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Fatty acids chemistry, synthesis, and applications
 9780128095218, 0128095210, 9780128095447, 012809544X

Table of contents :
Content: 1. History of Fatty Acids Chemistry 2. Naturally Occurring Fatty Acids: Source, Chemistry, and Uses 3. Epoxy Fatty Acids: Chemistry and Biological Effects 4. Acetylenic Epoxy Fatty Acids: Chemistry, Synthesis and their Pharmaceutical Applications 5. Carbocyclic Fatty Acids: Chemistry and Biological Properties 6. Modification of Oil Crops to produce Fatty Acids for Industrial Applications 7. Microbial Production of Fatty Acids 8. Chemical Derivatization of Castor Oil and Their Industrial Utilization 9. Chemical Modification of High Free Fatty Acid Oils for Biodiesel Production 10. Synthesis of Sugar-Fatty Acid Esters and their Industrial Utilization 11. Fatty Acids-based Surfactants and their Uses 12. The Role of Fatty Acids in Cosmetic Technology 13. Chemistry of Long Chain a, B-unsaturated Fatty Acid and Reactions Thereof 14. Estolides: Synthesis and Applications 15. An Efficient, Multigram Synthesis of Dietary Cis- and Trans-Octadecenoic (18:1) Fatty Acids 16. Advancement in Chromatographic and Spectroscopic Analyses of Dietary Fatty Acids 17. Mass Spectrometry in the Analysis of Fatty Acids and Derivatives 18. Crystallization of Fats and Fatty Acids in Edible Oils and Structure Determination

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Fatty Acids

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Fatty Acids Chemistry, Synthesis, and Applications

Edited by

Moghis U. Ahmad Jina Pharmaceuticals, Inc., Libertyville, IL, United States

Academic Press and AOCS Press Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1800, San Diego, CA 92101-4495, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom Copyright r 2017 AOCS Press. Published by Elsevier Inc. All rights reserved. Published in cooperation with American Oil Chemists’ Society www.aocs.org Director, Content Development: Janet Brown. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress ISBN: 978-0-12-809521-8 For Information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Andre´ G. Wolff Acquisition Editor: Nancy Maragioglio Editorial Project Manager: Billie Jean Fernandez Production Project Manager: Lisa Jones Cover Designer: Victoria Pearson Typeset by MPS Limited, Chennai, India

Contents List of Contributors Meet the Editor Preface

1.

xvii xix xxi

History of Fatty Acids Chemistry Gary R. List, James A. Kenar and Bryan R. Moser 1.1 Introduction 1.2 Early Fatty Acid History 1.3 Major Developments in the Oleochemical Industry 1.3.1 Fat Splitting 1.3.2 Catalytic Hydrogenation 1.3.3 Fatty Acid Distillation 1.3.4 Fatty Alcohols 1.3.5 Estolides 1.3.6 Dimer and Trimer Cyclic Fatty Acids 1.3.7 Hydroformylation of Fatty Acids 1.3.8 Ozonolysis of Fatty Acids and Triglycerides 1.4 Contributions of Analytical Chemistry to Fatty Acids 1.5 Recent Developments in Fatty Acids 1.6 Conclusion References

2.

2 2 9 9 10 11 11 12 13 14 14 15 16 17 18

Naturally Occurring Fatty Acids: Source, Chemistry, and Uses James A. Kenar, Bryan R. Moser and Gary R. List 2.1 Introduction 2.2 Production of Naturally Occurring Fatty Acids 2.2.1 Chemical Splitting 2.2.2 Lipase Splitting 2.3 Purification of Fatty Acids 2.3.1 Simple Distillation 2.3.2 Fractional Distillation 2.3.3 Molecular Distillation 2.3.4 Crystallization 2.3.5 Urea Fractionation

24 28 29 30 31 31 32 35 35 36

v

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Contents

2.4 Sources and Types of Naturally Occurring Fatty Acids 2.4.1 Saturated Fatty Acids 2.4.2 Unsaturated Fatty Acids 2.4.3 Hydroxy Fatty Acids 2.4.4 Acetylenic Fatty Acids 2.4.5 Allenic and Cumulenic Fatty Acids 2.5 Chemistry of Naturally Occurring Fatty Acids 2.5.1 Reactions at the Carboxylic Acid Group 2.5.2 Reactions at Unsaturated Sites 2.6 Conclusion References

3.

