger and Aspergillus cellulosae.

Mucor species also oxidized compound 25 to give three metabolites, 13-hydroxy-8,9dehydronootkatone (33, 13.2%), an epoxide (34, 5.1%), and a diol (35, 19.9%) (Furusawa et al., 2003). The same substrate was investigated with cultured suspension cells of the liverwort Marchantia polymorpha to afford 33 (Hegazy et al., 2005) (Figure 23.15). O

H 11

O

H

OH 23 (13.6%)

A. niger

22

O

H 11

12

OH 24 (9.9%)

FIGURE 23.12  Biotransformation of tetrahydronootkatone (22) by Aspergillus niger.

OH

780

Handbook of Essential Oils O

O 11

O

12

R

O

11 OH

O

A. niger

27 (0.2%)

26 (15.6%) 25

O

O 12 11 COOH

11

OH

OH

OH

28 (69.7%) C-11 mixtures (3:2)

29 (0.8%) O

12

O A. sojae

12

8 days 25

30 (15.8%)

13

OH

FIGURE 23.13  Biotransformation of 8,9-dehydronootkatone (25) by Aspergillus sojae. O O O

31 (6.0%)

A. cellulosae

H O OH O

HO 25

NH

O HO 32

O OCH3

FIGURE 23.14  Biotransformation of 8,9-dehydronootkatone (25) by Aspergillus cellulosae.

Although Mucor species could give nootkatone (21) from valencene (1), this fungus biotransformed the same substrate (25) to the same alcohol (30, 13.2%) obtained from the same starting compound (25) in A. sojae, a new epoxide (34, 5.1%), and a diol (35, 9.9%). The metabolites (3, 4, 20, 21) inhibited the growth of lettuce stem, and 3 and 4 inhibited germination of the same plant (Hashimoto and Asakawa, 2007). Valerianol (35a), from Valeriana officinalis whose dried rhizome is traditionally used for its carminative and sedative properties, was biotransformed by Mucor plumbeus, to produce three metabolites, a bridged ether (35b), and a triol (35c), which might be formed via C1–C10 epoxide, and 35d arises from double dehydration (Arantes et al., 1999). In this case, allylic oxidative compounds have not been found (Figure 23.16).

781

Biotransformation of Sesquiterpenoids and Their Related Compounds O

25 Marchantia polymorpha cells

Mucor sp.

O

O

O

12 13 33 (13.2%)

11 12

11

O OH

OH

OH

13 34 (5.1%)

35 (19.9%)

FIGURE 23.15  Biotransformation of 8,9-dehydronootkatone (25) by Marchantia polymorpha and Mucor species. OH

O

OH

OH

+H2O

–H2O

OH

OH

OH

35c –H2O

M. plumbeus

O OH

35a

OH

35b

35d

FIGURE 23.16  Biotransformation of valerianol (35a) by Mucor plumbeus.

23.2.5 Biotransformation of (+)-1(10)-Aristolene (36) from the Crude Drug Nardostachys chinensis by Chlorella fusca, Mucor species, and Aspergillus niger The structure of sesquiterpenoid (+)-1(10)-aristolene (=calarene) (36) from the crude drug Nardostachys chinensis was similar to that of nootkatone. 2-Oxo-1(10)-aristolene (38) shows antimelanin-inducing activity and excellent citrus fragrance. On the other hand, the enantiomer (37) of 36 and (+)-aristolone (41) were also found in the liverworts as the natural products. In order to obtain compound 38 and its analogues, compound 36 was incubated with Chlorella fusca var. vacuolata IAMC-28, Mucor species, and A. niger (Furusawa et al., 2006a) (Figure 23.17). C. fusca was inoculated and cultivated stationary in Noro medium (pH 8.0) at 25°C for 7 days, and (+)-1(10)-aristolene (36) (20 mg/50 mL) was added to the medium and further incubated for 10–14 days and cultivated stationary under illumination (pH 8.0) at 25°C for 7 days to afford

782

Handbook of Essential Oils 1 2 3

4 14

10 5

O

9

8 6 7

15 36

11 12

13 37

38 OH

O

O

O

39

40

41

FIGURE 23.17  Naturally occurring aristolane sesquiterpenoids.

1(10)-Aristolene (36)

C. fusca

OH O

O

38 (18.7%)

39 (7.1%)

O

40 (7.0%)

FIGURE 23.18  Biotransformation of 1(10)-aristolene (36) by Chlorella fusca.

1(10)-aristolen-2-one (38, 18.7%), (−)-aristolone (39, 7.1%), and 9-hydroxy-1(10)-aristolen-2-one (40). Compounds 38 and 40 were found in Aristolochia species (Figure 23.18). Mucor species was inoculated and cultivated rotatory (100 rpm) in Czapek-peptone medium (pH 7.0) at 30°C for 7 days. (+)-1(10)-Aristolene (36) (100 mg/200 mL) was added to the medium and further for 7 days. The crude metabolites contained 38 (0.9%) and 39 (0.7%) as very minor products (Figure 23.19). O

O

Mucor sp.

36

38 (0.9%)

FIGURE 23.19  Biotransformation of 1(10)-aristolene (36) by Mucor species.

39 (0.7%)

783

Biotransformation of Sesquiterpenoids and Their Related Compounds

C. fusca

HO

O

C. fusca

Mucor sp.

Mucor sp. Route a

38

36 Route c C. fusca Mucor sp.

H

C. fusca Mucor sp.

[O]

Route b

OH

C. fusca

OH

OH

O

40 C. fusca Mucor sp.

[O]

C. fusca Mucor sp.

O

C. fusca

39

FIGURE 23.20  Possible pathway of biotransformation of 1(10)-aristolene (36) by Chlorella fusca and Mucor species.

Although Mucor species produced a large amount of nootkatone (2) from valencene (1), however, only poor yield of similar products as those from valencene (1) was seen in the biotransformation of tricyclic substrate (36). Possible biogenetic pathway of (+)-1(10)-aristolene (36) is shown in Figure 23.20. A. niger was inoculated and cultivated rotatory (100 rpm) in Czapek-peptone medium (pH 7.0) at 30°C for 3 days. (+)-1(10)-Aristolene (36) (100 mg/200 mL) was added to the medium and further for 7 days. From the crude metabolites, four new metabolic products 42, 1.3%; 43, 3.2%; 44, 0.98%; and 45, 2.8% were obtained in very poor yields (Figure 23.21). Possible metabolic pathways of 36 by A. niger are shown in Figure 23.22. O

CO OH

HO

42 (1.3%)

A. niger

CO OH 43 (2.7%)

6 days HO 36

O

CO OH 44 (0.8%)

HO

O 45 (2.1%)

FIGURE 23.21  Biotransformation of 1(10)-aristolene (36) by Aspergillus niger.

O

784

Handbook of Essential Oils H+

H

HO A. niger [O] 36 Route a

Route b

[O]

[O]

route c

[O]

O HO 38

COOH

HO

COOH

43

H+ O

O

COOH

O

42

COOH

HO O

45

HO

COOH

44

HO HO

H

O

HO

HO +

O– O

HO

COOH H+

FIGURE 23.22  Possible pathway of biotransformation of 1(10)-aristolene (36) by Aspergillus niger.

Commercially available (+)-1(10)-aristolene (36) was treated with Diplodia gossypina and Bacillus megaterium. Both microorganisms converted 36–4 (46–49; 0.8%, 1.1%, 0.16%, 0.38%) and six metabolites (40, 50–55; 0.75%, 1.0%, 1.0%, 2.0%, 1.1%, 0.5%, 0.87%), together with 40 (0.75%), respectively (Abraham et al., 1992) (Figure 23.23). It is noteworthy that Chlorella and Mucor species introduce hydroxyl group at C2 of the substrate (36) as seen in the biotransformation of valencene (1), while D. gossypina and B. megaterium oxidize C2, C8, C9, and/or 1,1-dimethyl group on a cyclopropane ring. A. niger oxidizes not only C2 but also stereoselectively oxidized one of the gem-dimethyl groups on cyclopropane ring. Stereoselective oxidation of one of gem-dimethyl of cyclopropane and cyclobutane derivatives is observed in biotransformation using mammals (see later).

23.2.6 Biotransformation of Various Sesquiterpenoids by Microorganisms Aromadendrene-type sesquiterpenoids have been found not only in higher plants but also in liverworts and marine sources. Three aromadendrenes (56, 57, 58) were biotransformed by D. gossypina, B. megaterium, and Mycobacterium smegmatis (Abraham et al., 1992). Aromadendrene (56) (800 mg)

785

Biotransformation of Sesquiterpenoids and Their Related Compounds OH D. gossypina

O

HO OH

36

R 47: R = CH2OH 48: R = COOH

46 OH

49

OH O

R B. megaterium

HO

R

HO R2 R1

36

50: R = α-OH 51: R = β-OH

52: R = H 53: R = OH

40

54: R1=OH,R2=H 55: R1=H,R2=OH

FIGURE 23.23  Biotransformation of 1(10)-aristolene (36) by Diplodia gossypina and Bacillus megaterium.

was converted by B. megaterium to afford a diol (59) and a triol (60) of which 59 (7 mg) was the major product. The triol (60) was also obtained from the metabolite of (+)-(1R)-aromadendrene (56) by the plant pathogen Glomerella cingulata (Miyazawa et al., 1995d). allo-Aromadendrene (57) (1.2 g) was also treated in M. smegmatis to afford 61 (10 mg) (Abraham et al., 1992) (Figure 23.24). The same substrate was also incubated with G. cingulata to afford C10 epimeric triol (62) (Miyazawa et  al., 1995d). Globulol (58) (400 mg) was treated in M. smegmatis to give only a carboxylic acid (63) (210 mg). The same substrate (58) (1 g) was treated in D. gossypina and B. megaterium to give two diols, 64 (182 mg) and 65, and a triol (66) from the former and 67–69 from the latter organism among which 64 (60 mg) was predominant (Abraham et al., 1992). G. cingulata and

H

H

OH OH

H

OH OH

B. megaterium H

H

H

56

59

60

H

OH

HO M. smegmatis

H

H

57

61

H

H

OH

OH

M. smegmatis H

H

58

63

COOH

FIGURE 23.24  Biotransformation of aromadendrene (56), allo-aromadendrene (57), and globulol (58) by Bacillus megaterium and Mycobacterium smegmatis.

786

Handbook of Essential Oils OH H

H

O

H

OH

G. cingulata

H

H

OH

H

56

60

H

H

HO HO H

O

G. cingulata

H

OH

H

H

57

62

FIGURE 23.25  Biotransformation of aromadendrene (56) and allo-aromadendrene (57) by Glomerella cingulata.

Botrytis cinerea also bioconverted globulol (58) to diol (64) regio- and stereoselectively (Miyazawa et al., 1994) (Figures 23.25 and 23.26). Globulol (58) (1.5 g) and 10-epiglobulol (70) (1.2 mL) were separately incubated with Cephalosporium aphidicola in shake culture for 6 days to give the same diol 64 (780 mg) as obtained from the same substrate by B. megaterium mentioned earlier and 71 (720 mg) (Hanson et al., 1994). A. niger also converted globulol (58) and epiglobulol (70) to a diol (64) and three 13-hydroxylated globulol (71, 72, 74) and 4α-hydroxylated product (73). The epimerization at C4 is very rare example (Hayashi et al., 1998). H

OH

H

OH

H

OH OH

D. gossypina H

OH

B. megaterium

H

H HO

OH

64 H

H

65 OH

HO

H

OH

66 OH

HO

H

OH

B. megaterium

58

H

H

OH

A. niger G. cingulata

H

H

H

67

68

69

H

OH

Cephalosporium aphidicola H

H

58

64

OH

FIGURE 23.26  Biotransformation of globulol (58) by various microorganisms.

OH

787

Biotransformation of Sesquiterpenoids and Their Related Compounds HO H

HO H C. aphidicola A. niger H

H

70

71

OH

A. niger HO H

HO H

H HO

HO H

OH H

72

OH 73

HO H

OH

H

OH 74

HO H

HO H

G. cingulata H 75

H CHO

H COOH 76

FIGURE 23.27  Biotransformation of 10-epi-glubulol (70) and ledol (75) by Cephalosporium aphidicola, Aspergillus niger, and Glomerella cingulata.

Ledol (75), an epimer at C1 of globulol, was incubated with G. cingulata to afford C13 carboxylic acid (76) (Miyazawa et al., 1994) (Figure 23.27). Squamulosone (77), aromadendr-1(10)-en-9-one isolated from Hyptis verticillata (Labiatae), was reduced chemically to give 78–82, which were incubated with the fungus Curvularia lunata in two different growth media (Figure 23.28). From 78, two metabolites 80 and 83 were obtained. Compound 79 and 80 were metabolized to give ketone 81 as the sole product and 78 and 83, respectively. From compound 81, two metabolites, 79 and 84, were obtained (Figure 23.29). From the metabolite of the substrate (82), five products (84–88) were isolated (Collins et al., 2002a) (Figure 23.30). Squamulosone (77) was treated in the fungus M. plumbeus ATCC 4740 to give not only cyclopentanol derivatives (89, 90) but also C12-hydroxylated products (91–93) (Collins et al., 2002b) (Figure 23.31). Spathulenol (94), which is found in many essential oils, was fed by A. niger to give a diol (95) (Higuchi et al., 2001). ent-10β-Hydroxycyclocolorenone (96) and myli-4(15)-en-9-one (96a) isolated from the liverwort Mylia taylorii were incubated with A. niger IFO 4407 to give C10 epimeric product (97) (Hayashi et al., 1999) and 12-hydroxylated product (96b), respectively (Nozaki et al., 1996) (Figures 23.32 and 23.33). (+)-ent-Cyclocolorenone (98) [α]D − 405° (c = 8.8, EtOH), one of the major compounds isolated from the liverwort Plagiochila sciophila (Asakawa, 1982, 1995), was treated by A. niger to afford three metabolites, 9-hydroxycyclocolorenone (99, 15.9%) 12-hydroxy-(+)-cyclocolorenone (100, 8.9%) and a unique cyclopropane-cleaved metabolite, 6β-hydroxy-4,11-guaiadien-3-one (101, 35.9%), and 6β,7β-dihydroxy-4,11-guaiadien-3-one (102, trace), of which 101 was the major component.

788

Handbook of Essential Oils O

H

H

OH

77 H H

O

78

H H

80

C. lunata in growth medium 1 C. lunata in growth medium 2

OH

H 83

FIGURE 23.28  Biotransformation of aromadendra-9-one (80) by Curvularia lunata.

H

OH

H O

H

79

H

81 C. lunata in growth medium 1 C. lunata in growth medium 2

H

O

H HO

84

FIGURE 23.29  Biotransformation of 10-epi-aromadendra-9-one (81) by Curvularia lunata.

The enantiomer (103) [α]D + 402° (c = 8.8, EtOH) of 98 isolated from Solidago altissima was biotransformed by the same organism to give 13-hydroxycycolorenone (103a, 65.5%), the enantiomer of 100, 1β,13-dihydroxycyclocolorenone (103b, 5.0%), and its C11 epimer (103c) (Furusawa et al., 2005c, 2006a). It is noteworthy that no cyclopropane-cleaved compounds from 103 have been detected in the crude metabolites even in GC-MS analysis (Figure 23.34). Plagiochiline A (104) that shows potent insect antifeedant, cytotoxicity, and piscicidal activity is very pungent, 2,3-secoaromadendrane sesquiterpenoids having 1,1-dimethyl cyclopropane ring, isolated from the liverwort Plagiochila fruticosa. Plagiochilide (105) is the major component of this liverwort. In order to get more pungent component, the lactone (105, 101 mg) was incubated with A. niger to give two metabolites: 106 (32.5%) and 107 (9.7%). Compound 105 was incubated

789

Biotransformation of Sesquiterpenoids and Their Related Compounds

C. lunata

OH OH

H

OH OH

HO HO

HO

OH

H

H

H

85

86

87

HO

OH

HO

O

82

H

H

88

89

FIGURE 23.30  Biotransformation of aromadendr-1(10),9-diene (82) by Curvularia lunata. O

H 77

M. plumbeus HO

O

HO

O

O

H

H

H

89

90

91

HO

O

OH

O

H

H

92

HO

OH

93

OH

FIGURE 23.31  Biotransformation of squamulosone (77) by Mucor plumbeus.

in A. niger including 1-aminobenzotriazole, the inhibitor of CYP450, to produce only 106, since this enzyme plays an important role in the formation of carboxylic acid (107) from primary alcohol (106). Unfortunately, two metabolites show no hot taste (Hashimoto et al., 2003c; Furusawa et al., 2006a,b) (Figure 23.35). Partheniol, 8α-hydroxybicyclogermacrene (108) isolated from Parthenium argentatum × Parthenium tomentosum, was cultured in the media of Mucor circinelloides ATCC 15242 to afford six metabolites, a humulane (109), three maaliane (110, 112, 113), an aromadendrane (111), and a tricylohumulane

790

Handbook of Essential Oils OH H

H

OH

A. niger HO

HO

94

H 95

FIGURE 23.32  Biotransformation of spathulenol (94) by Aspergillus niger.

H

H A. niger

HO

HO

H

H

94 H

95 HO H

OH A. niger

O

O

96 H

97 H

O

O

A. niger

HO 96a

96b

FIGURE 23.33  Biotransformation of spathulenol (94), ent-10β-hydroxycyclocolorenone (96) and myli-4(15)en-9-one (96a) by Aspergillus niger.

triol (114), the isomer of compound 111. Compounds 110, 111, and 114 were isolated as their acetates (Figure 23.36). Compounds 110 might originate from the substrate by acidic transannular cyclization since the broth was pH 6.4 just before extraction (Maatooq, 2002a). The same substrate (108) was incubated with the fungus Calonectria decora to afford six new metabolites (108a–108f). In these reactions, hydroxylation, epoxidation, and transannular cyclization were evidenced (Maatooq, 2002c) (Figure 23.37). ent-Maaliane-type sesquiterpene alcohol, 1α-hydroxymaaliene (115), isolated from the liverwort M. taylorii, was treated in A. niger to afford two primary alcohols (116, 117) (Morikawa et al., 2000). Such an oxidation pattern of 1,1-dimethyl group on the cyclopropane ring has been found in aromadendrane series as described earlier and mammalian biotransformation of a monoterpene hydrocarbon, D3-carene (Ishida et al., 1981) (Figure 23.38).

791

Biotransformation of Sesquiterpenoids and Their Related Compounds OH

H 9

A. niger

O H 9 6 7

O

99 (15.9%) H O

(+)-Cyclocolorenone (98) from Plagiochila sciophila

H

100

H+

OH

H

1

O

O

OH 103a (65.5%)

(–)-Cyclocolorenone (103) from Solidago altissima

OH

102

HO

A. niger O

6 7 HO

101 (35.9%)

H

O

O

6 7 HO

HO–

H

O

6 7

H

OH 103b (5.0%)

HO 103c

FIGURE 23.34  Biotransformation of (+)-cyclocolorenone (98) and (−)-cyclocolorenone (103) by Aspergillus niger. Ac O

H

O

O H Ac O O

104 O

H A. niger

O

H

H

O

A. niger 1-aminobenzotriazole

H

105

106

OH Cyp-450

O

O

H

O

Cyp-450 H HOOC

H

O H OHC

107

FIGURE 23.35  Biotransformation of plagiochiline C (104) by Aspergillus niger.

792

Handbook of Essential Oils OH OAc OH HO HO 109

OH M. circinelloides

110

111

OH

OH

HO

OH

108 HO

OH

OH

OH

H 112

HO

113

114

FIGURE 23.36  Biotransformation of 8α-hydroxybicyclogermacrene (108) by Mucor circinelloides. OH

OH

OH

OH OH 108a

Calonectria decora

108b OH

108c

OH OH

108

OH O

HO 108d

108e

108f

FIGURE 23.37  Biotransformation of 8α-hydroxybicyclogermacrene (108) by Calonectria decora. OH

OH

O

A. niger H

H

H

115

HO 116

HO 117

FIGURE 23.38  Biotransformation of 1α-hydroxymaaliene (115) Aspergillus niger.

9(15)-Africanene (117a), a tricyclic sesquiterpene hydrocarbon isolated from marine soft corals of Simularia species, was biotransformed by A. niger and Rhizopus oryzae for 8 days to give 10α-hydroxy (117b) and 9α,15-epoxy derivative (117c) (Venkateswarlu et al., 1999) (Figure 23.39). Germacrone (118), (+)-germacrone-4,5-epoxide (119), and curdione (120) isolated from Curcuma aromatica, which has been used as crude drug, were incubated with A. niger. From compound 119 (700 mg), two naturally occurring metabolites, zedoarondiol (121) and isozedoarondiol (122), were obtained (Takahashi, 1994). Compound 119 was cultured in callus of Curcuma zedoaria and C. ­aromatica to give the same secondary metabolites 121, 122, and 124 (Sakui et al., 1988) (Figures 23.40 and 23.41).

793

Biotransformation of Sesquiterpenoids and Their Related Compounds H

H

A. niger HO R. oryzae

H

H

O +

H

117a

H 117b

117c

FIGURE 23.39  Biotransformation of 9(15)-africanene (117a) by Aspergillus niger and Rhizopus oryzae. OH

H

O HO O

O

A. niger

HO H

O 118

H 121

119

O HO

O

H 122

HO 123

FIGURE 23.40  Biotransformation of germacrone (118) by Aspergillus niger. HO H O

Curcumazedoaria and C. aromatica cells

O

HO

O

118

O

119

H

122 O

HO

123

FIGURE 23.41  Biotransformation of germacrone (118) by Curcuma zedoaria and Curcuma aromatica cells.

A. niger biotransformed germacrone (118, 3 g) to very unstβ-hydroxygermacrone (123) and 4,5-epoxygermacrone (119), which was further converted to two guaiane sesquiterpenoids (121) and (122) through transannular-type reaction (Takahashi, 1994). The same substrate was incubated in the microorganism Cunninghamella blakesleeana to afford germacrone-4,5-epoxide (119) (Hikino et al., 1971), while the treatment of 118 in the callus of C. zedoaria gave four metabolites 121, 122, 125, and 126 (Sakamoto et al., 1994) (Figure 23.42). The same substrate (118) was treated in plant cell cultures of S. altissima (Asteraceae) for 10 days to give various hydroxylated products (121, 127, 125, 128–132) (Sakamoto et al., 1994). Guaiane (121) underwent further rearrangement C4–C5, cleavage, and C5–C10 transannular cyclization to the bicyclic hydroxyketone (128) and diketone (129) (Sakamoto et al., 1994) (Figure 23.43).

794

Handbook of Essential Oils

O

O

p-TsOH

C. blakesleeana

O

O

OH

118

119 C. zedoaria OH H

OH

H

O O

O HO

124

H

H

HO

H

O

121

122

129

C. zedoaria O

O

O

O

C. zedoaria O 118

125

126

FIGURE 23.42  Biotransformation of germacrone (118) by Cunninghamella blakesleeana and Curcuma zedoaria cells.

OH– O

O

O

H

O

OH O

Cell culture (Solidago altissima)

118

HO

O

HO

O 127

H+

H 121

O R 130

H+ O

OH O

HO OH– 126

OH O

H2O H 131

O

H

OH

O

H OH 132

FIGURE 23.43  Biotransformation of germacrone (118) by Solidago altissima cells.

128: R = OH 129: R = O

795

Biotransformation of Sesquiterpenoids and Their Related Compounds O

A. niger

HO

O

O 120

O

HO

O

O

133

134

OH

H

+ O

O

–H+

O

OH O

H 2O

O

OH

135

FIGURE 23.44  Biotransformation of curdione (120) by Aspergillus niger.

Curdione (120) was also treated in A. niger to afford two allylic alcohols (133, 134) and a spirolactone (135). C. aromatica and Curcuma wenyujin produced spirolactone (135), which might be formed from curdione via transannular reaction in vivo and was biotransformed to spirolactone diol (135) (Asakawa et al., 1991; Sakui et al., 1992) (Figure 23.44). A. niger also converted shiromodiol diacetate (136) isolated from Neolitsea sericea to 2β-hydroxy derivative (137) (Nozaki et al., 1996) (Figure 23.45). Twenty strains of filamentous fungi and four species of bacteria were screened initially by thinlayer chromatography for their biotransformation capacity of curdione (120). Mucor spinosus, Mucor polymorphosporus, Cunninghamella elegans, and Penicillium janthinellum were found to be able to biotransform curdione (120) to more polar metabolites. Incubation of curdione with M. spinosus, which was most potent strain to produce metabolites, for 4 days using potato medium gave five metabolites (134, 134a–134d) among which compounds 134c and 134d are new products (Ma et al., 2006) (Figure 23.46). Many eudesmane-type sesquiterpenoids have been biotransformed by several fungi and various oxygenated metabolites obtained. β-Selinene (138) is ubiquitous sesquiterpene hydrocarbon of seed oil from many species of Apiaceae family, for example, Cryptotaenia canadensis var. japonica, which is widely used as vegetable for Japanese soup. β-Selinene was biotransformed by plant pathogenic fungus G. cingulata to give an epimeric mixtures (1:1) of 1β,11,12-trihydroxy product (139) (Miyazawa et al., 1997b). The same substrate was treated in A. wentii to give 2α,11,12-trihydroxy derivative (140) (Takahashi et al., 2007). Eudesm-11(13)-en-4,12-diol (141) was biotransformed by A. niger to give 3β-hydroxy derivative (142) (Hayashi et al., 1999). α-Cyperone (143) was fed by Collectotrichum phomoides (Lamare and Furstoss, 1990) to afford 11,12-diol (144) and 12-manool (145) (Higuchi et al., 2001) (Figure 23.47). OAc

HO

OAc A. niger O

OAc 136

O

OAc 137

FIGURE 23.45  Biotransformation of shiromodiol diacetate (136) by Aspergillus niger.

796

Handbook of Essential Oils O

O

O

O

O 134a O

O

134b

HO

O

O

M. spinosus

120

OH

O

O

O

134

134c O

HO

OH

O 134d

FIGURE 23.46  Biotransformation of curdione (120) by Mucor spinosus.

The filamentous fungi Gliocladium roseum and Exserohilum halodes were used as the bioreactors for 4β-hydroxyeudesmane-1,6-dione (146) isolated from Sideritis varoi subsp. ­cuatrecasasii. The former fungus transformed 146 to 7α-hydroxyl (147), 11-hydroxy (148), 7α,11-dihydroxy (149), 1α,11dihydroxy (150), and 1α,8α-dihydroxy derivatives (151), while E. halodes gave only 1α-hydoxy product (152) (Garcia-Granados et al., 2001) (Figure 23.48). OH

OH

G. cingulata

H

O

H

H

138

OH

OH

139 A. wentii

HO

H

H

138

140 A. niger

HO

H

OH

HO

HO

141

OH

H 142

C. phomoides O

O

O OH 143

144

OH

OH 145

FIGURE 23.47  Biotransformation of eudesmenes (138, 141, 143) by Aspergillus wentii, Glomerella ­cingulata, and Collectotrium phomoides.

797

Biotransformation of Sesquiterpenoids and Their Related Compounds R1

O

R4

Gliocladium roseum

HO

R2

O 146

R3 O 147: R1 = O, R2 = OH, R3 = R4 = H

Exserohilum halodes

149: R1 = O, R2 = R3 = OH, R4 = H

HO

148: R1 = O, R2 = R4 = H, R3 = OH 150: R1 = αOH, βH,R2 = R4 = H, R3 = OH 151: R1 = αOH, βH,R2 = R3 = H, R4 = OH OH

HO

O 152

FIGURE 23.48  Biotransformation of 4β-hydroxy-eudesmane-1,6-dione (146) by Gliocladium roseum and Exserohilum halodes.

Orabi (2000) reported that Beauveria bassiana is the most efficient microorganism to metabolize plectanthone (152a) among 20 microorganisms, such as Absidia glauca, Aspergillus flavipes, B. bassiana, Cladosporium resinae, and Penicillium frequentans. The substrate (152a) was incubated with B. bassiana to give metabolites 152b (2.1%), 152c (21.2%), 152d (2.5%), 152e (no data), and 152f (1%) (Figure 23.49). (–)-α-Eudesmol (153) isolated from the liverwort Porella stephaniana was treated by A. cellulosae and A. niger to give 2-hydroxy (154) and 2-oxo derivatives (155), among which the latter product OH

OAc O

O

H

OAc

OH

H

152b OH

OAc O

B. bassiana H

OAc

152d

152a

OAc O

H

OAc

152c

OH

O

H

OH

H

OH OH

152e

O

HO

OAc

OAc

152f

FIGURE 23.49  Biotransformation of eudesmenone (152a) by Beauveria bassiana.

798

Handbook of Essential Oils A. niger

O

HO

O

A. cellulosae H 153

OH

A. niger

H 157

H 154

OH

HO

OH

H 155 HO

H

OH

O

O

H 161

OH

OH

H 156

OH

H 160

OH

H

OH

HO

H 158

159

OH

OH

OH

O

H 162

OH

OH

156

FIGURE 23.50  Biotransformation of α-eudesmol (153) and β-eudesmol (157) by Aspergillus niger and Aspergillus cellulosae.

was predominantly obtained. This bioconversion was completely blocked by 1-aminobenzotriazole, CYP450 inhibitor. Compound 155 has been known as natural product, isolated from Pterocarpus santalinus (Noma et  al., 1996). Biotransformation of α-eudesmol (153) isolated from the dried Atractylodes lancea was reinvestigated by A. niger to give 2-oxo-11,12-dihydro-α-eudesmol (156) together with 2-hydroxy- (154) and 2-oxo-α-eudesmol (155). β-Eudesmol (157) was treated in A. niger, with the same culture medium, to afford 2α- (158) and 2β-hydroxy-α-eudesmol (159) and 2α,11,12-trihydroxy-β-eudesmol (160) and 2-oxo derivative (161), which was further isomerized to compound 162 (Noma et al., 1996, 1997a) (Figure 23.50). Three new hydroxylated metabolites (157b–157d) along with a known 158 and 157e–157 g were isolated from the biotransformation reaction of a mixture of β- (157) and γ-eudesmols (157a) by Gibberella suabinetii. The metabolites proved a super activity of the hydroxylase, dehydrogenase, and isomerase enzymes. The hydroxylation is a common feature; on the contrary, cyclopropyl ring formation like compound (158d) is very rare (Maatooq, 2002b) (Figure 23.51). A furanosesquiterpene, atractylon (163) obtained from Atractylodes rhizoma, was treated with the same fungus to yield atractylenolide III (164) possessing inhibition of increased vascular permeability in mice induced by acetic acid (Hashimoto et al., 2001a,b). The biotransformation of sesquiterpene lactones have been carried out by using different microorganisms. Costunolide (165), a very unstable sesquiterpene γ-lactone, from Saussurea radix, was treated in A. niger to produce three dihydrocostunolides (166–168) (Clark and Hufford, 1979). Costunolide is easily converted into eudesmanolides (169–172) in diluted acid, thus, 166–168 might be biotransformed after being cyclized in the medium including the microorganisms. If the crude drug including costunolide (165) is orally administered, 165 will be easily converted into 169–172 by stomach juice (Figure 23.52). (+)-Costunolide (165), (+)-cnicin (172a), and (+)-salonitgenolide (172b) were incubated with Cunninghamella echinulata and R. oryzae. The former fungus converted compound 165, to four metabolites, (+)-11β,13-dihydrocostunolide (165a), 1β-hydroxyeudesmanolide, (+)-santamarine (166a), (+)-reynosin (166b), and (+)-1β-hydroxyarbusculin A (168a), which might be formed from 1β,10α-epoxide (166c). Treatment of 172a with C. echinulata gave (+)-salonitenolide (172b) (Barrero et al., 1999) (Figure 23.53).

799

Biotransformation of Sesquiterpenoids and Their Related Compounds HO

OH

H G. suabinetii H

OH

H

157b

OH

157c

OH

HO

157

OH H

OH

OH

H 158

157d

HO

O

OH

G. suabinetii

OH 157f

157e OH 157a OH OH 157g

FIGURE 23.51  Biotransformation of β-eudesmol (157) and γ-eudesmol (157a) by Gibberella suabinetii.

OH

O

A. niger

O O

H

H 163

164 OH

OH

A. niger H

O O

165

H 169

O

166

H

O O

170

H

O

O

167

O

HO

O

O 171

O

O O

168

HO

O

H

H

172

O O

FIGURE 23.52  Biotransformation of atractylon (163) and costunolide (165) by Aspergillus niger.

800

Handbook of Essential Oils OH

O

165a

O

165

H

O

Cunninghamella echinulata

166a

OH

O

OH

O

H

HO

O

166b H

O

H

O

O

168a

O

O

O

166c

O

O

172a

O

OH

OH O

HO

O

OH

C. echinulata

HO

O

172b

O

FIGURE 23.53  Biotransformation of costunolide (165) and its derivative (172a) by Cunninghamella ­echinulata and Rhizopus oryzae.

α-Cyclocostunolide (169), β-cyclocostunolide (170), and γ-cyclocostunolide (171) prepared from costunolide were cultivated in A. niger. From the metabolite of 169, four dihydro lactones (173–176) were obtained, among which sulfur-containing compound (176) was predominant (Figure 23.54). The same substrate (169) was cultivated for 3 days by A. cellulosae to afford a sole metabolite, 11β,13-dihydro-α-cyclocostunolide (177). Possible metabolic pathways of 169 by both microorganisms were shown in Figure 23.55. A double bond at C11–C13 of 169 was firstly reduced stereoselectively to afford 177, followed by oxidation at C2 to give 173, and then further oxidation occurred to furnish two hydroxyl derivatives (174, 175) in A. niger. The sulfide compound (176) might be formed from 175 or by Michael condensation of ethyl 2-hydroxy-3-mercaptopropanate, which might originate from Czapek-peptone medium into exomethylene group of α-cyclocostunolide (Hashimoto et al., 1999c, 2001a,b). A. niger converted β-cyclocostunolide (170) to two oxygenated metabolites (173, 174, 178–181) of which 173 was predominant. It is suggested that compound 173 and 174 might be formed during biotransformation period since metabolite media after 7 days was acidic (pH 2.7). Surprisingly, A. cellulosae gave a sole 11β,13-dihydro-β-cyclocostunolide (182), which was abnormally folded in the mycelium of A. cellulosae as a crystal form after biotransformation of 170. On the other hand, the

801

Biotransformation of Sesquiterpenoids and Their Related Compounds

2

1

A. cellulosae 13 3 days

H

O 169

H

O 177

O

O

A. niger 3 days OH O

O

H

O 173

O

H

O 174

O O

2

2

13 H

OH

O

175

O

O

H

COOEt 13 CH2-S-CH2-CH

O

176

O

OH

FIGURE 23.54  Biotransformation of α-cyclocostunolide (169) by Aspergillus niger and Aspergillus cellulosae. OH O

21 H

O 174

O

O

2 13 H

O 169

H 177

O

H

O

173

O

O O

HS-CH2-CH-COOEt OH O

O 2

13 CH2-S-CH2-CH-COOEt

H

O 176

O

OH

2

13 H

A. niger A. cellulosae

175

OH

O O

FIGURE 23.55  Possible pathway of biotransformation of α-cyclocostunolide (169) by Aspergillus niger and Aspergillus cellulosae.

802

Handbook of Essential Oils

2

1 13

H

O 170

O A. niger (7 days)

OH O

O

2 H

2 H

O

H

O

174

O

173

HO

1

O

178

O

O

OH HO

HO

2 H 179

H

O O

OH O O

180

HO 2 H 181

13 CH2-S-CH2-CH-COOEt

O O

OH

FIGURE 23.56  Biotransformation of β-cyclocostunolide (170) by Aspergillus niger.

metabolites were normally liberated in medium outside of the mycelium of A. niger and B. dothidea (Hashimoto et al., 1999c, 2001a,b) (Figure 23.56). B. dothidea has no stereoselectivity to reduce the C11–C13 double bond of β-cyclocostunolide (170) since this organism gave two dihydro derivatives, 182 (16.7%) and 183 (37.8%), respectively, as shown in Figure 23.57. It is noteworthy that both α- and β-cyclocostunolides were biotransformed by A. niger to give the sulfur-containing metabolites (176, 181). Possible biogenetic pathway of 170 is shown in Figure 23.58. When γ-cyclocostunolide (171) was cultivated in A. niger to give dihydro-α-santonin (187%, 25%) and its related C11–C13, dihydro derivatives (184–186, 188, 189) were obtained as a small amount. Compound 186 was recultivated for 2 days by the same organism as mentioned earlier to afford 187 (25%) and 5β-hydroxy-α-cyclocostunolide (189%, 54%). Recultivation of 185 for 2 days by A. niger afforded compound 187 as a sole metabolite. During the biotransformation of 171, no sulfur-containing product was obtained. Both A. cellulosae and B. dothidea produced only dihydroγ-cyclocostunolide (184) from the substrate (171) (Hashimoto et al., 1999c, 2001a,b) (Figure 23.59). Santonin (190) has been used as vermicide against roundworm. C. blakesleeana and A. niger converted 190–187 (Atta-ur-Rahman et al., 1998). When 187 was fed by A. niger for 1 week to give

803

Biotransformation of Sesquiterpenoids and Their Related Compounds NOE H

H7

67 H

O 170

B. dothidea (16.7%) A. cellulosae (78.0%) NaBH4 /EtOAc (76.8%)

H 6 11 H 13 O 182 O 13

11

NOE H

H

O H

B. dothidea (37.8%) A. cellulosae (0.0%) NaBH4 /EtOAc (1.2%)

H

O 183

O

FIGURE 23.57  Biotransformation of β-cyclocostunolide (170) by Aspergillus cellulosae and Botryosphaeria dothidea.

2

HO

1 4 H

O 170

HO

2

13

11

13 CH2-S-CH2-CH-COOEt

H

O 181

O

O

O 182

A. niger A. cellulosae

O 180

O

OH HO

2 13

11

OH

H

OH

HO

H

2

21

H

O 178

O

H

O 179

O

O

OH O

O 4 H

O

2 4

O O

H

H

O

173

1

O

O

174

O

FIGURE 23.58  Possible pathway of biotransformation of β-cyclocostunolide (170) by Aspergillus niger and Aspergillus cellulosae.

2β-hydroxy-1,2-dihydro-α-santonin (188%, 39%) as well as 1β-hydroxy-1,2-dihydro-α-santonin (195, 6.5%), 9β-hydroxy-1,2-dihydro-α-santonin (196, 6.9%), and α-santonin (190, 5.4%), which might be obtained from dehydroxylation of 188, as a minor component (Hashimoto et al., 2001a,b), compound 188 was isolated from the crude metabolite of γ-cyclocostunolide (171) by A. niger as mentioned earlier (Figure 23.60). It was treated with A. niger for 7 days to give 191 (18.3%), 192 (2.3%), 193 (19.3%), and 194 (3.5%) of which 193 was the major metabolite. Compound 191 was isolated from dog’s urine after the oral administration of 190. The structure of compound 194 was established as lumisantonin obtained by the photoreaction of 190. α-Santonin 190 was not converted into 1,2-dihydro derivative by A. niger, whereas the other strain of A. niger gave a single product, 1,2-dihydro-α-santonin (187) (Hashimoto et al., 2001a,b) (Figure 23.61).

804

Handbook of Essential Oils HO HO

3

5

2

O O

185

O 188

O

11 13

O

O 171

184

O

O

O

3 O 187

O

O

A. niger A. cellulosae B. dothidea

HO

5

3

OH

O 186

O

189

O

O

FIGURE 23.59  Biotransformation of γ-cyclocostunolide (171) by Aspergillus niger, Aspergillus cellulosae, and Botryosphaeria dothidea. OH

A. niger O

HO O

O

O

187

O O

O O

188

195

O

OH

O O

196

O

FIGURE 23.60  Biotransformation of dihydro-α-santonin (187) by Aspergillus niger.

Ata and Nachtigall (2004) reported that α-santonin (190) was incubated with Rhizopus stolonifer to give (187a) and (183b), while with Cunninghamella bainieri, C. echinulata, and M. plumbeus to afford the known 1,2-dihydro-α-santonin (187) (Figure 23.62). α-Santonin (190) and 6-epi-α-santonin (198) were cultivated in Absidia coerulea for 2 days to give 11β-hydroxy- (191, 71.4%) and 8α-hydroxysantonin (197, 2.0%), while 6-epi-santonin (198) afforded four major products (199–201, 206) and four minor analogues (202, 203–205). Asparagus

805

Biotransformation of Sesquiterpenoids and Their Related Compounds OH– 10

O

9

9

OH

10

4

O 190

H+ O

13

11

HO

O 193

O

O

O

O

O

O

OH

O

4

O

O

O O

11

O

O

191

A. niger Dog

13

O O 192

OH

O O 194

O

O

FIGURE 23.61  Biotransformation of α-santonin (190) by Aspergillus niger and dogs.

b,c,d

O O 187

O

a O

O

O 190

O

187a

a a: b: c: d:

Rhizopus stolonifera Cunninghamella bainieri Cunninghamella echinulata Mucor plumbeus

O O

O O 187b

O

FIGURE 23.62  Biotransformation of α-santonin (190) by Rhizopus stolonifer, Cunninghamella bainieri, Cunninghamella echinulata, and Mucor plumbeus.

officinalis also biotransformed α-santonin (190) into three eudesmanolides (187, 207, 208) and a guaianolide (209) in a small amount. 6-epi-Santonin (198) was also treated in the same bioreactor as mentioned earlier to give 199 and 206, the latter of which was obtained as a major metabolite (44.7%) (Yang et al., 2003) (Figure 23.63).

806

Handbook of Essential Oils OH Absidia coerulea O

O

HO O

O O

198

199 (13.4%)

OH

HO

205

O

203

O

201

OH O CO2H

O

O

O O

O

O

200 (11.0%)

O

O O 202

O

OH

OH O

O

O

O

O

204

206

OH Asparagus officinalis O

O

O O

O 198

O

CO2H

O

199

206 (44.7%)

FIGURE 23.63  Biotransformation of α-epi-santonin (198) by Absidia coerulea and Asparagus officinalis.

α-Santonin (190) was incubated in the cultured cells of Nicotiana tabacum and the liverwort M. polymorpha. N. tabacum cells gave 1,2-dihydro-α-santonin (187) (50%) for 6 days. The latter cells also converted α-santonin to 1,2-dihydro-α-santonin, but conversion ratio was only 28% (Matsushima et al., 2004) (Figure 23.64). OH Absidia coerulea

OH

O

O O O

190

O O

O O

191 (71.4%)

197 (2.0%)

O

Asparagus officinalis O

O

O O 190

H

O O

O

187

O

207

Marchantia polymorpha cell Nicotiana tabacum

O

OH

H O O

O O 187

O

O

HO 208

O

O 209

O

FIGURE 23.64  Biotransformation of 6-epi-α-santonin (190) by Absidia coerulea, Asparagus officinalis, Marchantia polymorpha, and Nicotiana tabacum.

807

Biotransformation of Sesquiterpenoids and Their Related Compounds O 2 O

3

1

2

R. nigricans

4

HO

1

O

O

O

O

O

O

198

O

O

211

R. nigricans O

H

HO

H

O

H

O

O

210

4

HO O

O

212: (4S) 213: (4R)

O

FIGURE 23.65  Biotransformation of α-epi-santonin (198) and tetrahydrosantonin (210) by Rhizopus nigricans.

6-epi-α-Santonin (198) and its tetrahydro analogue (210) were also incubated with fungus Rhizopus nigricans to give 2α-hydroxydihydro-α-santonin (211) (Amate et al., 1991), the epimer of 188 obtained from the biotransformation of dihydro-α-santonin (187) by A. niger (Hashimoto et al., 2001a,b). The product 211 might be formed via 1,2-epoxide of 198. Compound 210 was converted through carbonyl reduction to furnish 212 and 213 under epimerization at C4 (Amate et al., 1991) (Figure 23.65). 1,2,4β,5α-Tetrahydro-α-santonin (214) prepared from α-santonin (190) was treated with A. niger to afford six metabolites (215–220) of which 219 was the major product (21%). When the substrate (214) was treated with CYP450 inhibitor, 1-aminobenzotriazole, only 215 was obtained without its homologues, 216–220, while the C4 epimer (221) of 214 was converted by the same microorganism to afford a single metabolite (222) (73%). Further oxidation of 222 did not occur. This reason might be considered by the steric hindrance of β-(axial) methyl group at C4 (Hashimoto et al., 2001a,b) (Figure 23.66). O

OH 2 3 O

1 4

1 H 214

HO

H

O

216

O

HO HO

H 215

1

3

HO

O

HO

3 HO

4

220

O O O

O

O O

219

O

H

H

O

2

O

2 3

O

218

O

O

217

O

H

H

O

2 3

O

A. niger A. niger + Cytochrome P-450 inhibitor

HO

3

2 3 H

O O

FIGURE 23.66  Biotransformation of 1,2,4β,5α-tetrahydro-α-santonin (214) by Aspergillus niger.

808

Handbook of Essential Oils

3 O

A. niger 3 days HO

4 H

O 221

3

4 H

O 222

O

OH

A. niger

OH

O

O O

223

O

224

O

A. quardilatus

O

O O

225

OH A. niger

O

O 226

O

227

O

FIGURE 23.67  Biotransformation of C4-epimer (221) of 214, 7α-hydroxyfrullanolide (223), and frullanolide (226) by Aspergillus niger and Aspergillus quardilatus.

7α-Hydroxyfrullanolide (223) possessing cytotoxicity and antitumor activity, isolated from Sphaeranthus indicus (Asteraceae), was bioconverted by A. niger to afford 13R-dihydro derivative (224). The same substrate was also treated in Aspergillus quardilatus (wild type) to give 13-acetyl product (225) (Atta-ur-Rahman et al., 1994) (Figure 23.67). Incubation of (−)-frullanolide (226), obtained from the European liverwort Frullania tamarisci subsp. tamarisci that causes a potent allergenic contact dermatitis, was incubated by A. niger to give dihydrofrullanolide (227), nonallergenic compound in 31.8% yield. In this case, C11–C13 dihydro derivative was not obtained (Hashimoto et al., 2005). Guaiane-type sesquiterpene hydrocarbon, (+)-γ-gurjunene (228), was treated in plant pathogenic fungus G. cingulata to give two diols, (1S,4S,7R,10R)-5-guaien-11,13-diol (229) and (1S,4S,7R,10S)5-guaien-10,11,13-triol (230) (Miyazawa et al., 1997a, 1998a) (Figure 23.68). G. cingulata converted guaiol (231) and bulnesol (232) to 5,10-dihydroxy (233) and 15-hydroxy derivative (234), respectively (Miyazawa et al., 1996a) (Figure 23.69). When Eurotium rubrum was used as the bioreactor of guaiene (235), rotundone (236) was obtained (Sugawara and Miyazawa, 2004). Guaiol (231) was also transformed by A. niger to give a cyclopentane derivative, pancherione (237), and two dihydroxy guaiols (238, 239) (Morikawa et al., 2000), of which 237 was obtained from the same substrate using E. rubrum for 10 days (Sugawara and Miyazawa, 2004; Miyazawa and Sugawara, 2006) (Figure 23.70).

809

Biotransformation of Sesquiterpenoids and Their Related Compounds H

H

OH

H

A. niger

228

OH OH

229

230

OH

OH

FIGURE 23.68  Biotransformation of (+)-γ-gurjunene (228) by Glomerella cingulata.

OH G. cingulata OH

OH

231

OH

233 OH

G. cingulata H

H

OH

232

234

OH

FIGURE 23.69  Biotransformation of guaiol (221) and bulnesol (232) by Glomerella cingulata.

O Eurotium rubrum

235

236 O Eurotium rubrum A. niger

231

OH A.

ni

237

OH

ge

r

OH HO OH 238

OH OH

239

OH

FIGURE 23.70  Biotransformation of guaiene (235) by Eurotium rubrum and guaiol (231) by Aspergillus niger and Eurotium rubrum.