37 38 39 43 45 47 49 50 57 71 71

Epoxy Fatty Acids: Chemistry and Biological Effects Arnis Kuksis and Waldemar Pruzanski 3.1 Introduction 3.2 Natural Occurrence and Structure of Epoxy Fatty Acids 3.2.1 Oleic and Linoleic Acid Monoepoxides and Hydroxides 3.2.2 Arachidonic Acid Monoepoxides 3.2.3 Eicosapentaenoic Acid and Docosahexaenoic Acid Monoepoxides 3.3 Chemical Synthesis 3.3.1 Direct Epoxidation 3.3.2 Chemo-Enzymatic Perhydrolysis 3.3.3 Other Chemo-Enzymatic Epoxidations 3.4 Biosynthesis of Epoxy Fatty Acids 3.4.1 Oxygenases and Lipoxygenases 3.4.2 Peroxygenases 3.4.3 Cytochrome P450-Like Oxygenases 3.5 Analysis of Epoxy Fatty Acids 3.5.1 Resolution of Regioisomers 3.5.2 Resolution of Enantiomers 3.5.3 GC/MS and LC/MS Identification of Lipid Epoxides 3.6 Biological Effects 3.6.1 Lipid Signaling 3.6.2 Cellular Effects 3.6.3 Systemic Effects 3.7 Pathological Effects 3.7.1 Toxicity 3.7.2 Inflammation and Pain 3.7.3 Angiogenesis and Cardiovascular Disease 3.7.4 Cancer 3.8 Conclusion Abbreviations References

83 84 84 85 85 88 88 89 90 90 91 91 92 94 95 97 103 104 104 105 107 108 108 108 110 111 112 112 113

Contents

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vii

Acetylenic Epoxy Fatty Acids: Chemistry, Synthesis, and Their Pharmaceutical Applications Valery M. Dembitsky and Dmitry V. Kuklev 4.1 4.2 4.3 4.4

Introduction Occurrence Epoxy Acetylenic Fatty Acids in Nature Lipids Containing Epoxy Acetylenic Fatty Acids Epoxy Acetylenic Furanoid and Thiophene Fatty Acid and Derivatives 4.5 Pyranone and Macrocyclic Epoxides 4.6 Acetylenic Cyclohexanoid Epoxy Fatty Acids 4.7 Determination or Epoxy Acetylenic Lipids 4.8 Synthesis of Epoxy Acetylenic Lipids 4.9 Concluding Remarks References Further Reading

5.

121 122 125 128 129 130 131 136 141 142 146

Carbocyclic Fatty Acids: Chemistry and Biological Properties Moghis U. Ahmad, Shoukath M. Ali, Ateeq Ahmad, Saifuddin Sheikh and Imran Ahmad 5.1 Introduction 5.2 Naturally Occurring Cyclopropene Fatty Acids 5.2.1 The Halphen Test 5.2.2 Isolation of Cyclopropene Fatty Acids From Seed Oils 5.2.3 Chemical Characterization 5.3 Synthesis and Characterization of Sterculic Acid 5.3.1 Characterization of Dihydrosterculic Acid 5.3.2 Total Synthesis of cis-Cyclopropane Fatty Acids 5.3.3 Deuterated Cyclopropene Fatty Acids 5.4 Biosynthesis of Cyclopropane and Cyclopropene Fatty Acids 5.5 Mass Spectrometry of Cyclopropene Fatty Acids 5.5.1 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropene Fatty Acids 5.5.2 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropane Fatty Acids 5.6 Physiological Properties of Cyclopropene Fatty Acids 5.7 Cyclopropaneoctanoic Acid 2-Hexyl in Human Adipose Tissue and Serum 5.7.1 Cyclopropaneoctanoic Acid 2-Hexyl in Patients With Hypertriglyceridemia 5.8 Leishmania Cyclopropane Fatty Acid Synthetase 5.8.1 Leishmania: A Fungal Infection 5.9 Conclusion References Further Reading

148 150 151 152 152 156 158 160 161 163 165 166 171 171 173 175 176 177 178 179 185

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Modification of Oil Crops to Produce Fatty Acids for Industrial Applications John L. Harwood, Helen K. Woodfield, Guanqun Chen and Randall J. Weselake 6.1 Introduction 6.2 Key Aspects of Plant Oil Biosynthesis 6.3 Major Oil Crops 6.3.1 Oil Palm (Elaeis guineensis) 6.3.2 Soybean (Glycine max) 6.3.3 Brassica Oilseed Species (Brassica napus, Brassica rapa, Brassica oleracea, Brassica carinata) 6.3.4 Sunflower (Helianthus annuus) 6.4 Minor Oil Crops 6.4.1 Alfalfa (Medicago sativa, Medicago falcata) 6.4.2 Almond (Prunus dulcis, Prunus amygdalus, Amygdalus communis) 6.4.3 Avocado (Persea americana, Persea gratissima) 6.4.4 Blackcurrant (Ribes niger) 6.4.5 Borage (Borago officinalis) 6.4.6 Borneo Tallow (Shorea stenoptera) 6.4.7 Camelina (Camelina sativa) (Section 6.5 Also) 6.4.8 Castor (Ricinus communis) 6.4.9 Cocoa (Theobroma cacao) 6.4.10 Coconut (Cocos nucifera) 6.4.11 Coriander (Coriandrum sativum) 6.4.12 Cottonseed (Gossypium hirsutum, Gossypium barbadense) 6.4.13 Crambe (Crambe abyssinica, Crambe hispanica) (Section 6.5 Also) 6.4.14 Cuphea spp. 6.4.15 Dimorphotheca (Dimorphotheca pluvialis) 6.4.16 Echium (Echium plantagineum) 6.4.17 Flax (Linum usitatissimum) 6.4.18 Hazelnut (Corylus avellana) 6.4.19 Jatropha curcas (See Section 6.5) 6.4.20 Jojoba (Simmondsia chinensis) 6.4.21 Lesquerella (Lesquerella fendleri) (See Section 6.5) 6.4.22 Maize (Corn; Zea mays) 6.4.23 Meadowfoam (Limnanthes alba) 6.4.24 Mustard (Brassica alba, Brassica carinata, Brassica hirta, Brassica juncea, Brassica nigra) 6.4.25 Oats (Avena sativa) 6.4.26 Olive (Olea europaea) 6.4.27 Peanut (Ground Nut, Arachis hypogaea) 6.4.28 Pine Nuts (Pinus spp.) 6.4.29 Poppy (Papaver somniferum)

188 189 194 194 197 201 206 208 209 209 209 209 209 209 211 211 211 212 212 212 212 212 213 213 213 213 213 214 214 215 215 215 215 215 216 216 216

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6.4.30 Rice (Oryza sativa) Bran Oil 216 6.4.31 Safflower (Carthamus tinctorius) 217 6.4.32 Shea (Butyrospermum parkii, Shea Butter, Karate Butter) 217 6.4.33 Tall 217 6.4.34 Tung (Aleurites fordii) 217 6.4.35 Vernonia Oils 218 6.5 Emerging Industrial Oil Crops 218 6.6 Prospects for Production of Industrial Oils in Vegetative Tissue 222 Acknowledgments 223 References 223 Further Reading 236

7.

Microbial Production of Fatty Acids Colin Ratledge and Casey Lippmeier 7.1 Introduction 7.2 The Process of Lipid Accumulation in Oleaginous Microorganisms 7.3 Economic Considerations—Heterotrophic Microorganisms 7.4 Economic Considerations—Phototrophic Microorganisms 7.5 Production of PUFAs 7.5.1 Nutritionally Important Fatty Acids—Background Information 7.5.2 Production of Gamma-Linolenic Acid (GLA 18:3 n-6) 7.5.3 Production of Arachidonic Acid (ARA 20:4 n-6) 7.5.4 Production of Docosahexaenoic Acid (DHA 22:6 n-3) 7.5.5 Production of Eicosapentaenoic Acid (EPA 20:5 n-3) 7.5.6 Production of EPA/DHA Mixtures as Alternatives to Fish Oils 7.6 Safety Aspects 7.7 Future Prospects References

8.

237 241 244 248 251 251 255 258 259 260 264 266 268 270

Chemical Derivatization of Castor Oil and Their Industrial Utilization Rachapudi B.N. Prasad and Bhamidipati V.S.K. Rao 8.1 Introduction 8.2 Derivatives of Castor Oil Based on Unsaturation of Ricinoleic Acid 8.2.1 Hydrogenated Castor Oil 8.2.2 Epoxy Castor Oil 8.2.3 Ozonolysis of Castor Oil 8.2.4 Preparation of 9,10,12-Trihydroxy Octadecanoic Acid 8.2.5 Halogenated Derivatives of Castor Oil 8.2.6 Novel Derivatives of Ricinoleic Acid Employing Metathesis Reaction