810

Handbook of Essential Oils H

OH

4

A. niger

10 5 O

3 days

H

H HO

OH

H

OH

O

O

244 (5.0%)

H

H HO

O

O

OH

H

OH

O

245 (9.6%)

O

243 (13.7%)

242 (11.3%)

OH

H

O O

O

OH

H

O

O

241 (12.3%)

O

O Parthenolide (240)

O

H

O

246 (5.1%)

O

FIGURE 23.71  Biotransformation of parthenolide (240) by Aspergillus niger.

Parthenolide (240), a germacrane-type lactone, isolated from the European feverfew (Tanacetum parthenium) as a major constituent, shows cytotoxic, antimicrobial, antifungal, anti-inflammatory, antirheumatic activity, apoptosis inducing, and NF-βB and DNA binding inhibitory activity. This substrate was incubated with A. niger in Czapek-peptone medium for 2 days to give six metabolites (241, 12.3%; 242, 11.3%; 243, 13.7%; 244, 5.0%; 245, 9.6%; 246, 5.1%) (Hashimoto et al., 2005) (Figure 23.71). Compound 244 was a naturally occurring lactone from Michelia champaca (Jacobsson et al., 1995). The stereostructure of compound 243 was established by x-ray crystallographic analysis. When parthenolide (240) was treated in A. cellulosae for 5 days, two new metabolites, 11β,13dihydro-(247, 43.5%) and 11α,13-dihydroparthenolides (248, 1.6%), were obtained together with the same metabolites (241, 5.3%; 243, 11.2%; 245, 10.4%) as described earlier (Figure 23.72). Possible metabolic root of 240 has been shown in Figure 23.73 (Hashimoto et al., 2005). Galal et al. (1999) reported that Streptomyces fulvissimus or R. nigricans converted parthenolide (240) into 11α-methylparthenolide (247) in 20%–30% yield, while metabolite 11β-hydroxyparthenolide (248) was obtained by incubation of 240 with R. nigricans and Rhodotorula rubra. In addition to the metabolite 247, S. fulvissimus gave a minor polar metabolite, 9β-hydroxy derivative (248a) in low yield (3%). The same metabolite (248a) was obtained from 247 by fermentation of S. fulvissimus as H

H OH O A. cellulosae O

OH

H

H HO

O

241 O (5.3%)

243 (11.2%)

O

O

H

O

5 days O

O Parthenolide (240)

O

247 (43.4%)

O

O

248 (1.6%)

O

FIGURE 23.72  Biotransformation of parthenolide (240) by Aspergillus cellulosae.

OH

H OH O 245 O (10.4%)

811

Biotransformation of Sesquiterpenoids and Their Related Compounds OH

H

OH

H

H –H2O

[H] H2O

10

A. n

5 4 O H+

O

11

246

13 A .n

O Parthenolide (240)

H OH O

H OH O

r ige

ige

r

H

O

O

245

OH

H [H]

O

O

244 Michampanolide

241

O

OH 10

4 H HO

OHH O

H HO

13

5 O

243

O

FIGURE 23.73  Possible pathway of biotransformation of parthenolide (240).

a minor constituent. Furthermore, 14-hydroxyparthenolide (248b) was obtained from 240 and 247 as a minor component (4%) by R. nigricans (Figure 23.74). Pyrethrosin (248c), a germacranolide, was treated in the fungus R. nigricans to afford five metabolites (248d–248 h). Pyrethrosin itself and metabolite 248e displayed cytotoxic activity against human malignant melanoma with IC50 4.20 and 7.5 mg/mL, respectively. Metabolite 248 h showed significant in vitro cytotoxic activity against human epidermoid carcinoma (KB cells) and against human ovary carcinoma with IC50 < 1.1 and 8.0 mg/mL, respectively. Compounds 248f and 248i were active against Cryptococcus neoformans with IC50 35.0 and 25 mg/mL, respectively, while 248a and 248 g showed antifungal activity against Candida albicans with IC50 30 and 10 mg/mL. Metabolites 248 g and its acetate (248i), derived from 248 g, showed antiprotozoal activity against Plasmodium falciparum with IC50 0.88 and 0.32 mg/mL, respectively, without significant toxicity. Compound 248i also exhibited pronounced activity against the chloroquine-resistant strain of P. falciparum with IC50 0.38 mg/mL (Galal, 2001) (Figure 23.75). (−)-Dehydrocostus lactone (249), inhibitors of nitric oxide synthases and TNF-α, isolated from Saussurea radix, was incubated with C. echinulata to afford (+)-11α,13-dihydro­dehydrocostuslactone (250a). The epoxide (251) and a C11 reduced compound (250) were obtained by the aforementioned microorganisms (Galal, 2001). C. echinulata and R. oryzae bioconverted 249 into C11/C13 dihydrogenated (250) and C10/C14 epoxidated product (251). Treatment of 252a in C. echinulata and R. oryzae gave (–)-16-(1-methyl1-propenyl)eremantholanolide (252b) (Galal, 2001) (Figure 23.76). The same substrate (249) was fed by A. niger for 7 days to afford four metabolites, costus lactone (250), and their derivatives (251–253), of which 251 was the major product (28%); while the same substrate was cultivated with A. niger for 10 days, two minor metabolites (254, 255) were newly obtained in addition to 252 and 253 of which the latter lactone was predominant (20.7%) (Hashimoto et al., 2001a,b) (Figure 23.77). When compound 249 was treated with A. niger in the presence of 1-aminobenzotriazole, 249 was completely converted into 11β,13-dihydro derivative (250) for 3 days; however, further biodegradation did not occur for 10 days (Hashimoto et al., 1999a, 2001a,b). The same substrate (249) was cultivated with A. cellulosae IFO to furnish 11,13-dihydro- (250) (82%) for only 1 day and then the product (250) slowly oxidized into 11,13-dihydro-8β-hydroxycostuslactone (256) (1.6%) for 8 days (Hashimoto et al., 1999a, 2001a,b) (Figure 23.78). The lactone (249) was biodegraded by the plant pathogen B. dothidea for 4 days to give the metabolites 250 (37.8%) and 257 (8.6%), while A. niger IFO-04049 (4 days) and A. cellulosae for

812

Handbook of Essential Oils

a O

O

O O

247

O

248 OH

O

b O

O

O O

240

O

O O

247

c

O

248a

O

OH O

O

248b

O

a OH O O

O

247

O

248b OH O

O

b

O a: Rhizopus nigricans b: Streptomyces fulvissimus c: Rhodotorula rubra

O

248a

O

FIGURE 23.74  Biotransformation of parthenolide (240) and its dihydro derivative (247) by Rhizopus nigricans, Streptomyces fulvissimus, and Rhodotorula rubra. OH

OH

O

O

O

O

R. nigricans

HO

O O

H

H

OAc 248d OH

OH

248c

O

O

O OAc

OH 248e

O

O H

HO

OH

H

OAc 248g

248f

OAc

OH

O

O

O

O H

H OAc 248h

HO

H

OAc 248i

FIGURE 23.75  Biotransformation of pyrethrosin (248c) by Rhizopus nigricans.

813

Biotransformation of Sesquiterpenoids and Their Related Compounds H a H

H

O

250a H

O

H

H

O

249

O

b

O

H a: Cunninghamella echinulata b: Rhizopus oryzae

H

O

250

O

251

O O

O

O

O

252a

OH

O

O O

a,b

O

O

O

O

O

a: Cunninghamella echinulata b: Rhizopus oryzae

252b

FIGURE 23.76  Biotransformation of (−)-dehydrocostuslactone (249) and rearranged guaianolide (252a) by Cunninghamella echinulata and Rhizopus oryzae.

1 day gave only 250. Thus, B. dothidea demonstrated low stereoselectivity to reduce the C11–C13 double bond (Hashimoto et al., 2001a,b). Furthermore, three Aspergillus species, A. niger IFO 4034, Aspergillus awamori IFO 4033, and Aspergillus terreus IFO6123, were used as bioreactors for compounds 249. A. niger IFO 4034 gave three products (250–252), of which 252 was predominant (56% in GC-MS). A. awamori IFO 4033 and A. terreus IFO 6123 converted 249 to give 250 (56% from A. awamori, 43% from A. terreus) and 252 (43% from A. awamori, 57% from A. terreus), respectively (Hashimoto et al., 2001a,b) (Figure 23.79). Vernonia arborea (Asteraceae) contains zaluzanin D (258) in high content. Ten microorganisms were used for the biotransformation of compound 258. B. cinerea converted 258 into 259 and 260 (85%:15%) and Fusarium equiseti gave 259 and 260 (33%:66%). C. lunata, Colletotrichum lindemuthianum, Alternaria alternata, and Phyllosticta capsici produced 259 as the sole metabolite in good yield, while Sclerotinia sclerotiorum and Rhizoctonia solani gave deactyl product (261) as a sole product and 260 and 262–264, among which 263 and 264 are the major products, respectively. Reduction of C11–C13 exocylic double bond is the common transformation of α-methylene-γbutyrolactone (Kumari et al., 2003) (Figure 23.80). Incubation of parthenin (264a) with the fungus B. bassiana in modified Richard’s medium gave C11–C13 reduced product (264b) in 37% yield, while C11 α-hydroxylated product (264c) was obtained in 32% yield from the broth of the fungus Sporotrichum pulverulentum using the same medium (Bhutani and Thakur, 1991) (Figure 23.81). Cadina-4,10(15)-dien-3-one (265) possessing insecticidal and ascaricidal activity, from the Jamaican medicinal plant H. verticillata, was metabolized by C. lunata ATCC 12017 in potato dextrose to give its 12-hydoxy- (266), 3α-hydroxycadina-4,10(15)-dien (267), and

814

Handbook of Essential Oils 14 H 10

3 H

11

O

249

13

O

A. niger 7 days

A. niger 10 days

H HO H

11

O

250

O

14 O

H

13

3 H

251

252 O HO H

H O

H

O

10

O

OH 10

10 H

OH

14 H

H

254

14 OH

O

253 O

O

O

H HO

O

10

3 H

O

255

O

FIGURE 23.77  Biotransformation of (−)-dehydrocostuslactone (249) by Aspergillus niger.

3α-hydroxy-4,5­dihydrocadinenes (268), while 265 was incubated by the same fungus in peptone, yeast, and beef extracts and glucose medium, only 267 and 268 were obtained. Compound 267 derived synthetically was treated in the same fungus C. lunata to afford three metabolites (269–271) (Collins and Reese, 2002) (Figure 23.82). The incubation of the same substrate (265) in M. plumbeus ATCC 4740 in high-iron-rich medium gave 270, which was obtained from C. lunata mentioned earlier, 268, 272, 273, 277, 278, and 279. In low-iron medium, this fungus converted the same substrate 265 into three epoxides (274–276), a tetraol (280) with common metabolites (268, 273, 277, 278), and 271, which was the same metabolite used by C. lunata (Collins et al., 2002a). It is interesting to note that only epoxides were obtained from the substrate (265) by Mucor fungus in low-iron medium (Figure 23.83). The same substrate (265) was incubated with the deuteromycete fungus B. bassiana, which is responsible for the muscardine disease in insects, in order to obtain new functionalized analogues with improved biological activity. From compound 265, nine metabolites were obtained. The insecticidal potential of the metabolites (267, 268, 268a–268f) were evaluated against Cylas formicarius. The metabolites (273, 268, 268d) showed enhanced activity compared with the substrate (265). The plant growth regulatory activity of the metabolites against radish seeds was tested. All the compounds showed inhibitory activity; however, their activity was less than colchicine (Buchanan et al., 2000) (Figure 23.84).

815

Biotransformation of Sesquiterpenoids and Their Related Compounds 14

14

H

H

10

3

4

8 6 7 H 11 O 249

HO

3

HO H

13

O

O

H 10

3 H

O

252 O

O

255

14

OH

14

O

H

O

OH

H

10

10

H H H

O

11

O

13

O

254

14

O

250

H

O

H

O

H

OH 10

10 H H 8 H

OH

O

H

O

14 OH

O

253 O

251 O A. niger A. niger + Cyt.-P450 inhibitor

O O

256

A. cellulosae

FIGURE 23.78  Possible pathway of biotransformation of (−)-dehydrocostuslactone (249) by Aspergillus niger and Aspergillus cellulosae. 14

H

O

H 10

A. niger A. cellulosae A. awamori A. terreus Botryosphaeria dothidea

14

H

10

H

O

249

11

H

13

O

250

13

O HO

11

O

H

O

251

H

H

H

H

O

3

O

252

O

13

O

257

11

O

FIGURE 23.79  Biotransformation of (−)-dehydrocostuslactone (249) by Aspergillus species and Botryosphaeria dothidea.

816

Handbook of Essential Oils H

H HO

H

O

259

O

H

260

O

O

HO

H

O

261

S.

B. cinerea F. equiseti C. lunata C. lindemuthianum B. theobromae F. oxysporum A. alternata P. capsici

H

sc

H

ler ot io ru m

AcO

O

AcO H

O

258

O

R. solani H

H

H

O

O

HO H

H

O O

262

H HO

H

O O

263

H

O

264

O

O

260

O

FIGURE 23.80  Biotransformation of zaluzanin D (258) by various fungi.

HO

HO

HO

S. pulverulentum

O

O

O 264c

B. bassiana

OH O

O

O 264a

O

O 264b

O

FIGURE 23.81  Biotransformation of parthenin (264a) by Sporotrichum pulverulentum and Beauveria bassiana.

Cadinane-type sesquiterpene alcohol (281) isolated from the liverwort M. taylorii gave a primary alcohol (282) by A. niger treatment (Morikawa et al., 2000) (Figure 23.85). Fermentation of (−)-α-bisabolol (282a) possessing an anti-inflammatory activity with the plant pathogenic fungus G. cingulata for 7 days yielded oxygenated products (282b–282e) of which compound 282e was predominant. 3,4-Dihydroxy products (282b, 282d, 282e) could be formed by hydrolysis of the 3,4-epoxide from 282a and 282c (Miyazawa et al., 1995c) (Figure 23.86). El Sayed et al. (2002) reported microbial and chemical transformation of (S)-(+)-curcuphenol (282 g) and curcudiol (282n), isolated from the marine sponges Didiscus axiata. Incubation of compound 282 g with Kluyveromyces marxianus var. lactis resulted in the isolation of six

817

Biotransformation of Sesquiterpenoids and Their Related Compounds

O

H C. lunata

H

O

H

HO

H

H

266

265

H

HO

H

H OH

267

268

C. lunata OH

HO

H

HO

H

HO

H

H

H

269

270

OH

OH

271

FIGURE 23.82  Biotransformation of cadina-4,10(15)-dien-3-one (265) by Curvularia lunata.

O

H

M. plumbeus

O

H

H

O

R1

H

H

H H

R1

R2

R1

R2

272: R1=OH, R2=OH 273: R1=H, R2=OH

270: R1=H, R2=OH 271a: R1=OH, R2=H

O

H

O

R2

274: R1=OH, R2=OH 275: R1=H, R2=OH

H 265

HO

H

O

HO

H

H H 276

HO

H

OH

H

HO

H

H R1

R2

268: R1=R2=H 277: R1=OH, R2=H 278: R1=H, R2=OH

OH OH

H

OH 279

OH 280

FIGURE 23.83  Biotransformation of cadina-4,10(15)-dien-3-one (265) by Mucor plumbeus.

metabolites (3–8, 282h–282j). The same substrate was incubated with Aspergillus alliaceus to give the metabolites (282p, 282q, 282 s) (Figure 23.87). Compounds 282 g and 282n were treated in Rhizopus arrhizus and Rhodotorula glutinus for 6 and 8 days to afford glucosylated metabolites, 1α-D-glucosides (282o) and 282r, respectively. The substrate itself showed antimicrobial activity against C. albicans, C. neoformans, and MRSAresistant Staphylococcus aureus and S. aureus with MIC and MFC/MBC ranges of 7.5–25 and 12.5–50 mg/mL, respectively. Compounds 282 g and 282 h also exhibited in vitro antimalarial activity against P. falciparum (D6 clone) and P. falciparum (W2 clone) of 3600 and 3800 ng/mL (selective index [S.I.] > 1.3), 1800 (S.I. > 2.6), and 2900 (S.I. > 1.6), respectively (El Sayed et al., 2002) (Figure 23.87).

818

Handbook of Essential Oils

O

H

H

O

O

B. bassiana

H

H

H

268a

HO

H

H

H

H

HO

H

H

268c

268 H

HO

H

H

HO

H

268d

OH

273

H

HO

267 HO

H OH

268b

H

265

H

O

H

OH

268e

268f

FIGURE 23.84  Biotransformation of cadina-4,10(15)-dien-3-one (265) by Beauveria bassiana.

OH

OH A. niger OH 282

281

FIGURE 23.85  Biotransformation of cadinol (281) by Aspergillus niger.

HO

H

OH

H O

H

HO

OH G. cingulata

282a

282c OH

282b

HO

H

HO

H O

O HO

HO

282d

OH

FIGURE 23.86  Biotransformation of β-bisabolol (282a) by Glomerella cingulata.

282e

OH

819

Biotransformation of Sesquiterpenoids and Their Related Compounds

Kluyveromyces marxianus

OH

OH

282g

OH

OH

OHC 282k

OGlc

OH HO

HO 282h

Rhizopus arrhizus OH

OH

OH

282j

282i

OH

OH

HO 282l

HO 2C

H

H 282m

OH

OH

282o

OH 282p OH

OH Aspergillus alliaceus

O

282n

OH

Rhodotorula glutinus

OH 282q OH

OGlc COOH 282s

OH 282r

FIGURE 23.87  Biotransformation of (S)-(+)-curcuphenol (282 g) by Kluyveromyces marxianus and Rhizopus arrhizus and curcudiol (282n) by Aspergillus alliaceus and Rhodotorula glutinus.

Artemisia annua is one of the most important Asteraceae species as antimalarial plant. There are many reports of microbial biotransformation of artemisinin (283), which is active antimalarial rearranged cadinane sesquiterpene endoperoxide, and its derivatives to give novel antimalarials with increased activities or differing pharmacological characteristics. Lee et  al. (1989) reported that deoxoartemisinin (284) and its 3α-hydroxy derivative (285) were obtained from the metabolites of artemisinin (283) incubated with Nocardia corallina and Penicillium chrysogenum (Figure 23.88).

820

Handbook of Essential Oils HO

H a,b,c

O O H

4

O O O H

6

10

9

7

8

5 O

O H

O

H

3

H

O H

H

O

O

O 285 (15%)

284 (38%)

H H

H O a

Artemisinin (283)

AcO O H

a: A. niger b: Nocardia sp. c: Penicillium sp.

O H

O

HO

H

H

O

O

O

286 (8%)

287 (5%)

FIGURE 23.88  Biotransformation of artemisinin (283) by Aspergillus niger, Nocardia corallina, and Penicillium chrysogenum.

Zhan et al. (2002) reported that incubation of artemisinin (283) with C. echinulata and A. niger for 4 days at 28°C resulted in the isolation of two metabolites, 10β-hydroxyartemisinin (287a) and 3α-hydroxydeoxyartemisinin (285), respectively. Compound 283 was also biotransformed by A. niger to give four metabolites, deoxyartemisinin (284%, 38%), 3α-hydroxydeoxyartemisinin (285%, 15%), and two minor products (286%, 8% and 287%, 5%) (Hashimoto et al., 2003d). Artemisinin (283) was also bioconverted by C. elegans. During this process, 9β-hydroxyartemisinin (287b, 78.6%), 9β-hydroxy-8α-artemisinin (287c, 6.0%), 3α-hydroxydeoxoartemisinin (285, 5.4%), and 10β-hydroxyartemisinin (287d, 6.5%) have been formed. On the basis of quantitative structure– activity relationship and molecular modeling investigations, 9β-hydoxy derivatization of artemisinin skeleton may yield improvement in antimalarial activity and may potentially serve as an efficient means of increasing water solubility (Parshikov et al., 2004) (Figure 23.89). Albicanal (288) and (−)-drimenol (289) are simple drimane sesquiterpenoids isolated from the liverwort, Diplophyllum serrulatum, and many other liverworts and higher plants. The latter compound was incubated with M. plumbeus and R. arrhizus. The former microorganism converted 289 to 6,7α-epoxy- (290), 3β-hydroxy- (291), and 6α-drimenol (292) in the yields of 2%, 7%, and 50%, respectively. On the other hand, the latter species produced only 3β-hydroxy derivative (291) in 60% yield (Aranda et al., 1992) (Figure 23.90). (−)-Polygodial (293) possessing piscicidal, antimicrobial, and mosquito-repellant activity is the major pungent sesquiterpene dial isolated from Polygonum hydropiper and the liverwort Porella vernicosa complex. Polygodial was incubated with A. niger; however, because of its antimicrobial activity, no metabolite was obtained (Sekita et al., 2005). Polygodiol (295) prepared from polygodial (293) was also treated in the same manner as described earlier to afford 3β-hyrdoxy (297), which was isolated from Marasmius oreades as antimicrobial activity (Ayer and Craw, 1989), and 6α-hydroxypolygodiol (298) in 66%–70% and 5%–10% yields, respectively (Aranda et al., 1992). The same metabolite (297) was also obtained from polygodiol (295) as a sole metabolite from the culture broth of A. niger in Czapek-peptone medium for 3 days in 70.5% yield (Sekita et al., 2005), while the C9 epimeric product (296) from isopolygodial (294) was incubated with M. plumbeus to afford 3β-hydroxy- (299) and 6α-hydroxy derivative (300) in low yields, 7% and 13% (Aranda et al.,

821

Biotransformation of Sesquiterpenoids and Their Related Compounds H O O H

OH

O H

O OH

287a

a H O O H

H c

O

O O H

H

O

b HO

HO

H

O H

H

O O 285

a: Cunninghamella echinulata b: Aspergillusniger c: Cunningham ellaelegans

OH

O H

O

O

O

287b

287c

H

H

OH

O

O

O O H

O O H

H

O

O 283

H

OH

O

O O H

H

O

H

O

O

O

285

287d

FIGURE 23.89  Biotransformation of artemisinin (283) by Cunninghamella echinulata, Cunninghamella elegans, and Aspergillus niger.

CHO

H

288 OH

OH

OH Mucor plumbeus

OH

O

H

H

289

290 (2%) Rhizopus arrhizus*

HO

H

H

291 (7%) (60%)*

292 (50%)

FIGURE 23.90  Biotransformation of drimenol (289) by Mucor plumbeus and Rhizopus arrhizus.

OH

822

Handbook of Essential Oils OH

CHO

OH M. plumbeus* OH R. arrhizus*

CHO LiAlH4

A. niger**

293

OH 298 (5%–10%)*

297 (66%–70%)* (70.5%)**

OH CHO

OH

OH

HO

295

CHO

OH

OH OH M. plumbeus

OH OH

OH

HO 296

294

OH 300 (13%)

299 (7%)

FIGURE 23.91  Biotransformation of polygodiol (295) by Mucor plumbeus, Rhizopus arrhizus, and Aspergillus niger. OAc

OAc

OH

OH OAc

A. niger 3 days

301

OAc HO 302 (47.2%)

FIGURE 23.92  Biotransformation of drim-9α-hydroxy-11,12-diacetoxy-7-ene (301) by Aspergillus niger.

1992). Drim-9α-hydroxy-11β,12-diacetoxy-7-ene (301) derived from polygodiol (295) was treated in the same manner as described earlier to yield its 3β-hydroxy derivative (302%, 42%) (Sekita et al., 2005) (Figures 23.91 and 23.92). Cinnamodial (303) from the Malagasy medicinal plant, Cinnamosma fragrans, was also treated in the same medium including A. niger to furnish three metabolites, respectively, in very law yields (304, 2.2%; 305, 0.05%; and 306, 0.62%). Compound 305 and 306 are naturally occurring cinnamosmolide, possessing cytotoxicity and antimicrobial activity, and fragrolide. In this case, the introduction of 3β-hydroxy group was not observed (Sekita et al., 2006) (Figure 23.93). Naturally occurring rare drimane sesquiterpenoids (307–314) were biosynthesized by the fungus Cryptoporus volvatus with isocitric acids. Among these compounds, in particular, cryptoporic acid E (312) possesses antitumor promoter, anticolon cancer, and very strong superoxide anion radical scavenging activities (Asakawa et al., 1992). When the fresh fungus allowed standing in moisture condition, olive fungus Paecilomyces variotii grows on the surface of the fruit body of this fungus. C. volvatus infected 2 kg of the fresh fungus for 1 month, followed by the extraction of methanol to give the crude extract and then purification using silica gel and Sephadex LH-20 to give five metabolites (316, 318–321), which were not found in the fresh fungus (Takahashi et al., 1993b). Compound 318 was also isolated from the liverworts, Bazzania and Diplophyllum species (Asakawa, 1982, 1995) (Figure 23.94). Liverworts produce a large number of enantiomeric mono-, sesqui-, and diterpenoids to those found in higher plants and lipophilic aromatic compounds. It is also noteworthy that some liverworts produce both normal and its enantiomers. The more interesting phenomenon in the chemistry of liverworts is that the different species in the same genus, for example, Frullania tamarisci subsp. tamarisci and Frullania dilatata, produce totally enantiomeric terpenoids. Various sesqui- and

823

Biotransformation of Sesquiterpenoids and Their Related Compounds 11 CHO OH CHO 9 12 6

HO HO A. niger

9

4 days

6

OAc

O 11 12

O 11 12

O

9

8

O

6 OAc

OAc 304 (2.2%)

Cinnamodial (303)

O

HO

O 306 (0.20%) Fragrolide

305 (0.05%) Cinnamosmolide

FIGURE 23.93  Biotransformation of cinnamodial (303) by Aspergillus niger. O

O

O COOMe CO COOH O

COOMe COOMe COOH

H Cryptoporic acid A (307)

H

H COOH COOR COOMe

O COOMe COOMe COOH

OH

COOMe COOH COOH

H Cryptoporic acid B (308)

OH

315

O R=Me Cryptoporic acid C (309) R=H Cryptoporic acid F (310)

OH

OH

O O

H

COOH COOH COOH COOH COOH COOH

H Albicanol (318)

H H Cryptoporic acid H (314)

OH

316

OH

319

OH

O O

O COOMe CO COOH O

H O COOH CO COOMe O Cryptoporic acid D (311)

COOH CO COOH O H

O

H 320 COOH

COOMe CO COOH O H

OH

OH

COOH COOR COOMe

COOH CO COOH O

317

O R=Me Cryptoporic acid E (312) R=H Cryptoporic acid G (313)

HO H 321 COOH

FIGURE 23.94  Biotransformation of cryptoporic acids (307–317, 316) by Paecilomyces variotii.

diterpenoids, bibenzyls, and bisbibenzyls isolated from several liverworts show a characteristic fragrant odor, intensely hot and bitter taste, muscle relaxation, allergenic contact dermatitis, and antimicrobial, antifungal, antitumor, insect antifeedant, superoxide anion release inhibitory, piscicidal, and neurotrophic activity (Asakawa, 1982, 1990, 1995, 1999, 2007, 2008; Asakawa and Ludwiczuk, 2008). In order to obtain the different kinds of biologically active products and to compare the metabolites of both normal and enantiomers of terpenoids, several secondary metabolites of specific liverworts were biotransformed by Penicillium sclerotiorum, A. niger, and A. cellulosae. (−)-Cuparene (322) and (−)-2-hydroxycuparene (323) have been isolated from the liverworts Bazzania pompeana and M. polymorpha, while its enantiomer (+)-cuparene (324) and (+)-2-hydroxycuparene (325) from the higher plant Biota orientalis and the liverwort Jungermannia rosulans. (R)-(−(-α-Cuparenone (326) and grimaldone (327) demonstrate intense flagrance. In order to obtain such compounds from both cuparene and its hydroxy compounds, both enantiomers mentioned earlier were cultivated with A. niger (Hashimoto et al., 2001a) (Figure 23.95). From (−)-cuparene (322), five metabolites (328–332) all of which contained cyclopentanediols or hydroxycyclopentanones were obtained. An aryl methyl group was also oxidized to give primary

824

Handbook of Essential Oils

S

R

(–)-Cuparene (322)

(+)-Cuparene (324)

Bazzania pompeana Marchantia polymorpha

Jungermannia rosulans Biota orientalis (higher plant)

HO (–)–2-Hydroxycuparene (323)

HO (+)–2-Hydroxycuparene (325)

Bazzania pompeana Marchantia polymorpha

Jungermannia rosulans Biota orientalis (higher plant)

O 326

O 327

FIGURE 23.95  Naturally occurring cuparene sesquiterpenoids (322–327).

alcohol, which was further oxidized to afford carboxylic acids (329–331) (Hashimoto et al., 2001a) (Figure 23.96). From (+)-cuparene, six metabolites (333–338) were obtained. These are structurally very similar to those found in the metabolites of (−)-cuparene, except for the presence of an acetonide (336), but they are not identical. All metabolites possess benzoic acid moiety. The possible biogenetic pathways of (+)-cuparene (324) have been proposed in Figure 23.97. Unfortunately, none of the metabolites show strong mossy odor (Hashimoto et  al., 2006). The presence of an acetonide in the metabolites has also been seen in those of dehydronootkatone (25) (Furusawa et al., 2003) (Figure 23.98). The liverworts Herbertus aduncus, Herbertus sakurai, and Mastigophora diclados produce (−)-herbertene, the C3 methyl isomer of cuparene, with its hydroxy derivatives, for example, herbertanediol (339), which shows no production inhibitory activity (Harinantenaina et al., 2007), and herbertenol (342). Treatment of compound (339) in P. sclerotiorum in Czapek-polypeptone medium gave two dimeric products, mastigophorene A (340) and mastigophorene B (341), which showed neurotrophic activity (Harinantenaina et al., 2005). When (−)-herbertenol (342) was biotransformed for 1 week by the same fungus, no metabolic product was obtained; however, five oxygenated metabolites (344–348) were obtained from its methyl ether (343). The possible metabolic pathway is shown in Figure 23.99. Except for the presence of the ether (348), the metabolites from 342 resemble those found in (−)- and (+)-cuparene (Hashimoto et al., 2006) (Figures 23.100 and 23.101).

825

Biotransformation of Sesquiterpenoids and Their Related Compounds OH

HO

OH

OH HOOC OH

328

329 HO

OH A. niger

HOOC

O

HOOC O

330

(–)-Cuparene (322)

331 OH

HO

332

OH

FIGURE 23.96  Biotransformation of (−)-cuparene (322) by Aspergillus niger.

O HOOC

HOOC

334

333

COOH

O A. niger

HOOC

HOOC

O

O 336

335

(+)-Cuparene (324)

HO

OH HOOC

HOOC

O 337

O 338

FIGURE 23.97  Biotransformation of (+)-cuparene (324) by Aspergillus niger.

Maalioxide (349), mp 65°–66°, [α]D21 − 34.4°, obtained from the liverwort P. sciophila was inoculated and cultivated rotatory (100 rpm) in Czapek-peptone medium (pH 7.0) at 30°C for 2 days. (−)-Maalioxide (349) (100 mg/200 mL) was added to the medium and further cultivated for 2 days to afford three metabolites, 1β-hydroxy- (350), 1β,9β-dihydroxy- (351), and 1β,12dihydroxymaalioxides (352), of which 351 was predominant (53.6%). When the same substrate was cultured with A. cellulosae in the same medium for 9 days, 7β-hydroxymaalioxide (353) was obtained as a sole product in 30% yield (Hashimoto et al., 2004). The same substrate (349) was also incubated with the fungus M. plumbeus to obtain a new metabolite, 9β-hydroxymaalioxide (354), together with two known hydroxylated products (350, 353) (Wang et al., 2006). Maalioxide (349) was oxidized by m-chloroperoxybenzoic acid to give a very small amount of 353 (1.2%), together with 2α-hydroxy- (355%, 2%) and 8α-hydroxymaalioxide (356, 1.5%), which have not been obtained in the metabolite of 349 in A. niger and A. cellulosae (Tori et al., 1990) (Figure 23.102).

826

Handbook of Essential Oils 9

15

[O]

HO HOOC

10

O 335

(+)-Cuparene (324)

[O] O 16 O

HOOC

–H2O OH

HOOC

336

HOOC 333

O 337

[O]

[O]

OH [O]

HOOC

[O]

HOOC

OH

OH

HOOC

O

FIGURE 23.98  Possible pathway of biotransformation of (+)-cuparene (324) by Aspergillus niger.

HO Penicillium sclerotiorum HO

OH

HO

OH

Mastigophorene A (340)

OH

(–)-Herbertenediol (339) HO

OH

HO

OH

Mastigophorene B (341)

FIGURE 23.99  Biotransformation of (−)-herbertenediol (339) by Penicillium sclerotiorum.

P. sciophila is one of the most important liverworts, since it produces bicyclohumulenone (357), which possesses strong mossy note and is expected to manufacture compounding perfume. In order to obtain much more strong scent, 357 was treated in A. niger for 4 days to give 4α,10βdihydroxybicyclohumulenone (358, 27.4%) and bicyclohumurenone-12-oic acid (359). An epoxide (360) prepared by m-chloroperoxybenzoic acid was further treated in the same fungus as described earlier to give 10β-hydoxy derivative (361, 23.4%). Unfortunately, these metabolites possess only faint mossy odor (Hashimoto et al., 2003c) (Figure 23.103). The liverwort Reboulia hemisphaerica biosynthesizes cyclomyltaylanoids like 362 and also ent-1α-hydroxy-β-chamigrene (367). Biotransformation of cyclomyltaylan-5-ol (362) in the same medium including A. niger gave four metabolites, 9β-hydroxy (363%, 27%), 9β,15-dihydroxy (364, 1.7%), 10β-hydroxy (365, 10.3%), and 9β,15-dihydroxy derivative (366, 12.6%). In this case, the stereospecificity of alcohol was observed, but the regiospecificity of alcohol moiety was not seen in this substrate (Furusawa et al., 2005c, 2006b) (Figure 23.104).

827

Biotransformation of Sesquiterpenoids and Their Related Compounds

A. niger

No metabolites

OH

HOOC

(–)–α-Herbertenol (342)

HO

OH O

(Me)2SO4

OH

OMe

OMe 345

344(159 mg) HOOC

HO

A. niger 1 week

O OMe

OMe (-)-Methoxy-α-herbertene (343)

OH OMe

OH

347

346

OH O 348

FIGURE 23.100  Biotransformation of (−)-methoxy-α-herbertene (343) by Aspergillus niger.

15

HO

9

[O]

10 OMe

HO [O] OH OMe

13

(–)-Methoxy-α-herbertene (343)

OM e

347

[O] HO

OH OMe 345

[O]

HOOC

HOOC

HOOC

–CH3

10

O

O OMe

OH HOOC

HOOC

OMe

O CH2OH

346 OH

OH O 348

13

[O]

[O]

[O]

O OMe

344

FIGURE 23.101  Possible pathway of biotransformation of (−)-methoxy-α-herbertene (343) by Aspergillus niger.

828

Handbook of Essential Oils HO

OH

H

O

355 (2.0%)

A

B CP

OH

H

O

H

O

353 (1.2%)

356 (1.5%)

m

OH

OH

OH

OH

A. niger O

H

H

O

M

O

s

u be

351 (53.6%)

350 (6.2%)

m lu .p

349

H

O

H

352 (11.0%)

OH

O

OH

OH

H

H

O

350

OH O

353

H

354

A. cellulosae

FIGURE 23.102  Biotransformation of maalioxide (349) by Aspergillus niger, Aspergillus cellulosae, and Mucor plumbeus. O

O H

2

4

O H

A. niger

10 12 1

H

OH

CO2H

HO

357

358 (27.4%)

359 (6.1%)

mCPBA O

O H

H

A. niger

3 O

360

2

10

OH

O

361 (23.4%)

FIGURE 23.103  Biotransformation of bicyclohumulenone (357) by Aspergillus niger.

The biotransformation of spirostructural terpenoids was not carried out. ent-1α-Hydroxy-βchamigrene (367) was inoculated in the same manner as described earlier to give three new metabolites (368–370), of which 370 was the major product (46.2% in isolated yield). The hydroxylation of vinyl methyl group has been known to be very common in the case of microbial and mammalian biotransformation (Furusawa et al., 2005a, 2006a,b) (Figure 23.105).

829

Biotransformation of Sesquiterpenoids and Their Related Compounds HO

HO OH

9 5 OH

15

9 10

363 (27.3%)

A. niger

362

O 364 (1.7%)

5 OH

HO OH

9 HO OH

OH

366 (12.6%)

365 (10.3%)

FIGURE 23.104  Biotransformation of cyclomyltaylan-5-ol (362) by Aspergillus niger. OH 9

1 8

15

367

A. niger

HO

9

OH

OH

OH

1

1

1

15

HO

HO 368 (4.8%)

8

8

OH

369 (25.0%)

370 (46.2%)

FIGURE 23.105  Biotransformation of ent-1α-hydroxy-β-chamigrene (367) by Aspergillus niger.

β-Barbatene (=gymnomitrene) (4), a ubiquitous sesquiterpene hydrocarbon, is from liverwort like P. sciophila and many others. Jungermanniales liverworts treated in the same manner using A. niger for 1 day gave a triol, 4β,9β,10β-trihydoxy-β-barbatene (27%, 8%) (Hashimoto et al., 2003c). Pinguisane sesquiterpenoids have been isolated from the Jungermanniales, Metzgeriales, and Marchantiales. In particular, the Lejeuneaceae and Porellaceae are rich sources of this unique type of sesquiterpenoids. One of the major furanosesquitepene (373) was biodegradated by A. niger to afford primary alcohol (375), which might be formed from 374 as shown in Figure 23.106 (Lahlou et al., 2000) (Figure 23.107). In order to obtain more pharmacologically active compounds, the secondary metabolites from crude drugs and animals, for example, nardosinone (376) isolated from the crude drug N. chinensis, which has been used for headache, stomachache, and diuresis, possesses antimalarial activity. Hinesol (384), possessing spasmolytic activity, obtained from A. lancea rhizoma, and animal perfume (−)-ambrox (391) from ambergris were biotransformed by A. niger, A. cellulosae, B. dothidea, and so on.

830

Handbook of Essential Oils H

H A. niger

OH

HO HO

371

372

FIGURE 23.106  Biotransformation of β-barbatene (371) by Aspergillus niger.

–H2O O

+H2O HO

O

O

OH H+ 373a

A. niger

O

O H

HO

OH 373

HO

OHC O

O 373c

HO 373b

O HO

HO O

O 374

375

FIGURE 23.107  Biotransformation of pinguisanol (373) by Aspergillus niger.

Nardosinone (376) was incubated in the same medium including A. niger as described earlier for 1 day to give six metabolites (377%, 45%; 378%, 3%; 379%, 2%; 380%, 5%; 381%, 6%; and 382%, 3%). Compounds 380–382 are unique trinorsesquiterpenoids although their yields are very poor. Compound 380 might be formed by the similar manner to that of phenol from cumene (383) (Figures 23.108 and 23.109) (Hashimoto et al., 2003d). From hinesol (384), two allylic alcohols (386, 387) and their oxygenated derivative (385) and three unique metabolites (388–390) having oxirane ring were obtained. The metabolic pathway is very similar to that of oral administration of hinesol since the same metabolites (395–387) were obtained from the urine of rabbits (Hashimoto et al., 1998a, 1999b, 2001a,b) (Figure 23.110). To obtain a large amount of ambrox (391), a deterrence, labda-12,14-dien-7α,8-diol obtained from the liverwort Porella pettottetiana as a major component, was chemically converted into (−)-ambrox via six steps in relatively high yield (Hashimoto et al., 1998b). Ambrox was added to Czapek-peptone medium including A. niger, for 4 days, followed by chromatography of the crude extract to afford four oxygenated products (392–395), among which the carboxylic acid (393, 52.4%) is the major product (Hashimoto et al., 2001a,b) (Figure 23.111). When ambrox (391) was biotransformed by A. niger for 9 days in the presence of 1-aminobenzotriazole, an inhibitor of CYP450, compounds 396 and 397 were obtained instead of the metabolites (392–395), which were obtained by incubation of ambrox in the absence of the inhibitor. Ambrox was cultivated by A. cellulosae for 4 days in the same medium to afford C1 oxygenated products (398 and 399), the former of which was the major product (41.3%) (Hashimoto et al., 2001a,b) (Figure 23.112).

831

Biotransformation of Sesquiterpenoids and Their Related Compounds O

O

7 11

67

OH 11

OH

378a (3%)

377 (45%)

O

H A. niger 7

6

O

O

10

1 day

OH

O 7

6

OH

OH

O 379 (2%)

Nardosinone (376)

HO

H 2

380 (5%)

O

H

HO

2

6 7

O

6 7

O

O

381 (6%)

382 (3%)

FIGURE 23.108  Biotransformation of nardosinone (376) by Aspergillus niger. OH

O O H O2

H3O+

(CH3)2CO

OH– Cumene (383) O

6 11

O 7

O

6

O O

Nardosinone (376)

11

O OH

OH 380

FIGURE 23.109  Possible pathway of biotransformation of nardosinone (376) to trinornardosinone (380) by Aspergillus niger.

The metabolite pathways of ambrox are quite different between A. niger and A. cellulosae. Oxidation at C1 occurred in A. cellulosae to afford 398 and 399, which was also afforded by John’s oxidation of 398, while oxidation at C3 and C18 and ether cleavage between C8 and C12 occurred in A. niger to give 392–395. Ether cleavage seen in A. niger is very rare. Fragrances of the metabolites (392–395) and 7α-hydroxy-(−)-ambrox (400) and 7-oxo-(−)ambrox (401) obtained from labdane diterpene diol were estimated. Only 399 demonstrated a similar odor to ambrox (391) (Hashimoto et al., 2001a,b) (Figure 23.113).

832

Handbook of Essential Oils

OH H Hinesol (384)

r

ige

A. n

A.

it bb

nig

er

Ra

O OH

O H

H HO

385

388

O HO

OH H

389

O

O OH

HO H

H

HO

387

390

FIGURE 23.110  Biotransformation of hinesol (384) by Aspergillus niger.

O

H (–)-Ambrox (391) A. niger at 30°C for 4 days

O

O 3 HO

HO

H OH

H 18COOH 393 (52.4%)

392 OH

OH

OH

OH

8

3 HO

H 394

O

OH

H

HO

386

OH

H 395

FIGURE 23.111  Biotransformation of (−)-ambrox (391) by Aspergillus niger.

OH

833

Biotransformation of Sesquiterpenoids and Their Related Compounds

O

H (–)-Ambrox Aspergillus niger A. niger + 1-Aminobenzotriazole at 30°C for 4 days

O HO

H 396

O HO

O OH

H

O

OH

OH

H 397

392 HO

H 394

OH OH

O HO

H COOH 393

O

H 395

FIGURE 23.112  Possible pathway of biotransformation of (−)-ambrox (391) by Aspergillus niger.

(−)-Ambrox (391) was also microbiotransformed with Fusarium lini to give mono-, di-, and trihydroxylated metabolites (401a–401d), while incubation of the same substrate with R. stolonifer afforded two metabolites (394, 396), which were obtained from 391 by A. niger as mentioned earlier, together with 397 and 401e (Choudhary et al., 2004) (Figure 23.114). The sclareolide (402), which is C12 oxo derivative of ambrox, was incubated with M. plumbeus to afford three metabolites, 3β-hydroxy- (403, 7.9%), 1β-hydroxy- (404, 2.5%), and 3-ketosclareolide (405, 7.9%) (Aranda et al., 1991) (Figure 23.115). A. niger in the same medium as mentioned earlier converted sclareolide (402) into two new metabolites (406, 407), together with known compounds (403, 405), of which 3β-hydroxy-sclareolide (403) is preferentially obtained (Hashimoto et al., 2007) (Figure 23.116). From the metabolites of sclareolide (402) incubated with C. lunata and A. niger, five oxidized compounds (403, 404, 405, 405a, 405b) were obtained. Fermentation of 402 with Gibberella fujikuroi afforded 403, 404, 405, and 405a. Metabolites 403 and 405a were formed from the same substrate by

834

Handbook of Essential Oils

O

H (–)-Ambrox (391) A. cellulosae 4 days

O

OH O

1

CrO3 - H2SO4

O

1

H

H

398 (41.3%)

399 (1.6%)

O

H 400

OH

O

H

O

401

FIGURE 23.113  Biotransformation of (−)-ambrox (391) by Aspergillus cellulosae.

the incubation of F. lini. No microbial transformation of 402 was observed with Pleurotus ostreatus (Atta-ur-Rahman et al., 1997) (Figure 23.117). Compound 391 treated in C. lunata gave metabolites 401e and 396, while C. elegans yielded compounds 401e and 396 and (+)-sclareolide (402) (Figure 23.113). The metabolites (401a–401e, 396) from 391 do not release any effective aroma when compared to 391. Compound 394 showed a strong sweet odor quite different from the amber-like odor (Choudhary et al., 2004). Sclareolide (402) exhibited phytotoxic and cytotoxic activity against several human cancer cell lines. C. elegans gave new oxidized metabolites (403, 404, 405a, 405c, 405d, 405e), resulting from the enantioselective hydroxylation. Metabolites 403, 404, and 405a have been known as earlier as biotransformed products of 402 by many different fungi and have shown cytotoxicity against various human cancer cell lines. The metabolites (403, 404, and 405a) indicated significant phytotoxicity at higher dose against Lemna minor L. (Choudhary et al., 2004) (Figure 23.117). Ambrox (391) and sclareolide (402) were incubated with the fungus C. aphidicola for 10 days in shake culture to give 3β-hydroxy- (396), 3β,6β-dihydroxy- (401 g), 3β,12-dihydroxy- (401 h), and sclareolide 3β,6β-diol (401f), and 3β-hydroxy- (403), 3-keto- (405), and sclareolide 3β,6β-diol (401f), respectively (Hanson and Truneh, 1996) (Figure 23.118). Zerumbone (408), which is easily isolated from the wild ginger Zingiber zerumbet, and its epoxide (409) were incubated with F. culmorum and A. niger in Czapek-peptone medium, respectively. The former fungus gave (1R,2R)-(+)-2,3-dihydrozerumbol (410) stereospecifically via either 2,3-dihydrozerumbone (408a) or zerumbol (408b) or both and accumulated 410 in the mycelium. The facile production of optically active 410 will lead a useful material of woody fragrance, 2,3-dihydrozerumbone. A. niger biotransformed 408 via epoxide (409) to several metabolites containing zerumbone-6,7-diol as a main product. The same fungus converted the epoxide (409) into three major metabolites containing (2R,6S,7S,10R,11S)-1-oxo7,9-dihydroxyisodaucane (413) via

835

Biotransformation of Sesquiterpenoids and Their Related Compounds HO OH

OH

O

O

H

H O

401b

401a

a

H

O

HO OH

OH

O

O

391

HO

H

H

OH

401c

OH

c,d

O HO

O

OH

401d

OH HO

H

H 394

396

b

O O

O

H 391

O

HO

H

H

397

OH

401e

c,d

a: Fusarium lini b: Rhizopus strolonifera c: Curvlaria lanata d: Cunninghamella elegans

FIGURE 23.114  Biotransformation of (−)-ambrox (391) by Fusarium lini and Rhizopus stolonifer. O O H

3

O

O

OH

H

O

O

O

M. plumbeus

O

H

H O

HO 18 Sclareolide (402)

403

404

405

FIGURE 23.115  Biotransformation of (+)-sclareolide (402) by Mucor plumbeus.

dihydro derivatives (411, 412). However, A. niger biotransformed 409 only into 412 in the presence of the CYP450 inhibitor 1-aminobenzotriazole (Noma et al., 2002). The same substrate was incubated in A. niger, Aspergillus oryzae, Candida rugosa, Candida tropicalis, Mucor mucedo, Bacillus subtilis, and Schizosaccharomyces pombe; however, any metabolites have been obtained. All microbes except for the last organism, zerumbone epoxide (409), prepared by mCPBA, bioconverted into two diastereoisomers, 2R,6S,7S-dihydro- (411) and 2R,6R,7R-derivative (412), whose ratio was determined by GC, and their enantio-excess was over 99% (Nishida and Kawai, 2007) (Figure 23.119).