280 282 282 282 284 285 285 285

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8.3 Derivatives of Castor Oil Based on Hydroxy Functionality of Ricinoleic Acid 8.3.1 Dehydrated Castor Oil and Dehydrated Castor Oil Fatty Acids 8.3.2 Sulfated Castor Oil (Turkey Red Oil) 8.3.3 Acetylated Castor Oil 8.3.4 Castor OilBased Estolides 8.3.5 Castor OilBased Polymer Products 8.3.6 Potent Hydroxy Derivatives of Ricinoleic Acid 8.4 Derivatives Based on Ester Functionality of Castor Oil 8.4.1 Hydroxy Fatty Acid Esters 8.4.2 Castor OilBased Biodiesel 8.4.3 Preparation of Ricinoleyl Alcohol 8.4.4 Ricinoleic AcidBased Amides 8.4.5 Ethanolamides of Castor Oil Fatty Acids 8.5 Unique Derivatives of Castor Oil 8.5.1 Castor OilBased Dimer Acids 8.5.2 10-Undecenoic Acid and Heptaldehyde 8.5.3 Sebacic Acid and 2-Octanol References

9.

286 286 288 288 289 289 291 291 291 292 293 293 293 293 293 294 295 296

Chemical Modification of High Free Fatty Acid Oils for Biodiesel Production Godlisten G. Kombe 9.1 Introduction 9.2 Production of Biodiesel 9.2.1 Types of Feedstocks 9.2.2 The Potential of High FFA Feedstocks in Biodiesel Production 9.2.3 Challenges of Processing High FFA Feedstocks 9.3 Chemical Modification of High FFA Feedstocks for Biodiesel 9.3.1 Potential Processes for Modification of High FFA Feedstocks 9.4 Conclusion and Recommendations References Further Reading

305 306 306 307 308 309 309 321 323 327

10. Synthesis of Sugar Fatty Acid Esters and Their Industrial Utilizations Bianca Pe´rez, Sampson Anankanbil and Zheng Guo 10.1 Introduction 10.2 Synthesis of Sugar Fatty Acid Esters 10.2.1 Chemical Synthesis of Sugar Fatty Acid Esters 10.2.2 Enzymatic Synthesis of Sugar Fatty Acid Esters

329 331 331 333

Contents

10.3 Physicochemical Properties of Sugar Fatty Acid Esters 10.3.1 Emulsifying Stability and Foaming Ability 10.3.2 Toxicity and Biodegradability 10.4 Industrial Applications of Sugar Fatty Acid Esters 10.5 Conclusion Acknowledgment Abbreviations References Further Reading

xi 343 344 345 346 347 348 348 348 354

11. Fatty AcidsBased Surfactants and Their Uses Douglas G. Hayes 11.1 Introduction 11.1.1 Biobased Surfactants: A Growing Market 11.2 Biobased Surfactants Are a Robust Product for an Oleochemical-Based Biorefinery 11.3 Oleochemical Feedstocks for Surfactant Synthesis 11.4 Sustainability of Oleochemical-Based Surfactants: Truths and Myths 11.5 Green Manufacturing of Biobased Surfactants 11.6 Ionic Surfactants 11.6.1 Methyl Ester Sulfonates 11.6.2 Esterquats 11.6.3 Amino AcidBased Surfactants 11.6.4 Others 11.7 Ester-Based Nonionic Surfactants 11.7.1 Glyceride Esters 11.7.2 Ethoxylates of Fatty Acids and Partial Glycerides 11.7.3 Sugar Esters 11.7.4 Polyol Esters 11.8 Ether and Amide-Based Nonionic Surfactants 11.8.1 Alkyl Polyglucosides 11.8.2 N-Alkyl N-Methyl Glucamine 11.8.3 Others 11.9 Zwitterionic (Amphoteric) Surfactants 11.9.1 Phospholipids 11.9.2 Betaines 11.10 Glycolipid Biosurfactants 11.11 Conclusion References

355 355 359 361 367 368 369 369 369 370 371 372 372 372 372 373 373 373 374 374 374 374 375 376 378 379

12. The Role of Fatty Acids in Cosmetic Technology Gary R. Kelm and Randall R. Wickett 12.1 Introduction 12.2 Cosmetic and Personal Care Product Formulation Types

385 386

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12.3 Cosmetic and Personal Care Product Categories 12.4 Reviewed Fatty Acid Derivatives and Overview of Uses in Cosmetic and Personal Care Products 12.4.1 Fatty Alcohols 12.4.2 Anionic and Nonionic Surfactants Based Upon Fatty Acids 12.4.3 Fatty Amines and Quaternary Ammonium Compounds 12.4.4 Esters of Fatty Acids 12.5 Cleansing 12.6 Vehicles/Solvents 12.7 Rheological Modification of Suspensions and Sticks 12.8 Stabilization of Emulsions 12.9 Skin Emollients and Hair Conditioners 12.10 Conclusion References

388 391 392 393 393 393 394 395 397 399 401 402 402

13. Chemistry of Long-Chain α,β-Unsaturated Fatty Acid and Reactions Thereof Abdul Rauf and Mohammad F. Hassan 13.1 Introduction 13.2 Synthesis of α,β-Unsaturated Fatty Acids 13.3 Reactions of α,β-Unsaturated Fatty Acids/Esters 13.3.1 BrominationDehydrobromination 13.3.2 Cyclopropanation 13.3.3 Hypohalogenation 13.3.4 Peracid Oxidation 13.3.5 Allylic Halogenations 13.3.6 Nitrogen, Oxygen, Sulfur Derivatives of α,β-Unsaturated Fatty Acids/Esters 13.3.7 Other Derivatives 13.3.8 α,β-Epoxy Compounds 13.4 Applications 13.5 Conclusion Acknowledgment References Abbreviations

405 406 407 407 408 409 410 412 414 422 425 425 426 427 427 430

14. Estolides: Synthesis and Applications Steven C. Cermak, Terry A. Isbell, Jakob W. Bredsguard and Travis D. Thompson 14.1 Introduction 14.2 Synthesis 14.2.1 Free-Acid Estolides 14.2.2 Estolide 2-Ethylhexyl Esters

432 435 436 438

Contents

14.2.3 Coco-Oleic Estolide 2-Ethylhexyl Esters (One-Step Process) 14.2.4 Coco-Oleic Dimer and Coco-Oleic Trimer Plus Estolides 14.2.5 Commercial Estolide 2-Ethylhexyl Ester (SE7B) 14.3 Identification 14.3.1 GC Analysis 14.3.2 Acid Value 14.3.3 Nuclear Magnetic Resonance (NMR) Spectroscopy 14.4 Basic Physical Properties of Oleic-Based Estolides and Esters 14.4.1 Gardner Color 14.4.2 Viscosity and Viscosity Index 14.4.3 Pour Point and Cloud Point 14.4.4 Oxidation Tests 14.4.5 NOACK Evaporative Loss 14.5 Estolides (SE7B), Base Oil, and Motor Oil Properties—Applications 14.5.1 Performance Properties 14.5.2 Estolide Application-Based Motor Oil SE7B—Field Test 14.6 Conclusion References

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440 440 443 444 444 447 447 449 449 451 454 456 465 466 467 471 472 473

15. An Efficient, Multigram Synthesis of Dietary cis- and trans-Octadecenoic (18:1) Fatty Acids Moghis U. Ahmad 15.1 Introduction 15.2 Organic Synthesis of Unsaturated Fatty Acids 15.3 Fatty Acids Containing One Acetylene Bond 15.3.1 Synthesis of Δ3-Acetylenic (Octadec-3-Ynoic) Acid 15.3.2 Synthesis of Δ4-Acetylenic (Octadec-4-Ynoic) Acid 15.3.3 Synthesis of Δ5-Acetylenic (Octadec-5-Ynoic) Acid 15.3.4 Synthesis of Δ6-Acetylenic (Octadec-6-Ynoic) Acid 15.3.5 Synthesis of Δ7-Acetylenic (Octadec-7-Ynoic) Acid 15.3.6 Synthesis of Δ8-Acetylenic (Octadec-8-Ynoic) Acid 15.3.7 Synthesis of Δ9-Acetylenic (Octadec-9-Ynoic) Acid 15.3.8 Synthesis of Δ10-Acetylenic (Octadec-10-Ynoic) Acid 15.3.9 Synthesis of Δ11-Acetylenic (Octadec-11-Ynoic) Acid 15.3.10 Synthesis of Δ12-Acetylenic (Octadec-12-Ynoic) Acid 15.3.11 Synthesis of Δ13-Acetylenic (Octadec-13-Ynoic) Acid 15.3.12 Synthesis of Δ14-Acetylenic (Octadec-14-Ynoic) Acid