836

Handbook of Essential Oils O

O O 3

O A. niger HO

O

3

18 402

O

[O] O

3 404

403

[O]

[O] O O HO

O O

[O]

3 18 OH 406

O

3 18 OH 407

FIGURE 23.116  Biotransformation of (+)-sclareolide (402) by Aspergillus niger.

Several microorganisms and a few mammals (see later) for the biotransformation of (+)-cedrol (414), which is widely distributed in the cedar essential oils, were used. Plant pathogenic fungus G. cingulata converted cedrol (414) into three diols (415–417) and 2α-hydroxycedrene (418) (Miyazawa et al., 1995a). The same substrate (414) was incubated with A. niger to give 416 and 417 together with a cyclopentanone derivative (419) (Higuchi et al., 2001). Human skin microbial flora, Staphylococcus epidermidis also converted (+)-cedrol into 2α-hydroxycedrol (415) (Itsuzaki et al., 2002) (Figure 23.120). C. aphidicola bioconverted cedrol (414) into 417 (Hanson and Nasir, 1993). On the other hand, Corynespora cassiicola produced 419 in addition to 417 (Abraham et al., 1987). It is noteworthy that B. cinerea that damages many flowers, fruits, and vegetables biotransformed cedrol into different metabolites (420–422) from those mentioned earlier (Aleu et al., 1999a). 4α-Hydroxycedrol (424) was obtained from the metabolite of cedrol acetate (423) by using G. cingulata (Matsui et al., 1999) (Figure 23.121). Patchouli alcohol (425) was treated in B. cinerea to give three metabolites two tertiary alcohols (426, 427), four secondary alcohols (428, 430, 430a), and two primary alcohols (430b, 430c) of which compounds 425, 427, and 428 are the major metabolites (Aleu et al., 1999a), while plant pathogenic fungus G. cingulata converted the same substrate to 5-hydroxy- (426) and 5,8-dihydroxy derivative (429) (Figure 23.122). In order to confirm the formation of 429 from 426, the latter product was reincubated in the same medium including G. cingulata to afford 429 (Miyazawa et al., 1997c) (Figure 23.123). Patchouli acetate (431) was also treated in the same medium to give 426 and 429 (Matsui and Miyazawa, 2000). 5-Hydroxy-α-patchoulene (432) was incubated with G. cingulata to afford 1α-hydroxy derivative (426) (Miyazawa et al., 1998b). (−)-α-Longipinene (433) was treated with A. niger to afford 12-hydroxylated product (434) (Sakata et al., 2007). Ginsenol (435), which was obtained from the essential oil of Panax ginseng, was incubated with B. cinerea to afford four secondary alcohols (436–439) and two cyclohexanone derivatives (440) from 437 and 441 from 438 or 439. Some of the oxygenated products were considered as potential antifungal agents to control B. cinerea (Aleu et al., 1999b) (Figures 23.124 and 23.125).

837

Biotransformation of Sesquiterpenoids and Their Related Compounds O O HO

H 405f

g

OH

g

g O

O

O

O OH

O

O

O

O

a,b d,e,f HO

H

O

H

402

403

H

H

404

405

O OH O

a,d,e,f HO

O OH O

HO

H 405a

H 405b O

O OH

O

O

O OH O

c a: Curvularia lunata b: Mucro plumbeus c: Cunnighamella elegans d: Aspergillus niger e: Gibberella fujikuroii f: Fusarium lini (only 403, 405a) g: Cephalosporium HO aphidicola

O

H

H

404

405 O O

H 405c

HO HO

H

HO

H 405a

O

O

O

O

HO

405d

H 405e

FIGURE 23.117  Biotransformation of (+)-sclareolide (402) by various fungi.

(+)-Isolongifolene-9-one (442), which was isolated from some cedar trees, was treated in G. cingulata for 15 days to afford two primary alcohols (443, 444) and a secondary alcohol (445) (Sakata and Miyazawa, 2006) (Figure 23.126). Choudhary et al. (2005) reported that fermentation of (−)-isolongifolol (445a) with F. lini resulted in the isolation of three metabolites, 10-oxo- (445b), 10α-hydroxy- (445c), and 9α-hydroxyisolongifolol (445d). Then the same substrate was incubated with A. niger to yield the products 445c and 445d. Both 445c and 445d showed inhibitory activity against butylcholinesterase enzyme in a concentrationdependent manner with IC50 13.6 and 299.5 mM, respectively (Figure 23.127).

838

Handbook of Essential Oils

O

O HO O

HO

H

H

a

OH

401g

396

O

OH

H

O

O

391 HO

HO

H

H

a: Cephalosporium aphidocola

OH

401f

401h

FIGURE 23.118  Biotransformation of (−)-ambrox (391) by Cephalosporium aphidicola. O

O 1

2

a

OH

408a

410 O

OH

10 3 9 7 6

O

a

HO H

b

O

408 O

408b

mPCBA

O

409

411

HO 413

b + 1-aminobenzotriazole

O

O

O

b-g

O

O

O

409

411

412

a: Fusarium culmorum b: Aspergillus niger c: Aspergillus orizae d: Candida rugosa e: Candida tropicalis f: Mucor mucedo g: Bacillus subtilis

FIGURE 23.119  Biotransformation of zerumbone (408) by various fungi.

(+)-Cycloisolongifol-5β-ol (445e) was fermented with C. elegans to afford three oxygenated metabolites: 11-oxo (445f), 3β-hydroxy (445 g), and 3β,11α-dihydroxy derivative (445 h) (Choudhary et al., 2006a) (Figure 23.128). A daucane-type sesquiterpene derivative, lancerroldiol p-hydroxybenzoate (446), was hydrogenated with cultured suspension cells of the liverwort M. polymorpha to give 3,4-dihydrolancerodiol (447) (Hegazy et al., 2005) (Figure 23.129). Widdrane sesquiterpene alcohol (448) was incubated with A. niger to give an oxo and an oxy derivative (449, 450) (Hayashi et al., 1999) (Figure 23.130). (−)-β-Caryophyllene (451), one of the ubiquitous sesquiterpene hydrocarbons found not only in higher plants but also in liverworts, was biotransformed by Pseudomonas cruciviae, D. gossypina, and Chaetomium cochliodes (Lamare and Furstoss, 1990). P. cruciviae gave a ketoalcohol (452)

839

Biotransformation of Sesquiterpenoids and Their Related Compounds

a c

OH

HO

a

HO H 418

H 415 HO a

OH

OH

b,f H 414

a: Glomerella cingulata b: Aspergillus niger c: Staphylococcus epidermidis d: Cephalosporium aphidicola e: Corynespora cassiicola f: Botrytis cinerea

H 416

a,b d,e

OH

HO H 417

e

OH

O H 419

FIGURE 23.120  Biotransformation of cedrol (414) by various fungi.

HO

B. cinerea

OH H 414

OH

OH

HO H 420

HO

H 421

HO OH H 422

OAc G. cingulata H 423

OAc HO

H 424

FIGURE 23.121  Biotransformation of cedrol (414) by Botrytis cinerea and Glomerella cingulata.

840

Handbook of Essential Oils

HO

HO

HO OH

OH 426 HO

H 427

HO OH H 428

H 425

OH

H 429

OH

HO

a,b

OH

HO

HO H 430

H 430a

HO HO

HO a: B. cinerea b: G. cingulata

OH H

H 430c

430b

FIGURE 23.122  Biotransformation of patchoulol (425) by Botrytis cinerea.

HO

RO

G. cingulata

HO

G. cingulata

OH

H

OH

OH

425: R=H 431: R=Ac

426

429

OH 432

FIGURE 23.123  Biotransformation of patchoulol (425) by Glomerella cingulata.

OH A. niger

433

434

FIGURE 23.124  Biotransformation of α-longipinene (433) by Aspergillus niger.

841

Biotransformation of Sesquiterpenoids and Their Related Compounds

10

OH 9

HO

8

10

HO OH 8

OH OH 9

OH

B. cinerea

OH

6 6 OH 436

435

437

438

439

OH O

O

OH 8

9 6 OH

441

440

FIGURE 23.125  Biotransformation of ginsenol (435) by Botrytis cinerea.

OH O

O

G. cingulata

OH O

O

OH

442

443

444

445

FIGURE 23.126  Biotransformation of (+)-isolongifolene-9-one (442) by Glomerella cingulata. O

OH

OH

Fusarium lini

445a

OH

445b

OH

445c

OH

OH

445d

Aspergillus niger

FIGURE 23.127  Biotransformation of (−)-isolongifolol (445a) by Aspergillus niger and Fusarium lini. OH

OH C. elegans OH

OH

OH O

445e

OH

HO 445f

445g

445h

FIGURE 23.128  Biotransformation of (+)-cycloisolongifol-5β-ol (445e) by Cunninghamella elegans.

842

Handbook of Essential Oils O

O Marchantia polymorpha cell HO

HO

OH

O

OH

O O

O

447

446

FIGURE 23.129  Biotransformation of lancerodiol p-hydroxybenzoate (446) by Marchantia polymorpha cells.

A. niger

OH

OH

OH O

HO 449

448

450

FIGURE 23.130  Biotransformation of widdrol (448) by Aspergillus niger.

O H

14

H 8 7 10 15 9 1 5 11 2 3 H

6

13 1) Pseudomonus ceuciviae 2) Diplodia gossypina

4

3) Chaetominum cochlioides

HO

12 451

452 OH H

H

O

O H

H

454

453

D. gossypina C. cochlioides

OH

HOOC

H

OH H

H O

O H

H

O H OH

457

455 (5%)

456 (12%)

FIGURE 23.131  Biotransformation of (−)-β-caryophyllene (451) by Pseudomonas cruciviae, Diplodia ­gossypina, and Chaetomium cochliodes.

843

Biotransformation of Sesquiterpenoids and Their Related Compounds

(Devi, 1979), while the latter two species produced the 14-hydroxy-5,6-epoxide (454), its carboxylic (455), and 3α-hydroxy- (456) and norcaryophyllene alcohol (457), all of which might be formed from caryophyllene C5,C6-epoxide (453). The oxidation pattern of (−)-β-caryophyllene by the fungi is very similar to that by mammals (see later) (Figure 23.131). Fermentation of (−)-β-caryophyllene (451) with D. gossypina afforded 14 different metabolites (453–457j), among which 14-hydroxy-5,6-epoxide (454) and the corresponding acid (455) were the major metabolites. Compound 457j is structurally very rare and found in Poronia punctata. The main reaction path is epoxidation at C5, C6 as mentioned earlier and selective hydroxylation at C4 (Abraham et al., 1990) (Figure 23.132). (−)-β-Caryophyllene epoxide (453) was incubated with C. aphidicola for 6 days to afford two metabolites (457 l, 457 m), while Macrophomina phaseolina biotransformed the same substrate to 14- (454) and 15-hydroxy derivatives (457k). The same substrate was treated in A. niger, G. fujikuroi, and R. stolonifer for 8 days and F. lini for 10 days to afford the metabolites 457n, 457o, 457p and HO H

H

HOOC

O

O

H

H 453

H 455

OH H

OH H

O

O

H

15

OH

H 11

451

Diplodia gossypina

5 2

H

457a

OH H

6

O

OH

H

457 H

O

454

OH H

14

H

OH 457b

OH H

OH H

O

O

H

HO

H 457c

O

OH H

H

OH

457d

OH H

OH

O

O

H

H OH

457f OH H

457g

OH H OH O

H 457i

457e

OH H

O H O

O

OH

OHC 457j

FIGURE 23.132  Biotransformation of (−)-β-caryophyllene (451) by Diplodia gossypina.

OH

457h

844

Handbook of Essential Oils

457q, and 457r, respectively. All metabolites were estimated for butyrylcholine esterase inhibitory activity, and compound 457k was found to show potency similar activity to galantamine HBr (IC50 10.9 vs. 8.5 mM) (Choudhary et al., 2006b) (Figure 23.133). The fermentation of (−)-β-caryophyllene oxide (453) using B. cinerea and the isolation of the metabolites were carried out by Duran et al. (1999). Kobuson (457w) was obtained with 14 products (457s–457u, 457x). Diepoxides 457t and 457u could be the precursors of epimeric alcohols 457q OH H

a

H O

14

H

H

H 6

15

5 H

O

HO

454

O

457k

2 H

H

11 453

OH a: M. phaseolina b: C. aphidicola

b

H

OH

HO

457l

457m

OH

H

a

OH HO

OH 457n H

b

OH 14

H

OH

15

HO

6 5 H

457o

O

2

11

453

HO H

H

c

O

O a: A. niger b: G. fujikuroii c: R. stolonifera d: F. lini

H O

H 457p

HO 457q

H

OH

d O H HO

457r

FIGURE 23.133  Biotransformation of (−)-β-caryophyllene epoxide (453) by various fungi.

845

Biotransformation of Sesquiterpenoids and Their Related Compounds

and 457y obtained through reductive opening of the C2,C11-epoxide. The major reaction paths are stereoselective epoxidation and introduction of hydroxyl group at C3. Compound 457ae has a caryolane skeleton (Figure 23.134). When isoprobotryan-9α-ol (458) produced from isocaryophyllene was incubated with B. cinerea, it was hydroxylated at tertiary methyl groups to give three primary alcohols (459–461) (Aleu et al., 2002) (Figure 23.135). Acyclic sesquiterpenoids, racemic cis-nerolidol (462), and nerylacetone (463) were treated by the plant pathogenic fungus G. cingulata (Miyazawa et al., 1995d). From the former substrate, a triol (464) was obtained as the major product. The latter was bioconverted to give the two methyl 14 15

6

5

H

OH H

H

H O

B. cinerea

OH

2 3

H

O H

457s

11 453

454

B. cinerea H

H

H

O

O

HO

H O

H 457k H

457t

O

O H O

H O

O 457w

457v

457x

H

H

H O

O H

H HO

H HO CHO 457z

H

H

O

457q

457y

H O

O

O H HO

H OH 457aa

H HO 457ab

H

457ac

H O

H HO

457u

H

O

HO

H O

H

H O

O

OH

O

H 457ad

HO

457ae

FIGURE 23.134  Biotransformation of (−)-β-caryophyllene epoxide (453) by Botrytis cinerea.

846

Handbook of Essential Oils H

HO

458 B. cinerea

H

HO

HO

HO

H

HO

H OH

OH 459

460

461

FIGURE 23.135  Biotransformation of isoprobotryan-9α-ol (458) by Botrytis cinerea.

ketones (465, 467) and a triol (468), among which 465 was the predominant. The C10,C11 diols (464, 465) might be formed from both epoxides of the substrates, followed by the hydration although no C10,C11-epoxides were detected (Figure 23.136). Racemic trans-nerolidol (469) was also treated in the same fungus to afford w2-hydroxylated product (471) and C10,C11-hydroxylated compounds (472) as seen in racemic cis-nerolidol (462) (Miyazawa et al., 1996b) (Figure 23.137). 12-Hydroxy-trans-nerolidol (472a) is an important precursor in the synthesis of interesting ­flavor of α-sinensal. Hrdlicka et al. (2004) reported the biotransformation of trans- (469) and c­ is-nerolidol (462) and cis-/trans-mixture of nerolidol using repeated batch culture of A. niger grown in computercontrolled bioreactors. Trans-nerolidol (469) gave 472a and 472 and cis-isomer (462) afforded 464a and 464. From a mixture of cis- and trans-nerolidol, 12-hydroxy-trans-neroridol 472a (8%) was 10

12 11 13

9 462

OH

5

8 7 6

4

3

2

OH

G. cingulata 1

HO

OH

OH

464

O

HO 465 OH

O G. cingulata

OH

HO 466

463

OH

468 O

HO 467

FIGURE 23.136  Biotransformation of cis-nerolidol (462) and cis-geranyl acetone (463) by Glomerella cingulata.

847

Biotransformation of Sesquiterpenoids and Their Related Compounds HO

OH 471

OH

G. cingulata

469

OH HO

OH

472

OH HO 473

OH

O

HO

G. cingulata 470

476

O

474

OH HO

OH

477 HO 475

OH

O

FIGURE 23.137  Biotransformation of trans-nerolidol (469) and trans-geranyl acetone (470) by Glomerella cingulata.

obtained in postexponential phase at high dissolved oxygen. At low dissolved oxygen condition, the mixture gave 472a (7%) and 464a (6%) (Figures 23.138 and 23.139). From geranyl acetone (470) incubated with G. cingulata, four products (473–477) were formed. It is noteworthy that the major compounds from both substrates (469, 470) were w2-hydroxylated OH

OH

OH A. niger

464a OH

462

OH

HO 464 OH

A. niger OH

OH

472a

OH

469 HO

OH 472

FIGURE 23.138  Biotransformation of cis- (462) and trans-nerolidol (469) by Aspergillus niger.

848

Handbook of Essential Oils a

OH

HO

OH

HO

479

481

OH OH

OH OH

a,b

OH

HO

HO

480

2E,6E-farnesol (478)

a: Cytochrome P-450 b: Aspergillus niger

482

OH

OH

b OH

HO

480a

FIGURE 23.139  Biotransformation of 2E,6E-farnesol (478) by cytochrome P-450 and Aspergillus niger.

products, but not C10,C11-dihydroxylated products as seen in cis-nerolidol (462) and nerylacetone (463) (Miyazawa et al., 1995b) (Figure 23.136). The same fungus bioconverted (2E,6E)-farnesol (478) to four products, w2-hydroxylated product (479), which was further oxidized to give C10,C11 dihydroxylated compound (480) and 5-hydroxy derivative (481), followed by isomerization at C2,C3 double bond to afford a triol (482) (Miyazawa et al., 1996c) (Figure 23.140). The same substrate was bioconverted by A. niger to afford two metabolites, 10,11-dihydroxy (480) and 5,13-hydroxy derivative (480a) (Madyastha and Gururaja, 1993). The same fungus also converted (2Z,6Z)-farnesol (483) to three hydroxylated products: 10,11-dihydroxy-(2Z,6Z)- (484), 10,11-dihydroxy-(2E,6Z)-farnesol (485), and (5Z)-9,10-dihydroxy6,10-dimethyl-5-undecen-2-one (486) (Nankai et al., 1996) (Figures 23.140 and 23.141). A linear sesquiterpene 9-oxonerolidol (487) was treated in A. niger to give w1-hydroxylated product (488) (Higuchi et al., 2001) (Figure 23.142). Racemic diisophorone (488a) dissolved in ethanol was incubated with the Czapek–Dox medium of A. niger to afford 8α- (488b), 10β- (488c), and 17-hydroxydiisophorone (488d) (Kiran et al., 2004). On the other hand, the same substrate was fed with Neurospora crassa and C. aphidicola to afford only 8β-hydroxydiisophorone (488e) in 20% and 10% yield, respectively (Kiran et al., 2005) (Figure 23.143). From the metabolites of 5β,6β-dihydroxypresilphiperfolane 2β-angelate (488f) using the fungus Mucor ramannianus, 2,3-epoxyangeloyloxy derivative (488 g) was obtained (Orabi, 2001) (Figure 23.144). OH OH HO 12

11

10 5

13

480

OH 478

1

A. niger OH HO

OH 480a

FIGURE 23.140  Biotransformation of 2E,6E-farnesol (478) by Aspergillus niger.

849

Biotransformation of Sesquiterpenoids and Their Related Compounds OH

OH

HO 484 OH A. niger OH

OH

HO 485

2E,6Z-farnesol (483) OH

O

HO 486

FIGURE 23.141  Biotransformation of 2Z,6Z-farnesol (478) by Aspergillus niger. OH

O

O

A. niger

OH

HO

487

488

FIGURE 23.142  Biotransformation of 9-oxo-trans-nerolidol (487) by Aspergillus niger. OH OH

a

O

O

OH 488a

OH

O

488b

OH 488c

b,c OH a: Aspergillus niger b: Cephalsporium aphidicola c: Neurospora crassa O

O

OH

OH 488d

OH 488e

FIGURE 23.143  Biotransformation of diisophorone (488a) by Aspergillus niger, Cephalosporium aphidicola, and Neurospora crassa. O

O

OH O

H 488f

OH

M. ramannianus

OH

O O 488g

OH

FIGURE 23.144  Biotransformation of 5β,6β-dihydroxypresilphiperfolane 2β-angelate (488f) by Mucor ramannianus.

850

Handbook of Essential Oils

23.3 BIOTRANSFORMATION OF SESQUITERPENOIDS BY MAMMALS, INSECTS, AND CYTOCHROME P-450 23.3.1 Animals (Rabbits) and Dosing Six male albino rabbits (2–3 kg) were starved for 2 days before experiment. Monoterpenes were suspended in water (100 mL) containing polysorbate 80 (0.1 g) and were homogenized well. This solution (20 mL) was administered to each rabbit through a stomach tube followed by water (20 mL). This dose of sesquiterpenoids corresponds to 400–700 mg/kg. Rabbits were housed in stainless steel metabolism cages and were allowed rabbit food and water ad libitum. The urine was collected daily for 3 days after drug administration and stored at 0°–5°C until the time of analysis. The urine was centrifuged to remove feces and hairs at 0°C and the supernatant was used for the experiments. The urine was adjusted to pH 4.6 with acetate buffer and incubated with β-glucuronidase–arylsulfatase (3/100 mL of fresh urine) at 37°C for 48 h, followed by continuous ether extraction for 48 h. The ether extracts were washed with 5% NaHCO3% and 5% NaOH to remove the acidic and phenolic components, respectively. The ether extract was dried over MgSO 4, followed by evaporation of the solvent to give the neutral crude metabolites (Ishida et al., 1981).

23.3.2 Sesquiterpenoids Wild rabbits (hair) and deer damage the young leaves of Chamaecyparis obtusa, one of the most important furniture and house-constructing tree in Japan. The essential oil of the leaves contains a large amount of (−)-longifolene (489). Longifolene (36 g) was administered to 18 of rabbits to obtain the metabolites (3.7 g) from which an aldehyde (490) (35.5%) was isolated as pure state. In the metabolism of terpenoids having an exomethylene group, glycol formation was often found, but in the case of longifolene such as a diol was not formed. Introduction of an aldehydes group in biotransformation is very remarkable. Stereoselective hydroxylation of the gem-dimethyl group on a seven-membered ring is first time (Ishida et al., 1982) (Figure 23.145). (−)-β-Caryophyllene (451) is one of the ubiquitous sesquiterpene hydrocarbons in the plant kingdom and the main component of beer hops and clove oil and is being used as a culinary ingredient and as a cosmetic in soaps and fragrances. (−)-β-Caryophyllene also has cytotoxic against breast carcinoma cells and its epoxide is toxic to Planaria worms. It contains unique 1,1-dimethylcyclobutane skeleton. (−)-β-Caryophyllene (3 g) was treated in the same manner as described earlier to afford the crude metabolite (2.27 g) from which (10S)-14-hydroxycaryophyllene-5,6-oxide (491) (80%) and a diol (492) were obtained (Asakawa et al., 1981). Later, compound 491 was isolated from the Polish mushroom Lactarius camphoratus (Basidiomycetes) as a natural product (Daniewski et al., 1981). 14-Hyroxy-βcallyophyllene and 1-hydroxy-8-keto-β-caryophyllene have been found in Asteraceae and Pseudomonas species, respectively. In order to confirm that caryophyllene epoxide (453) is the intermediate of both metabolites, it was treated in the same manner as described earlier to give the same metabolites, 491 and 492, of which 491 was predominant (Asakawa et al., 1981, 1986) (Figure 23.146). HO Rabbit

H H

CHO

O CHO

489

FIGURE 23.145  Biotransformation of longifolene (489) by rabbit.

490

851

Biotransformation of Sesquiterpenoids and Their Related Compounds H

H O

H

O H

O

OH

OH 492 H O 14 15

H 8

7

13

6 11 9 O 1 5 10 2 4 H 3

OH H

H Rabbit

12 451

5

453

14

6 O

H

OH H

491

H

FIGURE 23.146  Biotransformation of (−)-β-caryophyllene (451) by rabbit.

The grapefruit aroma, (+)-nootkatone (2), was administered into rabbits to give 11,12-diol (6, 7). The same metabolism has been found in that of biotransformation of nootkatone by microorganisms as mentioned in the previous paragraph. Compounds (6, 7) were isolated from the urine of hypertensive subjects and named urodiolenone. The endogenous production of 6 and 7 seem to occur interdentally from the administrative manner of nootkatone or grapefruit. Synthetic racemic nootkatone epoxide (14) was incubated with rabbit liver microsomes to give 11,12-diol (6, 7) (Ishida, 2005). Thus, the role of the epoxide was clearly confirmed as an intermediate of nootkatone (2). (+)-ent-Cyclocolorenone (98) and its enantiomer (103) were biotransformed by Aspergillus species to give cyclopropane-cleaved metabolites as described in the previous paragraph. In order to compare the metabolites between mammals and microorganisms, the essential oil (2 g/rabbit) containing (−)-cyclocolorenone (103) obtained from S. altissima was administered in rabbits to obtain two metabolites: 9β-hydroxycyclocolorenone (493) and 10-hydroxycyclocolorenone (494) (Asakawa et al., 1986). 10-Hydroxyaromadendrane-type compounds are well known as the natural products. No oxygenated compound of cyclopropane ring was found in the metabolites of cyclocolorenone in rabbit (Figure 23.147). From the metabolites of elemol (495) possessing the same partial structures of monoterpene hydrocarbon, myrcene, and nootkatone, one primary alcohol (496) was obtained from rabbit urine after the administration of 495 (Asakawa et al., 1986) (Figure 23.148). H

H Rabbit

O

103

O

OH

H O

493

FIGURE 23.147  Biotransformation of (+)-ent-cyclocolorenone (101) by rabbit.

494

OH

852

Handbook of Essential Oils

Rabbit

OH

H

OH

H HO

495

496

FIGURE 23.148  Biotransformation of elemol (495) by rabbit.

Components of cedar wood such as cedrol (414) and cedrene shorten the sleeping time of mice. In order to search for a relationship between scent, olfaction, and detoxifying enzyme induction, (+)-cedrol (414) was administered to rabbits and dogs. From the metabolites from rabbits, two C3-hydroxylated products (418 and 497) and a diol (415 or 416), which might be formed after the hydrogenation of double bond. Dogs converted cedrol (414) into different metabolite products, C2- (498) and C2/C14-hydroxylated products (499), together with the same C3- (415) and C15hydroxylated products (416) as those found in the metabolites of microorganisms and rabbits. The aforementioned species-specific metabolism is very remarkable (Bang and Ourisson, 1975). The microorganisms C. aphidicola, C. cassiicola, B. cinerea, and G. cingulata also biotransformed cedrol to various C2-, C3-, C4-, C6-, and C15-hydroxylated products as shown in the previous paragraph. The microbial metabolism of cedrol resembles that of mammals (Figure 23.149). Patchouli alcohol (425) with fungi static properties is one of the important essential oils in perfumery industry. Rabbits and dogs gave two oxidative products (500, 501) and one norpatchoulen1-one (502) possessing a characteristic odor. Plant pathogen B. cinerea causes many diseases for vegetables and flowers. This pathogen gave totally different five metabolites (426–430) from those found in the urine metabolites of mammals as described earlier (Bang et al., 1975) (Figure 23.150).

OH Rabbit

3 2

HO

OH

3

HO

HO

H

H 414

415 or 417

Dog HO

H

418

497

15

COOH OH

OH

HO

H

OH

OH

2 H

H 415

HO

416

H 498

HO

H 499

FIGURE 23.149  Biotransformation of cedrol (414) by rabbits or dogs.

HO

HO

HO

HO

Rabbit or Dog H

425

15

H OH 500

HOOC

H

501

FIGURE 23.150  Biotransformation of patchouli alcohol (425) by rabbits or dogs.

H

502

853

Biotransformation of Sesquiterpenoids and Their Related Compounds

Sandalwood oil contains mainly α-santalol (503) and β-santalol. Rabbits converted α-santalol to three diastereomeric primary alcohols (504–506) and dogs did carboxylic acid (507) (Zundel, 1976) (Figure 23.151). (2E,6E)-Farnesol (478) was treated in cockroach cytochrome P-450 (CYP4C7) to form regio- and diastereospecifically w-hydroxylated at the C12 methyl group to the corresponding diol (508) with 10E-configuration (Sutherland et al., 1998) (Figure 23.152). Juvenile hormone III (509) was also treated in cockroach CYP4C7 to the corresponding 12-hydroxylated product (510) (Sutherland et al., 1998).

OH 15

OH

504 10 OH Rabbit

14

14

OH

OH

15 503

505 13

Dog 10 COOH

OH OH

506 507

FIGURE 23.151  Biotransformation of santalol (503) by rabbits or dogs. 13 1 12

11

10

7 478

13 11

10

CYP450 (CYP4C7)

7 509

2

HO

OH

2

12

508 O

O

HO

O

O 12

OH

2

OMe

12 OMe

510

CYP450 (CYP4C7)

R1 12

12' O

R2

O 7

OMe

511: R1=H, R2=OH 512: R1=OH, R2=H

FIGURE 23.152  Biotransformation of 2E,6E-farnesol (478) by cockroach cytochrome P-450 and 10,11-epoxyfarnesic acid methyl ester (509) by African locust cytochrome P-450.

854

Handbook of Essential Oils

The African locust converted the same substrate (509) into a 7-hydroxy product (511) and a 13-hydroxylated product (512). It is noteworthy that the African locust and cockroach showed clear species specificity for introduction of oxygen function (Darrouzet et al., 1997).

23.4 BIOTRANSFORMATION OF IONONES, DAMASCONES, AND ADAMANTANES Racemic α-ionone (513) was converted to 4-hydroxy-α-ionone (514), which was further dehydrogenated to 4-oxo-α-ionone (515) by Chlorella ellipsoidea IAMC-27 and C. vulgaris IAMC-209. α-Ionone (513) was reduced preferentially to α-ionol (516) by Chlorella sorokiniana and Chlorella salina (Noma et al., 1997b). α-Ionol (516) was oxidized by C. pyrenoidosa to afford 4-hydroxy-α-ionol (524). The same substrate was fed by the same microorganism and A. niger to furnish α-ionone (513) (Noma and Asakawa, 1998) (Figure 23.153). 4-Oxo-α-ionone (515), which is one of the major product of α-ionone (513) by A. niger, was transformed reductively by Hansenula anomala, Rhodotorula minuta, Dunaliella tertiolecta, Euglena gracilis, C. pyrenoidosa C28 and other eight kinds of Chlorella species, B. dothidea, A. cellulosae IFO 4040, and A. sojae IFO 4389 to give 4-oxo-α-ionol (517), 4-oxo-7,8-dihydro-αionone (518), and 4-oxo-7,8-dihydro-α-ionol (519). Compound 515 was also oxidized by A. niger and A. sojae to give 1-hydroxy-4-oxo-α-ionone (520) and 7,11-oxido-4-oxo-7,8-dihydro-α-ionone (521). C7–C8 double bond of α-ionone (513), 4-oxo-α-ionone (515), and 4-oxo-α-ionol (517) were easily reduced to their corresponding dihydro products (522, 518, 519), respectively, by Euglena,

O

12 E. gracilis

522

5 4

A. sojae

13 6 3

E. gracilis OH

HO

523

O

1 2

O

7 8

9

11 513 A. niger C. ellipsoidea C. pyrenoidosa C. vulgaris O

514 A. niger C. ellipsoidea C. pyrenoidosa O

A. niger C. pyrenoidosa

OH O

O

520

515

O

O O 521

OH

A. niger C. pyrenoidosa

516 C. pyrenoidosa OH

HO B. dothidea C. pyrenoidosa D. tertiolecta E. gracilis H. anomala Rh. minuta A. cellulosae A. niger A. sojae

O

OH

O

FIGURE 23.153  Biotransformation of α-ionone (513) by various microorganisms.

517 C. pyrenoidosa OH

B. dothidea D. tertiolecta E. gracilis

518

524 C. pyrenoidosa

A. cellulosae, A sojae B. dothidea, E. gracilis D. tertiolecta, Rh. minuta

er nig e A. . soja A

O

10

A. cellulosae C. pyrenoidosa C. salina C. sorokiniana D. tertiolocta H. anomala

O

519

855

Biotransformation of Sesquiterpenoids and Their Related Compounds O

O

1R

O

O

A. niger HO

HO

513a

O

514a (46.2%)

514b (23.1%)

O

515a (15.6%)

O

1S

O

O

A. niger HO

HO

513a΄

O

514a΄ (17.0%)

514b΄ (6.5%)

515a΄ (69.2%)

FIGURE 23.154  Biotransformation of (1R)-α-ionone (513a) and (1S)-α-ionone (513a′) by Aspergillus niger.

Aspergillus, Botryosphaeria, and Chlorella species. The metabolite (522) was further reduced to 523 by E. gracilis (Noma and Asakawa, 1998). Biotransformation of (+)-1R-α-ionone (513a), [α]D + 386.5°, 99% ee, and (−)-1S-α-ionone (513a′), [α]D 361.6°, 98% ee, which were obtained by optical resolution of racemic α-ionone (513), was fed by A. niger for 4 days in Czapek-peptone medium. From 513a, 4α-hydroxy-α-ionone (514a), 4β-hydroxy-αionone (514b), and 4-oxo-α-ionone (515a) were obtained, while from compound 513a′, the enantiomers (514a′, 514b′, 515a′) of the metabolites from 513a were obtained; however, the difference of their yields was observed. In case of 513a, 4α-hydroxy-α-ionone (514a) was obtained as the major product, while 515a′ was predominantly obtained from 513a′. This oxidation was inhibited by 1-aminobenzotriazole, thus, CYP-450 is contributed to this oxidation process (Hashimoto et al., 2000a) (Figure 23.154). α-Damascone (525) was incubated with A. niger and A. terreus, in Czapek-peptone medium to give cis- (525) and trans-3-hydroxy-α-damascones (527) and 3-oxo-α-damscone (528), while the latter Aspergillus species afforded 3-oxo-8,9-dihydro-α-damascone (529). The hydroxylation process of 525–527 was inhibited by CYP-450 inhibitor. H. anomala reduced α-damascone (525) to α-damascol (530). Cis- (526) and trans-4-hydroxy-α-damascone (527) were fed by C. pyrenoidosa in Noro medium to give 4-oxodamascone (528) (Noma et al., 2001b) (Figure 23.155). 13 O

12 5 4

3

r

ige

A. n

HO 526

5 P4

r ito

17 8 2 525

yre

no

A. n

CY

530

ige

P4

r

50 X inh ib

O

ito

CY

C. p

10 H. anomala

11

hib

n

0i

O

6

OH 9

r

HO

sa

ido

no yre

ido

sa

O

527

C. p

O

A. terreus O

O 528

529

FIGURE 23.155  Biotransformation of α-damascone (525) by various microorganisms.

856

Handbook of Essential Oils O

O A. niger

HO

532

531 O

O

O O

A. terreus

er

ig A. n

534

533 OH

OH

O

O

537

OH

A. nig

er OH

535

O

536

O

FIGURE 23.156  Biotransformation of β-damascone (531) by Aspergillus niger and Aspergillus terreus.

β-Damascone (531) was also treated in A. niger to afford 5-hydroxy-β-damascone (532), 3-hydroxy-β-damascone (533), 5-oxo- (534), 3-oxo-β-damscone (535), and 3-oxo-1,9-dihydroxy1,2-dihydro-β-damascone (536) as the minor components. In case of A. terreus, 3-hydroxy-8,9dihydro-β-damascone (537) was also obtained (Figure 23.156). Adamantane derivatives have been used as many medicinal drugs. In order to obtain the drugs, adamantanes were incubated by many microorganisms, such as A. niger, A. awamori, A. cellulosae, Aspergillus fumigatus, A. sojae, A. terreus, B. dothidea, C. pyrenoidosa IMCC-28, C. sorokiniana, F. culmorum, E. gracilis, and H. anomala (Figure 23.157). Adamantane (538) was incubated with A. niger, A. cellulosae, and B. dothidea in Czapekpeptone medium. The same substrate was also treated in C. pyrenoidosa in Noro medium. Compound 538 was converted into both 1-hydroxy- (539) and 9α-hydroxyadamantane (540) by all four microorganisms, followed by oxidation oxidized to give 1,9α-dihydroxyadamantanol (541) by A. niger, which was further oxidized to 1-hydroxyadamantane-9-one (542), which was reduced to afford 1,9β-hydroxyadamantane (544). A. niger gave the metabolite (541) as the major product in 80% yield. A. cellulosae converted 538–539 and 540 in the ratio of 81:19. C. pyrenoidosa gave 539, 540, and adamantane-9-one (543) in the ratio 74:16:10. 4α-Adamantanol (540) was directly OH a

7 5 6

1

a

541

a-d

a 9 3

4

542

a a 540

HO 544

O

OH a-d

2 538

a

HO

539

8 10

a HO

HO

OH

O

543

a: A. niger b: A. cellulosae c: B. dothidea d: C. pyrenoidosa

FIGURE 23.157  Biotransformation of adamantane (538) by various microorganisms.

857

Biotransformation of Sesquiterpenoids and Their Related Compounds

converted by C. pyrenoidosa, A. niger, and A. cellulosae to afford 543, which was also reduced to 9α-adamantanol (540) by A. niger. The biotransformation of adamantane, however, did not occur by the microorganisms H. anomala, C. sorokiniana, D. tertiolecta, and E. gracilis (Noma et al., 1999). Adamantanes (538–543, 542) were also incubated with various fungi including with F. culmorum. 1-Hydroxyadamantane-9-one (542) was reduced stereoselectively to 541 by A. niger, A. cellulosae, B. dothidea, and F. culmorum. On the contrary, F. culmorum reduced 541–542. A. cellulosae and B. dothidea bioconverted 542 to 1,9β-hydroxyadamantane (544) stereoselectively. Adamantane-9-one (543) was treated by A. niger to give nonstereoselectively 545–547 that were further converted into diketone (548, 549) and a diol (550). It is noteworthy that oxidation and reduction reactions were observed between ketoalcohol (547) and diols (551, 552). The same phenomenon was also seen between 546 and 553. The latter diol was also oxidized by A. niger to furnish diketone (549) (Noma et al., 2001c, 2003). Direct hydroxylation at C3 of 1-hydroxyadamantane-9-one (542) was seen in the incubation of 539 with A. niger. 4-Adamantanone (543) showed promotion effect of cell division of the fungus, while 1-adamantanol (539) and adamantane-9-one (543) inhibited germination of lettuce seed. 1-Hydroxyadamantane-9one (542) inhibited the elongation of root of lettuce, while adamantane-1,4-diol (544) and adamantane itself (538) promoted root elongation (Noma et al., 1999, 2001c) (Figure 23.158). Stereoselective reduction of racemic bicyclo[3.3.1]nonane-2,6-dione (555a, 555a′) was carried out by A. awamori, A. fumigatus, A. cellulosae, A. sojae, A. terreus, A. niger, B. dothidea, and F. culmorum in Czapek-peptone, H. anomala in yeast, E. gracilis in Hunter, and D. tertiolecta in Noro

a-e HO

538

OH

a 539

a-d

HO

a

OH

554 OH

OH

HO

544

a HO

540

e

541

d

b,

a,

a,b,d

O

OH

a

O

HO

HO

550

O

a

a

546

O a HO

543

542

HO

548

O

O

a: A. niger b: A. cellulosae c: C. pyrenoidosa d: B. dothidea e: F. curmurorum

a O

HO

a O

O 545

a

a 546

OH

HO

a

a

553

549

551

a

a

a

OH

HO

OH

HO

O

HO

a a

552

547

FIGURE 23.158  Biotransformation of adamantane (538) and adamantane-9-one by various micro­organisms.

858

Handbook of Essential Oils Aspergillus sp. B. dothidea F. curmurorum

O

O 555

OH

OH

OH

H. anomala E. gracilis D. tertiolecta

O 556

O

557 HO

HO

HO O

555a

O

556a

557a

Piridium dichromate

FIGURE 23.159  Biotransformation of bicyclo[3.3.1]nonane-2,6-dione (555a, 555a′) by various microorganisms.

medium. All microorganisms reduced 555 and 555a′ to give corresponding monoalcohol (556, 556a) and optical-active (−)-diol (557a) ([α]D − 71.8° in the case of A. terreus), which was formed by enantioselective reduction of racemic monool, 556 and 556a (Noma et al., 2003) (Figure 23.159).

23.5  BIOTRANSFORMATION OF AROMATIC COMPOUNDS Essential oils contain aromatic compounds, such as p-cymene, carvacrol, thymol, vanillin, cinnamaldehyde, eugenol, chavicol, safrole, and asarone (558), among others. Takahashi (1994) reported that simple aromatic compounds, propylbenzene, hexylbenzene, decylbenzene, o- and p-hydroxypropiophenones, p-methoxypropiophenone, 4-hexylresorcinol, and methyl 4-hexylresorcionol, were incubated with A. niger. From hexyl- and decylbenzenes, w1-hydroxylated products were obtained, whereas from propylbenzene, w2-hydroxylated metabolites were obtained (Takahashi, 1994). Asarone (558) and dihydroeugenol (562) were not biotransformed by A. niger. However, dihydroasarone (559) and methyl dihydroeugenol (563) were biotransformed by the same fungus to produce a small amount of 2-hydroxy (560, 561) and 2-oxo derivatives (564, 565), respectively. The chirality at C2 was determined to be R and S mixtures (1:2) by the modified Mosher method (Takahashi, 1994) (Figure 23.160). Chlorella species are excellent microalgae as oxidation bioreactors as mentioned earlier. Treatment of monoterpene aldehydes and related aldehydes were reduced to the corresponding primary alcohols, indicating that these green algae possess reductase. A microalgae E. gracilis Z. also contains reductase. The following aromatic aldehydes were treated in this organism: benzaldehyde; 2-cyanobenzaldehyde; o-, m-, and p-anisaldehyde; o-, m-, and p-salicylaldehyde; o-, m-, and p-tolualdehyde; o-chlorobenzaldehyde; p-hydroxybenzaldehyde; o-, m-, and p-nitrobenzaldehyde; 3-cyanobenzaldehyde; vanillin; isovanillin; o-vanillin; nicotine aldehyde; 3-phenylpropionaldehyde; and ethyl vanillin. Veratraldehyde, 3-nitrosalicylaldehde, phenylacetaldehyde, and 2-phenylproanaldehyde gave their corresponding primary alcohols. 2-Cyanobenzaldehyde gave its corresponding alcohol with phthalate. m- and p-Chlorobenzaldehyde gave its corresponding alcohols and m- and p-chlorobenzoic acids. o-Phthalaldehyde and p-phthalate and iso- and terephthalaldehydes gave their corresponding monoalcohols and dialcohols. When cinnamaldehyde and α-methyl cinnamaldehyde were incubated in E. gracilis, cinnamyl alcohol and 3-phenylpropanol, and 2-methylcinnamyl alcohol, and 2-methyo-3-phenylpropanol were obtained in

859

Biotransformation of Sesquiterpenoids and Their Related Compounds OMe OMe MeO

558 OMe

OMe OMe

OMe OMe

OMe A. niger MeO

MeO

MeO O

OH 559

560

561

OH OMe

562 OMe

OMe

OMe OMe

OMe

OMe

OH

O

A. niger

563

564

565

FIGURE 23.160  Biotransformation of dihydroasarone (559) and methyl dihydroeugenol (563) by Aspergillus niger.

good yield. E. gracilis could convert acetophenone to 2-phenylethanol; however, its enantio-excess is very poor (10%) (Takahashi, 1994). Raspberry ketone (566) and zingerone (574) are the major components of raspberry (Rubus idaeus) and ginger (Zingiber officinale), and these are used as food additive and spice. Two substrates were incubated with the Phytolacca americana cultured cells for 3 days to produce two secondary alcohols (567, 568) as well as five glucosides (569–572) from 566 and a secondary alcohol (576) and four glycoside products (575, 577–579) from 574. In the case of raspberry ketone, phenolic hydroxyl group was preferably glycosylated after the reduction of carbonyl group of the substrate occurred. It is interesting to note that one more hydroxyl group was introduced into the benzene ring to give 568, which were further glycosylated by one of the phenolic hydroxyl groups, and no glycoside of the secondary alcohol at C2 were obtained (Figure 23.161). On the other hand, zingerone (574) was converted into 576, followed by glycosylation to give both glucosides (577, 578) of phenolic and secondary hydroxyl groups and a diglucoside (579) of both phenolic and secondary hydroxyl group in the molecule. It is the first report on the introduction of individual glucose residues onto both phenolic and secondary hydroxyl groups by cultured plant cells (Shimoda et al., 2007a) (Figure 23.162).

860

Handbook of Essential Oils

HO

GlcO

Raspberry (566)

569

O

O

HO

GlcO

HO

567

570

OH

GlcO

HO

HO 568

OGlc

HO

GlcO

HO

571

OH

572

OH

573

OH

OH

FIGURE 23.161  Biotransformation of raspberry ketone (566) by Phytolacca americana cells. HO

GlcO

H3CO

H3CO

Gingerone (574) O

575

O

576

OH

H3CO

H3CO

H3CO

H3CO

GlcO

HO

GlcO

HO

577

OH

578

OGlc

579

OGlc

FIGURE 23.162  Biotransformation of zingerone (574) by Phytolacca americana cells.

Thymol (580), carvacrol (583), and eugenol (586) were glucosylated by glycosyltransferase of cell-cultured Eucalyptus perriniana to each glucoside (581%, 3%; 584%, 5%; 587%, 7%) and gentiobioside (582%, 87%; 585%, 56%; 588%, 58%). The yield of thymol glycosides was 1.5 times higher than that of carvacrol and 4 times higher than that of eugenol. Such glycosylation is useful to obtain higher water-soluble products from natural and commercially available secondary metabolites for food additives and cosmetic fields (Shimoda et al., 2006) (Figure 23.163). Hinokitiol (589), which is easily obtained from cell suspension cultures of Thujopsis dolabrata and possesses potent antimicrobial activity, was incubated with cultured cells of E. perriniana for 7 days to give its monoglucosides (590, 591%, 32%) and gentiobiosides (592, 593) (Furuya et al., 1997; Hamada et al., 1998) (Figure 23.164). (−)-Nopol benzyl ether (594) was smoothly biotransformed by A. niger, A. cellulosae, A. sojae, Aspergillus usami, and Penicillium species in Czapek-peptone medium to give (−)-4-oxonopol-2′,4′dihydroxybenzyl ether (595%, 23% in the case of A. niger), which demonstrated antioxidant activity (ID50 30.23 m/M), together with a small amount of nopol (6.3% in A. niger). This is very rare direct introduction of oxygen function on the phenyl ring (Noma and Asakawa, 2006) (Figure 23.165).

861

Biotransformation of Sesquiterpenoids and Their Related Compounds

HO HO

HO

OH O

HO HO

OH O

O OH HO HO

O

OH

580

O O OH

581

HO HO

HO

582

HO HO

OH O

OH O O OH HO HO

O

OH

583

584

OCH3 HO HO

HO

OH O O

O

OH

585

HO HO

OCH3

OH

OH O O OH HO HO

OCH3

O O OH 588

587

586

O

FIGURE 23.163  Biotransformation of thymol (580), carvacrol (583), and eugenol (586) by Eucalyptus ­perriniana cells.

HO O HO O

HO HO

CH2OH O

O

590

E. perriniana O

Hinokitiol (589)

OH O

HO HO

HO

HO HO

CH2OH O O O HO 591

CH2OH O

O CH 2 OH O HO O HO OH O HO CH2OH O HO O HO CH2 HO O HO HO

592

O O

593

FIGURE 23.164  Biotransformation of hinokitiol (589) by Eucalyptus perriniana cells.