478 480 481 481 482 483 484 486 487 488 488 490 490 491 492

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15.3.13 Synthesis of Δ15-Acetylenic (Octadec-15-Ynoic) Acid 15.3.14 Synthesis of Δ16-Acetylenic (Octadec-16-Ynoic) Acid 15.4 Partial Hydrogenation of Acetylenic Acid and Structure Determination 15.5 Reduction of Acetylenic Acid to cis-Olefinic Acid 15.6 Reduction of Acetylenic Acid to trans-Olefinic Acid 15.7 High-Performance Liquid Chromatography Analyses 15.8 Conclusion References

494 495 495 496 497 498 501 502

16. Advancement in Chromatographic and Spectroscopic Analyses of Dietary Fatty Acids Magdi M. Mossoba, Sanjeewa R. Karunathilaka, Jin K. Chung and Cynthia T. Srigley 16.1 Introduction 16.2 Gas Chromatography With Flame Ionization Detection 16.3 Fourier-Transform Infrared Spectroscopy 16.3.1 Infrared Spectroscopy 16.3.2 Attenuated Total Reflection Spectroscopy 16.3.3 Negative Second Derivative ATR-FT-IR Official Method 16.3.4 Novel Portable ATR- and Transmission-Mode FT-IR Devices 16.4 FT-Near-Infrared Spectroscopy in Conjunction With Partial Least Squares 16.5 Conclusion References

505 506 510 510 510 511 513 514 525 525

17. Mass Spectrometry in the Analysis of Fatty Acids and Derivatives Yu Lin, Ming Guan, Lin Li, Yangyang Zhang and Zhenwen Zhao 17.1 17.2 17.3 17.4

Introduction Extraction of Fatty Acids (FAs) and Derivatives Fatty Acids (FAs) Analysis by Mass Spectrometry Arachidonic Acid (AA) and Its Derivatives Analysis by Mass Spectrometry 17.5 Triacylglycerols (TAGs) Analysis by Mass Spectrometry 17.6 Glycerophospholipids and Sphingolipids Analysis by Mass Spectrometry 17.7 Double Bounds Position Analysis by Mass Spectrometry 17.8 Future Perspective Acknowledgment References

529 531 532 532 533 534 535 536 536 536

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18. Crystallization of Fats and Fatty Acids in Edible Oils and Structure Determination Michael A. Rogers 18.1 Nucleation and Crystal Growth of Fatty Acids & TAGs 18.1.1 Super Cooling and Nucleation 18.1.2 Crystal Growth 18.2 Lipid Polymorphism 18.2.1 Lipid Mesophase Polymorphism 18.2.2 Crystalline Polymorphism 18.3 Nanostructure and Lipid Domains 18.4 Microstructure and Fractal Assembly 18.5 Modified Fatty Acids and Their Gels 18.6 Conclusion Acknowledgments References Index

541 542 544 546 546 548 549 552 553 555 555 555 561

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List of Contributors Ateeq Ahmad Jina Pharmaceuticals, Inc., Libertyville, IL, United States Imran Ahmad Jina Pharmaceuticals, Inc., Libertyville, IL, United States Moghis U. Ahmad Jina Pharmaceuticals, Inc., Libertyville, IL, United States Shoukath M. Ali Jina Pharmaceuticals, Inc., Libertyville, IL, United States Sampson Anankanbil Aarhus University, Aarhus, Denmark Jakob W. Bredsguard Biosynthetic Technologies, Irvine, CA, United States Steven C. Cermak USDA, Agricultural Research Service, Peoria, IL, United States Guanqun Chen University of Alberta, Edmonton, AB, Canada Jin K. Chung U.S. Food and Drug Administration, College Park, MD, United States Valery M. Dembitsky National Scientific Center of Marine Biology, Vladivostok, Russia Ming Guan Institute of Chemistry Chinese Academy of Sciences, Beijing, P.R. China; University of Chinese Academy of Sciences, Beijing, P.R. China Zheng Guo Aarhus University, Aarhus, Denmark John L. Harwood Cardiff University, Cardiff, United Kingdom Mohammad F. Hassan Aligarh Muslim University, Aligarh, Uttar Pradesh, India Douglas G. Hayes University of Tennessee, Knoxville, TN, United States Terry A. Isbell USDA, Agricultural Research Service, Peoria, IL, United States Sanjeewa R. Karunathilaka U.S. Food and Drug Administration, College Park, MD, United States Gary R. Kelm University of Cincinnati, Cincinnati, OH, United States James A. Kenar National Center for Agricultural Utilization Research, Peoria, IL, United States Godlisten G. Kombe The University of Dodoma, Dodoma, Tanzania Dmitry V. Kuklev University of Michigan Medical School, Ann Arbor, MI, United States Arnis Kuksis University of Toronto, Toronto, ON, Canada Lin Li Institute of Chemistry Chinese Academy of Sciences, Beijing, P.R. China; University of Chinese Academy of Sciences, Beijing, P.R. China Yu Lin Institute of Chemistry Chinese Academy of Sciences, Beijing, P.R. China

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Casey Lippmeier DSM Nutritional Products, Columbia, MD, United States Gary R. List G.R. List Consulting, Washington, IL, United States Bryan R. Moser National Center for Agricultural Utilization Research, Peoria, IL, United States Magdi M. Mossoba U.S. Food and Drug Administration, College Park, MD, United States Bianca Pe´rez Aarhus University, Aarhus, Denmark Rachapudi B.N. Prasad CSIR-Indian Institute of Chemical Technology, Hyderabad, Telangana, India Waldemar Pruzanski University of Toronto, Toronto, ON, Canada Bhamidipati V.S.K. Rao CSIR-Indian Institute of Chemical Technology, Hyderabad, Telangana, India Colin Ratledge University of Hull, Hull, United Kingdom Abdul Rauf Aligarh Muslim University, Aligarh, Uttar Pradesh, India Michael A. Rogers University of Guelph, Guelph, ON, Canada Saifuddin Sheikh Jina Pharmaceuticals, Inc., Libertyville, IL, United States Cynthia T. Srigley U.S. Food and Drug Administration, College Park, MD, United States Travis D. Thompson Biosynthetic Technologies, Irvine, CA, United States Randall J. Weselake University of Alberta, Edmonton, AB, Canada Randall R. Wickett University of Cincinnati, Cincinnati, OH, United States Helen K. Woodfield Cardiff University, Cardiff, United Kingdom Yangyang Zhang Institute of Chemistry Chinese Academy of Sciences, Beijing, P.R. China Zhenwen Zhao Institute of Chemistry Chinese Academy of Sciences, Beijing, P.R. China; University of Chinese Academy of Sciences, Beijing, P.R. China

Meet the Editor Dr. Moghis U. Ahmad has obtained his PhD in Chemistry (1978) from AMU, Aligarh, India; and did postdoctoral research at the Department of Biochemistry & Biophysics, Texas A&M University, College Station, Texas, United States and Department of Food Science and Technology, Oregon State University, Corvallis, Oregon, United States. He has extensive experience in basic and applied lipid research and development. He is an expert in the synthesis of lipids and their related products. He has developed and successfully marketed many novel lipid products for the chemical, pharmaceutical, and biotechnology industries. His research is detailed at great length in more than 60 research publications in peer-reviewed journals and book chapters, and in more than 30 patents and patent applications. Most of his contributions remain the company proprietary. He has recently edited two best seller AOCS books, namely Lipids in Nanotechnology (2011) and Polar Lipids: Biology, Chemistry, and Technology (2015). His stature is recognized internationally. He is an elected Fellow of the Royal Society of Chemistry (2011) and AOCS (2014), and the recipient of the prestigious Alton E. Bailey Award (2016) and Stephen S. Chang Award (2017). He chaired the AOCS Phospholipids Division (200911), and is a member of the AOCS Books and Special Publication Committee. He also serves the executive committee of the International Lecithin and Phospholipids Society (ILPS). Currently, he is Vice President of Chemical Technology & Manufacturing at Jina Pharmaceuticals, Inc., Libertyville, Illinois, United States.