Capsicum annuum contains capsaicin (596), and its homologues having an alkylvanillylamides possess various interesting biological activities such as anti-inflammatory, antioxidant, salivaand stomach juice–inducing activity, analgesic, antigenotoxic, antimutagenic, anticarcinogenic, antirheumatoid arthritis, and diabetic neuropathy and are used as food additives. On the other hand, because of potent pungency and irritation on skin and mucous membrane, it has not yet been permitted as medicinal drug. In order to reduce this typical pungency and application of nonpungent capsaicin metabolites to the crude drug, capsaicin (596) (600 mg) including 30% of dihydrocapsaicin (600) was incubated in Czapek-peptone medium including A. niger for 7 days to give three metabolites, w1-hydroxylated capsaicin (597, 60.9%), 8,9-dihydro-w1-hydroxycapsaicin (598%, 16%), and a carboxylic acid (599, 13.6%). All of the metabolites do not show pungency (Figure 23.166).

862

Handbook of Essential Oils

O

OH

HO

A. niger A. cellulosae A. sojae A. usami Penicillium sp. KCPYN

O

(–)-Nopol benzylether (594)

595 (23%)

FIGURE 23.165  Biotransformation of nopol benzyl ether (531) by Aspergillus and Penicillium species. O MeO HO

HO

N H Capsaicin (596)

9

A. niger

MeO HO

HO

11

N H

OH

598 (16.0%) 7 COOH

N H

OH

597 (60.9%)

O MeO

11

8

O

O MeO

N H

599 (13.6%)

FIGURE 23.166  Biotransformation of capsaicin (596) by Aspergillus niger.

Dihydrocapsaicin (600) was also treated in the same manner as described earlier to afford w1-hydroxydihyrocpsaicin (598, 80.9%) in high yield and the carboxylic acid (599, 5.0%). Capsaicin itself showed carbachol-induced contraction of 60% in the bronchus at a concentration of 1 mmol/L. 11-Hydroxycapsicin (85) retained this activity of 60% at a concentration of 30 mmol/L. Dihydrocapsaicin (600) showed the same activity of contraction in the bronchus, at the same concentration as that used in capsaicin. However, the activity of contraction in the bronchus of 11-hydroxy derivative (598) showed weaker (50% at 30 mmol/L) than that of the substrate. Since both metabolites (597 and 598) are tasteless, these products might be valuable for the crude drug although the contraction in the bronchus is weak. 2,2-Diphenyl-1-picrylhydrazyl radical scavenging activity test of capsaicin and dihydrocapsaicin derivatives was carried out. 11-Hydroxycapsaicin (597), 11-dihydrocapsaicin (598), and capsaicin (596) showed higher activity than (±)-α-tocopherol, and 11-dihydroxycapsaicin (598) displayed a strong scavenging activity (IC50 50 mmol/L) (Hashimoto and Asakawa, unpublished results) (Figure 23.167). Shimoda et al. (2007b) reported the bioconversion of capsaicin (596) and 8-nor-dihydrocapsaicin (601) by the cultured cell of Catharanthus roseus to give more water-soluble capsaicin derivatives. From capsaicin, three glycosides, capsaicin 4-O-β-D-glucopyranoside (602), which was one of the capsaicinoids in the fruit of Capsicum and showed 1/100 weaker pungency than capsaicin, 4-O-(6O-β-D-xylopyranosyl)-β-D-glucoside (603), and 4-O-(6-O-α-L-arbinosyl)-β-D-glucopyranoside (604), were obtained. 8-Nor-dihydrocapsaicin (601) was also incubated with the same cultured cell to afford the similar products (605–607) all of which reduced their pungency and enhanced water solubility. Since many synthetic capsaicin glycosides possess remarkable pharmacological activity, such as decrease of liver and serum lipids, the present products will be used for valuable prodrugs (Figure 23.168). Z. officinale contains various sesquiterpenoids and pungent aromatic compounds such as 6-shogaol (608) and 6-gingerol (613), and their pungent compounds that possess cardio tonic and sedative

863

Biotransformation of Sesquiterpenoids and Their Related Compounds O MeO

N H

HO

Dihydrocapsaicin (600)

Aspergillus niger

O MeO

N

OH

H

HO

598

O MeO

COOH

N H

HO

599

FIGURE 23.167  Biotransformation of dihydrocapsaicin (600) by Aspergillus niger.

O

HO HO HO H3CO

H N

HO HO

596 O

OH OO OH H3CO

H N 602 O

O OH O O HO HO OH H3CO OH O O HO OH O O HO OH HO H3CO

H3CO

H N

601 O

HO HO

OH OO OH H3CO

H N

605 O

603 O

H N 604 O

O

HO HO HO

H N

O OH O O HO HO OH H3CO OH

HO

H N

606 O

O

O OH O O HO HO OH H3CO

H N

607 O

FIGURE 23.168  Biotransformation of capsaicin (596) and 8-nor-dihydrocapsaicin (601) by Catharanthus roseus cells.

864

Handbook of Essential Oils O

MeO

O

6

A. niger

7

HO

MeO

2 days

8

HO

6-Shagol (608)

6

OH

7

609 (9.9%)

O MeO

MeO

COOH 9

OH

O 5 8

MeO

OH

5

HO

HO

HO

8

612 (48.5%)

610 (16.1%)

611 (22.4%)

FIGURE 23.169  Biotransformation of 6-shogaol (608) by Aspergillus niger.

activity. 6-Shogaol (608) was incubated with A. niger in Czapek-peptone medium for 2 days to afford w1-hydroxy-6-shagaol (609, 9.9%), which was further converted to 8-hydroxy derivative (610, 16.1%), a γ-lactone (611, 22.4%), and 3-methoxy-4-hydroxyphenylacetic acid (612, 48.5%) (Figure 23.169). 6-Gingerol (613) (1 g) was treated in the same condition as mentioned earlier to yield six metabolites, w1-hydroxy-6-gingerol (614, 39.8%), its carboxylic derivative (616, 14.5%), a γ-lactone (618) (16.9%) that might be formed from (616), its 8-hydoxy-γ-lactone (619, 12.1%), w2-hydroxy-6-­ gingerol (615, 19.9%), and 6-deoxy-gingerol (617, 14.5%) (Takahashi et al., 1993a). The metabolic pathway of 6-gingerol (613) resembles that of 6-shagaol (608). That of 6-shogaol and dihydrocapsaicin (600) is also similar since both substrates gave carboxylic acids as the final metabolites (Takahashi et al., 1993a) (Figure 23.170). O MeO

OH 6

HO

1

6-Gingerol (613)

A. niger

O

6

1

OH

O

MeO HO

614 (39.8%)

OH

OH

6

2

615 (19.9%)

O

OH

MeO HO

O

OH

MeO HO

A. niger

6

3

COOH

MeO HO

616 (14.5%)

OH

6

1

617 (14.5%) O

O O MeO HO

O 3 6

618 (16.9%)

OH MeO HO

8 619 (21.1%)

FIGURE 23.170  Biotransformation of 6-gingerol (613) by Aspergillus niger.

O

Biotransformation of Sesquiterpenoids and Their Related Compounds

865

In conclusion, a number of sesquiterpenoids were biotransformed by various fungi and mammals to afford many metabolites, several of which showed antimicrobial and antifungal, antiobesity, cytotoxic, neurotrophic, and enzyme inhibitory activity. Microorganisms introduce oxygen atom at allylic position to give secondary hydroxyl and keto groups. Double bond is also oxidized to give epoxide, followed by hydrolysis to afford a diol. These reactions precede stereo- and regiospecifically. Even at nonactivated carbon atom, oxidation reaction occurs to give primary alcohol. Some fungi like A. niger cleave the cyclopropane ring with a 1,1-dimethyl group. It is noteworthy that A. niger and A. cellulosae produce totally different metabolites from the same substrates. Some fungi occurs reduction of carbonyl group, oxidation of aryl methyl group, phenyl coupling, and cyclization of a 10-membered ring sesquiterpenoids to give C6/C6- and C5/C7-cyclic or spiro compounds. Cytochrome P-450 is responsible for the introduction of oxygen function into the substrates. The present methods are very useful for the production of medicinal and agricultural drugs as well as fragrant components from commercially available cheap, natural, and unnatural terpenoids or a large amount of terpenoids from higher medicinal plants and spore-forming plants like liverworts and fungi. The methodology discussed in this chapter is a very simple one-step reaction in water, nonhazard, and very cheap, and it gives many valuable metabolites possessing different properties from those of the substrates.

REFERENCES Abraham, W.R., L. Ernst, and B. Stumpf, 1990. Biotransformation of caryophyllene by Diplodia gossypina. Phytochemisrty, 29: 115–120. Abraham, W.-R., K. Kieslich, B. Stumpf, and L. Ernst, 1992. Microbial oxidation of tricyclic sesquiterpenoids containing a dimethylcyclopropane ring. Phytochemistry, 31: 3749–3755. Abraham, W.-R., P. Washausen, and K. Kieslich, 1987. Microbial hydroxylation of cedrol and cedrene. Z. Naturforsch., 42C: 414–419. Aleu, J., J.R. Hanson, R. Hernandez-Galan, and I.G. Collado, 1999a. Biotransformation of the fungistatic sesquiterpenoid patchoulol by Botrytis cinerea. J. Nat. Prod., 62: 437–440. Aleu, J., R. Hernandez-Galan, and I.G. Collad, 2002. Biotransformation of the fungistatic sesquiterpenoid isobotryan-9α-ol by Botrytis cinerea. J. Mol. Catal. B, 16: 249–253. Aleu, J., R. Hernandez-Galan, J.R. Hanson, P.B. Hitchcock, and I.G. Collado, 1999b. Biotransformation of the fungistatic sesquiterpenoid ginsenol by Botrytis cinerea. J. Chem. Soc. Perkin Trans., 1: 727–730. Amate, A., A. Garcia-Granados, A. Martinez et al. 1991. Biotransformation of 6α-eudesmanolides functionalized at C-3 with Curvularia lunata and Rhizopus nigricans cultures. Tetrahedron, 47: 5811–5818. Aranda G., I. Facon, J.-Y. Lallemand, and M. Leclaire, 1992. Microbiological hydroxylation in the drimane series. Tetrahedron Lett., 33, 7845–7848. Aranda, G., M.S. Kortbi, J.-Y. Lallemand et al. 1991. Microbial transformation of diterpenes: Hydroxylation of sclareol, manool and derivatives by Mucor plumbeus. Tetrahedron, 47: 8339–8350. Arantes, S.F., J.R. Hanson, and P.B. Hitchcok, 1999. The hydroxylation of the sesquiterpenoid valerianol by Mucor plumbeus. Phytochemistry, 52: 1063–1067. Asakawa, Y., 1982. Chemical constituents of the Hepaticae. In: Progress in the Chemistry of Organic Natural Products, Herz, W., H. Grisebach, and G.W. Kirby (eds.), Vol. 42, pp. 1–285. Vienna, Austria: Springer. Asakawa, Y., 1990. Terpenoids and aromatic compounds with pharmaceutical activity from bryophytes. In: Bryophytes: Their Chemistry and Chemical Taxonomy, Zinsmeister, D.H. and R. Mues (eds.), pp. 369– 410. Oxford, U.K.: Clarendon Press. Asakawa, Y., 1995. Chemical constituents of the bryophytes. In: Progress in the Chemistry of Organic Natural Products, Herz, W., G.W. Kirby, R.E. Moore, W. Steglich, and Ch. Tamm (eds.), Vol. 65, pp. 1–618. Vienna, Austria: Springer. Asakawa, Y., 1999. Phytochemistry of bryophytes. In: Phytochemicals in Human Health Protection, Nutrition, and Plant Defense, J. Romeo (ed.), Vol. 33, pp. 319–342. New York: Kluwer Academic, Plenum Publishers. Asakawa, Y., 2007. Biologically active compounds from bryophytes. Pure Appl. Chem., 75: 557–580. Asakawa, Y., 2008. Recent advances of biologically active substances from the Marchantiophyta. Nat. Prod. Commun., 3: 77–92. Asakawa, Y., T. Hashimoto, Y. Mizuno, M. Tori, and Y. Fukuzawa, 1992. Cryptoporic acids A-G, drimane-type sesquiterpenoid ethers of isocitric acid from the fungus Cryptoporus volvatus. Phytochemistry, 31: 579–592.

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Asakawa, Y., T. Ishida, M. Toyota, and T. Takemoto, 1986. Terpenoid biotransformation in mammals IV. Biotransformation of (+)-longifolene, (−)-caryophyllene, (−)-caryophyllene oxide, (−)-cyclocolorenone, (+)-nootkatone, (−)-elemol, (−)-abietic acid and (+)-dehydroabietic acid in rabbits. Xenobiotica, 6: 753–767. Asakawa, Y. and A. Ludwiczuk, 2008. Bryophytes-Chemical diversity, bioactivity and chemosystematics. Part 1. Chemical diversity and bioactivity. Med. Plants Poland World, 14: 33–53. Asakawa, Y., Z. Taira, T. Takemoto, T. Ishida, M. Kido, and Y. Ichikawa, 1981. X-ray crystal structure analysis of 14-hydroxycaryophyllene oxide, a new metabolite of (−)-caryophyllene in rabbits. J. Pharm. Sci., 70: 710–711. Asakawa, Y., H. Takahashi, and M. Toyota, 1991. Biotransformation of germacrane-type sesquiterpenoids by Aspergillus niger. Phytochemistry, 30: 3993–3997. Ata, A. and J.A. Nachtigall, 2004. Microbial transformation of α-santonin. Z. Naturforsch., 59C: 209–214. Atta-ur-Rahman, M.I. Choudhary, A. Ata et al., 1994. Microbial transformation of 7α-hydroxyfrullanolide. J. Nat. Prod., 57: 1251–1255. Atta-ur-Rahman, M.I. Choudhary, F. Shaheen, A. Rauf, and A. Farooq, 1998. Microbial transformation of some bioactive natural products. Nat. Prod. Lett., 12: 215–222. Atta-ur-Rahman, A. Farooq, and M.I. Choudhary, 1997. Microbial transformation of sclareolide. J. Nat. Prod., 60: 1038–1040. Ayer, W.A. and P.A. Craw, 1989. Metabolites of fairy ring fungus, Marasmius oreades. Part 2. Norsesquiterpenes, further sesquiterpenes, and argocybin. Can. J. Chem., 67: 1371–1380. Bang, L. and G. Ourisson, 1975. Hydroxylation of cedrol by rabbits. Tetrahedron Lett., 16: 1881–1884. Bang, L., G. Ourisson, and P. Teisseire, 1975. Hydroxylation of patchoulol by rabbits. Hemisynthesis of norpatchoulenol, the odour carrier of patchouli oil. Tetrahedron Lett., 16: 2211–2214. Barrero, A.F., J.E. Oltra, D.S. Raslan, and D.A. Sade, 1999. Microbial transformation of sesquiterpene lactones by the fungi Cunninghamella echinulata and Rhizopus oryzae. J. Nat. Prod., 62: 726–729. Bhutani, K.K. and R.N. Thakur, 1991. The microbiological transformation of parthenin by Beauveria bassiana and Sporotrichum pulverulentum. Phytochemistry, 30: 3599–3600. Buchanan, G.O., L.A.D. Williams, and P.B. Reese, 2000. Biotransformation of cadinane sesquiterpenes by Beauveria bassiana ATCC 7159. Phytochemistry, 54: 39–45. Choudhary, M.I., W. Kausar, Z.A. Siddiqui, and Atta-ur-Rahman, 2006a. Microbial metabolism of (+)-cycloisolongifol-5β-ol. Z. Naturforsch., 61B: 1035–1038. Choudhary, M.I., S.G. Musharraf, S.A. Nawaz et al., 2005. Microbial transformation of (−)-isolongifolol and butyrylcholinesterase inhibitory activity of transformed products. Bioorg. Med. Chem., 13: 1939–1944. Choudhary, M.I., S.G. Musharraf, A. Sami, and Atta-ur-Rahman, 2004. Microbial transformation of sesquiterpenes, (−)-ambrox® and (+)-sclareolide. Helv. Chim. Acta, 87: 2685–2694. Choudhary, M.I., Z.A. Siddiqui, S.A. Nawaz, and Atta-ur-Rahman, 2006b. Microbial transformation and butyrylcholinesterase inhibitory activity of (−)-caryophyllene oxide and its derivatives. J. Nat. Prod., 69: 1429–1434. Clark, A.M. and C.D. Hufford, 1979. Microbial transformation of the sesquiterpene lactone costunolide. J. Chem. Soc. Perkin Trans., 1: 3022–3028. Collins, D.O. and P.B. Reese, 2002. Biotransformation of cadina-4,10(15)-dien-3-one and 3α-hydroxycadina4,10(15)-diene by Curvularia lunata ATCC 12017. Phytochemistry, 59: 489–492. Collins, D.O., W.F. Reynold, and P.B. Reese, 2002a. Aromadendrane transformations by Curvularia lunata ATCC 12017. Phytochemistry, 60: 475–481. Collins, D.O., P.L.D. Ruddock, J. Chiverton, C. de Grasse, W.F. Reynolds, and P.B. Reese, 2002b. Microbial transformation of cadina-4,10(15)-dien-3-one, aromadendr-1(10)-en-9-one and methyl ursolate by Mucor plumbeus ATCC 4740. Phytochemistry, 59: 479–488. Daniewski, W.M., P.A. Grieco, J. Huffman, A. Rymkiewicz, and A. Wawrzun, 1981. Isolation of 12-hydroxycaryophyllene-4,5-oxide, a sesquiterpene from Lactarius camphoratus. Phytochemistry, 20: 2733–2734. Darrouzet, E., B. Mauchamp, G.D. Prestwich, L. Kerhoas, I. Ujvary, and F. Couillaud 1997. Hydroxy juvenile hormones: New putative juvenile hormones biosynthesized by locust corpora allata in vitro. Biochem. Biophys. Res. Commun., 240: 752–758. Devi, J.R., 1979. Microbiological transformation of terpenes: Part XXVI. Microbial transformation of caryophyllene. Ind. J. Biochem. Biophys., 16: 76–79. Dhavlikar, R.S. and G. Albroscheit, 1973. Microbiologische Umsetzung von Terpenen: Valencen. Dragoco Rep., 12: 250–258. Duran, R., E. Corrales, R. Hernandez-Galan, and G. Collado, 1999. Biotransformation of caryophyllene oxide by Botrytis cinerea. J. Nat. Prod., 62: 41–44.

Biotransformation of Sesquiterpenoids and Their Related Compounds

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El Sayed, K.A., M. Yousaf, M.T. Hamann, M.A. Avery, M. Kelly, and P. Wipf, 2002. Microbial and chemical transformation studies of the bioactive marine sesquiterpenes (S)-(+)-curcuphenol and -curcudiol isolated from a deep reef collection of the Jamaican sponge Didiscus oxeata. J. Nat. Prod., 65: 1547–1553. Furusawa, M., 2006. Microbial biotransformation of sesquiterpenoids from crude drugs and liverworts: Production of functional substances. PhD thesis. Tokushima Bunri University, Tokushima, Japan, pp. 1–156. Furusawa, M., T. Hashimoto, Y. Noma, and Y. Asakawa, 2005a. Biotransformation of Citrus aromatics nootkatone and valencene by microorganisms. Chem. Pharm. Bull., 53: 1423–1429. Furusawa, M., T. Hashimoto, Y. Noma, and Y. Asakawa 2005b. Highly efficient production of nootkatone, the grapefruit aroma from valencene, by biotransformation. Chem. Pharm. Bull., 53: 1513–1514. Furusawa, M., T. Hashimoto, Y. Noma, and Y. Asakawa, 2005c. The structure of new sesquiterpenoids from the liverwort Reboulia hemisphaerica and their biotransformation. Proceeding of 49th TEAC, pp. 235–237. Furusawa, M., T. Hashimoto, Y. Noma, and Y. Asakawa, 2006a. Biotransformation of aristolene- and 2,3-secoaromadendrane-type sesquiterpenoids having a 1,1-dimethylcyclopropane ring by Chlorella fusca var. vacuolata, Mucor species, and Aspergillus niger. Chem. Pharm. Bull., 54: 861–868. Furusawa, M., T. Hashimoto, Y. Noma, and Y. Asakawa, 2006b. Isolation and structures of new cyclomyltaylane and ent-chamigrane-type sesquiterpenoids from the liverwort Reboulia hemisphaerica. Chem. Pharm. Bull., 54: 996–1003. Furusawa, M., Y. Noma, T. Hashimoto, and Y. Asakawa, 2003. Biotransformation of Citrus oil nootkatone, dihydronootkatone and dehydronootkatone. Proceedings of 47th TEAC, pp. 142–144. Furuya, T., Y. Asada, Y. Matsuura, S. Mizobata, and H. Hamada, 1997. Biotransformation of β-thujaplicin by cultured cells of Eucalyptus perriniana. Phytochemistry, 46: 1355–1358. Galal, A.M., 2001. Microbial transformation of pyrethrosin. J. Nat. Prod., 64: 1098–1099. Galal, A.M., A.S. Ibrahim, J.S. Mossa, and F.S. El-Feraly, 1999. Microbial transformation of parthenolide. Phytochemistry, 51: 761–765. Garcia-Granados, A., M.C. Gutierrez, F. Rivas, and J.M. Arias, 2001. Biotransformation of 4β-hydroxyeudesmane1,6-dione by Gliocladium roseum and Exserohilum halodes. Phytochemistry, 58: 891–895. Hamada, H., F. Murakami, and T. Furuya, 1998. The production of hinokitiol glycoside. Proceedings of 42nd TEAC, pp. 145–147. Hanson, J.R. and H. Nasir, 1993. Biotransformation of the sesquiterpenoid, cedrol, by Cephalosporium aphidicola. Phytochemistry, 33: 835–837. Hanson, J.R. and A. Truneh, 1996. The biotransformation of ambrox and sclareolide by Cephalosporium aphidicola. Phytochemistry, 42: 1021–1023. Hanson, R.L., J.M. Wasylyk, V.B. Nanduri, D.L. Cazzulino, R.N. Patel, and L.J. Szarka, 1994. Site-specific enzymatic hydrolysis of tisanes at C-10 and C-13. J. Biol. Chem., 269: 22145–22149. Harinantenaina, L., Y. Noma, and Y. Asakawa, 2005. Penicillium sclerotiorum catalyzes the conversion of herbertenediol into its dimers: Mastigophorenes A and B. Chem. Pharm. Bull., 53: 256–257. Harinantenaina, L., D.N. Quang, T. Nishizawa et al., 2007. Bioactive compounds from liverworts: Inhibition of lipopolysaccharide-induced inducible NOS mRNA in RAW 264.7 cells by herbertenoids and cuparenoids. Phytomedicine, 14: 486–491. Hashimoto, T. and Y. Asakawa, 2007. Biological activity of fragrant substances from Citrus and herbs, and production of functional substances using microbial biotransformation. In: Development of Medicinal Foods, M. Yoshikawa (ed.), pp. 168–184. Tokyo, Japan: CMC Publisher. Hashimoto, T., Y. Asakawa, Y. Noma et al., 2003a. Production method of nootkatone. Jpn. Kokai Tokkyo Koho, 250591A. Hashimoto, T., Y. Asakawa, Y. Noma et al., 2003b. Production method of nootkatone. Jpn. Kokai Tokkyo Koho, 70492A. Hashimoto, T., M. Fujiwara, K. Yoshikawa, A. Umeyama, M. Tanaka, and Y. Noma, 2007. Biotransformation of sclareolide and sclareol by microorganisms. Proceedings of 51st TEAC, pp. 316–318. Hashimoto, T., S. Kato, M. Tanaka, S. Takaoka, and Y. Asakawa, 1998a. Biotransformation of sesquiterpenoids by microorganisms (4): Biotransformation of hinesol by Aspergillus niger. Proceedings of 42nd TEAC, pp. 127–129. Hashimoto, T., Y. Noma, Y. Akamatsu, M. Tanaka, and Y. Asakawa, 1999a. Biotransformation of sesquiterpenoids by microorganisms. (5): Biotransformation of dehydrocostuslactone. Proceedings of 43rd TEAC, pp. 202–204. Hashimoto, T., Y. Noma, and Y. Asakawa, 2001a. Biotransformation of terpenoids from the crude drugs and animal origin by microorganisms. Heterocycles, 54: 529–559.

868

Handbook of Essential Oils

Hashimoto, T., Y. Noma, and Y. Asakawa, 2006. Biotransformation of cuparane- and herbertane-type sesquiterpenoids. Proceedings of 50th TEAC, pp. 263–265. Hashimoto, T., Y. Noma, Y. Goto, S. Takaoka, M. Tanaka, and Y. Asakawa, 2003c. Biotransformation of sesquiterpenoids from the liverwort Plagiochila species. Proceedings of 47th TEAC, pp. 139–141. Hashimoto, T., Y. Noma, Y. Goto, M. Tanaka, S. Takaoka, and Y. Asakawa, 2004. Biotransformation of (−)-maalioxide by Aspergillus niger and Aspergillus cellulosae. Heterocycles, 62: 655–666. Hashimoto, T., Y. Noma, S. Kato, M. Tanaka, S. Takaoka, and Y. Asakawa, 1999b. Biotransformation of hinesol isolated from the crude drug Atractylodes lancea by Aspergillus niger and Aspergillus cellulosae. Chem. Pharm. Bull., 47: 716–717. Hashimoto, T., Y. Noma, H. Matsumoto, Y. Tomita, M. Tanaka, and Y. Asakawa, 2000a. Microbial biotransformation of optically active (+)-α-ionone and (−)-α-ionone. Proceedings of 44th TEAC, pp. 154–156. Hashimoto, T., Y. Noma, Y. Matsumoto, Y. Akamatsu, M. Tanaka, and Y. Asakawa, 1999c. Biotransformation of sesquiterpenoids by microorganisms. (6): Biotransformation of α-, β- and γ-cyclocostunolides. Proceedings of 43rd TEAC, pp. 205–207. Hashimoto, T., Y. Noma, C. Murakami, N. Nishimatsu, M. Tanaka, and Y. Asakawa, 2001b. Biotransformation of valencene and aristolene. Proceedings of 45th TEAC, pp. 345–347. Hashimoto, T., Y. Noma, C. Murakami, M. Tanaka, and Y. Asakawa, 2000b. Microbial transformation of α-santonin derivatives and nootkatone. Proceedings of 44th TEAC, pp. 157–159. Hashimoto, T., Y. Noma, N. Nishimatsu, M. Sekita, M. Tanaka, and Y. Asakawa, 2003d. Biotransformation of antimalarial sesquiterpenoids by microorganisms. Proceedings of 47th TEAC, pp. 136–138. Hashimoto, T., M. Sekita, M. Furusawa, Y. Noma, and Y. Asakawa, 2005. Biotransformation of sesquiterpene lactones, (−)-parthenolide and (−)-frullanolide by microorganisms. Proceedings of 49th TEAC, pp. 387–389. Hashimoto, T., K. Shiki, M. Tanaka, S. Takaoka, and Y. Asakawa, 1998b. Chemical conversion of labdanetype diterpenoid isolated from the liverwort Porella perrottetiana into (−)-ambrox. Heterocycles, 49: 315–325. Hayashi, K., H. Morikawa, A. Matsuo, D. Takaoka, and H. Nozaki, 1999. Biotransformation of sesquiterpenoids by Aspergillus niger IFO 4407. Proceedings of 43rd TEAC, pp. 208–210. Hayashi, K., H. Morikawa, H. Nozaki, and D. Takaoka, 1998. Biotransformation of globulol and epiglubulol by Aspergillus niger IFO 4407. Proceedings of 42nd TEAC, pp. 136–138. Haze, S., K. Sakai, and Y. Gozu, 2002. Effects of fragrance inhalation on sympathetic activity in normal adults. Jpn. J. Pharmacol., 90: 247–253. Hegazy, M.-E.F., C. Kuwata, Y. Sato et al., 2005. Research and development of asymmetric reaction using biocatalysts-biotransformation of enones by cultured cells of Marchantia polymorpha. Proceedings of 49th TEAC, pp. 402–404. Higuchi, H., R. Tsuji, K. Hayashi, D. Takaoka, A. Matsuo, and H. Nozaki, 2001. Biotransformation of sesquiterpenoids by Aspergillus niger. Proceedings of 45th TEAC, pp. 354–355. Hikino, H., T. Konno, T. Nagashima, T. Kohama, and T. Takemoto, 1971. Stereoselective epoxidation of germacrone by Cunninghamella blakesleeana. Tetrahedron Lett., 12: 337–340. Hrdlicka, P.J., A.B. Sorensen, B.R. Poulsen, G.J.G. Ruijter, J. Visser, and J.J.L. Iversen, 2004. Characterization of nerolidol biotransformation based on indirect on-line estimation of biomass concentration and physiological state in bath cultures of Aspergillus niger. Biotechnol. Prog., 20: 368–376. Ishida, T., 2005. Biotransformation of terpenoids by mammals, microorganisms, and plant-cultured cells. Chem. Biodivers., 2: 569–590. Ishida, T., Y. Asakawa, and T. Takemoto, 1982. Hydroxyisolongifolaldehyde: A new metabolite of (+)-longifolene in rabbits. J. Pharm. Sci., 71: 965–966. Ishida, T., Y. Asakawa, T. Takemoto, and T. Aratani, 1981. Terpenoid biotransformation in mammals. III. Biotransformation of α-pinene, β-pinene, pinane, 3-carene, carane, myrcene and p-cymene in rabbits. J. Pharm. Sci., 70: 406–415. Itsuzaki, Y., K. Ishisaka, and M. Miyazawa, 2002. Biotransformation of (+)-cedrol by using human skin microbial flora Staphylococcus epidermidis. Proceedings of 46th TEAC, pp. 101–102. Jacobsson, U., V. Kumar, and S. Saminathan, 1995. Sesquiterpene lactones from Michelia champaca. Phytochemistry, 39: 839–843. Kaspera, R., U. Krings, T. Nanzad, and R.G. Berger, 2005. Bioconversion of (+)-valencene in submerged cultures of the ascomycete Chaetomium globosum. Appl. Microbiol. Biotechnol., 67: 477–583. Kiran, I., T. Akar, A. Gorgulu, and C. Kazaz, 2005. Biotransformation of racemic diisophorone by Cephalosporium aphidicola and Neurospora crassa. Biotechnol. Lett., 27: 1007–1010.

Biotransformation of Sesquiterpenoids and Their Related Compounds

869

Kiran, I., H.N. Yildirim, J.R. Hanson, and P.B. Hitchcock, 2004. The antifungal activity and biotransformation of diisophorone by the fungus Aspergillus niger. J. Chem. Technol. Biotechnol., 79: 1366–1370. Kumari, G.N.K., S. Masilamani, R. Ganesh, and S. Aravind, 2003. Microbial biotransformation of zaluzanin D. Phytochemistry, 62: 1101–1104. Lahlou, E.L., Y. Noma, T. Hashimoto, and Y. Asakawa, 2000. Microbiotransformation of dehydropinguisenol by Aspergillus sp. Phytochemistry, 54: 455–460. Lamare, V. and R. Furstoss, 1990. Bioconversion of sesquiterpenes. Tetrahedron, 46: 4109–4132. Lee, I.-S., H.N. ElSohly, E.M. Coroom, and C.D. Hufford, 1989. Microbial metabolism studies of the antimalarial sesquiterpene artemisinin. J. Nat. Prod., 52: 337–341. Ma, X.C., M. Ye, L.J. Wu, and D.A. Guo, 2006. Microbial transformation of curdione by Mucor spinosus. Enzyme Microb. Technol., 38: 367–371. Maatooq, G.A., 2002b. Microbial transformation of a β- and γ-eudesmols mixture. Z. Naturforsch., 57C: 654–659. Maatooq, G.A., 2002c. Microbial conversion of partheniol by Calonectria decora. Z. Naturforsch., 57C: 680–685. Maatooq, G.T., 2002a. Microbial metabolism of partheniol by Mucor circinelloides. Phytochemistry, 59: 39–44. Madyastha, K.M. and T.L. Gururaja, 1993. Utility of microbes in organic synthesis: Selective transformations of acyclic isoprenoids by Aspergillus niger. Ind. J. Chem., 32B: 609–614. Matsui, H., Y. Minamino, and M. Miyazawa, 1999. Biotransformation of (+)-cedryl acetate by Glomerella cingulata, parasitic fungus. Proceedings of 43rd TEAC, pp. 215–216. Matsui, H. and M. Miyazawa, 2000. Biotransformation of pathouli acetate using parasitic fungus Glomerella cingulata as a biocatalyst. Proceedings of 44th TEAC, pp. 149–150. Matsushima, A., M.-E.F. Hegazy, C. Kuwata, Y. Sato, M. Otsuka, and T. Hirata, 2004. Biotransformation of enones using plant cultured cells—The reduction of α-santonin. Proceedings of 48th TEAC, pp. 396–398. Meyer, P. and C. Neuberg 1915. Phytochemische reduktionen. XII. Die umwandlung von citronellal in citronelol. Biochem. Z, 71: 174–179. Mikami, Y., 1988. Microbial Conversion of Terpenoids. Biotechnology and Genetic Engineering Reviews, Vol. 6, pp. 271–320. Wimborne, UK: Intercept Ltd. https://www.nottingham.ac.uk/ncmh/documents/ bger/volume-6/bger6-7.pdf Accessed date: 16 March 2020. Miyazawa, M., S. Akazawa, H. Sakai, and H. Kameoka, 1997a. Biotransformation of (+)-γ-gurjunene using plant pathogenic fungus, Glomerella cingulata as a biocatalyst. Proceedings of 41st TEAC, pp. 218–219. Miyazawa, M., Y. Honjo, and H. Kameoka, 1996a. Biotransformation of guaiol and bulnesol using plant pathogenic fungus Glomerella cingulata as a biocatalyst. Proceedings of 40th TEAC, pp. 82–83. Miyazawa, M., Y. Honjo, and H. Kameoka, 1997b. Biotransformation of the sesquiterpenoid β-selinene using the plant pathogenic fungus Glomerella cingulata. Phytochemistry, 44: 433–436. Miyazawa, M., Y. Honjo, and H. Kameoka, 1998a. Biotransformation of the sesquiterpenoid (+)-γ-gurjunene using a plant pathogenic fungus Glomerella cingulata as a biocatalyst. Phytochemistry, 49: 1283–1285. Miyazawa, M., H. Matsui, and H. Kameoka, 1997c. Biotransformation of patchouli alcohol using plant parasitic fungus Glomerella cingulata as a biocatalyst. Proceedings of 41st TEAC, pp. 220–221. Miyazawa, M., H. Matsui, and H. Kameoka, 1998b. Biotransformation of unsaturated sesquiterpene alcohol using plant parasitic fungus, Glomerella cingulata as a biocatalyst. Proceedings of 42nd TEAC, pp. 121–122. Miyazawa, M., H. Nakai, and H. Kameoka, 1995a. Biotransformation of (+)-cedrol by plant pathogenic fungus, Glomerella cingulata. Phytochemistry, 40: 69–72. Miyazawa, M., H. Nakai, and H. Kameoka, 1995b. Biotransformation of cyclic terpenoids, (±)-cis-nerolidol and nerylacetone, by plant pathogenic fungus, Glomerella cingulata. Phytochemistry, 40: 1133–1137. Miyazawa, M., H. Nakai, and H. Kameoka, 1996b. Biotransformation of acyclic terpenoid (±)-trans-nerolidol and geranylacetone by Glomerella cingulata. J. Agric. Food Chem., 44: 1543–1547. Miyazawa, M., H. Nakai, and H. Kameoka, 1996c. Biotransformation of acyclic terpenoid (2E,6E)-farnesol by plant pathogenic fungus Glomerella cingulata. Phytochemistry, 43: 105–109. Miyazawa, M., H. Nankai, and H. Kameoka, 1995c. Biotransformation of (−)-α-bisabolol by plant pathogenic fungus, Glomerella cingulata. Phytochemistry, 39: 1077–1080. Miyazawa, M. and A. Sugawara, 2006. Biotransformation of (2)-guaiol by Euritium rubrum. Nat. Prod. Res., 20: 731–734. Miyazawa, M., T. Uemura, and H. Kameoka, 1994. Biotransformation of sesquiterpenoids, (−)-globulol and (+)-ledol by Glomerella cingulata. Phytochemistry, 37: 1027–1030.

870

Handbook of Essential Oils

Miyazawa, M., T. Uemura, and H. Kameoka, 1995d. Biotransformation of sesquiterpenoids, (+)-aromadendrene and (−)-alloaromadendrene by Glomerella cingulata. Phytochemistry, 40: 793–796. Morikawa, H., K. Hayashi, K. Wakamatsu et al., 2000. Biotransformation of sesquiterpenoids by Aspergillus niger IFO 4407. Proceedings of 44th TEAC, pp. 151–153. Nankai, H., M. Miyazawa, and H. Kameoka, 1996. Biotransformation of (Z,Z)-farnesol using plant pathogenic fungus, Glomerella cingulata as a biocatalyst. Proceedings of 40th TEAC, pp. 78–79. Nishida, E. and Y. Kawai, 2007. Bioconversion of zerumbone and its derivatives. Proceedings of 51st TEAC, pp. 387–389. Noma, Y. and Y. Asakawa, 1998. Microbiological transformation of 3-oxo-α-ionone. Proceedings of 44th TEAC, pp. 133–135. Noma, Y. and Y. Asakawa, 2006. Biotransformation of (−)-nopol benzyl ether. Proceedings of 50th TEAC, pp. 434–436. Noma, Y., M. Furusawa, C. Murakami, T. Hashimoto, and Y. Asakawa, 2001a. Formation of nootkatol and nootkatone from valencene by soil microorganisms. Proceedings of 45th TEAC, pp. 91–92. Noma, Y., T. Hashimoto, Y. Akamatsu, S. Takaoka, and Y. Asakawa, 1999. Microbial transformation of adamantane (Part 1). Proceedings of 43rd TEAC, pp. 199–201. Noma, Y., T. Hashimoto, and Y. Asakawa, 2001b. Microbiological transformation of damascone. Proceedings of 45th TEAC, pp. 93–95. Noma, Y., T. Hashimoto, and Y. Asakawa, 2001c. Microbial transformation of adamantane. Proceedings of 45th TEAC, pp. 96–98. Noma, Y., T. Hashimoto, S. Kato, and Y. Asakawa, 1997a. Biotransformation of (+)-β-eudesmol by Aspergillus niger. Proceedings of 41st TEAC, pp. 224–226. Noma, Y., T. Hashimoto, A. Kikkawa, and Y. Asakawa, 1996. Biotransformation of (−)-α-eudesmol by Asp. niger and Asp. cellulosae M-77. Proceedings of 40th TEAC, pp. 95–97. Noma, Y., T. Hashimoto, S. Sawada, T. Kitayama, and Y. Asakawa, 2002. Microbial transformation of zerumbone. Proceedings of 46th TEAC, pp. 313–315. Noma, Y., K. Matsueda, I. Maruyama, and Y. Asakawa, 1997b. Biotransformation of terpenoids and related compounds by Chlorella species. Proceedings of 41st TEAC, pp. 227–229. Noma, Y., Y. Takahashi, and Y. Asakawa, 2003. Stereoselective reduction of racemic bicycle[33.1]nonane2,6-dione and 5-hydroxy-2-adamantanone by microorganisms. Proceedings of 46th TEAC, pp. 118–120. Nozaki, H., K. Asano, K. Hayashi, M. Tanaka, A. Masuo, and D. Takaoka, 1996. Biotransformation of shiromodiol diacetate and myli-4(15)-en-9-one by Aspergillus niger IFO 4407. Proceedings of 40th TEAC, pp. 108–110. Okuda, M., K. Sonohara, and H. Takikawa, 1994. Production of natural flavors by laccase catalysis. Jpn. Kokai Tokkyo Koho, 303967. Orabi, K.Y., 2000. Microbial transformation of the eudesmane sesquiterpene plectranthone. J. Nat. Prod., 63: 1709–1711. Orabi, K.Y., 2001. Microbial epoxidation of the tricyclic sesquiterpene presilphiperfolane angelate ester. Z. Naturforsch., 56C: 223–227. Otsuka, S. and K. Tani, 1991. Catalytic asymmetric hydrogen migration of ally amines. Synthesis, (9) 665–680. https://www.thieme-connect.com/products/ejournals/pdf/10.1055/s-1991-26541.pdf. Accessed date: 16 March 2020. Parshikov, I.A., K.M. Muraleedharan, and M.A. Avery, 2004. Transformation of artemisinin by Cunning­ hamella elegans. Appl. Microbiol. Biotechnol., 64: 782–786. Sakamoto, S., N. Tsuchiya, M. Kuroyanagi, and A. Ueno, 1994. Biotransformation of germacrone by suspension cultured cells. Phytochemistry, 35: 1215–1219. Sakata, K., I. Horibe, and M. Miyazawa, 2007. Biotransformation of (+)-α-longipinene by microorganisms as a biocatalyst. Proceedings of 51st TEAC, pp. 321–322. Sakata, K. and M. Miyazawa, 2006. Biotransformation of (+)-isolongifolen-9-one by Glomerella cingulata as a biocatalyst. Proceedings of 50th TEAC, pp. 258–260. Sakui, N., M. Kuroyamagi, Y. Ishitobi, M. Sato, and A. Ueno, 1992. Biotransformation of sesquiterpenes by cultured cells of Curcuma zedoaria. Phytochemistry, 31: 143–147. Sakui, N., M. Kuroyanagi, M. Sato, and A. Ueno, 1988. Transformation of ten-membered sesquiterpenes by callus of Curcuma. Proceedings of 32nd TEAC, pp. 322–324. Salvador, J.A.R. and J.H. Clark, 2002. The allylic oxidation of unsaturated steroids by tert-butyl hydroperoxide using surface functionalized silica supported metal catalysts. Green Chem., 4: 352–356.

Biotransformation of Sesquiterpenoids and Their Related Compounds

871

Sekita, M., M. Furusawa, T. Hashimoto, Y. Noma, and Y. Asakawa, 2005. Biotransformation of pungent tasting polygodial from Polygonum hydropiper and related compounds by microorganisms. Proceedings of 49th TEAC, pp. 380–381. Sekita, M., T. Hashimoto, Y. Noma, and Y. Asakawa, 2006. Biotransformation of biologically active terpenoids, sacculatal and cinnamodial by microorganisms. Proceedings of 50th TEAC, pp. 406–408. Shimoda, K., T. Harada, H. Hamada, N. Nakajima, and H. Hamda, 2007a. Biotransformation of raspberry ketone and zingerone by cultured cells of Phytolacca americana. Phytochemistry, 68: 487–492. Shimoda, K., Y. Kondo, T. Nishida, H. Hamada, N. Nakajima, and H. Hamada, 2006. Biotransformation of thymol, carvacrol, and eugenol by cultured cells of Eucalyptus perriniana. Phytochemistry, 67: 2256–2261. Shimoda, K., S. Kwon, A. Utsuki et al., 2007b. Glycosylation of capsaicin and 8-norhydrocapsaicin by cultured cells of Catharanthus roseus. Phytochemistry, 68: 1391–1396. Shoji, N., A. Umeyama, Y. Asakawa, T. Takeout, K. Nocoton, and Y. Ohizumi, 1984. Structure determination of nootkatol, a new sesquiterpene isolated form Alpinia oxyphylla Miquel possessing calcium antagonist activity. J. Pharm. Sci., 73: 843–844. Sowden, R.J., S. Yasmin, N.H. Rees, S.G. Bell, and L.-L. Wong, 2005. Biotransformation of the sesquiterpene (+)-valencene by cytochrome P450cam and P450BM-3. Org. Biomol. Chem., 3: 57–64. Sugawara, A. and M. Miyazawa, 2004. Biotransformation of guaiene using plant pathogenic fungus, Eurotium rubrum as a biocatalyst. Proceedings of 48th TEAC, pp. 385–386. Sutherland, T.D., G.C. Unnithan, J.F. Andersen et al., 1998. A cytochrome P450 terpenoid hydroxylase linked to the suppression of insect juvenile hormone synthesis. Proc. Natl. Acad. Sci. USA, 95: 12884–12889. Takahashi, H., 1994. Biotransformation of terpenoids and aromatic compounds by some microorganisms. PhD thesis. Tokushima Bunri University, Tokushima, Japan, pp. 1–115. Takahashi, H., T. Hashimoto, Y. Noma, and Y. Asakawa, 1993a. Biotransformation of 6-gingerol, and 6-shogaol by Aspergillus niger. Phytochemistry, 34: 1497–1500. Takahashi, T., I. Horibe, and M. Miyazawa, 2007. Biotransformation of β-selinene by Aspergillus wentii. Proceedings of 51st TEAC, pp. 319–320. Takahashi, T. and M. Miyazawa, 2005. Biotransformation of (+)-nootkatone by Aspergillus wentii, as biocatalyst. Proceedings of 49th TEAC, pp. 393–394. Takahashi, T. and M. Miyazawa, 2006. Biotransformation of sesquiterpenes which possess an eudesmane skeleton by microorganisms. Proceedings of 50th TEAC, pp. 256–257. Takahashi, H., M. Toyota, and Y. Asakawa, 1993b. Drimane-type sesquiterpenoids from Cryptoporus volvatus infected by Paecilomyces variotii. Phytochemistry, 33: 1055–1059. Tani, K., T. Yamagata, S. Otsuka et al. 1982. Cationic rhodium (I) complex-catalyzed asymmetric isomerization of allylamines to optically active enamines. J. Chem. Soc. Chem. Commun., (11) 600–601. Tori, M., M. Sono, and Y. Asakawa, 1990. The reaction of three sesquiterpene ethers with m-chloroperbenzoic acid. Bull. Chem. Soc. Jpn., 63: 1770–1776. Venkateswarlu, Y., P. Ramesh, P.S. Reddy, and K. Jamil, 1999. Microbial transformation of D9(15)-africane. Phytochemistry, 52: 1275–1277. Wang, Y., T.-K. Tan, G.K. Tan, J.D. Connolly, and L.J. Harrison, 2006. Microbial transformation of the sesquiterpenoid (−)-maalioxide by Mucor plumbeus. Phytochemistry, 67: 58–61. Wilson, C.W. III and P.E. Saw, 1978. Quantitative composition of cold-pressed grapefruit oil. J. Agric. Food Chem., 26: 1430–1432. Yang, L., K. Fujii, J. Dai, J. Sakai, and M. Ando, 2003. Biotransformation of α-santonin and its C-6 epimer by fungus and plant cell cultures. Proceedings of 47th TEAC, pp. 148–150. Zhan, J., H. Guo, J. Dai, Y. Zhang, and D. Guo, 2002. Microbial transformation of artemisinin by Cunninghamella echinulata and Aspergillus niger. Tetrahedron Lett., 43: 4519–4521. Zundel, J.-L., 1976. PhD thesis. Universite Louis Pasteur, Strasbourg, France.

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Use of Essential Oils in Agriculture Catherine Regnault-Roger, Susanne Hemetsberger, and Gerhard Buchbauer

CONTENTS 24.1 Introduction........................................................................................................................... 874 24.1.1 Essential Oils: Very Complex Natural Mixes........................................................... 875 24.2 Essential Oils as Antipests.................................................................................................... 876 24.2.1 Health and Environmental Impact of Botanical Antipests....................................... 876 24.2.2 Pesticidal and Repellent Action of Essential Oils..................................................... 877 24.2.2.1 Insecticidal Activities of Essential Oils...................................................... 877 24.2.3 Development and Commercialization of Botanical................................................... 883 24.2.3.1 Examples of Essential Oils Used as Antipests........................................... 885 24.3 Essential Oils as Herbicides.................................................................................................. 892 24.3.1 Phytotoxicity.............................................................................................................. 892 24.3.2 Prospects of Organic Weed Control.......................................................................... 892 24.3.3 Examples of Essential Oils in Weed Control............................................................ 892 24.3.3.1 Thymus vulgaris.......................................................................................... 892 24.3.3.2 Mentha sp.................................................................................................... 898 24.3.3.3 Cymbopogon sp.......................................................................................... 898 24.3.3.4 Eucalyptus sp.............................................................................................. 898 24.3.3.5 Lavandula sp.............................................................................................. 899 24.3.3.6 Origanum sp............................................................................................... 899 24.3.3.7 Artemisia scoparia...................................................................................... 899 24.3.3.8 Zataria multiflora....................................................................................... 899 24.3.3.9 Tanacetum sp.............................................................................................. 899 24.4 Essential Oils as Inhibitors of Various Pests......................................................................... 899 24.4.1 Effect on Bacteria...................................................................................................... 901 24.4.2 Effect on Fungi..........................................................................................................902 24.4.3 Effect on Viruses.......................................................................................................902 24.4.4 Effect on Nematodes..................................................................................................903 24.5 Effect of Essential Oils on the Condition of the Soil............................................................903 24.5.1 Effects of Essential Oils on Microorganisms and Soil..............................................903 24.5.2 Examples of Essential Oils with an Effect on Soil Conditions.................................905 24.5.2.1 Mentha spicata...........................................................................................905 24.5.2.2 Lavandula sp..............................................................................................906 24.5.2.3 Salvia sp......................................................................................................906 24.5.2.4 Myrtus communis.......................................................................................906 24.5.2.5 Laurus nobilis.............................................................................................906 24.5.2.6 Cymbopogon sp..........................................................................................906 24.6 EOs Used in Post-Harvest Disease Control...........................................................................907 24.6.1 Effects of Essential Oils on Stored-Product Pests.....................................................907 24.6.2 Examples of Essential Oils Used on Stored Products...............................................907 24.6.2.1 Thymus zygis...............................................................................................907 873

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24.6.2.2 Cinnamomum sp.........................................................................................907 24.6.2.3 Cymbopogon citratus..................................................................................907 24.6.2.4 Laurus nobilis.............................................................................................909 24.7 Conclusion.............................................................................................................................909 References....................................................................................................................................... 910

24.1 INTRODUCTION Essential oils (EOs) play a major role in nature for the plants that produce them in order to protect the plants against bacteria, fungi, and viruses, as well as against herbivores and pests (Bakkali et al., 2008). Since pollution of our environment and intoxication of mammalians with some pesticides and herbicides is an acute problem all over the world, healthier alternatives must be researched. Natural products such as EOs offer some solutions for agricultural pests as far as they are efficient but nontoxic to vertebrates and do not harm our ambience. Even though there are so many advantages of EOs, also some disadvantages must be mentioned, like the slow action and short duration of effectiveness as well as the high quantities needed. The object of this compilation is to review the chances and problems of the use of EOs in agriculture. Crop protection has been an important topic in agriculture for thousands of years. Many Greek and Roman authors—for example, Theophrastus (371−287 B.C.), Cato the Censor (234−149 B.C.), Varro (116−27 B.C.), Vergil (70−19 B.C.), Columella (4−70 A.D.), and Pliny the elder (23−79 A.D.)—wrote about different substances, such as EOs, to protect plants against pests. Also, Chinese literature, such as a survey of the Shengnong Ben Tsao Jing era (25−220 A.D.) tells about pesticidal activity of plants. Famously, Linnaeus (1752 A.D.) did research about natural pesticides which could be used on caterpillars. During the 19th century, compounds of plant origins were identified and frequently used as repellents or toxic compounds. Among them are alkaloids like nicotine and its isomer anabasine extracted from tobacco, a Solanaceae (Nicotiana tabacum, N. rustica, and N. galuca) or isolated from a plant growing in the Russian steppes and high plateaux of North Africa, Anabasis aphylla; nornicotine from Duboisia hopwoodi (an Australian plant); veratrine extracted from a Liliaceae of the Balkans, Veratrum album; ryanodine identified from Rynia spp. (chiefly R. speciose) in Amazonia; and also other families of molecules represented by rotenone and rotenoids and by pyrethrines, which were made of Chrysanthemum (Pyrethrum) cinerariaefolium flower heads and Lonchocarpus nicou or Derris elliptica roots. Most of these compounds are not used today because of toxicity to humans (e.g., neurotoxic activity of nicotine or Parkinson disease linked with rotenone) (Philogène et al., 2005). Consequently and according to the societal claim for a friendly environment fort use of pesticides today, more plant-derived solutions to control agricultural pests have been actively explored since 1960. Among these plant extracts, EOs became important, because they have a wide range of activities against microorganisms, weeds, and insects (insecticides, antifeedants, or repellents), as previously mentioned (Bakkali et al., 2008). Grieneisen and Isman (2018) have counted more than 1300 published articles per year devoted to botanical research efforts since 2012, with 20% focused on EOs, including numerous reviews (as examples, Arnason et al., 2012; Regnault-Roger et al., 2012a,b; Regnault-Roger, 2013; Buchbauer and Hemetberger, 2016; Mossa, 2016; Sparagano et al., 2016; Isman and Tak, 2017a). EOs uses have a long history. They have been used for many industrial applications for a long time. The first distillation of these products was mentioned during the 13th century in Andalusia (Spain) by Ibn al-Baitar, after which their pharmacological properties induced their inclusion into the very early pharmacopoeias of several European countries (Regnault-Roger, 2013). These oils were first used largely for medicinal or ceremonial purposes, and according to Isman and Tak (2017a), anecdotal records suggest that “certain aromatic oils were burned during the Black Death (1346–1353) ‘to ward off evil spirits’ (healingscents.net), but may have inadvertently reduced the spread of bubonic plague by repelling or killing fleas that vectored the pathogen.” They have been used since ancient times as cosmetics and pharmaceuticals and also since the last century

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in fine chemistry and aromatics for the food industry. Today, they still play a major role in our life (Ammon et al., 2010). The plant protection in agriculture is now one very important focus of EOs activities.

24.1.1  Essential Oils: Very Complex Natural Mixes EOs appear to be very complex natural mixes. EOs are biosynthesized by around 17,500 species of aromatic plants belonging to a few families. They are particularly abundant in Conifers, Rutaceae, Umbelliferae, Myrtaceae, Lamiaceae, and Lauraceae. Depending on the species and families, they are localized in specialized histological structures: glandular trichomes (Lamiaceae), secretory canals (Myrtaceae), or resin ducts (Apiaceae). They could be stored in different parts of the plant such as flowers, leaves, wood, roots, or seeds (Bruneton, 2009). Terpenoids are major constituents of EOs and, to a lesser amount, phenylpropanoids. EO constituents belonging to terpenoids are mainly monoterpenes (ten atoms of carbon) and sesquiterpenes (15 atoms of carbon) of low molecular weight. They generally consist of several tens of constituents, of which the great majority possess an isoprenoid skeleton. Monoterpenes present in EOs may contain terpenes that are hydrocarbons (alpha-pinene), alcohols (menthol, geraniol, linalool, terpinen-4-ol, p-menthane-3,8-diol), aldehydes (cinnamaldehyde, cuminaldehyde), ketones (thujone), ethers (1,8-cineole or eucalyptol), and lactones (nepetalactone). As the elongation of the chain to 15 carbons increases the number of possible cyclizations, sesquiterpenes have a wide variety of structures (over 100 skeletons). Aromatic compounds are less common and are derived mainly from the shikimate pathway. Some compounds identified in EOs result from the degradation of fatty acids (jasmonic acid) or are glycosylated volatile compounds (e.g., linalool glucoside) (RegnaultRoger et al., 2012a,b). The majority of EOs contains a limited number of main compounds, but some of the minor compounds play an important role as vectors of fragrance and make up the richness of an extract. It is thus well established that EO composition is very variable depending of the species and of chemotypes within the species, as well as physiological parameters. Thyme (Thymus vulgaris L.) is a species with numerous chemotypes, named according to the major compound; for example, thyme with chemotype thymol or chemotype carvacrol or terpineol or linalool. A typical EO may contain 20–80 phytochemicals. Physiological expression of secondary metabolism of the plant may be different at all stages of its development. The proportions of monoterpenes depend on temperature and circadian rhythm and vary according to plant stage. For example, Gershenzon et  al. (2000) showed that limonene and menthone are the major monoterpenes present in the youngest leaves of peppermint, but limonene content declines rapidly with development whereas menthone increases and then declines at later stages; consequently, menthol becomes the dominant constituent. Soil acidity and climate (heat, photoperiod, humidity) directly affect the secondary metabolism of the plant and EO composition (Muller-Riebau et al., 1997). Isman and Machial (2006) reported that rosemary oil extracted from plants harvested in two different areas of Italy contained 1,8-cineole concentrations ranging from 7% to 55% and α-pinene concentrations ranging from 11% to 36%. The variability of the composition of an EO is also impacted by the choice of the method of extraction of EOs. One characteristic of EOs is the volatility of their compounds which allows them to be easily extracted by water vapors, in contrast to fixed lipid oils and essences (concrete, absolute, oleoresins, and resinoids) which are extracted by solvents and alcohol. More recently, extraction and separation methods using supercritical fluids, steam distillation, dry distillation, or mechanical cold pressing of plants are also mentioned (RegnaultRoger et al. 2012a,b). All these factors lead to the complexity of EO phytochemistry and to a certain inconsistency of the chemical composition of an EO. The biological significance of EOs has long been discussed. First considered as wastes of phytometabolism, it appears today that EOs present multiple actions. They prevent plants from losing water by excessive evaporation and some of their components react as donors of hydrogen in

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oxydation-reduction reactions. They also seem to be important agents of interspecific communication as they favor pollination by attracting insects and they also play a part in the defense of plants against herbivores, microorganisms, and fungi (Regnault-Roger, 1997). Therefore, all these activities could be taken into account in the development of pest-management strategies.

24.2  ESSENTIAL OILS AS ANTIPESTS Insects form the largest population in the animal kingdom and many of them are harmful toward human beings since they act as pathogenic vectors and devaster crops. Pest control, both by synthetics and biopesticides, bear a huge market in the regular growth. The massive use of oil-based synthetic pesticides since the middle of the 20th century, with limited knowledge of properties of some compounds, produced unexpected effects with negative sides, such as environmental persistence (organochloride compounds classified as POPs), chronic toxicity toward human or mammalians and increased insect resistance (Regnault-Roger, 1997). Because of that, it is important to find alternatives which do not damage our environment.

24.2.1 Health and Environmental Impact of Botanical Antipests EOs are natural products, which are excellent alternatives to synthetic products because they reduce negative impacts on human health and the environment (Koul et al., 2008). Even though there are a lot of botanical insecticides available, from 1980 to 2000, only one single product was registered in the United States and Europe, which is called Neem (now considered an endocrine disruptor). It is obtained by the seeds of the Indian tree Azadirachta indica, Meliaceae. In the last few years, EOs also obtained influence as botanical insecticides in the United States. But, due to the relatively slow action, variable efficacy, lack of persistence and inconsistent availability of natural products, they still cannot compete against synthetic pesticides. On the other hand, it is not necessary to kill harmful insects since botanical insecticides, like EOs, can also be used as repellents against animals. Moreover, those natural pesticides can also be mixed with synthetic products which could lessen the needed quantities of pesticides and improve the environmental problems which were caused by excessive use of synthetic pesticides. Especially, developing countries could benefit from the increased use of botanical pesticides, such as EOs, since farmers are more familiar with plant extracts and EOs they can do by themselves. They often use pesticides and do not know about the dangers because many of them do not receive any information and they are often unable to read the official languages in which the instructions are written (Isman, 2008). In static water, eugenol appears to be 1500 times less toxic than pyrethrum, which is also a botanical insecticide, and not more than 15,000 times less toxic than the organophosphate insecticide azinphosmethyl. Moreover, eugenol is volatile, which means that its half-life is extremely short. After about two days, there will be no eugenol left, which also avoids rare side effects of EOs (Isman, 2000). Also, the tobacco cutworm, which is a severe problem to vegetable and tobacco crops in Asia, can be killed by compounds of EOs, namely thymol, carvacrol, pulegone, eugenol, and trans-anethole, even though it is quite resistant (Hummelbrunner et al., 2001). Many monoterpenes like (+)-limonene, pinene, and Δ3-carene show acaricidal activity. Carvomenthenol and terpinen-4-ol proved to cause highest toxicity against mites (Ibrahim et  al., 2008). Eucalyptus EO was tested on several parasites, such as Varroa destructor (Varroa mite), Tetranychus urticae, Phytoseiulus perisimilis, and Boophilus microplus. The studies concluded that several Eucalyptus sp. (Myrtaceae) EOs can be used as acaricides (Batish et al., 2008). T. urticae can be killed by EOs obtained from Satureja hortensis L. (Lamiaceae), Ocimum basilicum L. (Lamiacae), and Thymus vulgaris L. (Lamiaceae) (Aslan et al., 2004). EOs do not only act as deterrents but also as attractants toward insects. Ethanolic extracts of Rosmarinus officinalis L. (Lamiaceae) EO attracted the moth Loberia botrana, a pest of grape berries (Katerinopoulos et al., 2005).

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24.2.2  Pesticidal and Repellent Action of Essential Oils EOs are neurotoxic to some pests as some of them interfere with the neuromodulator octopamine and others with GABA-gated chloride channels (Isman, 2006). Octopamine “controls and modulates neuronal development, circadian rhythm, locomotion ‘fight or flight’ responses, as well as learning and memory” (Balfanz et al. 2005). It activates GTP-binding protein G-coupled receptors, which leads to cAMP production or calcium release (Balfanz et al., 2005). Interrupting its function, “results in total break down of nervous system in insects” (Tripathi et  al., 2009). GABAA receptors of insects showed similarities as well as differences to vertebrate ones. Activation of insect GABA channels can increase the chloride ion conductance across cell membranes. Insect GABA receptors are also sensitive to benzodiazepines and barbiturates, but they are insensitive to antagonists like bicuculline and pitrazepin. The differences of insect and vertebrate GABA channels can be used as targets for pesticides (Anthony et al., 1993). At high concentrations of EOs, they can also inhibit the acetylcholinesterase activity, but it cannot explain the low-dose pesticidal effect (Kostyukovsky et al., 2002). Moreover, the acetylcholinesterase-inhibiting effect could not be seen in vivo for all monoterpenes (Rajendran et al., 2008). It seems to be a very good idea to use nontoxic substances that only act as deterrents or antifeedants toward insects. This concept obtained influence with the discovery of the antifeedant azadirachtin in the 1970s and 1980s. But even though deterrents and antifeedants were hyped a lot during that time, a few problems remain, such as the variability of response—even closely related insects vary in their behavior toward these substances; some may even get attracted by substances that other insects may find detesting. Furthermore, the deterrent action may change during a period of time, and insects can get used to it (Isman, 2006). The exact mode of action of repellents still cannot be explained completely. It is also unclear if there are common mechanisms in different anthropods. Just as insects do on the antennae, “tick detects repellents on the tarsi of the first pair of legs” (Tripathi et al., 2009). It is known that hairs on the antennae of mosquitos are used to detect temperature and moisture. Repellents target female mosquito olfactory receptors and block them. As for cockroaches and the method of defense, is just known that oleic and linoleic acid cause death recognition and aversion (Tripathi et al., 2009). Even though single compounds of the EO can be isolated and tested for repellent activity, it has been reported that the synergistic effect of the whole EO is more effective (Nerio et al., 2010) (Table 24.1). 24.2.2.1  Insecticidal Activities of Essential Oils An abundant literature, more than 2000 scientific papers, is devoted to study the EO–insect relationships over the past 20 years, with some major reviews (Regnault-Roger, 1997; Isman, 2000, 2006; Regnault-Roger and Philogène, 2008; Regnault-Roger et al., 2008; Adorjan and Buchbauer, 2010; Isman et al., 2011; and reviews mentioned above). The majority of this scientific literature focuses on the characterization of bioactivities of oils in the laboratory from previously uninvestigated plant species or untested insect pests. Most papers document the immediate effects (acute toxicity or repellency) of given EOs on a number of arthropod taxa, frequently on the basis of assays lasting less than 48 h, but few papers investigate the modes or mechanisms of action of EOs and the real efficacy in the field and the effective protection of a crop. If the complexity in the way EOs act on insects justifies this rich literature, some more consistent results regarding situations out of laboratories, as well as more papers devoted to a better understanding of mechanisms of action, ought to be produced. In this context, we just intend to give some keys to understand the diversities of EO activities on insects in regard to the routes of exposure and their cellular targets. Insect biocontrol by EOs results from several kinds of modes of action and depends on the routes of the exposure. An activity observed in earlier studies was the toxicity by ingestion or by contact through cuticle or by inhalation for volatile compounds. Insects are very sensitive to topical applications of EOs; for example, Citrus spp. EOs on the maize weevil Sitophilus zeamaïs (Motschulsky) or the larger grain borer Prostephanus Americana (Horn) (Haubruge et al., 1989) and the bruchid Acanthoscelides obtectus (Say) to fumigant toxicity of a large range of Mediterranean

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TABLE 24.1 Examples of Harmful Pests Name Family Acanthoscelides obtectus Bean weevil Coleoptera: Bruchidae Acrolepiopsis assectella Leek moth Lepidoptera: Yponomeutidae Acyrthosiphon pisum Pea aphid Hemiptera: Aphididae Aedes aegypti Yellow fever mosquito Diptera: Culicidae Anopheles stephensi Diptera: Anophelinae Bemisia tabaci Sweetpotato white fly Homoptera: Aleyrodidae Boophilus microplus Ixodida, Ixodidae Callosobruchus chinensis Adzuki bean weevil Coleoptera: Bruchidae Callosobruchus maculatus Cowpea weevil Coleoptera: Bruchidae Cryptolestes pusillus Flat grain beetle Coleoptera: Laemophloeidae Culex pipiens House mosquito Diptera: Culicidae Culex quinquefasciatus Diptera: Culicidae Delia radicum Cabbage fly Diptera: Anthomyiidae Dendroctonus rufipennis Spruce beetle Coleoptera: Scolytidae Dendroctonus simplex Eastern larch beetle Coleoptera: Scolytidae Dermanyssus gallinae Roost mite Gamasida, Dermanyssoidea, Dermanyssidae Drosophila auraria

Notes

Source

Native to Africa, Europe, America; eats fruits of crops

(a)

Native to Asia, Europe; eats foliage of crops

(a)

Pest of legumes, e.g., alfalfa, clover and peas

(a)

Lives in tropics and subtropics; pathogenous

(a)

Vector of malaria disease

(b)

Polyphageous, pathogenous vector (i.e. cassava mosaic disease); lives in America, Africa, Asia

(a)

(a) Storage pest; i.e., in Syria

(a)

Damages cowpeas

(a)

Stored grain pest; in temperate climate

(a)

Pathogenous vector of diseases

(a)

Tropical regions; pathogenous vector of diseases

(a)

Native to Asia, Europe and North America; damages crucifers

(a)

Damages spruce trees

(c)

Damages spruce trees

(c)

Worldwide; poultry pest; can reduce egg production

(a)

Subgroup of D. melanogaster

(d) (Continued)

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TABLE 24.1 (Continued) Examples of Harmful Pests Name Family Ephestia kuehniella Mediterranean flour moth Lepidoptera: Pyralidae Helicoverpa armigera Cotton bollworm Lepidoptera: Noctuideae Hyalomma marginatum Acari: Ixodidae Hylobius abietis Large pine weevil Coleoptera: Curculionidae Lasioderma serricorne Cigarette beetle Coleoptera: Anobiidae Leptinotarsa decemlineata Colorado potato beetle Coleoptera: Chrysomelidae Limantria dispar Gypsy moth Lepidoptera: Lymantridae Listronotus oregonensis Carrot weevil Coleoptera: Curculionidae Lobesia botrana Grape berry moth Lepidoptera: Tortricidae Megastigmus pinus Hymenoptera: Torymidae Megastigmus rafini Hymenoptera: Torymidae Musca domestica Housefly Diptera: Muscidae Oryzaephilus surinamensis Sawtoothed grain beetle Coleoptera: Silvavidae Papilio demoleus Common lime butterfly Lepidoptera: Papilionidae Periplaneta americana American cockroach Blattaria: Blattidae Phytoseiulus persimilis Acarina: Phytoseiidae Pissodes strobi White pine weevil Coleoptera: Curculionidae

Notes

Source

Pest of industrial flour mills; lives in temperate climate

(a), (e)

Pest of maize and other crops; native to Asia and Africa

(a)

In Europe and Asia

(a)

Pest of conifers; in Europe and Asia

(a)

Pest of stored food, hoursehold pest; native to America, Asia, and Africa

(a)

Destroys solanaceous crops; in Europe, North America

(a)

Pest of cork oak forests

(f)

Damages mainly carrots

(a)

Pest of grapes; for example, in Jordan and Iran

(a)

Destroys white fir trees

(c)

Destroys white fir trees

(c)

Feeds on garbage and feces; worldwide

(a)

Stored-product pest

(a)

Southeast Asia; pest of citrus

(a)

Household pest; omnivore

(a)

Is used to control mite pests in greenhouses; predatory mite

(a)

Destroys pines, for example, Eastern White pine

(a), (c)

(Continued)

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TABLE 24.1 (Continued) Examples of Harmful Pests Name Family Plodia interpunctella Indian meal moth Lepidoptera: Pyralidae Prays citri (Citrus blossom moth) Lepidoptera: Hyponomentidae Prostephanus truncatus Larger grain borer Coleoptera: Bostrichidae Rhyzopertha dominica Lesser grain Borer Coleoptera: Bostrichidae Sitophilus granarius Granary weevil Coleoptera: Curculionidae Sitophilus granarius Granary weevil Coleoptera: Curculionidae Sitophilus oryzae Rice weevil Coleoptera: Curculionidae Sitophilus zeamays Maize weevil Coleoptera: Curculionidae Spodoptera frugiperda Fall armyworm Lepidoptera: Noctuideae Spodoptera litura Rice cutworm Lepidoptera: Noctuidae Stegobium paniceum Drugstore beetle Coleoptera: Anobiidae Stephanitis pyri Azalea lace bug Heteroptera: Tingidae Tenebrio molitor Yellow mealworm Coleoptera: Tenebrionidae Tetranychus cinnabarinus Carmine mite Acari: Tetranychidae Tetranychus urticae Two-spotted spider mite Acari: Tetranychidae

Notes

Source

Stored-grain pest; worldwide

(a)

For example, in Egypt

(a)

Pest of maize; especially in tropical regions

(a)

Native to America, Africa, Asia; pest of grain

(a)

Pest of stored grain

(a)

Pest of stored grain

(a)

Pest of stored grain

(a)

Pest of stored grain

(a)

Pest of sugarcane fields and other crops; North and South America

(a)

Native to Australia and Asia; damages legumes and solanaceous crops

(a)

Temperate and subtropical areas; eats pharmaceutical drugs

(a)

Native to Europe; damages trees

(a)

Stored-product pest

(a)

Damages soybeans, cotton, plums, citrus, vegetables; in Syria, Saudi Arabia, Lebanon, Jordan

(a)

Worldwide in temperate and subtropical areas; greenhouse pest

(a), (g)

(Continued)

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TABLE 24.1 (Continued) Examples of Harmful Pests Name Family Thaumetopoea pityocampa Pine processionary Lepidoptera: Thaumetopoeidae Triatoma infestans Kissing bug Hemiptera: Reduviidae: Triatominae Tribolium castaneum Red flour beetle Coleoptera: Tenebrionidae Tribolium confusum Confused flour beetle Coleoptera: Tenebrionidae Tyrophagus putrescentiae Mold mite Acaridida: Acaroidea: Acaridae Varroa destructor (Varroa jacobsoni) Acari: Varroidae

Notes

Source

Pest of Pinaceae

(a)

Pathogeous vectors of Chagas disease in South America

(a)

Native to America, Africa and Asia; pest of stored food

(a)

Stored food pest, especially flour; difficult to control

(a)

Eats processed cereals

(a)

Pest of honey bees

(a)

Source: (a) Capinera (2008), (b) Prajapazi et al. (2005), (c) Ibrahim et al. (2008), (d) Konstantopoulou et al. (1992), (e) Ayvaz et al. (2010), (f) Moretti et al. (2002), (g).

EOs (Regnault-Roger et al., 1993). The first observations were mainly focused on insects of the stored products, but EOs could control a large range of flying insects as well: the Mediterranean fruit fly Ceratitis capitata, the greenhouse white fly Trialeurodes vaporariorum, and also the green peach aphid Myzus persicae (Regnault-Roger, 1997). They also repel or act as deterrents or antifeedants that affect insect fitness (Regnault-Roger and Hamraoui, 1994). All these activities the EOs exert on an insect could occur on several physiological targets at the same or at different stages of the insect development. As examples, EOs of Artemisia vulgaris develop a combined activity as repellent and fumigant upon Tribolium castaneum (Wang et al., 2006). Beside these activities on adult, EOs also have an influence on the level of reproduction. EOs disturb oviposition or disrupt the larvae growth or modify the imago’s behavior or physiology of the insect. The beetle Acanthoscelides obtectus (Say) has been shown to be a convenient model to point out the different ways the EOs impact the fitness of adult insects and their reproduction, in particular, which reproductive stage (oviposition, eggs hatching, larvae) is targeted and the speed of the activity of EOs. The inhibition of reproduction and the toxicity on adults can be quite different. As examples, parsley (Petroselinum sativum L., Umbelliferae) did not have significant fumigant toxicity on the beetle adults but inhibited strongly its reproduction, whereas the summer savory (Satureia hortensis L., Lamiaceae) presented a high toxicity on adults but inhibited poorly the bruchid reproduction. Mint (Mentha piperita L. Lamiaceae) developed fumigant toxicity and very strong inhibition of reproduction but no antifeedant effect; dill (Anethum graveolens L., Umbelliferae) has little fumigant toxicity and inhibited poorly the reproduction but had a significant antifeedant effect (Regnault-Roger, 2013). The complexity of the large range of EO activities on insects is reinforced by a pronounced variability of physiological responses of insect species for the same EO. Tansy oil (Tanacetum vulgare L.) is attractive and paralyzing for Rhizoperta dominica, repulsive for Tribolium confusum, and toxic for Sitophilus americana (Koul et al., 1990).

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From all these observations, it could be deduced that EOs present a widespread range of activities on insects that necessitate to be sharpened on a case-by-case study before application in pest management. These biological effects on pests involve both physical and chemical mechanisms, as the following considerations underline: • Creating a physical barrier, a topical application of EOs laid a film on insect cuticle that modifies the physiology of the insect. Because oils are lipophilic, the film changes the conditions of penetration of the air and other substances inside the body of the insect and causes the disruption of lipid bilayers of cell walls. • Because of the volatility of monoterpenes, several families of proteins including OBPs (odorant binding proteins) and CSPs (chemosensory proteins) are involved in the detection of bouquets of fragrant and chemosensory-active compounds of EOs. OBPs and CSPs are found on the periphery of the sensory receptors and function in the capture and transport of molecular stimuli. In the sensilla of insects like trichoid sensilla of the female silkworm, Bombyx mori, specialized OBPs respond to the volatile monoterpene linalool (Picimbon and Regnault-Roger, 2008). In moths, the protein GOBP2, identified in tobacco hornworm, Manduca sexta, preferentially interacts with floral aromas and green plant odors such as geraniol, geranyl acetate, and limonene (Feng and Prestwich, 1997). These proteins play an important role in the response of the insect to an EO blend. • According to the toxicity of monoterpenes, several main compounds identified in EOs (thymol, α-terpineol, linalool, geraniol, eugenol) induce a neurotoxicity through different receptors. Thymol binds to GABA receptors associated with chloride channels located on the membrane of postsynaptic neurons and disrupts the functioning of GABA synapses (Priestley et al., 2003). Eugenol acts through the concentration of cAMP and the octopaminergic system. It mimics the action of octopamine with the consequence of increasing intracellular calcium levels in the cockroach Periplaneta americana. Tyramine (a precursor of octopamine) receptors are involved in the recognition of monoterpenes (Enan, 2005). Thymol, carvacrol, and α-terpineol influence the production of cellular cAMP and calcium in D. melanogaster (Kostyukovsky et al., 2002). Another mechanism of neurotoxicity was identified through the transmission of the nervous impulse. Electrophysiological experiments showed that eugenol inhibits deeply neuronal activity, whereas citral and geraniol have a biphasic effect that is dose dependent. At low doses, these compounds induce an increase in spontaneous electrical activity but, at high doses, cause a decrease (Price and Berry, 2006). Using a similar electrophysiological experimentation, Huignard et al. (2008) observed that the Ocimum basilicum EO has a complex neurotoxic activity. The EO inhibited fully the neuronal electrical activity by decreasing the magnitude of nerve-action current, then reducing the post-hyperpolarization phase and firing the frequency of nerve-action current. The authors hypothesized that this effect could be the result of the combined action of linalool and estragole, two major components of the O. basilicum EO. They demonstrated that the mere application of pure linalool produced a reduction in the amplitude of nerve-action current and decreased the posthyperpolarization while estragole specifically induced a reduction of post-hyperpolarization. Another well-known target for neurotoxicity is the enzyme acetylcholinesterase (AChE). Tea tree oil inhibited acetylcholinesterase (Mills et al., 2004). Several mechanisms involving monoterpenes were identified through a reversible competitive inhibition at the enzyme’s hydrophobic active site or a mixed inhibition for this enzyme by linking to a different site from the active site where the substrate was bound (Lopez and Pascual-Villalobos, 2010). However, a systematic study of bioactivity of plant EOs and their AChE inhibitory activity in vitro led to the conclusion “that AChE is not a principal target site for insecticidal monoterpenoids naturally occurring in plant EOs, nor is inhibition of AChE an important mode of action for these substances in insects” (Isman and Tak, 2017b).

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These studies confirm that the insecticidal activity of EOs is the consequence of several mechanisms that affect multiple cellular and physiological targets.

24.2.3 Development and Commercialization of Botanical The utilization of insecticides, both botanicals and synthetics, are handled differently in various countries (Isman, 2006): In the US, many EOs are registered for agricultural use. Canada has a slightly different law as there are not as many EOs allowed. Mexico has a similar utilization of EOs as the US, but there is no exception of EOs. The European Union allows the utilization of components of EOs. Even though, the individual law differs for countries in the European Union. In India, more biological pesticides are used than in other countries. In South America, pesticide use varies in every region. In Africa there are no real restrictions by law for its local use of various pesticides. In general, countries tend to make laws for use of pesticides stricter than they were, because the majority of the population and media are aware of the environmental and health impacts of pesticides. This is especially the case in wealthier countries; but since developing countries are dependent on selling their products to so-called “first-world countries,” they also have to meet up with stricter criteria, even though it might not make sense for them (Isman, 2006). According to Isman, three aspects are important for successful commercialization of pesticides: sustainability of the botanical resource, standardization of the chemically complex extracts, and regulatory approval. Sustainability means that the base of the product must be available all the time and not only seasonally. Standardization means that there must be a method to investigate analytically the product so that the product obtained from plants will not vary. It is a problem if the pesticidal action of a substance differs even though it has been cultivated under the same circumstances. Regulatory control is—among those—the most difficult aspect. The profit made by selling botanicals will not be very high in industrialized countries. That is why “green pesticides” might not even reach the market. In industrialized countries, it is hard to imagine that botanicals such as EOs will play a bigger role in the future since synthetic pesticides are affordable and very potent against various insects (Isman, 2006). While the global market for synthetic and biopesticides pooled together should reach USD 79.3 billion by 2022 from USD 61.2 billion in 2017 at a compound annual growth rate (CAGR) of 5.3% during 2017–2022 (https://www.bccresearch.com), in 2016, the global biopesticides market was valued at USD 2.83 billion and, in 2017, at USD 3.3 billion with a projection of USD 9.5 billion by 2025, which means a CAGR of 13.9% during 2017–2022. North America, with USD 1300.25 million is the leading region followed by Asia, Europe, Latin America, Middle East, and Africa (TRM, 2018). Botanicals (including EOs) value for 10% of the biopesticides market. This market is dynamic and full of opportunities. However, to paraphrase Isman (2008) speaking of botanical insecticides, the EO market has been considered for two categories of countries: for richer and for poorer. For the richer, in developed countries, EOs are used for high-value crops (orchard, greenhouse vegetable production). For example, in California, 37 EOs have been registered as active ingredients. Among them, clove oil against post-harvest potato sprouting, tagetes, and wintergreen oil are newly registered as agricultural insecticides, and many crude EOs as repellent (eucalyptus, cedarwood, citronella, lemon grass, peppermint, rosemary) (Grieneisen and Isman, 2018). Arnason (2011) indicated that about 90 insect-repellent products sold in the US market contain an EO as one of the active ingredients in the formulation. The most commonly used ingredient is citronella oil (45 products), followed by geranium oil (geraniol) (33), lemongrass oil (24), cedar oil (22), peppermint oil (16), rosemary oil (15), soybean oil (15), and eucalyptus oil (14). EOs are now considered to be the most important commercial application of botanical insecticides in North America. The enhancement of using EOs to control insect pests in orchards and to protect high value crops is probably the result of the regulation rules in the US. Biopesticides are subject to special procedures outlined in Title 40, Code of Federal Regulations, of FIFRA. A number of natural substances, such as EOs of mint, thyme, rosemary, and lemon grass that did not benefit from this simplified procedure, however, were classified as GRAS (generally regarded as safe). They were placed on a list

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(FIFRA Section 25[b]) exempting them from the registration process (EPA, 2018). This exemption has become a marketing strategy to promote these products (Isman, 2004; Regnault-Roger et al., 2005). The Company EcoSMARTTM was a pioneer, 15 years ago, to develop a line of products on this exemption. These products are named “Ecoexempt® Minimum-Risk & EcoPCO® Products” and are commercialized on a large scale by several suppliers inside the US and abroad. Nowadays, these products are currently approved for use in seven other countries, including Mexico, Peru, Uruguay, Singapore, and the United Arab Emirates (Isman and Tak, 2017a). In the European Union, the procedure for reevaluation of plant protection products (PPPs) ended in 2008. To be authorized on market, all PPPs derived from biological as well as chemical sources have to be listed. Because they meet the purposes of Directive 2009/128/CE to promote Integrated Pest Management (IPM) obligatory in all European Union member states from the beginning of 2014, several plants oils have been authorized for plant protection on a special procedure. French Ministry of Agriculture listed natural products for biocontrol, among them orange and mint EOs, but also eugenol, geraniol, and thymol (DGAL, 2017). Mint oil is also currently used as a PPP in Belgium, Germany, and Italy, and fennel oil is used in organic farming in Czech Republic and Slovakia, and gillyflower oil in Italy (Szulc and Sobczak, 2017). Nevertheless, Isman and Tak (2017a) underlined that very few PPPs based on EOs are available in the European Union (EU) compared to the US. French Organic Farming Technical Institute (ITAB), looking for new solutions to expand the range of organic pesticides, indicated only two commercial formulations based on orange oil and mint EOs as insecticide and fungicide for legumes, fruits, and vines for the first one and potato sprout inhibition for the second. Some experimental research in the field was conducted between 2014 and 2017 in France to test the efficacy of seven EOs against mildew of potatoes, vines and salads, and apple scab. The EOs and their main compounds were eucalyptus (citronellal), clove (eugenol), mint (D-carvone and L-carvone), oregano (carvacrol), tea tree (terpinen-4-ol), and thyme (thymol). If these EOs showed some efficacy in vitro on the two fungi implicated in mildew and apple scab diseases, Phytophthora infestans and Venturia inaequalis, results were very poor in the field trials; these EOs biocontrol abilities were less effective than synthesized fungicides and also copper hydroxide. Moreover, the beneficial mites Typhlodromus spp. were impacted by these EOs (Vidal et al., 2018). This very interesting experiment showed that there is a gap between promising results in the laboratory and the real ones in the field. It also could explain the limited success for registration of new PPP substances based on EOs in the EU. In fact, to be registered as a PPP in the EU, contrary to US registration, the active substance has to be proved to have a real efficacy on the target species. In other developed or emerging countries like India, Brazil, or Australia, where an abundant plant biodiversity grows and where a “prolific research on botanicals” is developed, Isman and Tak (2017a) also noted the lack of approved active substances based on EOs. For poorer countries, the situation for using EOs is quite different. The tropical climate develops numerous species with high potentiality for pest management (Arnason et al., 2008). Aromatic plants are traditionally and widely used for stored-product insect biocontrol or to repel harmful insects in fields (Glitho et al., 2008). Currently, there is a move to enhance the use of steam-distilled EOs, but the lack of technologic resources leads to use of crafty solutions like a domestic pressure cooker for extracting EOs by steam distillation. Another point that impedes the proper development of EOs in Africa is that it is unsupported by scientific experimentation (Regnault-Roger, 2013). Isman (2017), analyzing the gap between the abundance of tropical pesticide plants and the lack of practical solutions for farmers in these countries, argued to develop simple methods for local cultivation of a relevant plant species, simple methods for extraction of active substances at minimal cost, and simple methods to do bioassays and then field trials, thus, to demonstrate and to learn strategies to optimize efficacy. He concluded that EOs are “well suited for exploitation by smallholder farmers in sub-Saharan Africa, whether prepared on an individual (‘do-it-yourself’) basis.” According to him, the use of low-cost botanical insecticides with consistent efficacy would enhance local productions. Appropriate regulations and relevant public policies for sustainable agriculture are also needed in these countries.

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24.2.3.1  Examples of Essential Oils Used as Antipests 24.2.3.1.1  Rosmarinus officinalis Rosmarinus officinalis L. (Lamiaceae) can be found in the Mediterranean area (Tables 24.2–24.4). The used parts are the fresh flowering tops. The EO can be used as an antipest due to its effect on beetles, caterpillar larvae, and many other insects (Dayan et al., 2009) like Drosophila auraria (Konstantopoulou et  al., 1992), Sitophilus oryzae (Lee et  al., 2004), or Rhyzopertha dominica (Shaaya et al., 1991). The microencapsulated EO also showed larvicidal effects on Limantria dispar (Moretti et al., 2002). In Acanthoscelides obtectus males and females, R. officinalis volatile oil also caused high mortality rates (Papachristos et al., 2002). Repellent activity was reported against Listronotus oregonensis (Niepel, 2000).

TABLE 24.2 Essential Oils that can be Used as Insecticide and Acaricide Name Family

Constituents

Acorus sp. Acoraceae

β-asarone, acorenone, acoragermacrone

Artemisia sp. Asteraceae

Limonene, myrcene, α-thujone, β-thujone, caryophellene, sabinylacetate

Baccharis salicifolia (Ruiz & Parvon) Pers Asteraceae, Garcia et al. (2005)

β-Pinene, pulegone, camphene, limonene, α-pinene, pulegone, pulegol, germacrol, germacrone

Carum sp. Apiaceae

D-Carvone, limonene

Chenopodium ambrosioides L. Amaranthaceae

α-Terpinene, cymol, cis-β-farnesene, acaridole, carvacrol

Cinnamomum sp. Lauraceae

Cinnamaldehyde, linalol, β-caryophyllene, eugenol

Citrus sinensis (L.) Osbeck Rutaceae, Loebe (2001), Njoroge et al. (2005)

Limonene, α-pinene, sabinene, α-terpinene

Coriandrum sativum L. Apiaceae

Linalol, α-terpinyl acetate, 1,8-cineole, linalyacetate, geranyl acetate, camphor

Cuminum cyminum L. Umbelliferae, Li and Jiang (2004)

Cuminal, cuminic alcohol, γ-terpinene, safranal, p-cymene, β-pinene

Cymbopogon sp. Poaceae

Citronellal, geraniol, citronellol, citral, limonene

Elettaria cardamomum (L.) Maton Zingiberaceae

α-Terpinyl acetate, 1,8-cineole, linalyl acetate

Eucalyptus sp. Myrtaceae

1,8-Cineole, limonene, α-pinene

Evodia rutaecarpa Hook f. et Thomas Rutaceae, Liu and Ho (1999)

Limonene, β-elemene, linalool, myrcene, valencene, linalyl acetate, β-caryophyllene

Foeniculum vulgare L. Apiaceae

Anethol, fenchone, estragol (Continued)

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TABLE 24.2 (Continued) Essential Oils that can be Used as Insecticide and Acaricide Name Family

Constituents

Hedomea mandonianum Wedd. Lamiaceae

Eucalyptol, pulegone, menthone

Juniperus sp. Cupressaceae

α-Pinene, myrcene, β-pinene, terpinen-4-ol, germacrene D

Lavandula sp. Lamiaceae

Linalol, linalylacetate, fenchone, camphor

Leptospermum scoparium J.R. et G. Forst. Myrtaceae, Douglas et al. (2004)

Trans-calamenene, δ-cadinene, β-caryophyllene, leptospermone

Lippia sp. Verbenaceae

Piperitenone oxide, limonene, thymol, carvacrol, p-cymene, β-caryophyllene and γ-terpinene

Melaleuca sp. Myrtaceae

Terpinene-4-ol, γ-terpinene, α-terpinene

Mentha sp. Lamiaceae

Carvone, pulegone, menthol, menthone, menthylacetate

Micromeria fruticosa L. Lamiaceae

Pulegone, β-caryophyllene, isomenthol, limonene

Minthostachys andina (Britton) Eppling Lamiaceae

Menthone, pulegone, iso-menthone

Myristica fragrans Houtt. Myristicaceae

Sabinene, α-pinene, β-pinene, myristicine

Myrtus communis Myrtaceae

1,8-Cineole, α-pinene, limonene, linalol, myrtenol

Nepeta racemosa L. Lamiaceae

Nepetalactone, myrtenol, terpinen-4-ol

Ocimum sp. Lamiaceae

γ-Terpinene, α-terpinene, thymol, carvacrol, linalool, eugenol

Origanum sp. Lamiaceae

Carvacrol, thymol, γ-terpinene, p-cymene, linalool

Pelargonium graveolens L’Hér Geraniaceae

Geraniol, citronellol

Pimpinella anisum L. Apiaceae

cis-Anethole

Piper nigrum L. Piperaceae

β-Caryophyllene, α-pinene, sabinene, limonene

Rosmarinus officinalis L. Lamiaceae

Limonene, cineole, borneol, terpineol

Salvia sp. Lamiaceae

Cineole, thujone, camphor, α-pinene, β-pinene (Continued)

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TABLE 24.2 (Continued) Essential Oils that can be Used as Insecticide and Acaricide Name Family

Constituents

Satureja sp. Lamiaceae

Carvacrol, thymol, γ-terpinene, p-cymene

Syzygium aromaticum (L.) Merr. Et L.M. Perry Myrtaceae

Eugenol, aceteugenol, β-caryophyllene

Syzygium aromaticum (L.) Merr. Et L.M. Perry Myrtaceae Tagetes sp. Asteraceae

Eugenol, aceteugenol, β-caryophyllene Limonene, (Z)-β-ocimene, (Z)- and (E)-ocimenone

 

Teuscher et al. (2004)

Toxic to V. destructor, no toxicity toward honey bees; causes repellence to S. zeamais

Eguaras et al. (2005); Nerio et al. (2009)

Tanacetum vulgare L. Asteraceae

α-Pinene, α-terpinene, γ-terpinene, carvone, borneol, β-thujone, camphor Carvacrol, thymol, γ-terpinene, p-cymene Thymol, carvacrol, p-cymene

Antifeedant against L. decemlineata and acaricide toward T. urticae

Chiasson et al. (2001)

High mortality rates in D. gallinae and T. cinnabarinus; Insecticidal activity on D. auraria  

Konstantopoulou et al. (1992); Ghrabi-Gammar et al. (2009); Sertkaya et al. (2010)

Thymbra sp. Lamiaceae

Thymus sp. Lamiaceae

Teuscher et al. (2004)

24.2.3.1.2 Thymus sp. Thymus vulgaris L. (Lamiaceae) also grows in the Mediterranean area. Parts used are partial dried or fresh aerial parts. The main components are “available for broad-spectrum insect control in organic farming” (Dayan et al., 2009). The EO was tested against Bemisia tabaci and caused its death (Aslan et al., 2004). T. vulgaris EO also showed toxicity against Dermanyssus gallinae, a pathogenic mite on hens (George et al., 2009a, 2010; Ghrabi-Gammar et al., 2009). It also caused high mortality in Tenebrio molitor (George et al., 2009b) and Tyrophagus putrescentiae (Kim et al., 2003a,b). The EO of Thymus herba-barona Loisel was microencapsulated and tested on Limantria dispar, on which it caused mortality (Moretti et al., 2002). 24.2.3.1.3  Syzygium aromaticum Syzygium aromaticum (L.) Merr et L. M. Perry (Myrtaceae) (Teuscher et al., 2004) grows in tropical areas. Parts used are flower buds, which have been dried. Its field of application is insect pest management as eugenol is toxic to many common pests (Dayan et  al., 2009). In another study conducted by Huang et al. (2002), toxicity of eugenol to Sitophilus zeamais and Tribolium castaneum could be shown. Acaricidal activity against Tyrophagus putrescentiae (Kim et al., 2003a,b) and Dermanyssus gallinae (Kim et al., 2007) could also be confirmed for clove EO.

Pine

Pine

Hylobius abietis (Large pine weevil)

 

Citrus limonum, C. decuminata, C. aurantium  

Prays citri (Citrus blossom moth)

Papilio demoleus (Common lime butterfly)

Polyphageous moth (e.g., cotton, tomatoes, conifers, etc) Fabaceae

White fir

Megastigmus pinus, M. rafini (Silver fir seed wasp) Thaumatopoea pityocampa (Pine processionary)

Helicoverpa armigera (Cotton bollworm)

Pine, Spruce

Hylobius abietis (Large pine weevil)

 

Emulsified with water and sprayed on the foliage of pine seedlings

Spruce spp

Dendroctonus rufipennis, D. simplex (Spruce beetle, eastern larch beetle)

 

Olfactory responses to pure α-pinene and limonene

Brassicaceae

Delia radicum (Cabbage fly)

Exposure to limonene vapors

Electroantennogram response to pure limonene No effect

Response

(+)-Limonene reduces the number of egg clusters on plants sprayed with it

(+)-Limonene at 60 ppm kills 100% of the pests after 24 hours High limonene concentrations show signs of poisoning within a few hours Limonene significally acts as repellent

Limonene from the surface part of plant host acts as a repellent

Limonene shows repellent activity

Low limonene levels does not affect feeding activity

(−)-Limonene shows Maximum attraction to the larvae Limonene activates oviposition

Attractive to 1–2 days old moths

Increased Activity of Pest Insect Electroantennography used to investigate electrophysiological responses Orientation responses to different odors in Olfactory

Exposure to limonene vapors

Bioassayed for their toxicity

Olfactometer with two parallel air currents containing a Y-shaped nylon fibre Olfactory stimuli for orientation behavior

Leek & onion, all Allium crops

Acrolepiopsis assectella (Leek moth)

Reduced Activity of Pest Insect

Limonene Application

 

Host Plant

 

Insect Species

TABLE 24.3 Effect of Limonene Application on Various Pests (Ibrahim et al., 2008)

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TABLE 24.4 Effect of Eucalyptus Essential Oil on Various Pests (Batish et al., 2008) Eucalyptus sp. E. camaldulensis (River red gum E.) E. citriodora (Lemon scented E.) E. globulus (Tasmanian blue gum E.) E. intertexta E. Sargentii E. camaldulensis E. saligna E. tereticornis Eucalyptus sp.

Tested Organism Repels adult females of Culex pipiens (common house mosquito); mortality of eggs in Tribolium confusum amd Ephestia kuehniella Toxicity against Sitophilus zeamais (greater rice weevil) Toxic to pupae of Musca domestica (housefly) as well as to female Pediculus humanus (louse) Causes death of adult Callosobruchus maculatus, Sitophilus oryzae and Tribolium castaneum Repelled Sitophilus zeamais and Tribolium confusum (confused flour beetle) Pesticidal activity against Anopheles stephensi Toxic to Thaumetopoea pityocampa (pine processionary)

24.2.3.1.4 Muña Muña are Bolivian medical plants and include species of Satureja, Minthostachys, Mentha, and Hedomea. They derive their name muña by the Kechuas people, who live in the Andean mountains where these plants grow at an altitude between 2500 and 5000 meters. These indigenous people of the Andes use these plants among others because of their insecticide and repellent activity in order to protect the crops of potatoes against insects and also to prevent Chagas disease of which an insect acts as the pathogenous vector. Plants used in the study against Triatoma infestans, which was conducted by Fournet et al. (1996), were Minthostachys andina (Britton) Eppling (Lamiaceae) and Hedomea mandonianum Wedd. (Lamiaceae). The EO of M. andina could significantly decrease the number of insects: after 28 days, 10 out of 20 insects were found dead. H. mandonianum had no such effect (Fournet et al., 1996). 24.2.3.1.5 Eucalyptus sp. The EO of Eucalyptus is obtained from the Australian tree Eucalyptus sp. that belongs to the Myrtaceae family. It has been known for its antibacterial, antifungal, and antiseptic action for hundreds of years (Batish et al., 2008). Toxicity of eucalyptus EO toward Sitophilus oryzae was reported by Lee et al. (2001), and it proves to be a promising fumigant to control that pest. Lee et al. (2004) reported in another study that several Eucalyptus species were toxic to S. oryzae, namely E. nicholii, E. codonocarpa, and E. blakelyi. The same species cause mortality in Tribolium castaneum and Rhyzopertha dominica. 24.2.3.1.6 Satureja sp. Aslan et al. tested the EO of Satureja hortensis L. (Lamiaceae) against Bemisia tabaci. Of the three oils (also, T. vulgaris and O. basilicum) tested, S. hortensis EO was most toxic to it (Aslan et al., 2004). S. thymbra L. (Lamiaceae) was tested against Hyalomma marginatum and was able to kill almost all of the ticks after 24 hours (Cetin et al., 2010). It also showed toxicity to Drosophila auraria (Konstantopoulou et al., 1992). According to Ayvaz et al., savory EO caused mortality against the pests Ephestia kuehniella and Plodia interpunctella (Ayvaz et al., 2010). 24.2.3.1.7 Ocimum sp. Basil (Ocimum basilicum L., Lamiaceae) EO was tested against B. tabaci and proved to be effective against it (Aslan et  al., 2004). Keita et  al. (2001) tested the fumigant and contact toxicity of O. basilicum and O. gratissimum L. on Callosobruchus maculatus beetles. Both EOs showed a high toxicity toward the beetles compared to the control. Also, the egg hatch rate

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decreased. Another study reports methyleugenol, methylcinnamate, linalool, and estragol as main compounds. Estragole was identified as one of the effective compounds against Cryptolestes pusillus and R. dominica, while it was variable against Sitophilus oryzae (López et al., 2008). Basil EO showed also very promising results in Oryzaephilus surinamensis (Shaaya et  al., 1991). O. kilimandscharicum Wild. EO, with camphor as the major compound, was proved to be toxic to four pests, namely, Sitophilus granarius, Sitophilus zeamais, Tribolium castaneum, and Prostephanus truncatus, while it also caused repellency in them. Compared with the other three, T. castaneum is the strongest among them in tolerating fumigation, as it has the lowest mortality rate (Obeng-Ofori et al., 1998). 24.2.3.1.8 Origanum sp. Calmasur et  al. tested the pesticidal effect of Origanum vulgare L. (Lamiaceae) EO on two pests, namely Tetranychus urticae and Bemisia tabaci. They showed that after 120 hours and at a concentration of 2 µL/L air, the mortality rate on B. tabaci was 100% and on Tetranychus urticae, it was 95% (Çalmaşur et al., 2006). Oregano EO can also be used as an acaricide against Tyrophagus putrescentiae (Kim et al., 2003a,b) and as insecticide against Oryzaephilus surinamensis (Shaaya et  al., 1991) and Drosophila auraria. O. dictamnus L. and O. majorana L. were also effective against D. auraria (Konstantopoulou et al., 1992). O. acutidens EO can be used against Sitophilus granarius and Tribolium confusum (Kordali et al., 2008), and the oil of O. onites L. (with carvacrol, linalool, and thymol as main constituents) is successful in controlling T. cinnabarinus, as reported by Sertkaya et al. (2010). Oregano EO showed high activity against Plodia interpunctella and Ephestia kuehniella (Ayvaz et al., 2010). 24.2.3.1.9 Artemisia sp. The main constituents of Artemisia absinthium L. (Asteraceae) EO are said to be effective against several fleas, flies, and mosquitoes. In a study conducted by Chiasson et al. (2001), A. absinthium EO showed toxicity to T. urticae and T. putrescentiae (Kim et al., 2003a,b). When used as fumigant, A. sieberi Besser EO caused mortality in Callosobruchus maculatus, Sitophilus oryzae, and Tribolium castaneum. Of all three insects, it was most toxic to C. maculatus, as it caused 100% mortality at a concentration of 37 µL/L after 12 h (Negahban et al., 2007). In another study, this author group tested the effect of A. scoparia Waldst et Kit EO on the same three stored-product pests as mentioned above. At a concentration of 37 µL/L 100% mortality of the pests can be observed after one day. It also causes repellent effects on all three insects (Negahban et al., 2006). 24.2.3.1.10 Mentha sp. Mentha pulegium L. (Lamiaceae) EO was tested on Dermanyssus gallinae, it showed high toxicity as fumigant against the mite (George et  al., 2009a). The same EO proved to be active as an acaricide against Tyrophagus putrescentiae. The same effect can be observed with M. spicata L. (Kim et al., 2003a,b). M. pulegium L. and M. spicata were very effective against Drosophila auraria (Konstantopoulou et  al., 1992). Mentha arvensis var. piperascens also showed high mortality rates on Dermanyssus gallinae (Kim et al., 2007). Mentha microphylla and M. viridis proved to be toxic to Acanthoscelides obtectus, and they also decreased the number of eggs (Papachristos et al., 2002). 24.2.3.1.11 Cinnamomum sp. Cinnamomum aromaticum Nees, Lauraceae proved to be a contact insecticide to Tribolium castaneum and Sitophilus zeamais (Huang et al., 1998). C. cassia and C. sieboldii proved to have pesticidal action against Sitophilus oryzae and Callosobruchus chinensis (Kim et al., 2003a,b). Cinnamaldehyde causes acaricidal effects on Tyrophagus putrescentiae, especially in closed containers (Kim et al., 2004). C. camphora (L.) J. Presl causes mortality to Dermanyssus gallinae and can be used as an acaricide against them (Kim et al., 2007).

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891

24.2.3.1.12 Acorus sp. Acorus calamus var. angustatus L. and Acorus gramineus Sol. showed 100% mortality at concentrations of 3.5 mg/cm3 after three days treatment with Sitophilus oryzae. EO obtained from A. calamus var. angustatus was also very potent against Callosobruchus chinensis. A mortality rate of 100% after one day could be realized at a concentration of 3.5 mg/cm3 (Kim et al., 2003a,b). Another pest that can be controlled by A. calamus var. angustatus is Dermanyssus gallinae (Kim et al., 2007). A. gramineus EO shows insecticidal action against Sitophilus oryzae and Callosobruchus chinensis when directly applied. It can also be used against Lasioderma serricorne, but it is more potent on the first two insects. As fumigant, it worked best in closed containers (Park et al., 2003). 24.2.3.1.13  Foeniculum vulgare Fennel EO caused 100% toxicity to Sitophilus oryzae after exposition for 3 days at concentrations of 3.5 mg/cm3. The EO was even more potent against Callosobruchus chinensis because 100% mortality can be realized at the same concentration after one day (Kim et al., 2003a,b). Compounds of fennel EO were also toxic, as proved in the previous study, to S. oryzae and C. chinensis and also to Lasioderma serricorne. At concentrations of 0.168 mg/cm2, estragole was most toxic to S. oryzae after one day, followed by (+)-fenchone and (E)-anethol. Against C. chinensis, (E)-anethole was most effective, followed by estragol and (+)-fenchone. After one day, (E)-anethole was lethal to L. serricorne, whereas estragole and (+)-fenchone showed lower toxicity (Kim et al., 2001). F. vulgare EO is also potent in controlling Dermanyssus gallinae (Kim et al., 2007). Fennel EO also showed toxic effects to Tyrophagus putrescentiae (Lee et al., 2006), which can possibly be attributed to fenchone, as the isolated compound also showed high mortality rates (Sánchez-Ramos and Castañera , 2001). 24.2.3.1.14 Lavandula sp. Lavandula stoechas L. (Lamiaceae) EO is effective in killing Drosophila auraria in a study conducted by Konstantopoulou et  al. In the same study, the insecticidal effect of Lavandula angustifolia Miller (Lamiaceae), with linalool and linalylacetate as main compounds, could be confirmed as well (Konstantopoulou et al., 1992). Lavandula hybrida volatile oil caused fumigant toxicity to Acanthoscelides obtectus males and females and a decreased number of eggs (Papachristos et al., 2002). Lavender EO also showed high activity against Rhyzopertha dominica (Shaaya et al., 1991). 24.2.3.1.15 Carum sp. Carum carvi L. (Apiaceae) EO is able to kill Sitophilus oryzae; carvone seems to be the active compound against it. (E)-anethole is proved to be effective against Rhyzopertha dominica. Limonene and fenchone, for example were active against Cryptolestes pusillus (López et al., 2008). C. copticum C. B. Clarke volatile oil constituents are thymol, α-terpineol and p-cymene. Especially S. oryzae was weak against the fumigant action of the EO, but also mortality on Tribolium castaneum can be observed (Sahaf et al., 2007). 24.2.3.1.16  Chenopodium ambrosioides At a concentration of 0.4%, the EO of Chenopodium ambrosioides was able to kill Callosobruchus chinensis, Callosobruchus maculatus, and Acanthoscelides obtectus at a high percentage. Sitophilus granarius and Sitophilus zeamais were totally killed at 6.4%, Prostephanus truncatus was the least sensitive, as 56% survived (Tapondjou et al., 2002). The use of EOs as an antipest is presented in this chapter. It could be shown that several EOs, like Rosmarinus officinalis, Thymus vulgaris, Eucalyptus species, and Origanum species show great potential in controlling several pests—for example, Sitophilus sp. or Tetranychus sp., among many others—both on stored products and in protection of livestock, bees, and crops. Since synthetical pesticides bear a lot of risk toward environmental and mammalian health there is a need for natural and healthier alternatives, just like EOs. Of course, the high volatility is a limiting factor in the use of volatile oils on the field, but in glass houses or on stored food, they bear a great chance to change the current situation.

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24.3  ESSENTIAL OILS AS HERBICIDES Weed control in agriculture is very important as a high percentage of weed causes a high crop reduction, even more than insect pests and pathogens. While in ancient times, people were dependent on removing weeds by hand like several famous painters (Emile Klaus, Vincent Van Gogh, or Jules Breton) illustrated, today’s weed control is managed by synthetic and natural substances but also by machine weeding. In order to diversify the approaches to control weeds, natural substances such as EOs have to be considered (Dayan et al., 2009).

24.3.1  Phytotoxicity Phytotoxicity means the harmful impact of chemical substances on plants which can be seen in many different ways: their parts can appear to be burned; they can suffer from chlorosis, which colors the leaves yellow; or they can also result in a lack of growth or even an excessive growth. When phytotoxicity of various components of EOs was tested, pulegone was least, and D-carvone most, phytotoxic to maize plants. (+)-Limonene appeared most toxic to sugar beet seedlings. Limonene showed toxicity toward strawberry seedlings in preliminary screenings, as well as toward cabbage and carrot seedlings (Ibrahim et al., 2008; de Almeida et al., 2010) found that EOs of balm, caraway, hyssop, thyme, and vervain showed a 100% inhibition of germination of Lepidium sativum (Brassicaceae). Raphanus sativus’ (Brassicaceae) germination was inhibited by 100% by vervain oil at all concentrations. At low concentrations, anise and basil EOs promoted the germination and radicle growth, while caraway, sage, vervain, and marjoram EOs inhibited the growth of radish. Lactuca sativa’s (Asteraceae) growth was inhibited by thyme EO. Vervain, balm, and caraway also had the ability to suppress the growth of lettuce. Generally said, a high level of oxygenated monoterpenes is most harmful toward weeds. When it comes to inhibit the seed germination and subsequent growth, ketones and alcohol showed the highest activity followed by aldehydes and phenols (de Almeida et al., 2010). Especially plants from the Lamiaceae family can inhibit the growth of several weeds by releasing phytotoxic monoterpenes, namely α-pinene, β-pinene, camphene, limonene, α-phellandrene, p-cymene, 1,8-cineole, borneol, pulegone, and camphor (Angelini et al., 2003). The herbicide effect of 1,4-cineole and 1,8-cineole is also described by Dayan et al. (2012). Plants that are exposed to EOs often metabolize them, and when citral was added, geraniol, nerol, and their acids appeared. When citronellal metabolization was tested, citronellol and citronellic acid were formed, with pulegone (iso)-menthone, isopulegol, and menthofuran found (Dudai et al., (2000). It was also reported that EOs lead to accumulation of H2O2 in other plants. That way they increase oxidative stress, which leads to “disruption of metabolic activities in the cell,” (Mutlu et al 2010) Another cause for phytotoxicity is that EO destroys the cell membrane of plants and inhibits their enzymes (Mutlu et al., 2010) (Table 24.5).

24.3.2  Prospects of Organic Weed Control The problem with organic weed control is the huge quantities that must be applied to harm the weeds. This may also cause a large negative impact on the environment and the microbes in the soil. Also, the volatility is a problem of EOs, on account of which they also have to be applied very often, which could be avoided by using microencapsulation of EOs, because it will decrease their volatility. The third problem is that EOs can also harm the desired crop and lead to part destruction of the culture. Altogether, organic weed control using EOs does not seem too promising as a future replacement of synthetic products (Dayan et al., 2009).

24.3.3  Examples of Essential Oils in Weed Control 24.3.3.1  Thymus vulgaris Thymus vulgaris L. belongs to the Lamiaceae family as well (Angelini et al., 2003; de Almeida et al., 2010) (Table 24.6). The latter author group observed that thyme EO can inhibit the growth of the

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TABLE 24.5 Examples of Tested Plants Name

Common Name

Achyranthes aspera Ageratum conyzoides Agrostis stolonifera Alcea pallida Amaranthus hybridus Amaranthus retroflexus Amaranthus viridis Ambrosia artemisiifolia Arabidopsis thaliana Bromus danthoniae Bromus intermedius Cassia occidentalis Centaurea solstitialis Chenopodium album Cirsium arvense Cynodon dactylon Cyperus rotundus Echinochloa crus-galli Hordeum spontaneum Lactuca sativa Lactuca serriola Lepidium sativum Lolium multiflorum Lolium rigidum Parthenium hysterophorus Phalaris canariensis Phalaris minor Portulaca oleracea Ranunculus repens Raphanus raphanistrum Raphanus sativus Rumex crispus Rumex nepalensis Rumex obtusifolius Secale cereale Senecio jacobaea Sinapis arvensis Sonchus oleraceus Sorghum halepense Taraxacum sp. Trifolium campestre Urtica dioica Source:

Family

Source

Prickly chaff flower Billygoat weed Creeping bentgrass Hollyhock Smooth amaranth Red-root amaranth Slender amaranth Common ragweed Thale cress

Amaranthaceae Asteraceae Poaceae Malvaceae Amaranthaceae Amaranthaceae Amaranthaceae Asteraceae Brassicaceae Poaceae Poaceae

Commmon lambsquarters Creeping thistle Bermuda grass Purple nut sedge Common barnyard grass Wild barley Lettuce Prickly lettuce Garden cress Annual ryegrass Ryegrass Whitetop weed Canarygrass Littleseed canarygrass Common purslane Creeping buttercup Wild radish Radish Curled dock

Amaranthaceae Asteraceae Poaceae Cyperaceae Poaceae Poaceae Asteraceae Asteraceae Brassicaceae Poaceae Poaceae Asteraceae Poaceae Poaceae Portulacaceae Ranunculaceae Brassicaceae Brassicaceae Polygonaceae Polygonaceae Polygonaceae Poaceae Asteraceae Brassicaceae Asteraceae Poaceae Asteraceae Fabaceae Urticaceae

(a) (a) (a) (a) (a) (a) (a) (a), (b) (a) (a) (a) (a) (a) (a), (b) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a) (a), (c) (a) (a) (a), (b) (a) (a), Teuscher et al. (2004) Teuscher et al. (2004)

Broad-leaved dock Rye Common ragwort Wild mustard Common sowthistle Johnsongrass Dandelion Hop trefoil Common nettle

(a) Plant Encyclopedia (2012), (b) Tworkoski (2002), (c) Clay et al. (2005).

α-Terpineol, myrtenol, limonene, trans-carveol and γ-eudesmol

     

     

1,8-Cineole, limonene, α-pinene citronellal, citronellol, linalool, α-terpinene Anethol, fenchone, estragol

Linalol, α-terpinylacetate, 1,8-cineole, linalyacetate Citronellal, geraniol, citronellol, citral, limonene

Spathulenol, germacrene B, viridiflorol, globulol D-Carvone, limonene

p-Cymene, β-myrcene, (+)-limonene

Precocene I and II, β-caryophyllene, γ-bisabolene, fenchyl acetate Isobornyl-acetate, isothujone, nerolidol, camphene, eugenol

Camphor, 1,8-cineole, piperitone, borneol, α-terpineol

Constituents

Foeniculum vulgare L. Apiaceae Hibiscus cannabinus L. Malvaceae

Cymbopogon sp. Poaceae Eucalyptus sp. Myrtaceae

Coriandrum sativum L. Apiaceae

Anisomeles indica L. Lamiaceae Artemisia scoparia Waldst et Kit. Asteraceae Callicarpa japonica Thunb. Verbenaceae Carum carvi L. Apiaceae

Achillea sp. Asteraceae Ageratum conyzoides L. Asteraceae

Name Family

TABLE 24.6 Essential Oils that can be Used in Weed Control

(Continued)

Teuscher et al. (2004); Azirak et al. (2008) Teuscher et al. (2004); Dayan et al. (2009) Teuscher et al. (2004); Batish et al. (2006)

 

 

 

 

 

Teuscher et al. (2004); Azirek et al. (2008); de Almeida et al. (2010)

Kobaisy et al. (2002)

Kaur et al. (2010)

Batish et al. (2007b); Ushir et al. (2010)

Kong et al. (1999); Plant Encyclopedia (2012)

 

Reduces germination rate (under 25%) of C. salsotitialis, S. arvensis and R. raphanistrum Controls various weeds e.g., A. retroflexus and L. multiflorum, at higher concentration effective against lettuce, bentgrass, and against one cyanobacterium      

Source Kordali et al. (2009)

 

Toxic to A. stolonifera, but had no such effect on lettuce Inhibits germination of A. retroflexus, C. salsotitialis, S. arvensis, S. oleraceus, R. raphanistrum and R. nepalensis and A. pallida Effective against C. salsotitialis, S. arvensis, S. oleraceus, R. raphanistrum, and R. nepalensis

Herbicide against P. minor, positive effects on growth of wheat  

Inhibitory effect on germination and seedling growth of A. retroflexus, C. arvense, and L. serriola Causes phytotoxic effects on radish, mungbean, and tomatoes

Notes

894 Handbook of Essential Oils

   

   

   

 

Inhibits germination of wheat seedlings

Pulegone, β-caryophyllene, isomenthol, limonene Nepetalactone, myrtenol, terpinen-4-ol

 

 

 

Carvone, pulegone, menthol, menthone, menthylacetate

γ-Terpinene, α-terpinene, thymol, carvacrol, linalool, eugenol, carvone    

 

Inhibitory effect on germination of wheat seeds

Citral, citronellal, carvacrol, iso-menthone

   

 

   

(Continued)

de Almeida et al. (2010) Dudai et al. (1999); Teuscher et al. (2004); de Almeida et al. (2010) Konstantopoulou et al. (1992); Teuscher et al. (2004) Dudai et al. (1999) Mutlu et al. (2010)

 

 

 

Konstantopoulou et al. (1992); de Almeida et al. (2010)  

 

Teuscher et al. (2004); Ismail et al. (2012)

1,8-Cineole, α-pinene, limonene

Inhibits germination of seeds of B. danthoniae, B. intermedius, A. retroflexus, C. dactylon, C. album, and L. serriola  

Source Teuscher et al. (2004); Azirak et al. (2008) Kobaisy et al. (2001) Dudai et al. (1999); de Almeida et al. (2010)

 

Effective against S. arvensis, T. campestre, L. rigidum and P. canariensis causing electrolyte leakage in them

    Inhibitory effect on germination of wheat seeds

Notes

α-Pinene, myrcene, β-pinene, terpinene4-ol, germacrene D Linalol, linalyacetate, fenchone, camphor

β-Pinene, iso-pinocamphone, trans-pinocamphone

   

Constituents

Ocimum sp. Lamiaceae

Nepeta meyeri Benth. Lamiaceae

Lavandula sp. Lamiaceae Majorana hortensis L. Lamiaceae Melissa officinalis L. Lamiaceae Mentha sp. Lamiaceae Micromeria fruticosa Lamiaceae

Juniperus sp. Cupressaceae

    Hyssopus officinalis L. Lamiaceae

Name Family

TABLE 24.6 (Continued) Essential Oils that can be Used in Weed Control

Use of Essential Oils in Agriculture 895

Satureja sp. Lamiaceae

Salvia officinalis L. Lamiaceae

Ruta graveolens L. Rutaceae

Rosa damascena Mill. Rosaceae Rosmarinus officinalis L. Lamiaceae

Peumus boldus Mol. Monimiaceae Pimpinella anisum L. Apiaceae Pinus sp. Pinaceae Pistacia sp. Anacardiaceae

Carvacrol, thymol, γ-terpinene, p-cymene

α-Pinene, limonene, 1,8-cineole, nonan-2-one, undecan-2-one 1,8-Cineole, thujone, camphor, borneol

Limonene, 1,8-cineole, borneol, terpineol

Damages several weeds, e.g., dandelion leaves, C. album, A. artemisiifolia, S. halepense

Lowers germination rate of S. arvensis and R. raphanistrum; inhibits germination of wheat seeds

Selective toxicity against C. album, P. oleracea, E. crus-galli, C. annuum; good inhibition rates against C. salsotitialis, S. arvensis and R. raphanistrum and wheat seeds Inhibits germination and radicle elongation of radish

Completely inhibits germination of dicotyles S. arvensis and T. campestre, partially germination of monocotyles L. rigidum and P. canariensis  

α-Terpinene, limonene

(Continued)

Dudai et al. (1999); Teuscher et al. (2004); Azirak et al. (2008); de Almeida et al. (2010) Konstantopoulou et al. (1992); Tworkoski (2002); Angelini et al. (2003)

de Feo et al. (2002)

Konstantopoulou et al. (1992); Dudai et al. (1999); Angelini et al. (2003); Azirak et al. (2008)

Aridoğan et al. (2002); Teuscher et al. (2004)

Ismail et al. (2012)

Herbicide against various weeds

α-Pinene, β-pinene

Citronellol, geraniol, nerol, linalool

Teuscher et al. (2004); Dayan et al. (2009)

Herbicidal effect against R. raphanistrum

Dudai et al. (1999); Aslan et al. (2004); de Almeida et al. (2010) Konstantopoulou et al. (1992); de Almeida et al. (2010)

Source

Teuscher et al. (2004); Ibrahim et al. (2008); Verdeguer et al. (2011) Azirak et al. (2008); de Almeida et al. (2010)

 

Inhibits germination of wheat seeds

Notes

Herbicidal activity against A. hybridus and P. oleracea

Origanum sp. Lamiaceae

Constituents

Carvacrol, thymol, γ-terpinene, p-cymene, linalool Limonene, 1,8-cineole, p-cymene, ascaridol cis-anethol

 

 

Name Family

TABLE 24.6 (Continued) Essential Oils that can be Used in Weed Control

896 Handbook of Essential Oils

Verbena officinalis L. Verbenaceae Zataria multiflora Boiss. Lamiaceae

Thymbra sp. Lamiaceae Thymus vulgaris L. Lamiaceae

Tanacetum sp. Asteraceae

Syzygium aromaticum (L.) Merr et L.M. Perry Myrtaceae Tagetes minuta L. Asteraceae

Name Family

Saharkhiz et al. (2010)

 

Citral, isobornyl formate, linalol, geranial

Carvacrole, linalool, α-pinene, p-cymene

Thymol, p-cymene, carvacrol

Azirak et al. (2008); Ghrabi-Gammar et al. (2009) Kim et al. (2001); Tworkoski (2002); Teuscher et al. (2004); de Almeida et al. (2010) de Almeida et al. (2010)

Chiasson et al. (2001)

Lowers germination rate to under 5% of A. retroflexus, S. arvensis, S. oleraceus, and R. raphanistrum Inhibits the growth of P. oleracea, E. crus-galli, and C. album, lettuce and pepper crops; damages Taraxacum sp., C. album, A. artemisiifolia, and S. halepense  

 

Eguaras et al. (2005); Batish et al. (2007a); Scrivanti et al. (2003)

Inhibits germination of E. crus-galli and C. rotundus; inhibits root-growth of Zea mays

Limonene, (Z)-β-ocimene, (Z)- and (E)-ocimenone

α-Pinene, α-terpinene, γ-terpinene, carvone, borneol, β-thujone, camphor Thymol, carvacrol

Tworkoski (2002); Teuscher et al. (2004); Dayan et al. (2009)

Source

Damages Taraxacum sp., S. halepense, C. album, and A. artemisiifolia

Notes

Eugenol, aceteugenol, β-caryophyllene

Constituents

TABLE 24.6 (Continued) Essential Oils that can be Used in Weed Control

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weeds Portulaca oleracea, Echinochloa crus-galli, and Chenopodium album, but also of lettuce and pepper crops. T. vulgaris EO also causes damage to dandelion (Taraxacum sp., Asteraceae) leaves, common lambsquarters (Chenopodium album L.), common ragweed (Ambrosia artemisiifolia), and Johnsongrass (Sorghum halepense L.) (Tworkoski 2002). 24.3.3.2  Mentha sp. 2-Phenylethyl propionate can be obtained from peppermint oil (Mentha x piperita, L., Lamiaceae); it also contains menthol and menthone (Dayan et al., 2009) Mentha spicata L. EO contains (−)-carvone and limonene. M. spicata shows good effects against several weeds. It is able to inhibit germination (under 15%) of Amaranthus retroflexus, Centaurea solstitialis, Sinapis arvensis, Sonchus oleraceus, Raphanus raphanistrum, and Rumex nepalensis Spreng (Azirak et al., 2008). Mentha longifolia (L.) Huds and Mentha officinalis can be used as herbicides as they are able to inhibit germination of wheat seeds. This must be considered if the crop happens to be wheat (Dudai et al., 1999). 24.3.3.3  Cymbopogon sp. Besides its mosquito-repellent actions, lemongrass oil can also be used as an herbicide and Cymbopogon citratus (DC) Stapf (Poaceae) as weed control and an antipest. It is used as contact herbicide, since limonene is able to “remove the waxy cuticular layer from leaves”(Dayan et al. 2009) which leads to death by dehydration. Limonene can be used as a pesticide as well as an herbicide (Dayan et al., 2009). Cymbopogon winterianus EO shows toxicity toward several weeds, namely Senecio jacobaea L., Ranunculus repens L., Rumex obtusifolius L., and Urtica dioica L. When sprayed on trees, it causes damage to the foliage, but it does not inhibit their growth (Clay et al., 2005). The mode of action of citral is probably that it causes disruption of microtubule polymerization in weeds, as could be proved in Arabidopsis thaliana seedlings (Dayan et al., 2012). The effect of C. citratus EO was also proved by Dudai et al. The EO proves to be one of the most effective ones, as it is able to inhibit germination sooner (0% germination at 80 nL/mL) than most other EOs tested (Dudai et al., 1999). 24.3.3.4  Eucalyptus sp. Species of Eucalyptus (Myrtaceae), such as Eucalyptus citriodora and Eucalyptus tereticornis, can be used for weed control, because they could reduce growth, chlorophyll and water content when used as fumigants. They also have been shown to reduce the cellular respiration of Parthenium hysterophorus (Asteraceae). After about two weeks, injuries like necrosis or wilting could be observed (Batish et al., 2008; Dayan et al., 2009). Moreover, the EO of E. citriodora can also inhibit its seed germination. The suspected mode of action is the inhibition of mitosis (Singh et al., 2005). The EO obtained from E. tereticornis showed a higher toxicity than that of E. citriodora, because of a slightly different composition of the EO. In another study, E. citriodora EO was tested on crops—namely Triticum aestivum, Zea mays, and Raphanus sativus—and weeds—namely Cassia occidentalis, Amaranthus viridis, and Echinochloa crus-galli. It could be shown that the EO was more toxic to small-seeded crops such as A. viridis. When Eucalyptus EO was tested on P. hysterophorus and Phalaris minor, the herbicidal effect could also be confirmed (Batish et al., 2008). The herbicidal effect of E. citriodora on P. minor was examined in another study by Batish et al. They found that the concentration of lemon-scented eucalyptus oil is important for inhibiting the growth. The inhibition is higher at higher concentrations. The mode of action is more the inhibition of growth of seedlings than inhibition of germination; E. citriodora also lessens the chlorophyll content and the respiratory activity of the weed and causes an ion leakage in membranes (Batish et al., 2007c). The herbicidal effect of E. citriodora was also mentioned by Dudai et al. (1999) when they examined the inhibition of germination of wheat seeds. E. camaldulensis also showed promising effects in controlling Amaranthus hybridus and Portulaca oleracea, because it inhibits seedling growth as well as germination in both of them. Its main compound is identified as spathulenol (Verdeguer et al., 2009).

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24.3.3.5  Lavandula sp. When tested on various weeds by Azirak et al., Lavandula stoechas was not able to significantly inhibit the germination (under 25%) of most of the weed plants. It showed an effect on a few of them, namely Centaurea solstitialis, Sinapis arvensis, and Raphanus raphanistrum (Azirak et al., 2008). Lavandula x intermedia cv Grosso was tested on Lolium rigidum by Haig et al. It showed a high phytotoxicity when it comes to inhibiting root growth (Haig et al., 2009). 24.3.3.6  Origanum sp. An author group found out that the composition of oregano was different compared to the literature because other authors named p-cymene as main compound (de Almeida et al., 2010). O. onites L. EO proved to be effective against several weeds, namely Amaranthus retroflexus, Centaurea solstitialis, Sinapis arvensis, Sonchus oleraceus, and Raphanus raphanistrum, as they lower the germination rate to less than 20% (Azirak et al., 2008). O. majorana L. is also able to significantly inhibit germination of wheat seeds (Dudai et al., 1999). O. acutidens is able to inhibit germination and seedling growth of Amaranthus retroflexus, Chenopodium album, and Rumex crispus (Kordali et al., 2008). 24.3.3.7  Artemisia scoparia Artemisia scoparia Waldst. and Kit. (Asteraceae) was tested on Achyranthes aspera L., Cassia occidentalis L., Echinochloa crus-galli (L.) P. Beauv., Parthenium hysterophorus L. and Ageratum conyzoides L. The inhibitory effect on seedling growth was greatest in the latter two weeds. When the EO was used on the weeds, it caused wilting and necrosis of sprayed parts. Also, a decreasing chlorophyll amount and ion leakage can be observed. This effect was greatest in E. crus-galli and P. hysterophorus. A. scoparia EO’s main constituents are p-cymene, β-myrcene, and (+)-limonene (Kaur et al., 2010). 24.3.3.8  Zataria multiflora Two different ecotypes of Zataria multiflora Boiss, Lamiaceae, were tested in the study conducted by Saharkhiz et al. (2010). Ecotype B had similar constituents but in lower concentrations; this also explains why ecotype A was more successful in inhibiting the germination and growth of the tested weeds, which were Hordeum spontaneum Koch, Secale cereale, Amaranthus retroflexus, and Cynodon dactylon. 24.3.3.9  Tanacetum sp. Tanacetum aucheranum and Tanacetum chiliophyllum var. chiliophyllum, Asteraceae, are common in Europe and western Asia. The EOs were tested for their potential to inhibit germination and seedling growth of Amaranthus retroflexus, Chenopodium album, and Rumex crispus, in which they were very successful. Besides their herbicidal effect, they also have antibacterial and antifungal activity (Salamci et al., 2007). Weeds are a severe problem for agriculture today as they lead to decreased yields in crops. EOs, like those obtained from Cymbopogon sp., Eucalyptus sp., Origanum sp., and Thymus vulgaris have proved to be very potent inhibitors of both germination and seedling growth of various weeds. However, two limiting factors must be mentioned: first, the high volatiliy of EOs and, second, the fact that some EOs do not only inhibit the growth of weeds, but also that of crops. More research needs to be done on which weeds and crops are influenced by different volatile oils.

24.4  ESSENTIAL OILS AS INHIBITORS OF VARIOUS PESTS Volatile oils can also be used in controlling various other pests like bacteria, viruses, fungi, and nematodes. Their application and potential will be discussed in the following section (Table 24.7).

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TABLE 24.7 Essential Oils with Effect on Bacteria, Viruses, Fungi, and Nematodes Name Family Aloysia triphyllata Verbenaceae Artemisia sp. Asteraceae Calamintha nepeta Lamiaceae Carum carvi L. Apiaceae Cinnamomum zeylanicum Lauraceae Coridothymus sp. Lamiaceae Cotinus coggygria Scop. Anacardiaceae Cymbopogon sp. Poaceae

Constituents

Notes

Source

α-Thujone, cis-carveol, carvone, limonene

Nematicide

Duschatzky et al. (2004)

Limonene, myrcene, α-thujone, β-thujone, caryophyllyne, sabinyl-acetate Limonene, menthone, pulegone, menthol D-Carvone, limonene

Fungicide Nematicide

Chiasson et al. (2001)

Bactericide

Ibrahim et al. (2008)

Nematicide

Teuscher et al. (2004)

Cinnamaldehyde, linalool, β-caryophyllene, eugenol

Fungicide

Teuscher et al. (2004)

Carvacrol, thymol, γ-terpinene, p-cymene

Nematicide

Konstantopoulou et al. (1992)

Limonene, (Z)-β-ocimene, (E)-β-ocimene, β- caryophyllene Citronellal, geraniol, citronellol

Bactericide

Demirci et al. (2003)

Virucide Bactericide Fungicide Bactericide

Teuscher et al. (2004)

Ibrahim et al. (2008)

Ducrosia anethifolia (DC.) Boiss Apiaceae

α-Pinene, myrcene, limonene, terpinolene, (E)-β-ocimene

Echinophora tenuifolia Apiaceae

α-Phellandrene, eugenol, p-cymene

Bactericide

Aridoğan et al. (2002)

Elettaria cardamomum (L.) Maton Zingiberaceae Eucalyptus sp. Myrtaceae

α-Terpinylacetate, 1,8-cineole, linalylacetate

Fungicide

Teuscher et al. (2004)

1,8-Cineole, limonene, α-pinene

Bactericide Fungicide Nematicide Bactericide Fungicide Nematicide Fungicide

Teuscher et al. (2004)

Teuscher et al. (2004)

Bactericide

Teuscher et al. (2004)

Fungicide

de Corato et al. (2010)

Bactericide

Konstantopoulou et al. (1992)

Nematicide

Duschatzky et al. (2004)

Virucide

Teuscher et al. (2004)

Foeniculum vulgare L. Apiaceae

Anethol, fenchone, estragol

Hibiscus cannabinus L. Malvaceae

α-Terpineol, myrtenol, limonene, trans-carveol and γ-eudesmol

Juniperus sp. Cupressaceae

α-Pinene, myrcene, β-pinene, terpinen-4-ol, germacrene D 1,8-Cineole, linalool, terpineol acetate, methyleugenol, linalylacetate, eugenol Linalool, linalylacetate, fenchone, camphor Piperitenone-oxide, limonene

Laurus nobilis L. Lauraceae Lavandula sp. Lamiaceae Lippia sp. Verbenaceae Melaleuca alternifolia (Maiden et Betche) Cheel Myrtaceae

Terpinene-4-ol, γ-terpinene, α-terpinene

Kobaisy et al. (2001)

(Continued)

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TABLE 24.7 (Continued) Essential Oils with Effect on Bacteria, Viruses, Fungi, and Nematodes Name Family

Constituents

Notes

Source

Mentha sp. Lamiaceae

Carvone, pulegone, menthol, menthone, menthylacetate

Nematicide Bactericide

Micromeria fruticosa Lamiaceae

Pulegone, β-caryophyllene, isomenthol, limonene

Nematicide

Konstantopoulou et al. (1992); Teuscher et al. (2004) Dudai et al. (1999)

Ocimum sp. Lamiaceae

γ-Terpinene, α-terpinene, thymol, carvacrol, linalol, and eugenol

Nematicide

Aslan et al. (2004)

Origanum sp. Lamiaceae

Carvacrol, thymol, γ-terpinene, p-cymene, linalool

Konstantopoulou et al. (1992)

Pelargonium graveolens L’Hér. Geraniaceae Peumus boldus Mol. Monimiaceae Pinus sp. Pinaceae Rosa damascena Mill. Rosaceae Rosmarinus officinalis L. Lamiaceae Salvia fruticosa Miller Lamiaceae Satureja thymbra L. Lamiaceae

Geraniol, citronellol

Fungicide Bactericide Nematicide Nematicide

Limonene, 1,8-cineole, p-cymene

Bactericide

α-Pinene, β-pinene

Fungicide

Citronellol, geraniol, nerol, linalool Limonene, cineole, borneol, terpineol Cineole, thujone, camphor, α-pinene, β-pinene

Bactericide

Syzygium aromaticum (L.) Merr. et L.M. Perry Myrtaceae Tagetes minuta L. Asteraceae Thymbra sp. Lamiaceae Thymus sp. Lamiaceae Xylopia longifolia A. DC. Annonaceae

Ghrabi-Gammar et al. (2009) Teuscher et al. (2004); Ibrahim et al. (2008) Teuscher et al. (2004)

Bactericide

Aridoğan et al. (2002); Teuscher et al. (2004) Konstantopoulou et al. (1992)

Fungicide

Konstantopoulou et al. (1992)

Fungicide

Konstantopoulou et al. (1992)

Fungicide

Teuscher et al. (2004)

Limonene, (Z)-β-ocimene, (Z)- and (E)-ocimenone Thymol

Bactericide

Eguaras et al. (2005)

Fungicide

Ghrabi-Gammar et al. (2009)

Thymol, carvacrol, p-cymene

Bactericide Fungicide Bactericide

Teuscher et al. (2004)

Carvacrol, thymol, γ-terpinene, p-cymene Eugenol, aceteugenol, β-caryophyllene

α-Phellandrene, limonene, p-cymene, spathulenol

Fournier et al. (1993)

24.4.1  Effect on Bacteria EOs also have the potential to kill bacteria. Since they are lipophiles themselves, they can pass through the membrane of cells and cause cytotoxicity (Bakkali et al., 2008). On one hand, they “inhibit the respiration and increase the permeability of bacterial cytoplasmatic membranes” of bacteria, and on the other hand, they cause potassium leakage (Ibrahim et al., 2008). According to Burt (2004), other mechanisms that cause the bactericidal effects are depletion of proton motive force and damage to membrane proteins, as well as to the to the cytoplasmatic membrane and degradation of the cell wall. For example, the EO of Cymbopogon densiflorus (Poaceae), whose main compounds are limonene, cymenene, p-cymene, carveol, and carvone, were active against Gram-negative, as well as Grampositive, bacteria (Ibrahim et al., 2008). Another EO with effect against bacteria is that of Eucalyptus sp.

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(Myrtaceae), which mostly contains 1,8-cineole, linalool, citronellal, and limonene (Batish et al., 2008). The EO of Calamintha nepeta (Lamiaceae), which contains limonene, menthone, and pulegone, showed high activity against Salmonella. Of all the constituents, pulegone was the most effective. The EO from Peumus boldus (Monimiaceae) contains mainly monoterpenes, especially limonene, and was also highly efficient against Gram-negative and Gram-positive bacteria. Also the EOs of Foeniculum vulgare (Apiaceae)—mostly sweet fennel, Rosmarinus officinalis, and Thymus vulgaris (Lamiaceae)—showed high activity against bacteria. Other EOs with activity against bacteria are those obtained from Tagetes minuta (Asteraceae), Xylopia longifolia (Annonaceae), Cotinus coggygria Scop. (Anacardiaceae), and Ducrosia anethifolia (Ibrahim et al., 2008). EOs obtained from Origanum onites (Lamiaceae), Rosa damascena (Rosaceae), Mentha piperita (Lamiaceae), Echinophora tenuifolia (Apiaceae), Lavandula hybrida (Lamiaceae), and Juniperus exalsa (Cupressaceae) were tested on various bacteria. The EO of Origanum onites was reported to elicit the biggest effect (Aridoğan et al., 2002).

24.4.2  Effect on Fungi Volatile oils are also capable of killing fungi that harm crops, because they cause the increasing of plasma membrane permeability. Several monoterpenes, such as isopulegol, (R)-carvone, and isolimonene, were very efficient against Candida albicans. The EOs obtained from Foeniculum vulgare (Apiaceae) proved to be very active against Aspergillus niger. Also, EOs of citrus fruits (other than limonene) were tested against several Penicillum species, and they were highly efficient. Unlike many other EOs, volatile compounds that can be found in tomato leaves were able to inhibit the growth of hyphes of Alternaria alternata. Cardamom oil, linalool, limonene, and cineole showed the highest toxicity against fungi (Ibrahim et al., 2008). Another EO that is able to kill harmful fungi like Aspergillus sp., Penicillum sp., Fusarium sp., and Mucor sp. is that of Eucalyptus sp., which mostly contains 1,8-cineole, citronellal, limonene, and linalool. It is able to inhibit the growth of mycelium as well as spore production and germination (Batish et al., 2008). Pinus roxburghii (Pinaceae) EO was able to kill Aspergillus sp. and Cymbopogon pendulus (Poaceae) and can inhibit the mycelium growth of Microsporum gypseum and Trichophyton mentagrophytes. Moreover, EOs of Eucalyptus amygdalia (Myrtaceae) and Eucalyptus panciflora showed activity against Erysiphe cichoracearum. EOs obtained from Thymbra spicata, Satureja thymbra, Salvia fruticosa, Eucalyptus sp., and Origanum minutiflorum (Lamiaceae) were tested on several fungi. Most toxic to those fungi were the EOs of T. spicata, S. thymbra, and O. minutiflorum (Blaeser et al., 2002). Origanum, cassia, and red thyme EOs are able to inhibit fungi that destroy wood (Chao et al., 2000). The EO of Origanum acutidens (Lamiaceae) shows antifungal activity as well, for example, against several fungi species of Fusarium and Botrytis (Kordali et al., 2008). Against Botrytis cinereal, the EO from Cymbopogon martini (Poaceae), Thymus zygis (Lamiaceae), Cinnamomum zeylanicum (Lauraceae), and Syzygium aromaticum (Myrtaceae) showed high activity as well. Especially C. martini and T. zygis were very effective in inhibiting spore germination, followed by S. aromaticum EO (Wilson et al., 1997). Laurel oil (Laurus nobilis L., Lauraceae) showed antifungal activity against Rhizoctonia solani and Sclerotinia sclerotiorum (de Corato et al., 2010). EO of Hibiscus cannabinus L. (Malvaceae) was effective against fungi of Colletotrichum species, even though the antifungal effect was weak (Kobaisy et al., 2001). EOs from Artemisia dranunculus, Artemisia absinthium, Artemisia santonicum, and Artemisia spicigera, all of them Asteraceae, were tested against various fungi and shown to be effective against them (Kordali et al., 2005).

24.4.3  Effect on Viruses EOs were also found to be active against viruses. The tobacco mosaic virus, which is an important pest in agriculture, is weakly resistant to lemongrass EO (Chao et  al., 2000). But, also, EO of Melaleuca alternifolia (Myrtaceae) resulted in fewer lesions caused by tobacco mosaic virus for at least 10 days (Bishop, 1995).

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24.4.4  Effect on Nematodes The population of nematodes (Heterodera schachtii) could be decreased down to less than 3% of the control within 3 months using (+)-limonene. Also, menthol proved to be very efficient against nematodes (Ibrahim et al., 2008). EO of Ocimum sanctum L. (Lamiaceae) and eugenol as the main compound were tested on Caenorhabditis elegans and showed an anthelmintic effect (Asha et al., 2001). Another EO that is very promising in nematode control is Eucalyptus sp. EO. It proved to be toxic to Meloidogyne incognita and Meloidogyne exigua (Batish et al., 2008). Ocimum basilicum L. (Lamiaceae) and Ocimum sanctum L. (Lamiaceae), as well as their main compounds linalol and eugenol, are effective in killing larvae of M. incognita (Chatterjee et al., 1982). Pandey et al. (2000) reported that several Eucalyptus and Mentha species, as well as O. basilicum, as proved in the previous study, and Pelargonium graveolens L’Hér (Geraniaceae), volatile oils act as nematicides on M. incognita. Another study conducted on Meloidogyne sp. showed that Aloysia triphyllata (Verbenaceae), Lippia juneliana (Verbenaceae), and Lippia turbinata (Verbenaceae) have a nematicidal effect. The main components are α-thujone, cis-carveole, carvone, and limonene in A. triphyllata; piperitenone oxide, limonene, and camphor in L. junelia and limonene and piperitenone oxide in L. turbinata (Duschatzky et al., 2004). Oka et al. (2000) examined the effect of EO on M. javanica. EOs that significantly inhibited its mobility (more than 80%) were Artemisia judaica, Carum carvi, Coridothymus capitatus, Coridothymus citratus, Foeniculum vulgare, Mentha rotundifolia, Mentha spicata, Micromeria fruticosa, Origanum syriacum, and Origanum vulgare. EOs that lowered the egg hatching rate to less than 2% were Artemisia judaica, Carum carvi, Foeniculum vulgare, and Mentha rotundifolia. Ocimum gratissimum, with eugenol and 1,8-cineole as main compounds, was reported to have anthelmintic activity on Haemonchus contortus, as it decreases the egg hatch rate (Pessoa et al., 2002) (Table 24.8). This section gave a short overview about various pests that play a role in agriculture. The bactericidal and fungicidal effect of various EOs, like Cymbopogon sp., Thymus sp., Eucalyptus sp., and Foeniculum vulgare, among many others, has been known for quite a long time. This effect has been studied in many different publications, and many facts about the mode of action and the different effects are known already. EOs already show virucidal and nematocidal effects as well. An agrochemical relevant virus is the tobacco mosaic virus, for example, that causes big losses every year. An EO that could be of used in killing the virus is Cymbopogon sp. Also, nematodes, like the root-knot nematode, can be controlled by EOs—for example, by several Mentha species.

24.5  EFFECT OF ESSENTIAL OILS ON THE CONDITION OF THE SOIL 24.5.1  Effects of Essential Oils on Microorganisms and Soil Successful biodegradation is influenced by several factors, namely soil water, oxygen, redox potential, pH, nutrient status, and temperature (Holden et al., 1997). EOs are not only capable of destroying microorganisms, but these can also use the EOs as carbon and energy sources if they have been exposed to them recently. Thus, some microorganisms can actually be activated by EOs. This applies mostly to bacteria living in Mediterranean areas where many EO-containing plants also exist (Vokou et al., 1999). Especially, bacteria like Arthrobacter sp. and Nocardia sp. have been reported to use EOs as a carbon source. Also, several Pseudomonas species have been detected to use α-pinene as an energy source (Hassiotis, 2010; Hassiotis and Lazari, 2010b). EOs tested by Vokou et al. (1999) were Origanum vulgare subsp. hirtum (Lamiaceae), Rosmarinus officinalis (Lamiaceae), Mentha spicata (Lamiaceae), and Coridothymus capitatus (Lamiaceae), as well as Lavandula angustifolia (Lamiaceae). In another study, Vokou et al. (2002) observed the effect of EOs obtained from Lavandula stoechas (Lamiaceae), Satureia thymbra (Lamiaceae), or fenchone. They were all able to increase the CO2 emissions of the soil and the microbial population, even though some substances had not been in previous contact with the soil sample before (Vokou et al., 1999). According to Hassiotis and Dina (2010a), the deciding factor if EOs act as inhibitors or not is the concentration of the EO applied. At lower concentrations, they can induce bacterial growth; at higher concentrations, most likely they

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TABLE 24.8 List of Various Pests in Agriculture, Including Post-Harvest Food Pathogens and Soil Microbes Name Aeromonas hydrophila Alternaria alternata Arthrobacter sp. Aspergillus flavus Aspergillus niger Aspergillus parasiticus Bacillus cereus Botrytis cinerea Caenorhabditis elegans Candida albicans Cladosporium herbarum Clostridium botulinum Colletotrichum coccodes Erysiphe cichoracearum Eurotium sp. Fusarium oxyporum f. sp. dianthi Haemonchus contortus Heterodera schachtii Liseria monocytogenes Meloidogyne exigua Root-knot nematode Meloidogyne incognita Root-knot nematode Meloidogyne javanica Root-knot nematode Methylosinus trichosporium Microsporum gypseum Monilinia laxa Mucor sp. Nitrosomonas sp. Nitrosospira sp. Nocardia sp. Penicillium digitatum Polysphondylium pallidum Pseudomonas sp. Rhizoctonia solani Rhizopus stolonifer Salmonella enteritidis Sclerotinia sclerotiorum Sclerotium cepivorum Trichophyton mentagrophytes

Notes

Group

Source

Spoils meat   Can use EO as carbon source Storage fungus, can produce toxic aflatoxines Post-harvest pathogen Can produce toxic aflatoxines Food pathogen Post-harvest pathogen     Post-harvest pathogen Spoils food, produces neurotoxins Post-harvest pathogen Phytopathogenous funghus Spoils food, i.e., bakery products Phytopathogenous fungus Gastrointestinal parasite of ruminants Infects various plants, i.e., sugarbeet Spoils food Damages plant roots

Bacterium Fungus Bacterium Fungus Fungus Fungus Bacterium Fungus Nematode Fungus Fungus Bacterium Fungus Fungus Fungus Fungus Nematode

Burt (2004) Burt (2004) Hassiotis (2010) Blaeser et al. (2002); RazzaghiAbyaneh et al. (2008) Tzortzakis et al. (2007) Razzaghi-Abyaneh et al. (2008) Burt (2004) Tzortzakis et al. (2007) Tzortzakis et al. (2007) Tzortzakis et al. (2007) Razzaghi-Abyaneh et al. (2008) Jobling (2000) Tzortzakis et al. (2007) Blaeser et al. (2002) Guynot et al. (2005) Pitarokili et al. (2002) Pessoa et al. (2002)

Nematode

Ibrahim et al. (2008)

Bacterium Nematode

Jobling (2000) Batish et al. (2008)

Damages plant roots

Nematode

Batish et al., (2008)

Damages plant roots

Nematode

Oka et al. (2000)

Consumes methane in soils   Post-harvest pathogen   Oxydize ammonia Oxydize ammonia Can use EOs as carbon source Spoils food, i. e. bakery products Cellular slime mold, feeds on bacteria in forest soils   Post-harvest fungus Spoils food Spoils food, especially raw food Phytopathogenous fungus Phytopathogenous fungus  

Bacterium Fungus Fungus Fungus Bacterium Bacterium Bacterium Fungus Fungus

Amaral et al. (1998) Amaral et al. (1998) de Corato et al. (2010) de Corato et al. (2010) Chalkos et al. (2010) Chalkos et al. (2010) Hassiotis (2010) Guynot et al. (2005) Hwang et al. (2004)

Bacterium Fungus Fungus Bacteria Fungus Fungus Fungus

Hwang et al. (2004) de Corato et al. (2010) Tzortzakis et al. (2007) US-FDA (2000) Pitarokili et al. (2002) Pitarokili et al. (2002) Pitarokili et al. (2002)

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inhibit bacterial action. The only difference between the EOs was the delay they showed before an effect. While some were immediately effective, others needed more time to be efficient. Grampositive, as well as Gram-negative, bacteria were able to use monoterpenes or oxygenated products (Vokou et al., 2002). This effect is also enhanced by the ability of EOs to kill harmful fungi and their spores. As one can see, plants that contain EOs may influence the condition of the soil by activating the soil respiration through increasing the microbial growth. The plants have to be native to the soil’s place to achieve that effect. If EOs are tested on soil where they do not belong, they appear to be less or non-effective (Vokou et al., 1999). Amaral et al. (1998) found that several monoterpenes, especially (−)-α-pinene, were able to inhibit the atmospheric methane consumption by forest soils. EOs can kill Methylosinus trichosporium, a methanotrope bacterium, and they are also said to be able to inhibit the nitrification of forest soils. In the same way as the nitrogen cycle, also the carbon cycle is affected by EOs (Amaral et al., 1998). Generally said, terpenes, alcohols and esters can be degraded by bacteria in a very fast way, while it takes a lot longer for ketones (Hassiotis, 2010). Another interesting question is how EOs affect Polysphondylium, a cellular slime mold that feeds on bacteria in forest soils. (R)-(−)limonene, (−)-camphene, and (S)-(+)-carvone, as well as (−)-menthone and (+)-β-pinene, inhibited growth of Polysphondylium pallidum (Hwang et al., 2004).

24.5.2  Examples of Essential Oils with an Effect on Soil Conditions 24.5.2.1  Mentha spicata The main compounds of Mentha spicata(Lamiaceae) are (R)-(−)-carvone, limonene, and 1,8-cineole (Vokou et al., 2002) (Table 24.9). Chalkos et al. (2010) evaluated the ability of M. spicata-composted plants to increase growth of tomato crops and avoid weed growth. It also has TABLE 24.9 Essential Oils that can Influence Condition of the Soil Name Family Cymbopogon sp. Poaceae Laurus nobilis L. Lauraceae Lavandula sp. Lamiaceae Mentha spicata L. Lamiaceae Myrtus communis L. Myrtaceae Origanum sp. Lamiaceae Rosmarinus officinalis L. Lamiaceae Salvia sp. Lamiaceae Satureja thymbra L. Lamiaceae Thymbra capitata (L.) Cav. Lamiaceae

Constituents

Notes

Source

Citronellal, geraniol, citronellol 1,8-Cineole, linalool, terpineol acetate, methyleugenol, linalylacetate, eugenol Linalool, linalylacetate, fenchone, camphor Carvone, pulegone, menthol, menthone, menthylacetate

 

Teuscher et al. (2004)

 

de Corato et al. (2010)

 

Konstantopoulou et al. (1992)

 

1,8-Cineole, α-pinene, limonene, linalool, myrtenol Carvacrol, thymol, γ-terpinene, p-cymene, linalool Limonene, 1,8-cineole, borneol, terpineol, camphor Cineole, thujone, camphor, α-pinene, β-pinene Carvacrol, thymol, γ-terpinene, p-cymene Carvacrol, thymol, γ-terpinene, p-cymene

 

Konstantopoulou et al. (1992), (Teuscher et al. (2004) Teuscher et al. (2004)

Activates soil respiration

Konstantopoulou et al. (1992); Vokou et al. (1999, 2002)

Activates soil respiration

Konstantopoulou et al. (1992); Vokou et al. (1999, 2002) Konstantopoulou et al. (1992)

  Activates soil respiration Activates soil respiration

Konstantopoulou et al. (1992); Vokou et al. (2002) Vokou et al. (2002); Ghrabi-Gammar et al. (2009)

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a positive effect on the bacterial and fungal population in the soil. The bacteria Nitrosomonas and Nitrosospira are still oxidizing ammonia in the soil despite adding the compost. Among all these effects, the compost of M. spicata was able to increase the pH. A long time after adding M. spicata to the soil, compounds of the EO can be found, but in another composition than the original EO. Carvone, for example, makes up 50% of the EO in the beginning, but after some time in the soil, the concentration lowers to 1%, while other monoterpenes are gone completely. Sesquiterpenes, which act as allelochemicals, on the other hand, can be found at the same or even higher concentrations than in the original oil. 24.5.2.2  Lavandula sp. Lavender oil (Lavandula angustifolia Mill., Lamiaceae) mainly contains linalool and linalyl acetate. Its natural habitat is also the Mediterranean area. Spanish or French lavender oil (L. stoechas L., Lamiaceae), as it is also called, is obtained from L. stoechas and consists of fenchone (Vokou et al., 2002). Other compounds are camphor, p-cymene, and 1,8-cineole. In a study by Hassiotis and Dina (2010a), the bacterial growth-stimulating effect could be shown. In another study, L. stoechas’ EO degradation was examined. The highest EO degradation was in October, November, and December. During the hot summer months, the EO decrease was not very high. After 17 months, only camphene, 1,8-cineole, and camphor remained (Hassiotis, 2010). 24.5.2.3  Salvia sp. Adding Salvia fruticosa Mill. (Lamiaceae) compost to soil decreases the high C:N rates and raises the pH of the soil. Moreover, it increases the number of microbes in the soil, such as bacteria and fungi (Chalkos et al., 2010). S. sclarea (Lamiaceae) volatile oil’s compounds are linalyl acetate, linalool, geranyl acetate, and α-terpineol. The EO is able to cause fungicidal activity on Sclerotinia sclerotiorum, a soil-borne pathogen. It also inhibits growth of Sclerotium cepivorum and Fusarium oxysporum f. sp. Dianthi (Pitarokili et al., 2002). 24.5.2.4  Myrtus communis Hassiotis and Lazari examined the bacterial population growth during degradation of Myrtus communis L. (Myrtaceae) EO. It could be shown that the bacterial population was able to grow and use myrtle EO, for which the compounds were decreasing over the time, as an energy source. At the end of the study, only low percentages of 1,8-cineole and camphene could be detected (Hassiotis and Lazari, 2010b). 24.5.2.5  Laurus nobilis Laurus nobilis showed inhibition of bacterial activity and lowered the number of colonies as well as the soil respiration rate. This is because 1,8-cineole and eugenol are known for their bactericidal effects (Hassiotis and Dina, 2010a). 24.5.2.6  Cymbopogon sp. Malkomes (2006) tested the dehydrogenase activity in soil with and without dung and the addition of citronella (Cymbopogon sp., Poaceae) oil. A low concentration of citronella oil caused a decreasing stimulation of the activity of dehydrogenases. In the soil in which no dung was added, citronella oil also stimulated the CO2 production from the very beginning; in the soil in which dung was added, the CO2 production decreased at first, but after three weeks, it started to increase. The bactericidal action of EOs has been known for quite a long time, but they can also be used as energy sources for bacteria and activate them. That way, they can increase the quality of the soil because CO2 production is stimulated and pathogenous microorganisms are killed. Several EOs, namely Mentha spicata, Lavandula sp., and Cymbopogon sp., bear great potential to be used as dung and control of soil microbes in the future, but not expected effects on non-targeted species have to be also carefully explored.

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24.6  EOS USED IN POST-HARVEST DISEASE CONTROL Food safety is a very important topic. Even though there is increased hygiene during food production, still, 30% of people suffer from food poisoning every year, and as a result, two million people worldwide still die. A lot of people nowadays also want their food to have less synthetic food additives in it; therefore, there is a need to find natural substances which can replace them (Burt, 2004). Insects and mites that damage food and act as pathogens are also a major problem in food storage, both in developed and developing countries. EOs can be used as acaricides, and insecticides are explained in the Section 24.2.

24.6.1  Effects of Essential Oils on Stored-Product Pests In order to keep the concentration of health-damaging effects low, it is important to find out on the benefits of EOs on stored products. EOs that have proved themselves to be useful against post-harvest pathogens, namely the fungi Botrytis cinerea, are red thyme oil, clove oil, and cinnamon oil. EOs of Monarda citriodora (Lamiaceae) and Melaleuca alternifolia (Myrtaceae) showed an antifungal effect against many different fungi. Tea tree oil also works as an inhibitor on bacteria (Jobling, 2000). The effect of anethole, carvacrol, cinnamaldehyde, eugenol, and safrole were tested against fungi of Mucor, Aspergillus, and Rhizopus species. Against most Rhizopus and Mucor species, carvacrol proved itself to be most effective. Against Aspergillus species, both carvacrol and eugenol showed best results (Thompson, 1989). Carvacrol is also able to inhibit toxins which are produced by Bacillus cereus in various dishes (Burt, 2004). Against bacteria that damage stored products, cedar, eucalyptus, thyme, and chamomile oils proved themselves useful, especially when used against pathogenous bacteria B. cereus, Clostridium botulinum, and Listeria monocytogenes (Jobling, 2000). Generally said, EOs with a high percentage of phenolic compounds seem to be more effective against pathogens that damage food. Moreover, EOs have a bigger effect on Grampositive bacteria than on Gram-negative bacteria (Burt, 2004). But, it must be mentioned that the effect of EOs was tested in laboratories and not in the commercial field, and that many EOs only show a fungistatic effect, which means that the fungi will start growing again after the EO vapor is gone (Jobling, 2000). Moreover, a higher concentration of EOs—up to 100 times as much in soft cheese—is needed to inhibit bacteria in food than in vitro. The chemical environment of the food is also important for the effect; for example, the bactericide effect increases when the pH decreases; also the concentration of the fat in the food plays a major role because EOs lose their ability to harm bacteria with increasing fat concentration (Burt, 2004).

24.6.2  Examples of Essential Oils Used on Stored Products 24.6.2.1  Thymus zygis The main components of thyme (Thymus zygis L., Lamiaceae) oil are thymol and p-cymene (Table 24.10). Its home is the Mediterranean area (Jobling, 2000). Thyme oil can be used to protect meat products from two bacteria, namely Listeria monocytogenes and Aeromonas hydrophila. Thymol also showed activity against several Salmonella species (Burt, 2004). 24.6.2.2  Cinnamomum sp. Cinnamon oil (Cinnamomum zeylanicum J. Presl, Lauraceae) consists mainly of cinnamaldehyde (Jobling, 2000). It can be used to preserve dairy products, but it also proved to be successful in defeating several Salmonella bacteria on vegetables (Burt, 2004). Guynot et  al. (2005) tested Cinnamon EO on Eurotium sp., Aspergillus sp., and Penicillium sp. to preserve bakery products. 24.6.2.3  Cymbopogon citratus Cymbopogon citratus (Poaceae) was able to kill Aspergillus flavus, a fungus that can produce very toxic aflatoxins and harms stored products, to 100%. It has to be applied at about 1000 ppm to

Linalool, α-terpinyl acetate, 1,8-cineole, linalyl acetate Citronellal, geraniol, citronellol

Coriandrum sp. Apiaceae

Inhibits bacterial growth on meat, fish and vegetable products

Carvacrole, thymol, γ-terpinene, p-cymene, linalool Carvacrol, thymol

Eugenol, aceteugenol, β-caryophyllene

Thymol, carvacrol, p-cymene

Eugenol, isoeugenol, caryophyllene, eucalyptol, caryophyllene oxide

Syzygium aromaticum (L.) Merr. Et L.M. Perry Myrtaceae

Thymus zygis L. Lamiaceae

Zizyphus jujuba Rhamnaceae

Satureja hortensis Lamiaceae

Inhibitor of Aspergillus parasiticus (can produce aflatoxines) Eugenol shows high activity against A. flavus; is antibacterial; inhibits L. monocytogenes and A. hydrophila Protects meat products against L. monocytogenes and A. hydrophila; also against Salmonella species Active acainst Listeria monocytogenes

Konstantopoulou et al. (1992), Burt (2004)

Antifungal activity on most post-harvest fungi, e.g., Alternaria, Fusarium and Rhizopus species

Thymol, thymol methylester, α-terpinene, p-cymene

Monarda citriodora Lamiaceae Origanum sp. Lamiaceae

Konstantopoulou et al. (1992); Tassou et al. (1995); Teuscher et al. (2004) Bishop et al. (1997); Jobling (2000); Rozzi et al. (2002)

Is able to kill Salmonella enteritidis in several foods

Carvone, pulegone, menthol, menthone, menthylacetate

Jobling (2000); Burt (2004); Teuscher et al. (2004) Al-Reza et al. (2009)

Jobling (2000); Blaeser et al. (2002); Burt (2004); Teuscher et al. (2004)

Razzaghi-Abyaneh et al. (2008)

Jobling (2000); Teuscher et al. (2004)

At 500 ppm it is able to kill 100% of the fungi Botrytis sp. on grapes

Terpinene-4-ol, γ-terpinene, α-terpinene

Melaleuca alternifolia (Maiden et Betche) Cheel Myrtaceae Mentha piperita L. Lamiaceae

de Corato et al. (2010)

Teuscher et al. (2004) Teuscher et al. (2004); Jobling (2000)

Burt (2004); Teuscher et al. (2004)

Teuscher et al. (2004)

Chiasson et al. (2001); Kordali et al. (2005)

Source

1,8-Cineole, linalool, terpineol acetate, methyleugenol, linalyacetate, eugenol

  Against browning of mushrooms in plastic bags, antibacterial  

Can be used to preserve meat products from bacteria

 

Very strong antifungal activity

Notes

Laurus nobilis L. Lauraceae

1,8-Cineole, limonene, α-pinene

Cinnamaldehyde, linalol, β-caryophyllene, eugenol

Cinnamomum sp. Lauraceae

Cymbopogon sp. Poaceae Eucalyptus sp. Myrtaceae

Limonene, myrcene, α-thujone, β-thujone, caryophellene, sabinyl-acetate, camphor, 1,8-cineole, borneol, terpinen-4-ol, bornyl acetate

Constituents

Artemisia sp. Asteraceae

Name Family

TABLE 24.10 Essential Oils that can be Used to Preserve Stored Products

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inhibit Aspergillus flavus. Besides A. flavus, it has the ability to kill many other fungi, as well, for about 210 days in storage. The EO of Cymbopogon citratus, especially if it was produced between May and November, also appeared to be more successful in fighting fungi than several synthetic products (Mishra et al., 1994). Lemongrass EO also showed activity against colony development of Cladosporium herbarum and Rhizopus stolonifer. A concentration of 500 ppm was lethal for Colletotrichum coccodes, Cladosporium herbarum, Rhizopus stolonifera, and Aspergillus niger. The fungal colony growth of Botrytis cinerea was only inhibited up to 60%. The fungal spore production could be inhibited in all five fungi (Tzortzakis et al., 2007). Lemongrass EO can also be used against A. flavus in parboiled rice. It causes fungistatic and fungicidal effects (Paraganama et al., 2003). 24.6.2.4  Laurus nobilis EO of Laurus nobilis L., Lauraceae, showed good effects in the protection of the harvested fruits of kiwi and peach against the post-harvest fungi Monilinia laxa, Botrytis cinerea, and Penicillium digitatum. The fungistatic effect is very high in M. laxa and B. cinerea, but P. digitatum cannot be completely killed (de Corato et al., 2010). The protection of stored food against pathogens like bacteria, fungi, insects, and mites is a big challenge, especially in developing countries. Especially volatile oils obtained from Cinnamomum sp., Cymbopogon sp., and Syzygium aromaticum (earlier known as Eugenia caryophyllata) bear great potential. One thing that should be considered is that they have a strong taste themselves, therefore not every EO can be used in every dish as the flavors might not match very well. But overall, adding EOs to food helps preserve it and seems to be a good idea since many EO plants are already used as spices throughout the world, and most of them do not have any known toxic effects. But of course, studies must be conducted to make sure adding volatile oils does not lead to yet-unknown side effects.

24.7 CONCLUSION EOs can be used in various fields of application, and one of them is in agriculture. Until now, agriculture has been dominated by synthetic products in order to control various pests, weed, and soil condition, but EOs bear great potential to replace or complement those substances. In insect pest control, EOs proved to be very promising, as they have great potential for killing various harmful insects. Especially in developing countries, they might play a bigger role in the future as they are very cheap and can be combined with synthetic products to minimize their toxicity. When it comes to repellent activity, EOs might not be that useful as only some insects will be deterred while others can be attracted by them. When it comes to weed control, EOs do not seem as promising because of the high quantities needed and the frequent application, which might also lead to negative environmental effects. Moreover, the possibility that the applied EOs might damage the wanted crops cannot be excluded. Volatile oils are also able to kill various pests, namely bacteria, fungi, and viruses, as well as mites and nematodes. Especially in the control of tobacco mosaic virus, EOs might become very important. More research needs to be done in this field of application. Furthermore, EOs are also able to improve the condition of the soil because if exposed to the oils earlier, bacteria can use them as a carbon source. That way, the respiration of the soil can be activated. Finally, a very interesting aspect of EOs is to use them in post-harvest pest management and against fungi and bacteria on stored products. Especially in developing countries, insect-, and other pest-, management on stored products might become very useful. But, also in industrialized countries, there is some use for them; for example, in supermarkets on fruits and vegetables to make them less perishable. Because they have a natural origin, they are biodegradable and diverse physiological targets to control pests that may delay the evolution of pest resistance, EOs are considered to be friendly environmental products and have been embraced by the public. However, the development of EOs as PPPs for a sustainable agriculture needs to have more data on environmental features of EOs and their components, and also on plant–pest interrelationships and mechanisms. More fundamental and applied studies are needed to better know the impact of

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these compounds on non-target organisms (unintended affects) or on the biodiversity if some plant endemic resources are hugely used to provide EOs. It is not because they are natural that the EOs have no adverse effect, even they have no history of adverse effects. So, a benefit-to-risk assessment on a case-by-case basis must to be required to prevent undesirable situations and to have a reasonable management of real risk when there is some. Most government policies in all the world are now seeking low-risk plant protection products. The US EPA is the first administration that has considered reduced regulatory regime for about 20 years and, as a result, has allowed a significant number of EOs for commercial use. The results of this American position must be checked carefully to determine positive and negative inputs of using EOs in such a way. In EU, the Directive 2009/128/CE makes an obligation to develop Integrated Pest Management (IPM) as the strategy of agricultural pests control. It entered into force in January 2014. Because of their numerous advantages in controlling harmful pests, and because they are especially suited to organic farming as well as to Integrated Pest Management in a complementary approach to synthetic insecticides, EOs certainly have a room in biocontrol approaches to promote sustainable development.

REFERENCES Adorjan, B., and G. Buchbauer, 2010. Biological properties of EOs: An updated review. Flavour Fragance J., 25: 407–426. Al-Reza, S.M., V.K. Bajpai, and S.C. Kang, 2009. Antioxidant and antilisterial effect of seed EO and organic extracts from Zizyphus jujuba. Food Chem. Toxicol., 47: 2374–2380. Amaral, J.A., and R. Knowles, 1998. Inhibition of methane consumption in forest soils by monoterpenes. J. Chem. Ecol., 24: 723–734. Ammon, H.P.T., C. Hunnius, and A. Bihlmayer, 2010. Hunnius pharmazeutisches Wörterbuch. de Gruyter. Angelini, L.G., G. Carpanese, P.L. Cioni, I. Morelli, M. Macchia, and G. Flamini, 2003. EOs from Mediterranean Lamiaceae as weed germination inhibitors. J. Agric. Food Chem., 51: 6158–6164. Anthony, N.M., J.B. Harrison, and D.B. Sattelle, 1993. GABA receptor molecules of insects. EXS, 63: 172–209. Aridoğan, B.C., H. Baydar, S. Kaya, M. Demirci, D. Ozbaşar, and E. Mumcu, 2002. Antimicrobial activity and chemical composition of some EOs. Arch. Pharm. Res., 25: 860–864. Arnason, J.T., 2011. Natural products from plants as insecticides in agriculture and human health (Chapter 13). In Photochemistry and pharmacognosy in Encyclopedia of Life Support Systems (EOLSS), developed under the Auspices of the UNESCO, J.M. Pezzuto, and M.J. Kato (eds.). Oxford, UK: Eolss Publishers. Arnason, J.T., T. Durst, J.R. Philogène, and I.A. Scott, 2008. Prospection d’insecticides phytochimiques de plantes tempérées et tropicales communes ou rares. In Biopesticides d’origine végétale, C. RegnaultRoger, B.J.R. Philogène, and C. Vincent (eds.). Paris: Lavoisier. Arnason, J.T., S.R. Sims, and I. Scott. 2012. Natural products from plants as insecticides. In Encyclopedia of Life Support Systems (EOLSS) EOLSS-publishers (editors). Developed under the Auspices of the UNESCO. Oxford, UK: Eolss Publishers, [http://www.eolss.net]. Asha, M.K., D. Prashanth, B. Murali, R. Padmaja, and A. Amit, 2001, Anthelmintic activity of EO of Ocimum sanctum and eugenol. Fitoterapia, 72: 669–670. Aslan, İ., H. Özbek, Ö. Çalmaşur, and F. Şahin, 2004. Toxicity of EO vapours to two greenhouse pests, Tetranychus urticae Koch and Bemisia tabaci Genn. Ind. Crops Prod., 19: 167–173. Ayvaz, A., O. Sagdic, S. Karaborklu, and I. Ozturk, 2010. Insecticidal activity of the EOs from different plants against three stored-product insects. J. Insect Sci., 10: 21. Azirak, S., and S. Karaman, 2008. Allelopathic effect of some EOs and components on germination of weed species. Acta Agric. Scand. Section B—Soil Plant Sci., 58: 88–92. Bakkali, F., S. Averbeck, D. Averbeck, and M. Idaomar, 2008. Biological effects of EOs—a review. Food Chem. Toxicol., 46: 446–475. Balfanz, S., T. Strünker, S. Frings, and A. Baumann, 2005. A family of octopamine [corrected] receptors that specifically induce cyclic AMP production or Ca2+ release in Drosophila melanogaster. J. Neurochem., 93: 440–451. Batish, D.R., K. Arora, H.P. Singh, and R.K. Kohli, 2007a. Potential utilization of dried powder of Tagetes minuta as a natural herbicide for managing rice weeds. Crop Prot., 26: 566–571. Batish, D.R., M. Kaur, H.P. Singh, and R.K. Kohli, 2007b. Phytotoxicity of a medicinal plant, Anisomeles indica, against Phalaris minor and its potential use as natural herbicide in wheat fields. Crop Prot., 26: 948–952.

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Batish, D.R., H.P. Singh, R.K. Kohli, and S. Kaur, 2008. Eucalyptus EO as a natural pesticide. Forest Ecol. Manage., 256: 2166–2174. Batish, D.R., H.P. Singh, N. Setia, S. Kaur, and R.K. Kohli, 2006. Chemical composition and phytotoxicity of volatile EO from intact and fallen leaves of Eucalyptus citriodora. Z. Naturforsch. C. J. Biosci., 61: 465–471. Batish, D.R., H.P. Singh, N. Setia, R.K. Kohli, S. Kaur, and S.S. Yadav, 2007c. Alternative control of littleseed canary grass using eucalypt oil. Agron. Sustain. Dev., 27: 171–177. Bishop, C.D., 1995. Antiviral activity of the EO of Melaleuca alternifolia (Maiden amp; Betche) Cheel (Tea Tree) against tobacco mosaic virus. J. Essent. Oil Res., 7: 641–644. Bishop, C.D., and I.B. Thornton, 1997. Evaluation of the antifungal activity of the EOs of Monarda citriodora var. citriodora and Melaleuca alternifolia on post-harvest pathogens. J. Essent. Oil Res., 9: 77–82. Blaeser, P., U. Steiner, and H.W. Dehne, 2002. Pflanzeninhaltsstoffe mit fungizider Wirkung. Schriftenreihe des Lehr- und Forschungsschwerpunktes USL, Landwirtschaftliche Fakultät der Universität Bonn, 97, 1–143. Bruneton, J., 2009. Pharmacognosie, 4ème éd. Paris: Tec & Doc Lavoisier. Buchbauer, G., S. Hemetsberger, 2016. Use of EOs in agriculture. In Handbook of EOs: Science, Technology, and Applications, 2nd ed, K. Husnu Can Baser and G. Buchbauer (eds.), pp. 669–706. CRC Press Book. Burt, S., 2004. EOs: Their antibacterial properties and potential applications in foods—a review. Intern. J. Food Microbiol., 94: 223–253. Çalmaşur, Ö., İ. Aslan, and F. Şahin, 2006. Insecticidal and acaricidal effect of three Lamiaceae plant EOs against Tetranychus urticae Koch and Bemisia tabaci Genn. Indus. Crops Prod., 23: 140–146. Capinera, J.L., 2008. Encyclopedia of Entomology. Springer Science & Business Media. Cavalcanti, S.C.H., E. dos S. Niculau, A.F. Blank, C.A.G. Câmara, I.N. Araújo, and P.B. Alves, 2010. Composition and acaricidal activity of Lippia sidoides EO against two-spotted spider mite (Tetranychus urticae Koch). Bioresour. Technol., 101: 829–832. Cetin, H., J.E. Cilek, E. Oz, L. Aydin, O. Deveci, and A. Yanikoglu, 2010. Acaricidal activity of Satureja thymbra L. EO and its major components, carvacrol and γ-terpinene against adult Hyalomma marginatum (Acari: Ixodidae). Vet. Parasitol., 170: 287–290. Chalkos, D., K. Kadoglidou, K. Karamanoli, C. Fotiou, A.S. Pavlatou-Ve, I.G. Eleftherohorinos, H.-I.A. Constantinidou, and D. Vokou, 2010. Mentha spicata and Salvia fruticosa composts as soil amendments in tomato cultivation. Plant Soil, 332: 495–509. Chao, S.C., D.G. Young, and C.J. Oberg, 2000. Screening for inhibitory activity of EOs on selected bacteria, fungi and viruses. J. Essent. Oil Res., 12: 639–649. Chatterjee, A., N.C. Sukul, S. Laskar, and S. Ghoshmajumdar, 1982. Nematicidal principles from two species of Lamiaceae. J. Nematol., 14: 118–120. Chiasson, H., A. Bélanger, N. Bostanian, C. Vincent, and A. Poliquin, 2001. Acaricidal properties of Artemisia absinthium and Tanacetum vulgare (Asteraceae) EOs obtained by three methods of extraction. J. Econom. Entomol., 94: 167–171. Clay, D.V., F.L. Dixon, and I. Willoughby, 2005. Natural products as herbicides for tree establishment. Forestry, 78: 1–9. Dayan, F.E., C.L. Cantrell, and S.O. Duke, 2009. Natural products in crop protection. Bioorg. Med. Chem., 17: 4022–4034. Dayan, F.E., D.K. Owens, and S.O. Duke, 2012. Rationale for a natural products approach to herbicide discovery. Pest Manag. Sci., 68: 519–528. de Almeida, L.F.R., F. Frei, E. Mancini, L. de Martino, and V. de Feo, 2010. Phytotoxic activities of Mediterranean EOs. Molecules, 15: 4309–4323. de Corato, U., O. Maccioni, M. Trupo, and G. Di Sanzo, 2010. Use of EO of Laurus nobilis obtained by means of a supercritical carbon dioxide technique against post harvest spoilage fungi. Crop Prot., 29: 142–147. de Feo, V., F. de Simone, and F. Senatore, 2002. Potential allelochemicals from the EO of Ruta graveolens. Phytochemistry, 61: 573–578. DGAL (Direction générale de l’alimentation du Ministère de l’agriculture et de l’alimentation). 2017. Note de service DGAL/SDQSPV/2017-635 du 19/07/2017. Demirci, B., F. Demirci, and K.H.C. Başer, 2003. Composition of the EO of Cotinus coggygria Scop. from Turkey. Flavour Fragr. J., 18: 43–44. Douglas, M.H., J.W. van Klink, B.M. Smallfield, N.B. Perry, R.E. Anderson, P. Johnstone, and R.T. Weavers, 2004. EOs from New Zealand manuka: Triketone and other chemotypes of Leptospermum scoparium. Phytochemistry, 65: 1255–1264. Dudai, N., O. Larkov, E. Putievsky, H.R. Lerner, U. Ravid, E. Lewinsohn, and A.M. Mayer, 2000. Biotransformation of constituents of EOs by germinating wheat seed. Phytochemistry, 55: 375–382.

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Dudai, N., A. Poljakoff-Mayber, A.M. Mayer, E. Putievsky, and H.R. Lerner, 1999. EOs as allelochemicals and their potential use as bioherbicides. J. Chem. Ecol., 25: 1079–1089. Duschatzky, C.B., A.N. Martinez, N.V. Almeida, and S.L. Bonivardo, 2004. Nematicidal activity of the EOs of several Argentina plants against the root-knot nematode. J. Essent. Oil Res., 16: 626–628. Eguaras, M.J., S. Fuselli, L. Gende, R. Fritz, S.R. Ruffinengo, G. Clemente, A. Gonzalez, P.N. Bailac, and M.I. Ponzi, 2005. An in vitro evaluation of tagetes minuta EO for the control of the honeybee pathogens Paenibacillus larvae and ascosphaera apis, and the parasitic mite Varroa destructor. J. Essent. Oil Res., 17: 336–340. Enan, E.E., 2005. Molecular and pharmacological analysis of an octopamine receptor from American cockroach and fruit fly in response to EOs. Arch. Insect. Biochem. Physiol., 59: 161–171. Environ Protection Agency (EPA). 2018. Regulating biopesticides. http://www.epa.gov/pesticides/biopesticides (accessed 15 August 2018). Feng, L., and Prestwich, G.D. 1997. Expression and characterization of a lepidopteran general odorant binding protein. Insect. Biochem. Mol. Biol., 27: 405–412. Fournet, A., A. Rojas de Arias, B. Charles, and J. Bruneton, 1996. Chemical constituents of EOs of muña, Bolivian plants traditionally used as pesticides, and their insecticidal properties against Chagas’ disease vectors. J. Ethnopharmacol., 52: 145–149. Fournier, G., A. Hadjiakhoondi, M. Leboeuf, A. Cavé, J. Fourniat, and B. Charles, 1993. Chemical and biological studies of Xylopia longifolia A. DC. EOs. J. Essent. Oil Res., 5: 403–410. García, M., O.J. Donadel, C.E. Ardanaz, C.E. Tonn, and M.E. Sosa, 2005. Toxic and repellent effects of Baccharis salicifolia EO on Tribolium castaneum. Pest. Manag. Sci., 61: 612–618. George, D.R., G. Olatunji, J. H. Guy, and O. A. E. Sparagano, 2010. Effect of plant EOs as acaricides against the poultry red mite, Dermanyssus gallinae, with special focus on exposure time. Vet. Parasitol., 169: 222–225. George, D.R., T.J. Smith, R.S. Shiel, O.A.E. Sparagano, and J.H. Guy, 2009a. Mode of action and variability in efficacy of plant EOs showing toxicity against the poultry red mite, Dermanyssus gallinae. Vet. Parasitol., 161: 276–282. George, D.R., O.A.E. Sparagano, G. Port, E. Okello, R.S. Shiel, and J.H. Guy, 2009b. Repellence of plant EOs to Dermanyssus gallinae and toxicity to the non-target invertebrate Tenebrio molitor. Vet. Parasitol., 162: 129–134. Gershenzon, J., M.E. McConkey, and R.B. Croteau, 2000. Regulation of monoterpene accumulation in leaves of peppermint. Plant Physiol., 122: 205–213. Ghrabi-Gammar, Z., D.R. George, A. Daoud-Bouattour, I. Ben Haj Jilani, S.B. Saad-Limam, and O.A.E. Sparagano, 2009. Screening of EOs from wild-growing plants in Tunisia for their yield and toxicity to the poultry red mite, Dermanyssus gallinae. Indus. Crops Prod., 30: 441–443. Glitho, A.I., G.K. Ketoh, P.Y. Nuto, S.K. Amevoin, and J. Huignard, 2008. Approches non Centre et de l’Ouest. In Biopesticides d’origine végétale, C. Regnault-Roger, B.J.R. Philogène, and C. Vincent (eds.). Paris: Lavoisier. Grieneisen, M.L., and M.B. Isman, 2018. Recent developments in the registration and usage of botanical pesticides in California in managing and analyzing pesticide use data for pest management, environmental monitoring, public health, and public policy. American Chemical Society Symposium Series. vol. 1283, DOI: 10.1021/bk-2018-1283.ch008 Guynot, M.E., S. MarÍn, L. SetÚ, V. Sanchis, and A.J. Ramos, 2005. Screening for antifungal activity of some EOs against common spoilage fungi of bakery products. Food Sci. Technol. Intern., 11: 25–32. Haig, T.J., T.J. Haig, A.N. Seal, J.E. Pratley, M. An, and H. Wu, 2009. Lavender as a source of novel plant compounds for the development of a natural herbicide. J. Chem. Ecol., 35: 1129–1136. Hassiotis, C.N., 2010. Chemical compounds and EO release through decomposition process from Lavandula stoechas in Mediterranean region. Biochem. Systematics Ecol., 38: 493–501. Hassiotis, C.N., and E.I. Dina, 2010a. The influence of aromatic plants on microbial biomass and respiration in a natural ecosystem. Israel J. Ecol. Evol., 56: 181–196. Hassiotis, C.N., and D.M. Lazari, 2010b. Decomposition process in the Mediterranean region. Chemical compounds and EO degradation from Myrtus communis. Int. Biodeter. Biodegr., 64: 356–362. Haubruge, E., G. Lognay, M. Marlier, P. Danhier, J.-C. Gilson, and C. Gaspar, 1989. The toxicity of five EOs extracted from Citrus species with regards to Sitophilus zeamais Motsch (Col., Curculionidae), Prostephanus truncatus (Horn) (Col., Bostrychidae) and Tribolium castaneum Herbst (Col., Tenebrionidae). Medelingen van de Fac. Landbouwwet. Rijksuniv. Gent, 54: 1083–1093. Holden, P.A., and M.K. Firestone, 1997. Soil microorganisms in soil cleanup: How can we improve our understanding? J. Environm. Qual., 26: 32.

Use of Essential Oils in Agriculture

913

Huang, Y., and S.H. Ho, 1998. Toxicity and antifeedant activities of cinnamaldehyde against the grain storage insects, Tribolium castaneum (Herbst) and Sitophilus zeamais Motsch. J. Stored Prod. Res., 34: 11–17. Huang, Y., S.-H. Ho, H.-C. Lee, and Y.-L. Yap, 2002. Insecticidal properties of eugenol, isoeugenol and methyleugenol and their effects on nutrition of Sitophilus zeamais Motsch. (Coleoptera: Curculionidae) and Tribolium castaneum (Herbst) (Coleoptera: Tenebrionidae). J. Stored Prod. Res., 38: 403–412. Huang, Y., S.L. Lam, and S.H. Ho, 2000. Bioactivities of EO from Elletaria cardamomum (L.) Maton. to Sitophilus zeamais Motschulsky and Tribolium castaneum (Herbst). J. Stored Prod. Res., 36: 107–117. Huignard, J., B. Lapied, S. Dugravot, M. Magnin-Robert, G.K. Ketoh, 2008. Modes d’action neurotoxiques des dérivés soufrés et de certaines huiles essentielles et risques liés à leur utilisation. In Biopesticides d’Origine Végétale, 2nd ed, C. Regnault-Roger, B.J.R. Philogène, and C. Vincent (eds.). Paris: Lavoisier Tech & Doc. Hummelbrunner, L.A., and M.B. Isman, 2001. Acute, sublethal, antifeedant, and synergistic effects of monoterpenoid EO compounds on the tobacco cutworm, Spodoptera litura (Lep., Noctuidae). J. Agric. Food Chem., 49: 715–720. Hwang, J.-Y., and J.-H. Kim, 2004. The effect of monoterpenoids on growth of a cellular slime mold, Polysphondylium pallidum. J. Plant Biol., 47: 8–14. Ibrahim, M.A., P. Kainulainen, and A. Aflatuni, 2008. Insecticidal, repellent, antimicrobial activity and phytotoxicity of EOs: With special reference to limonene and its suitability for control of insect pests. Agric. Food Sci., 10: 243–259. Ismail, A., H. Lamia, H. Mohsen, and J. Bassem, 2012. Herbicidal potential of EOs from three Mediterranean trees on different weeds. Curr. Bioactive Compd., 8: 3–12. Isman, M.B., 2000. Plant EOs for pest and disease management. Crop Prot., 19: 603–608. Isman, M.B., 2004. Plant EOs as green pesticides for pest and diseases management. ACS Symp. Ser., 887: 41–51 Isman, M.B., 2006. Botanical insecticides, deterrents, and repellents in modern agriculture and an increasingly regulated world. Annu. Rev. Entomol., 51: 45–66. Isman, M.B., 2008. Botanical insecticides: For richer, for poorer. Pest Manag. Sci., 64: 8–11. Isman, M.B., 2017. Bridging the gap: Moving botanical insecticides from the laboratory to the farm. Indus. Crops Prod., 110: 10–14. Isman, M.B., and C.M. Machial, 2006. Pesticides based on plant EOs: From traditional practice to commercialization. In Naturally Occurring Bioactive Compounds, M. Rai and M.C. Carpinella (eds.). Amsterdam: Elsevier BV. Isman, M.B., S. Miresmailli, and C. Machial, 2011. Commercial opportunities for pesticides based on plant EOs in agriculture, industry and consumer products. Phytochem. Rev., 10, 197–204. Isman, M.B., and J.-H. Tak, 2017a. Commercialization of insecticides based on plant EOs: Past, present, and future. In Green Pesticides Handbook: EOs for pest Control, L.M.L. Nollet and H.S. Rathore (eds.), pp. 27–39. CRC Press. Isman, M.B., and J.-H. Tak, 2017b. Inhibition of acetylcholinesterase by EOs and monoterpenoids: A relevant mode of action for insecticidal EOs? Biopestic. Int., 13(2): 71–78. Jobling, J., 2000. EOs: A new idea for postharvest disease control. Good Fruit Vegetables Mag., 11: 50. Katerinopoulos, H.E., G. Pagona, A. Afratis, N. Stratigakis, and N. Roditakis, 2005. Composition and insect attracting activity of the EO of Rosmarinus officinalis. J. Chem. Ecol., 31: 111–122. Kaur, S., H.P. Singh, S. Mittal, D.R. Batish, and R.K. Kohli, 2010. Phytotoxic effects of volatile oil from Artemisia scoparia against weeds and its possible use as a bioherbicide. Indus. Crops Prod., 32: 54–61. Kéita, S.M., C. Vincent, J.-P. Schmit, J.T. Arnason, and A. Bélanger, 2001. Efficacy of EO of Ocimum basilicum L. and O. gratissimum L. applied as an insecticidal fumigant and powder to control Callosobruchus maculatus (Fab.) [Coleoptera: Bruchidae]. J. Stored Prod. Res., 37: 339–349. Kim, D.-H., and Y.-J. Ahn, 2001. Contact and fumigant activities of constituents of Foeniculum vulgare fruit against three coleopteran stored-product insects. Pest. Manag. Sci., 57: 301–306. Kim, E.-H., H.-K. Kim, and Y.-J. Ahn, 2003a. Acaricidal activity of plant EOs against Tyrophagus putrescentiae (Acari: Acaridae). J. Asia-Pacific Entomol., 6: 77–82. Kim, H.-K., J.-R. Kim, and Y.-J. Ahn, 2004. Acaricidal activity of cinnamaldehyde and its congeners against Tyrophagus putrescentiae (Acari: Acaridae). J. Stored Prod. Res., 40: 55–63. Kim, S.-I., Y.-E. Na, J.-H. Yi, B.-S. Kim, and Y.-J. Ahn, 2007. Contact and fumigant toxicity of oriental medicinal plant extracts against Dermanyssus gallinae (Acari: Dermanyssidae). Vet. Parasitol., 145: 377–382. Kim, S.-I., J.-Y. Roh, D.-H. Kim, H.-S. Lee, and Y.-J. Ahn, 2003b. Insecticidal activities of aromatic plant extracts and EOs against Sitophilus oryzae and Callosobruchus chinensis. J. Stored Prod. Res., 39: 293–303.

914

Handbook of Essential Oils

Kobaisy, M., M.R. Tellez, F.E. Dayan, and S.O. Duke, 2002. Phytotoxicity and volatile constituents from leaves of Callicarpa japonica Thunb. Phytochemistry, 61: 37–40. Kobaisy, M., M.R. Tellez, C.L. Webber, F.E. Dayan, K.K. Schrader, and D.E. Wedge, 2001. Phytotoxic and fungitoxic activities of the EO of kenaf (Hibiscus cannabinus L.) leaves and its composition. J. Agric. Food Chem., 49: 3768–3771. Kong, C., F. Hu, T. Xu, and Y. Lu, 1999. Allelopathic potential and chemical constituents of volatile oil from Ageratum conyzoides. J. Chem. Ecol., 25: 2347–2356. Konstantopoulou, I., L. Vassilopoulou, P. Mavragani-Tsipidou, and Z.G. Scouras, 1992. Insecticidal effects of EOs. A study of the effects of EOs extracted from eleven Greek aromatic plants on Drosophila auraria. Experientia, 48: 616–619. Kordali, S., A. Cakir, T.A. Akcin, E. Mete, A. Akcin, T. Aydin, and H. Kilic, 2009. Antifungal and herbicidal properties of EOs and n-hexane extracts of Achillea gypsicola Hub-Mor. and Achillea biebersteinii Afan. (Asteraceae). Indus. Crops Prod., 29: 562–570. Kordali, S., A. Cakir, H. Ozer, R. Cakmakci, M. Kesdek, and E. Mete, 2008. Antifungal, phytotoxic and insecticidal properties of EO isolated from Turkish Origanum acutidens and its three components, carvacrol, thymol and p-cymene. Bioresour. Technol., 99: 8788–8795. Kordali, S., R. Kotan, A. Mavi, A. Cakir, A. Ala, and A. Yildirim, 2005. Determination of the chemical composition and antioxidant activity of the EO of Artemisia dracunculus and of the antifungal and antibacterial activities of Turkish Artemisia absinthium, A. dracunculus, Artemisia santonicum, and Artemisia spicigera EOs. J. Agric. Food Chem., 53: 9452–9458. Kostyukovsky, M., A. Rafaeli, C. Gileadi, N. Demchenko, and E. Shaaya, 2002. Activation of octopaminergic receptors by EO constituents isolated from aromatic plants: Possible mode of action against insect pests. Pest Manag. Sci., 58: 1101–1106. Koul, O., M.J. Smirle, and M.B. Isman, 1990. Asarones from Acorus Calamus oil: Their effect on feeding behavior and dietary utilization in Peridroma saucia. J. Chem. Ecol., 16: 1911–1920. Koul, O., S. Walia, and G. S. Dhaliwal, 2008. EOs as green pesticides: Potential and constraints. Biopestic. Int., 4: 63–84. Lee, B.-H., P.C. Annis, F. Tumaalii, and W.-S. Choi, 2004. Fumigant toxicity of EOs from the Myrtaceae family and 1,8-cineole against 3 major stored-grain insects. J. Stored Prod. Res., 40: 553–564. Lee, B.-H., W.-S. Choi, S.-E. Lee, and B.-S. Park, 2001. Fumigant toxicity of EOs and their constituent compounds towards the rice weevil, Sitophilus oryzae (L.). Crop Prot., 20: 317–320. Lee, C.-H., B.-K. Sung, and H.-S. Lee, 2006. Acaricidal activity of fennel seed oils and their main components against Tyrophagus putrescentiae, a stored-food mite. J. Stored Prod. Res., 42: 8–14. Li, R., and Z.-T. Jiang, 2004. Chemical composition of the EO of Cuminum cyminum L. from China. Flavour Fragr. J., 19: 311–313. Liu, Z.L., and S.H. Ho, 1999. Bioactivity of the EO extracted from Evodia rutaecarpa Hook f. et Thomas against the grain storage insects, Sitophilus zeamais Motsch. and Tribolium castaneum (Herbst). J. Stored Prod. Res., 35: 317–328. Loebe, L., 2001. Olfactory remedies for the evaluation of repellent and attractive properties of EOs against the Pea Aphid Acyrthosiphon pisum. Master Thesis, University of Vienna. López, M.D., M.J. Jordán, and M.J. Pascual-Villalobos, 2008. Toxic compounds in EOs of coriander, caraway and basil active against stored rice pests. J. Stored Prod. Res., 44: 273–278. Lopez, M.D., and M.J. Pascual-Villalobosa, 2010. Mode of inhibition of acetylcholinesterase by monoterpenoids and implications for pest control. Indus. Crops Prod., 31: 284–288. Malkomes, H.-P., 2006. Einfluss von herbizidem Citronella-Öl und Neem (Azadirachtin) auf mikrobielle Aktivitäten im Boden. Gesunde Pflanzen, 58: 205–212. Mills, C., B.J. Cleary, J.F. Gilmer, and J.J. Walsh, 2004. Inhibition of acetylcholinesterase by tea tree oil. J. Pharm. Pharmacol., 56: 375–379. Mishra, A.K., and N.K. Dubey, 1994. Evaluation of some EOs for their toxicity against fungi causing deterioration of stored food commodities. Appl. Environ. Microbiol., 60: 1101–1105. Moretti, M.D.L., G. Sanna-Passino, S. Demontis, and E. Bazzoni, 2002. EO formulations useful as a new tool for insect pest control. AAPS Pharm. Sci. Tech., 3: E13. Mossa, A.-T.H., 2016. Green pesticides: EOs as biopesticides in insect-pest management. J. Environ. Sci. Technol., 9: 354–378. Muller-Riebau, F.J., B.M. Berger, O. Yegen, and C. Cakir, 1997. Seasonal variations in the chemical compositions of EOs of selected aromatic plants growing wild in Turkey. J. Agric. Food Chem., 45: 4821–4825. Mutlu, S., Ö. Atici, N. Esim, and E. Mete, 2010. EOs of catmint (Nepeta meyeri Benth.) induce oxidative stress in early seedlings of various weed species. Acta Physiol. Plant., 33: 943–951.

Use of Essential Oils in Agriculture

915

Negahban, M., S. Moharramipour, and F. Sefidkon, 2006. Chemical composition and insecticidal activity of artemisia scoparia EO against three coleopteran stored-product insects. J. Asia-Pacific Entomol., 9: 381–388. Negahban, M., S. Moharramipour, and F. Sefidkon, 2007. Fumigant toxicity of EO from Artemisia sieberi Besser against three stored-product insects. J. Stored Prod. Res., 43: 123–128. Nerio, L.S., J. Olivero-Verbel, and E.E. Stashenko, 2009. Repellent activity of EOs from seven aromatic plants grown in Colombia against Sitophilus zeamais Motschulsky (Coleoptera). J. Stored Prod. Res., 45: 212–214. Nerio, L. S., J. Olivero-Verbel, and E. Stashenko, 2010. Repellent activity of EOs: A review. Bioresour. Technol., 101: 372–378. Niepel, D., 2000. Repellent properties of EOs against the carrot weevil. Master Thesis, University of Vienna. Njoroge, S.M., H. Koaze, P.N. Karanja, and M. Sawamura, 2005. EO constituents of three varieties of Kenyan sweet oranges (Citrus sinensis). Flavour Fragr. J., 20: 80–85. Obeng-Ofori, D., C.H. Reichmuth, A.J. Bekele, and A. Hassanali, 1998. Toxicity and protectant potential of camphor, a major component of EO of Ocimum kilimandscharicum, against four stored product beetles. Intern. J. Pest Manage., 44: 203–209. Oka, Y., S. Nacar, E. Putievsky, U. Ravid, Z. Yaniv, and Y. Spiegel, 2000. Nematicidal activity of EOs and their components against the root-knot nematode. Phytopathology, 90: 710–715. Pandey, R., A. Kalra, S. Tandon, N. Mehrotra, H.N. Singh, and S. Kumar, 2000. EOs as potent source of nematicidal compounds. J. Phytopathol., 148: 501–502. Papachristos, D.P., and D.C. Stamopoulos, 2002. Repellent, toxic and reproduction inhibitory effects of EO vapours on Acanthoscelides obtectus (Say) (Coleoptera: Bruchidae). J. Stored Prod. Res., 38: 117–128. Paranagama, P.A., K.H.T. Abeysekera, K. Abeywickrama, and L. Nugaliyadde, 2003. Fungicidal and antiaflatoxigenic effects of the EO of Cymbopogon citratus (DC.) Stapf. (lemongrass) against Aspergillus flavus Link. isolated from stored rice. Lett. Appl. Microbiol., 37: 86–90. Park, C., S.-I. Kim, and Y.-J. Ahn, 2003. Insecticidal activity of asarones identified in Acorus gramineus rhizome against three coleopteran stored-product insects. J. Stored Prod. Res., 39: 333–342. Pellati, F., S. Benvenuti, F. Yoshizaki, D. Bertelli, and M.C. Rossi, 2005. Headspace solid-phase microextractiongas chromatography—mass spectrometry analysis of the volatile compounds of Evodia species fruits. J. Chromatogr. A, 1087: 265–273. Pessoa, L.M., S.M. Morais, C.M.L. Bevilaqua, and J.H.S. Luciano, 2002. Anthelmintic activity of EO of Ocimum gratissimum Linn. and eugenol against Haemonchus contortus. Vet. Parasitol., 109: 59–63. Picimbon, J.F., and C. Regnault-Roger, 2008. Composés sémiochimiques volatils, phytoprotection et olfaction: Cibles moléculaires pour la lutte intégrée. In Biopesticides d’Origine Végétale, 2nd ed, C. RegnaultRoger, B.J.R. Philogène, and C. Vincent (eds.). Paris: Lavoisier Tech & Doc. Philogène, B.J.R., C. Regnault-Roger, and C. Vincent, 2005. Botanicals: yesterday and today’s promises. In Biopesticides of Plant Origin, C. Regnault-Roger, B.J.R. Philogène, and C. Vincent (eds.), pp. 1–16. Andover, UK: Intercept. Pitarokili, D., M. Couladis, N. Petsikos-Panayotarou, and O. Tzakou, 2002. Composition and antifungal activity on soil-borne pathogens of the EO of Salvia sclarea from Greece. J. Agric. Food Chem., 50: 6688–6691. Plant Encyclopedia, Aden Earth Zone. 2012. https://www.theplantencyclopedia.org Prajapati, V., A.K. Tripathi, K.K. Aggarwal, and S.P.S. Khanuja, 2005. Insecticidal, repellent and ovipositiondeterrent activity of selected EOs against Anopheles stephensi, Aedes aegypti and Culex quinquefasciatus. Bioresour. Technol., 96: 1749–1757. Price, D.N., and M.S. Berry, 2006. Comparison of effects of octopamine and insecticidal EOs on activity in the nerve cord, foregut and dorsal unpaired median neurons of cockroaches. J. Insect. Physiol., 52: 309–319. Priestley C.M., E.M. Williamson, K.A. Wafford, and D.B. Satelle, 2003. Thymol, a constituent of thyme EOs, is a positive modulator of human GABA and a homo-oligosteric GABA receptor from Drosophila melanogaster. Br. J. Pharmacol., 140: 1363–1372. Rajendran, S., and V. Sriranjini, 2008. Plant products as fumigants for stored-product insect control. J. Stored Prod. Res., 44: 126–135. Razzaghi-Abyaneh, M., M. Shams-Ghahfarokhi, T. Yoshinari, M.-B. Rezaee, K. Jaimand, H. Nagasawa, and S. Sakuda, 2008. Inhibitory effects of Satureja hortensis L. EO on growth and aflatoxin production by Aspergillus parasiticus. Intern. J. Food Microbiol., 123: 228–233. Regnault-Roger, C., 1997. The potential of botanical EOs for insect pest control. Integr. Pest. Manage. Rev., 2: 25–34 Regnault-Roger, C., 2013. EOs in insect control. In Natural Products, K.G. Ramawat, and J.M. Merillon (eds.), pp. 4088–4105. Berlin Heidelberg: Springer-Verlag.

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Regnault-Roger, C., and A. Hamraoui, 1994. Antifeedant effect of Mediterranean plant EOs upon Acanthoscelides obtectus Say (Coleoptera), bruchid of kidneybeans, Phaseolus vulgaris L. In Stored Product Protection, E. Highley, E.J. Wright, H.J. Banks, and B.R. Champs (eds.), vol. 2, pp. 837–840. Wallingford, UK: CAB International. Regnault-Roger, C., A. Hamraoui, M. Holeman, E. Theron, and R. Pinel, 1993. Insecticidal effect of EOs from mediterranean aromatic plants upon Acanthoscelides obtectus Say, Coleopterea, bruchid of kidney bean (Phaseolus vulgaris L.). J. Chem. Ecology, 19(6): 1233–1244. Regnault-Roger, C., and B.J.R. Philogène, 2008. Past and current prospects for the use of botanicals and plant allelochemicals in integrated pest management. Pharm. Biol., 46: 1–12. Regnault-Roger, C., B.J.R. Philogène, and C. Vincent, 2008. Biopesticides d’Origine Végétale, 2nd ed. Paris: Lavoisier Tech & Doc. Regnault-Roger, C., C. Silvy, and C. Alabouvette, 2005. Biopesticides: Réalités et perspectives commerciales. In Enjeux Phytosanitaires pour l’Agriculture et l’Environnement, C. Regnault-Roger (ed.). Paris: Lavoisier Tech & Doc. Regnault-Roger, C., C. Vincent, and J.T. Arnason, 2012a. EOs in insect control: Low-risk products in a highstakes world. Annu. Rev. Entomol., 57: 405–424. Regnault-Roger, C., C. Vincent, and J.T. Arnason, 2012b. Low-risk products in a high-stakes world. Annu. Rev. Entomol., 57: 405–424. Rozzi, N.L., W. Phippen, J.E. Simon, and R.K. Singh, 2002. Supercritical fluid extraction of EO components from lemon-scented botanicals. LWT – Food Sci. Technol., 35: 319–324. Sahaf, B.Z., S. Moharramipour, and M.H. Meshkatalsadat, 2007. Chemical constituents and fumigant toxicity of EO from Carum copticum against two stored product beetles. Insect Sci., 14: 213–218. Saharkhiz, M.J., S. Smaeili, and M. Merikhi, 2010. EO analysis and phytotoxic activity of two ecotypes of Zataria multiflora Boiss. growing in Iran. Nat. Prod. Res., 24: 1598–1609. Salamci, E., S. Kordali, R. Kotan, A. Cakir, and Y. Kaya, 2007. Chemical compositions, antimicrobial and herbicidal effects of EOs isolated from Turkish Tanacetum aucheranum and Tanacetum chiliophyllum var.\chiliophyllum. Biochem. Systematics Ecol., 35: 569–581. Sánchez-Ramos, I., and P. Castañera, 2000. Acaricidal activity of natural monoterpenes on Tyrophagus putrescentiae (Schrank), a mite of stored food. J. Stored Prod. Res., 37: 93–101. Scrivanti, L.R., M.P. Zunino, and J.A. Zygadlo, 2003. Tagetes minuta and Schinus areira EOs as allelopathic agents. Biochem. Systematics Ecol., 31: 563–572. Sertkaya, E., K. Kaya, and S. Soylu, 2010. Acaricidal activities of the EOs from several medicinal plants against the carmine spider mite (Tetranychus cinnabarinus Boisd.\) (Acarina: Tetranychidae). Indus. Crops Prod., 31: 107–112. Shaaya, E., U. Ravid, N. Paster, B. Juven, U. Zisman, and V. Pissarev, 1991. Fumigant toxicity of EOs against four major stored-product insects. J. Chem. Ecol., 17: 499–504. Singh, H.P., D.R. Batish, N. Setia, and R.K. Kohli, 2005. Herbicidal activity of volatile oils from Eucalyptus citriodora against Parthenium hysterophorus. Ann. Appl. Biol., 146: 89–94. Sparagano, O.A.E., J. Pritchard, and D. George, 2016. The future of EOs as pest biocontrol method. In BioBased Plant Oil Polymers and Composites, S.A. Madbouly, C. Zhang, and M.R. Kessler (eds.), pp. 207–211. William Andrew Publishing. Szulc, M., and J. Sobczak, 2017. Formulations of plant oils used in crop protection in selected member states. IX International Scientific Symposium Farm Machinery and Processes Management in Sustainable Agriculture, Lublin, Poland, DOI: 10.24326/fmpmsa.2017.66. Tapondjou, L.A., C. Adler, H. Bouda, and D.A. Fontem, 2002. Efficacy of powder and EO from Chenopodium ambrosioides leaves as post-harvest grain protectants against six-stored product beetles. J. Stored Prod. Res., 38: 395–402. Tassou, C., K. Koutsoumanis, and G.-J.E. Nychas, 2000. Inhibition of Salmonella enteritidis and Staphylococcus aureus in nutrient broth by mint EO. Food Res. Intern., 33: 273–280. Teuscher, E., M.F. Melzig, and U. Lindequist, 2004. Biogene Arzneimittel: Ein Lehrbuch der pharmazeutischen Biologie; 14 Tabellen. Wiss. Verlag-Ges: Stuttgart. Thompson, D.P., 1989. Fungitoxic activity of EO components on food storage fungi. Mycologia, 81: 151–153. Tripathi, A.K., S. Upadhyay, M. Bhuiyan, and P. Bhattacharya, 2009. A review on prospects of EOs as biopesticide in insect-pest management. J. Pharmacog. Phytother., 1: 52–63. TRM Market Biopesticides, 2018. Global Biopesticides Market: Rising Awareness about Environmentally Viable Agriculture Spikes Demand, TMR finds (Feb 2018), https://www.transparencymarketresearch. com/pressrelease/biopesticides-market.htm (accessed 15 August 2018).

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Tunç, İ., B.M. Berger, F. Erler, and F. Dağlı, 2000. Ovicidal activity of EOs from five plants against two storedproduct insects. J. Stored Products Res., 36: 161–168. Tworkoski, T., 2002. Herbicide effects of EOs. Weed Sci., 50: 425–431. Tzortzakis, N.G., and C.D. Economakis, 2007. Antifungal activity of lemongrass (Cympopogon citratus L.\) EO against key postharvest pathogens. Innov. Food Sci. Emer. Technol., 8: 253–258. United states of Food and Drug Administration (US-FDA). 2012. Bad Bug Book: Foodborne Pathogenic Microorganisms and Natural Toxins. Lampel K.A., Al-Khaldi S., Cahill S.M. eds., 2nd edition. Ushir, Y., A. Tatiya, S. Surana, and U. Patil, 2010. Gas chromatography-mass spectrometry analysis and antibacterial activity of EO from aerial parts and roots of Anisomeles indica Linn. Intern. J. Green Pharmacy, 4: 98. Verdeguer, M., M.A. Blázquez, and H. Boira, 2009. Phytotoxic effects of Lantana camara, Eucalyptus camaldulensis and Eriocephalus africanus EOs in weeds of Mediterranean summer crops. Biochem. Syst. Ecol., 37: 362–369. Verdeguer, M., D. García-Rellán, H. Boira, E. Pérez, S. Gandolfo, and M.A. Blázquez, 2011. Herbicidal activity of Peumus boldus and Drimys winterii EOs from Chile. Molecules, 16: 403–411. Vidal R., J. Muchembled, C., Deweer, L. Tournant, N. Corroyer, S. Flammier, 2018. Évaluation de l’intérêt de l’utilisation d’huiles essentielles dans des stratégies de protection des cultures. Innov. Agron., 63(2018), 191–210. Vokou, D., D. Chalkos, G. Karamanlidou, and M. Yiangou, 2002, Activation of soil respiration and shift of the microbial population balance in soil as a response to Lavandula stoechas EO. J. Chem. Ecol., 28: 755–768. Vokou, D., and S. Liotiri, 1999. Stimulation of soil microbial activity by EOs. Chemo. Ecol., 9: 41–45. Walderhaug, M., 2014. Bad Bug Book: Foodborne Pathogenic Microorganisms and Natural Toxins Handbook. Createspace Independent Pub. Wang, J., F. Zhua, X.M. Zhoua, C.Y. Niua, C.L. Leia, 2006. Repellent and fumigant activity of EO from Artemisia vulgaris to Tribolium castaneum (Herbst) (Coleoptera: Tenebrionidae). J. Stored Prod. Res., 42(3): 339–347. Wilson, C.L., J.M. Solar, A. El Ghaouth, and M.E. Wisniewski, 1997. Rapid evaluation of plant extracts and EOs for antifungal activity against botrytis cinerea. Plant Dis., 81: 204–210.

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Essential Oils Used in Veterinary Medicine K. Hüsnü Can Başer and Chlodwig Franz

CONTENTS 25.1 Introduction........................................................................................................................... 919 25.2 Oils Attracting Animals........................................................................................................ 921 25.3 Oils Repelling Animals......................................................................................................... 921 25.4 Oils against Pests................................................................................................................... 922 25.4.1 Insecticidal, Pest Repellent, and Antiparasitic Oils.................................................. 922 25.4.2 Fleas and Ticks.......................................................................................................... 922 25.4.3 Mosquitoes................................................................................................................. 922 25.4.4 Moths......................................................................................................................... 923 25.4.5 Aphids, Caterpillars, and Whiteflies......................................................................... 923 25.4.5.1 Garlic Oil.................................................................................................... 923 25.4.6 Ear Mites................................................................................................................... 923 25.4.7 Antiparasitic............................................................................................................... 923 25.5 Essential Oils Used in Animal Feed.....................................................................................924 25.5.1 Ruminants..................................................................................................................924 25.5.2 Poultry.......................................................................................................................925 25.5.2.1 Studies with CRINA® Poultry....................................................................925 25.5.2.2 Studies with Herbromix®............................................................................926 25.5.3 Pigs............................................................................................................................ 927 25.6 Essential Oils Used in Treating Diseases in Animals...........................................................928 References....................................................................................................................................... 929

25.1 INTRODUCTION Essential oils are volatile constituents of aromatic plants. These liquid oils are generally complex mixtures of terpenoid and/or nonterpenoid compounds. Mono-, sesqui-, and sometimes diterpenoids, phenylpropanoids, fatty acids and their fragments, benzenoids, and so on may occur in various essential oils (Baser and Demirci, 2007). Except for citrus oils obtained by cold pressing, all other essential oils are obtained by distillation. Products obtained by solvent extraction or supercritical fluid extraction are not technically considered as essential oils (Baser, 1995). Essential oils are used in perfumery, food flavoring, pharmaceuticals, and sources of aromachemicals. Essential oils exhibit a wide range of biological activities and 31 essential oils have monographs in the latest edition of the European Pharmacopoeia (Table 25.1). Antimicrobial activities of many essential oils are well documented (Bakkali et al., 2008). Such oils may be used singly or in combination with one or more oils. For the sake of synergism this may be necessary. Although many are generally regarded as safe, essential oils are generally not recommended for internal use. However, their much diluted forms (e.g., hydrosols) obtained during oil distillation as a by-product may be taken orally. 919

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TABLE 25.1 Essential Oil Monographs in the European Pharmacopoeia (6.5 Edition, 2009) English Name

Latin Name

Anise oil Bitter-fennel fruit oil

Anisi aetheroleum Foeniculi amari fructus aetheroleum

Bitter-fennel herb oil

Foeniculi amari herba aetheroleum

Caraway oil Cassia oil

Carvi aetheroleum Cinnamomi cassiae aetheroleum

Cinnamon bark oil, Ceylon

Citronella oil Clary sage oil Clove oil

Cinnamomi zeylanici corticis aetheroleum Cinnamomi zeylanici folium aetheroleum Citronellae aetheroleum Salviae sclareae aetheroleum Caryophylli aetheroleum

Coriander oil Dwarf pine oil Eucalyptus oil Juniper oil Lavender oil

Coriandri aetheroleum Pini pumilionis aetheroleum Eucalypti aetheroleum Juniperi aetheroleum Lavandulae aetheroleum

Lemon oil Mandarin oil Matricaria oil

Lemonis aetheroleum Citri reticulatae aetheroleum Matricariae aetheroleum

Mint oil, partly dementholized

Menthae arvensis aetheroleum partim mentholi privum

Neroli oil (formerly bitter-orange flower oil) Nutmeg oil Peppermint oil

Neroli aetheroleum (formerly Aurantii amari floris aetheroleum) Myristicae fragrantis aetheroleum Menthae piperitae aetheroleum

Pine silvestris oil Rosemary oil Spanish sage oil Spike lavender oil Star anise oil Sweet orange oil

Pini silvestris aetheroleum Rosmarini aetheroleum Salviae lavandulifoliae aetheroleum Spicae aetheroleum Anisi stellati aetheroleum Aurantii dulcis aetheroleum

Tea tree oil

Melaleucae aetheroleum

Thyme oil Turpentine oil, Pinus pinaster type

Thymi aetheroleum Terebinthini aetheroleum ab pinum pinastrum

Cinnamon leaf oil, Ceylon

Plant Name Pimpinella anisum L. fruits Foeniculum vulgare Miller subsp. vulgare var. vulgare Foeniculum vulgare Miller subsp. vulgare var. vulgare Carum carvi L. Cinnamomum cassia Blume (Cinnamomum aromaticum Nees) Cinnamomum zeylanicum Nees Cinnamomum verum J.S. Presl. Cymbopogon winterianus Jowitt Salvia sclarea L. Syzygium aromaticum (L.) Merill et L.M. Perry (Eugenia caryophyllus C.S. Spreng. Bull. et Harr.) Coriandrum sativum L. Pinus mugo Turra. Eucalyptus globulus Labill. Juniperus communis L. meyvesi Lavandula angustifolia P. Mill. (Lavandula officinalis Chaix.) Citrus limon (L.) Burman fil. Citrus reticulata Blanco Matricaria recutita L. (Chamomilla recutita (L.) Rauschert) Mentha canadensis L. (Mentha arvensis L. var. glabrata (Benth.) Fern, Mentha arvensis L. var. piperascens Malinv. ex Holmes) Japanese mint Citrus aurantium L. subsp. aurantium (Citrus aurantium L. subsp. amara Engl.) Myristica fragrans Houtt. Mentha × piperita L. Pinus silvestris L. Rosmarinus officinalis L. Salvia lavandulifolia Vahl. Lavandula latifolia Medik. Illicium verum Hooker fil. Citrus sinensis (L.) Osbeck (Citrus aurantium L. var. dulcis L.) Melaleuca alternifolia (Maiden et Betch) Cheel, Melaleuca linariifolia Smith, Melaleuca dissitiflora F. Mueller, and other species Thymus vulgaris L., T. zygis L. Pinus pinaster Aiton. (Maritime pine)

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Topical applications of some essential oils (e.g., oregano and lavender) in wounds and burns bring about fast recovery without leaving any sign of cicatrix. By inhalation, several essential oils act as a mood changer and have effect especially on respiratory conditions. Several essential oils (e.g., citronella oil) have been used as pest repellents or as insecticides and such uses are frequently encountered in veterinary applications. In recent years, especially after the ban on the use of antibiotics in animal feed in the European Union since January 2006, essential oils have emerged as a potential alternative to antibiotics in animal feed. Essential oils used in veterinary medicine may be classified as follows:

1. Oils attracting animals 2. Oils repelling animals 3. Insecticidal, pest repellent, and antiparasitic oils 4. Oils used in animal feed 5. Oils used in treating diseases in animals.

25.2  OILS ATTRACTING ANIMALS Valeriana oils (and valerianic and isovalerianic acids) and nepeta oils (and nepetalactones) are wellknown feline-attractant oils. Their odor attracts male cats. Douglas fir oil and its monoterpenes have been claimed to attract deer and wild boar (Buchbauer et al., 1994). Dogs are normally drawn to floral oils and usually choose to take these by inhalation only. Monoterpene-rich oils are usually too strong for dogs, with the exception of bergamot, Citrus bergamia. Cats also usually select only floral oils for inhalation. Cats do not have metabolic mechanism to break down essential oils due to the lack of the enzyme glucuronidase. Therefore, they should not be taken by mouth and should not be generally applied topically (Ingraham, 2008).

25.3  OILS REPELLING ANIMALS Peppermint oil (Mentha piperita) repels mice. It can be applied under the sink in the kitchen or applied in staples to prevent mice annoying horses and livestock. A few drops of peppermint oil in a bucket of water used to scrub out a stall and sprinkling a few drops around the perimeter and directly on straw or bedding are said to eliminate or severely curtail the habitation of mice (Scents and Sensibility, 2001). A patent (U.S. Patent 4,961,929) claims that a mixture of methyl salicylate, birch oil, wintergreen oil, eucalyptus oil, pine oil, and pine-needle oil repels dogs. Another patent (U.S. Patent 4,735,803) claims the same using lemon oil and α-terpinyl methyl ether. Another similar formulation (U.S. Patent 4,847,292) claims that a mixture of citronellyl nitrile, citronellol, α-terpinyl methyl ether, and lemon oil repels dogs. A mixture of black pepper and capsicum oils and the oleoresin of rosemary is claimed to repel animals (U.S. Patent 6,159,474). Citronella oil repels cats and dogs (Moschetti, 2003). Repellents alleged to repel cats include allyl isothiocyanate (oil of mustard), amyl acetate, anethole, capsaicin, cinnamaldehyde, citral, citronella, citrus oil, eucalyptus oil, geranium oil, lavender oil, lemongrass oil, menthol, methyl nonyl ketone, methyl salicylate, naphthalene, nicotine, paradichlorobenzene, and thymol. Oil of mustard, cinnamaldehyde, and methyl nonyl ketone are said to be the most potent. Essential oils comprised of 10 g/L solutions of cedarwood, cinnamon, sage, juniper berry, lavender, and rosemary; all of these were potent snake irritants. Brown tree snakes exposed to a 2s burst of aerosol of these oils exhibited prolonged, violent undirected locomotory behavior. In contrast, exposure

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to a 10 g/L concentration of ginger oil aerosol caused snakes to locomote, but in a deliberate, directed manner. The 10 g/L solutions delivered as aerosols of m-anisaldehyde, trans-anethole, 1,8-cineole, cinnamaldehyde, citral, ethyl phenylacetate, eugenol, geranyl acetate, or methyl salicylate acted as potent irritants for brown tree snakes (Boiga irregularis) (Clark and Shivik, 2002).

25.4  OILS AGAINST PESTS 25.4.1 Insecticidal, Pest Repellent, and Antiparasitic Oils The essential oil of bergamot (C. bergamia), anise (Pimpinella anisum), sage (Salvia officinalis), tea tree (Melaleuca alternifolia), geranium (Pelargonium sp.), peppermint (M. piperita), thyme (Thymus vulgaris), hyssop (Hyssopus officinalis), rosemary (Rosmarinus officinalis), and white clover (Trifolium repens) can be used to control certain pests on plants. They have been shown to reduce the number of eggs laid and the amount of feeding damage by certain insects, particularly lepidopteran caterpillars. Sprays made from tansy (Tanacetum vulgare) have demonstrated a repellent effect on imported cabbageworm on cabbage, reducing the number of eggs laid on the plants. Teas made from wormwood (Artemisia absinthium) or nasturtiums (Nasturtium spp.) are reputed to repel aphids from fruit trees, and sprays made from ground or blended catnip (Nepeta cataria), chives (Allium schoenoprasum), feverfew (Tanacetum parthenium), marigolds (Calendula, Tagetes, and Chrysanthemum spp.), or rue (Ruta graveolens) have also been used by gardeners against pests that feed on leaves (Moschetti, 2003).

25.4.2 Fleas and Ticks Dogs, cats, and horses are plagued by fleas and ticks. One to two drops of citronella or lemongrass oils added to the shampoo will repel these pests. Alternatively, 4–5 drops of cedarwood oil and pine oil is added to a bowl of warm water, and a bristle hairbrush is soaked with this solution to brush the pet down with it. Eggs and parasites gathered in the brush are rinsed out. This is repeated several times. This solution can be used similarly for livestock after adding citronella and lemongrass oils to this mixture. Flea collar can be prepared by a mixture of cedarwood (Juniperus virginiana), lavender (Lavandula angustifolia), citronella (Cymbopogon winterianus [Java]), thyme oils, and 4–5 garlic (Allium sativum) capsules. This mixture is thinned with a teaspoonful of ethanol and soaked with a collar or a cotton scarf. This is good for 30 days (Scents and Sensibility, 2001). Ticks can be removed by applying 1 drop of cinnamon or peppermint oil on Q-tip by swabbing on it. Carvacrol-rich oil (64%) of Origanum onites and carvacrol was found to be effective against the tick Rhipicephalus turanicus. Pure carvacrol killed all the ticks following 6 h of exposure, while 25% and higher concentrations of the oil were effective in killing the ticks by the 24 h posttreatment (Coskun et al., 2008).

25.4.3 Mosquitoes Catnip oil (N. cataria) containing nepetalactones can be used effectively as a mosquito repellent. It is said to be 10 times more effective than DEET (N,N-diethyl-meta-toluamide) (Moschetti, 2003). Juniperus communis berry oil is a very good mosquito repellent. Ocimum volatile oils including camphor, 1,8-cineole, methyl eugenol, limonene, myrcene, and thymol strongly repelled mosquitoes (Regnault-Roger, 1997). Citronella oil repels mosquitoes, biting insects, and fleas. Essential oils of Zingiber officinale and R. officinalis were found to be ovicidal and repellent, respectively, toward three mosquito species (Prajapati et al., 2005). Root oil of Angelica sinensis and ligustilide was found to be mosquito repellent (Wedge et al., 2009).

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25.4.4 Moths Cedarwood oil is used in mothproofing. A large number of patents have been assigned to the preservation of cloths from moths and beetles: application of a solution containing clove (Syzygium aromaticum) essential oil on woolen cloth, filter paper containing Juniperus rigida oil, and tablets of p-dichlorobenzene mixed with essential oils to be placed in wardrobe.

25.4.5 Aphids, Caterpillars, and Whiteflies 25.4.5.1  Garlic Oil Essential oils are effective in insect pest control (Regnault-Roger, 1997).

25.4.6  Ear Mites Peppermint oil is applied to a Q-tip and swabbed inside of the ear.

25.4.7 Antiparasitic There is a patent (U.S. Patent 6,800,294) on an antiparasitic formulation comprising eucalyptus oil (Eucalyptus globulus), cajeput oil (Melaleuca cajuputi), lemongrass oil, clove bud oil (S. aromaticum), peppermint oil (M. piperita), piperonyl, and piperonyl butoxide. The formulation can be used for treating an animal body, in the manufacture of a medicament for treating ectoparasitic infestation of an animal, or for repelling parasites. Two essential oils derived from L. angustifolia and Lavandula × intermedia were investigated for any antiparasitic activity against the human protozoal pathogens Giardia duodenalis and Trichomonas vaginalis and the fish pathogen Hexamita inflata, all of which have significant infection and economic impacts. The study has demonstrated that low (≤1%) concentrations of L. angustifolia and Lavandula × intermedia oil can completely eliminate T. vaginalis, G. duodenalis, and H. inflata in vitro. At 0.1% concentration, L. angustifolia oil was found to be slightly more effective than Lavandula × intermedia oil against G. duodenalis and H. inflata (Moon et al., 2006). The antiparasitic properties of essential oils from A. absinthium, Artemisia annua, and Artemisia scoparia were tested on intestinal parasites Hymenolepis nana, Lamblia intestinalis, Syphacia obvelata, and Trichocephalus muris (Trichuris muris). Infested white mice were injected with 0.01 mL/g of the essential oils (6%) twice a day for 3 days. The effectiveness of the essential oils was observed in 70%–90% of the tested animals (Chobanov et al., 2004). Parasites, such as head lice and scabies, as well as internal parasites, are repelled by oregano oil (86% carvacrol). The oil can be added to soaps, shampoos, and diluted in olive oil for topical applications. By taking a few drops daily under the tongue, one can gain protection from waterborne parasites, such as Cryptosporidium and Giardia. Internal dosages also are effective in killing parasites in the body (http://curingherbs.com/wild_oregano_oil.htm) (Foster, 2002). Essential oils from Pinus halepensis, Pinus brutia, Pinus pinaster, Pinus pinea, and Cedrus atlantica were tested for molluscicidal activity against Bulinus truncatus. The oil from C. atlantica was found the most active (LC 50 = 0.47 ppm). Among their main constituents, α-pinene, β-pinene, and myrcene exhibited potent molluscicidal activity (LC 50 = 0.49, 0.54, and 0.56 ppm, respectively). These findings have important application of natural products in combating schistosomiasis (Lahlou, 2003). Origanum essential oils have exhibited differential degrees of protection against myxosporean infections in gilthead and sharpsnout sea bream tested in land-based experimental facilities (Athanassopoulou et al., 2004a,b).

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25.5  ESSENTIAL OILS USED IN ANIMAL FEED Essential oils can be used in feed as appetite stimulant, stimulant of saliva production, gastric and pancreatic juice production enhancer, and antimicrobial and antioxidant to improve broiler performance. Antimicrobial effects of essential oils are well documented. Essential oils due to their potent nature should be used as low as possible levels in animal nutrition. Otherwise, they can lead to feed intake reduction, gastrointestinal tract (GIT) microflora disturbance, or accumulation in animal tissues and products. Odor and taste of essential oils may contribute to feed refusal; however, encapsulation of essential oils could solve this problem (Gauthier, 2005). Generally, Gram-positive bacteria are considered more sensitive to essential oils than Gramnegative bacteria because of their less complex membrane structure (Lis-Balchin, 2003). Carvacrol, the main constituent of oregano oils, is a powerful antimicrobial agent (Baser, 2008). It asserts its effect through the biological membranes of bacteria. It acts through inducing a sharp reduction of the intercellular adenosine triphosphate (ATP) pool through the reduction of ATP synthesis and increased hydrolysis. Reduction of the membrane potential (transmembrane electrical potential), which is the driving force of ATP synthesis, makes the membrane more permeable to protons. A high level of carvacrol (1 mM) decreases the internal pH of bacteria from 7.1 to 5.8 related to ion gradients across the cell membrane. One millimolar of carvacrol reduces the internal potassium (K) level of bacteria from 12 mmol/mg of cell protein to 0.99 mmol/mg in 5 min. K plays a role in the activation of cytoplasmic enzymes and in maintaining osmotic pressure and in the regulation of cytoplasmic pH. K efflux is a solid indication of membrane damage (Ultee et al., 1999). It has been shown that the mode of action of oregano oils is related to an impairment of a variety of enzyme systems, mainly involved in the production of energy and the synthesis of structural components. Leakage of ions, ATP, and amino acids also explains the mode of action. Potassium and phosphate ion concentrations are affected at levels below the minimum inhibitory concentration (MIC) concentration (Lambert et al., 2001).

25.5.1 Ruminants A recent review compiled information on botanicals including essential oils used in ruminant health and productivity (Rochfort et al., 2008). Unfortunately, there are few reports on the effects of essential oils and natural aromachemicals on ruminants. It was demonstrated that the consumption of terpene volatiles such as camphor and α-pinene in “tarbush” (Flourensia cernua) effected feed intake in sheep (Estell et al., 1998). In vitro and in vivo antimicrobial activities of essential oils have been demonstrated in ruminants (Elgayyar et al., 2001; Wallace et al., 2002; Cardozo et al., 2005; Moreira et al., 2005). Synergistic antinematodal effects of essential oils and lipids were demonstrated (Ghisalberti, 2002). Other nematocidal volatiles reported are as follows: benzyl isothiocyanate (goat), ascaridole (goat and sheep) (Ghisalberti, 2002; Githiori et al., 2006), geraniol, eugenol (Chitwood, 2002; Githiori et al., 2006), menthol, and 1,8-cineole (Chitwood, 2002). Methylsalicylate, the main component of the essential oil of Gaultheria procumbens (wintergreen), is topically used as emulsion in cattle, horses, sheep, goats, and poultry in the treatment of muscular and articular pain. The recommended dose is 600 mg/kg bw twice a day. The duration of treatment is usually less than 1 week (EMA, 1999). It is included in Annex II of Council Regulation (EEC) No. 2377/90 as a substance that does not need an maximum residue limit (MRL) level. G. procumbens should not be used as flavoring in pet food since salicylates are toxic to dogs and cats. As cats metabolize salicylates much more slowly than other s­pecies, they are more likely to be overdosed. Use of methylsalicylate in combination with anticoagulants such as warfarin can result in adverse interactions and bleedings (Chow et al., 1989; Yip et al., 1990; Ramanathan, 1995; Tam et al., 1995). The essential oil of L. angustifolia (Lavandulae aetheroleum) is used in veterinary medicinal products for topical use together with other plant extracts or essential oils for antiseptic and healing purposes. The product is used in horses, cattle, sheep, goats, rabbits, and poultry. It is included in

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Annex II of Council Regulation (EEC) No. 2377/90 as a substance that does not need an MRL level (EMEA, 1999; Franz et al., 2005). The outcomes of in vitro studies investigating the potential of P. anisum essential oil as a feed additive to improve nutrient use in ruminants are inconclusive, and more and larger preferably in vivo studies are necessary for evaluation of efficacy (Franz et al., 2005). Carvacrol, carvone, cinnamaldehyde, cinnamon oil, clove bud oil, eugenol, and oregano oil have resulted in a 30%–50% reduction in ammonia N concentration in diluted ruminal fluid with a 50:50 forage concentrate diet during the 24 h incubation (Busquet et al., 2006). Carvacrol has been suggested as a potential modulator of ruminal fermentation (Garcia et al., 2007).

25.5.2  Poultry 25.5.2.1  Studies with CRINA® Poultry Dietary addition of essential oils in a commercial blend (CRINA Poultry) showed a decreased Escherichia coli population in ileocecal digesta of broiler chickens. Furthermore, in high doses, a significant increase in certain digestive enzyme activities of the pancreas and intestine was observed in broiler chickens (Jang et al., 2007). In another study, CRINA Poultry was shown to control the colonization of the intestine of broilers with Clostridium perfringens, and the stimulation of animal growth was put down to this development (Losa, 2001). Commercial essential oil blends CRINA Poultry and CRINA Alternate were tested in broilers infected with viable oocysts of mixed Eimeria spp. It was concluded that these essential oil blends may serve as an alternative to antibiotics and/or ionophores in mixed Eimeria infections in noncocci-vaccinated broilers, but no benefit of essential oil supplementation was observed for vaccinated broilers against coccidia (Oviedo-Rondon et al., 2006). 25.5.2.1.1  Other Studies Supplementation of 200 ppm essential oil mixture (EOM) that included oregano, clove, and anise oils (no species name or composition given!) in broiler diets was said to significantly improve the daily live weight gain and feed conversion ratio (FCR) during a growing period of 5 weeks (Ertas et al., 2006). Similar results were obtained with 400 mg/kg anise oil (composition not known!) (Ciftci et al., 2005). A total of 50 and 100 mg/kg of feed of oregano oil* were tested on broilers. No growth-promoting effect was observed. At 100 mg/kg of feed, antioxidant effect was detected on chicken tissues (Botsoglou et al., 2002a). Positive results were also reported for oregano oil added in poultry feed (Bassett, 2000). Antioxidant activities of rosemary and sage oils on lipid oxidation of broiler meat have been shown. Following dietary administration of rosemary and sage oils to the live birds, a significant inhibition of lipid peroxidation was reported in chicken meat stored for 9 days (Lopez-Bote et al., 1998). A dietary supplementation of oregano essential oil (300 mg/kg) showed a positive effect on the performance of broiler chickens experimentally infected with Eimeria tenella. Throughout the experimental period of 42 days, oregano essential oil exerted an anticoccidial effect against E. tenella, which was, however, lower than that exhibited by lasalocid. Supplementation with dietary oregano oil to E. tenella–infected chickens resulted in body weight gains and FCRs not differing from the noninfected group, but higher than those of the infected control group and lower than those of chickens treated with the anticoccidial lasalocid (Giannenas et al., 2003). Inclusion of oregano oil at 0.005% and 0.01% in chicken diets for 38 days resulted in a significant antioxidant effect in raw and cooked breast and thigh muscle stored up to 9 days in refrigerator (Botsoglou et al., 2002b). * Oregano essential oil was in the form of a powder called Orego-Stim. This product contains 5% oregano essential oil (Ecopharm Hellas S.A., Kilkis, Greece) and 95% natural feed grade inert carrier. The oil of Origanum vulgare subsp. hirtum used in this product contains 85% carvacrol and thymol.

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Oregano oil (55% carvacrol) exhibited a strong bactericidal effect against lactobacilli and following the oral administration of the oil MIC values of amikacin, apramycin, and streptomycin and neomycin against E. coli strains increased (Horosova et al., 2006). An in vitro assay measuring the antimicrobial activity of essential oils of Coridothymus capitatus, Satureja montana, Thymus mastichina, Thymus zygis, and Origanum vulgare was carried out against poultry origin strains of E. coli, Salmonella enteritidis, and Salmonella essen and pig origin strains of enterotoxigenic E. coli, Salmonella choleraesuis, and Salmonella typhimurium. O. vulgare (MIC ≤ 1% v/v) oil showed the highest antimicrobial activity against the four strains of Salmonella. It was followed by T. zygis oil (MIC ≤ 2% v/v). T. mastichina oil inhibited all the microorganisms at the highest concentration, 4% (v/v). Monoterpenic phenols carvacrol and thymol showed higher inhibitory capacity than the monoterpenic alcohol linalool. The results confirmed potential application of such oils in the treatment and prevention of poultry and pig diseases caused by Salmonella (Penalver et al., 2005). In another study, groups of male, 1-day-old Lohmann broilers were given maize–soya bean meal diets, with oils extracted from thyme, mace, and caraway or coriander, garlic, and onion (0, 20, 40, and 80 mg/kg) for 6 weeks. The average daily gain and FCR were not different between the broilers fed with the different oils; meat was not tainted with flavor or smell of the oils (Vogt and Rauch, 1991). 25.5.2.2  Studies with Herbromix® Essential oils from oregano herb (O. onites), laurel leaf (Laurus nobilis), sage leaf (Salvia fruticosa), fennel fruit (Foeniculum vulgare), myrtle leaf (Myrtus communis), and citrus peel (rich in limonene) were mixed and formulated as feed additive after encapsulation. It is marketed in Turkey as poultry feed under the name Herbromix. The following three in vivo experiments with this product were recently accomplished. 25.5.2.2.1  In Vivo Experiment 1 In this study, 1250 sexed 1-day-old broiler chicks obtained from a commercial hatchery were randomly divided into five treatment groups of 250 birds each (negative control, antibiotic, and essential oil combination [EOC] at 3 levels). Each treatment group was further subdivided into five replicates of 50 birds (25 males and 25 females) per replicate. Commercial EOC at three different levels (24, 48, and 72 mg) and antibiotic (10 mg avilamycin) per kg were added to the basal diet. There were significant effects of dietary treatments on body weight, feed intake (except at day 42), FCR, and carcass yield at 21 and 42 days. Body weights were significantly different between the treatments. Birds fed on diet containing 48 mg essential oil/kg being the highest, and this treatment was followed by chicks fed on the diet containing 72 mg essential oil/kg, antibiotic, negative control, and 24 mg essential oil/kg at day 42. Supplementation with 48 mg EOC/kg to the broiler diet significantly improved the body weight gain, FCR, and carcass yield compared to other dietary treatments on 42 days of age. EOC may be considered as a potential growth promoter in the future of the new era, which agrees with producer needs for increased performance and today’s consumer demands for environment-friendly broiler production. The EOC can be used cost-effectively when its cost is compared with antibiotics and other commercially available products in the market. 25.5.2.2.2  In Vivo Experiment 2 In this study, 1250 sexed 1-day-old broiler chicks were randomly divided into five treatment groups of 250 birds each (negative control, organic acid, probiotic, and EOC at two levels). Each treatment group was further subdivided into five replicates of 50 birds (25 males and 25 females) per replicate. The oils in the EOC were extracted from different herbs growing in Turkey. The organic acid at 2.5 g/ kg diet, the probiotic at 1 g/kg diet, and the EOC at 36 and 48 mg/kg diet were added to the basal diet. The results obtained from this study indicated that the inclusion of 48 mg EOC/kg broiler diet significantly improved the body weight gain, FCR, and carcass yield of broilers compared to organic

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acid and probiotic treatments after a growing period of 42 days. The EOC may be considered as a potential growth promoter like organic acids and probiotics for environment-friendly broiler production. 25.5.2.2.3  In Vivo Experiment 3 The aim of this study was to examine the effect of essential oils and breeder age on growth performance and some internal organs’ weight of broilers. A total of 1008 unsexed 1-day-old broiler chicks (Ross-308) originating from young (30 weeks) and older (80 weeks) breeder flocks were randomly divided into three treatment groups of 336 birds each, consisting of control and two EOMs at a level of 24 and 48 mg/kg diet. There were no significant effects of dietary treatments on body weight gain of broilers at days 21 and 42. On the other hand, there were significant differences on the feed intake at days 21 and 42. The addition of 24 or 48 mg/kg EOM to the diet reduced significantly the feed intake compared to the control. The groups fed with the added EOM had significantly better FCR than the control at days 21 and 42. Although there was no significant effect of broiler breeder age on body weight gain at day 21, significant differences were observed on body weight gain at 42 days of age. Broilers originating from young breeder flock had significantly higher body weight gain than those originating from old breeder flock at 42 days of age. No difference was noticed for carcass yield, liver, pancreas, proventriculus, gizzard, and small intestine weight. Supplementation with EOM to the diet in both levels significantly decreased mortality at days 21 and 42. The results indicated that the Herbromix may be considered as a potential growth promoter. However, more trials are needed to determine the effect of essential oil supplementation to diet on the performance of broilers with regard to variable management conditions including different stress factors, essential oils and their optimal dietary inclusion levels, active substances of oils, dietary ingredients, and nutrient density (Alcicek et al., 2003, 2004; Bozkurt and Baser, 2002a,b; Cabuk et al., 2006a,b).

25.5.3  Pigs CRINA® Pigs was tested on pigs. The results for the first 21-day period showed that males grew faster, ate less, and exhibited superior FCR compared to females. Although female carcass weight was higher, males had a significantly lower carcass fat than females (Losa, 2001). The addition of fennel (F. vulgare) and caraway (Carum carvi) oils was not found beneficial for weaned piglets. In feed choice conditions, fennel oil caused feed aversion (Schoene et al., 2006). Oregano oil was found to be beneficial for piglets (Molnar and Bilkei, 2005). In a preliminary investigation, the effects of low-level dietary inclusion of rosemary, garlic, and oregano oils on pig performance and pork quality were carried out. Unfortunately, no information on the species from which the oils were obtained and their composition existed in the paper. The pigs appeared to prefer the garlic-treated diet, and the feed intake and the average daily gain were significantly increased although no difference in the feed efficiency was observed. Carcass and meat quality attributes were unchanged, although a slight reduction of lipid oxidation was noted in oregano-fed pork. Since the composition of the oils is not clear, it is not possible to evaluate the results (Janz et al., 2007). A study revealed that the inclusion of essential oil of oregano in pigs’ diet significantly improved the average daily weight gain and FCR of the pigs. Pigs fed with the essential oils had higher carcass weight, dressing percentage, and carcass length than those fed with the basal and antibioticsupplemented diet. The pigs that received the essential oil supplementation had a significantly lower fat thickness. Also lean meat and ham portions from these pigs were significantly higher. Therefore, the use of Origanum essential oil as feed additive improves the growth of pigs and has greater positive effects on carcass composition than antibiotics (Onibala et al., 2001). Ropadiar®, an essential oil of the oregano plant, was supplemented in the diet of weaning pigs as alternative for antimicrobial growth promoters (AMGPs), observing its efficacy on the performance of the piglets. Ropadiar liquid contains 10% oregano oil and has been designed to be added to water. Compared to the negative control (without AMGP), Ropadiar improved performance only during the

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first 14 days after weaning. Based on the results of this trial, it cannot be argued about the usefulness of Ropadiar as an alternative for AMGP in diets of weanling pigs. However, its addition in prestarter diets could improve performance of these animals (Krimpen and Binnendijk, 2001). The objective of another trial was to ascertain the effect on nutrient digestibilities and N-balance, as well as on parameters of microbial activity in the GIT of weaned pigs after adding oregano oil to the feed. The apparent digestibility of crude nutrients (except fiber) and the N-balance of the weaned piglets in this study were not influenced by feeding piglets restrictively with this feed additive. By direct microbiological methods, no influence of the additive on the gut flora could be found (Moller, 2001). The inclusion of essential oil of spices in the pigs’ diet significantly improved the average daily weight gain and FCR of the pigs in Groups 3, 4, and 5, as compared to Groups 1 and 2 (P