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Preface Fatty Acids, esterified to glycerol, are the main constituents of oils and fats. The oils and fats are the renewable resources for the chemical industry. The industrial exploitation of oils and fats, both for food and oleochemicals, is based on chemical modification of both the carboxyl group and unsaturation present in fatty acids. The oleochemicals could add value to existing crops and provide market for new crops, and research leads to novel fatty acids derivatives. The oleochemical production involves reaction at the carboxyl group, with the chain length, and at unsaturation of the fatty acid chain to give products of the desired structure and properties. Introducing functionality to the alkyl chain through known chemical reactions leads to novel compounds with commercial potential. The carboxyl groups and the unsaturated centers generally react independently, but when they are in proximity, they might react through neighboring group participation. In enzymatic reactions, the reactivity of the carboxyl group can be influenced by the presence of double bond in close proximity. The coverage in this book is selective, focusing on industrially important fatty acids, their chemistry and synthesis. Historical perspective of important developments in the chemistry of fatty acids in the last 100 years is presented. The main emphasis of this book is on enzymatic and chemical synthesis of fatty acids and derivatives; naturally occurring fatty acids, their purification and preparation for various applications; presence of unusual cyclic fatty acids like epoxy fatty acids and carbocyclic fatty acids in seed oils and their chemical and biological properties; natural and synthetic acetylenic epoxide and their industrial importance; microbial production of fatty acids; biosynthesis of vegetable oils and process improvement, new plant sources to meet future world needs of fatty acids; industrial importance of castor oil and derivatives; crystallization of fatty acids in edible oils and their structure; free fatty acid oils for biodiesel production; advancement in synthesis of sugar fatty acid esters and their applications; fatty acidsbased surfactants; fatty acids in Cosmetic Technology; chemistry of long-chain α,β-unsaturated fatty acid and derivatives; synthesis of different types of estolides as next generation of high-performance synthetic lubricant; synthesis of dietary cis- and trans-octadecenoic (18:1) fatty acids present in partially hydrogenated vegetable oils; chromatographic and spectroscopic

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analyses of dietary fatty acids; mass spectrometrybased methods for the analyses of fatty acids and derivatives. This book serves as reference manual to new generation of lipid scientists and researchers, useful for oleochemical industries, a valuable teaching resources for undergraduate and graduate students interested in the field of chemistry of oils, fats, and fatty acids, food chemistry, cosmetics and personal care products, and pharmaceuticals. This book also serves as a valuable reference and resource for those interested moving in the field of chemistry and technology of fatty acids. The goal in writing this book is to gather writing from many of the leaders in the field who had published one or several articles in various aspects of fatty acids chemistry. The authors have publications in the field of oils, fats, and fatty acids and are imminently qualified to summarize their own work and related work in their field of expertise. It is hoped that the readers will find it valuable to read and this will help them to understand the field of oils, fats, and fatty acids, and their utilization in oleochemical industries. I would like to thank all contributors for their magnificent work in the collection of research publications and their devotion to presenting accurate and detailed scientific information. The assistance from Academic Press (Elsevier) and AOCS Press is greatly appreciated with special thanks to Billie Jean Fernandez and Janet Brown. Moghis U. Ahmad

Chapter 1

History of Fatty Acids Chemistry Gary R. List1, James A. Kenar2 and Bryan R. Moser2 1

G.R. List Consulting, Washington, IL, United States, 2National Center for Agricultural Utilization Research, Peoria, IL, United States

Chapter Outline 1.1 Introduction 1.2 Early Fatty Acid History 1.3 Major Developments in the Oleochemical Industry 1.3.1 Fat Splitting 1.3.2 Catalytic Hydrogenation 1.3.3 Fatty Acid Distillation 1.3.4 Fatty Alcohols 1.3.5 Estolides 1.3.6 Dimer and Trimer Cyclic Fatty Acids

2 2 9 9 10 11 11 12

1.3.7 Hydroformylation of Fatty Acids 1.3.8 Ozonolysis of Fatty Acids and Triglycerides 1.4 Contributions of Analytical Chemistry to Fatty Acids 1.5 Recent Developments in Fatty Acids 1.6 Conclusion References

14 14 15 16 17 18

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 Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture (USDA). USDA is an equal opportunity provider and employer. USDA prohibits discrimination in all its programs and activities on the basis of race, color, national origin, age, disability, and where applicable, sex, marital status, familial status, parental status, religion, sexual orientation, genetic information, political beliefs, reprisal, or because all or part of an individual’s income is derived from any public assistance program. (Not all prohibited bases apply to all programs.) Persons with disabilities who require alternative means for communication of program information (Braille, large print, audiotape, etc.) should contact USDA’s TARGET Center at (202) 720-2600 (voice and TDD). To file a complaint of discrimination, write to USDA, Director, Office of Civil Rights, 1400 Independence Avenue, S.W., Washington, DC. 20250-9410, or call (800) 795-3272 (voice) or (202) 720-6382 (TDD). USDA is an equal opportunity provider and employer.

Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00001-5 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

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Fatty Acids

1.1 INTRODUCTION The history of fatty acids is a complex one with discoveries often coinciding. However, a number of landmark developments that had significant impact are covered here. The historian is often challenged by who discovered what and when. Fortunately, most of the major events and discoveries surrounding the chemistry of fatty acids are well documented in journal literature. The application of these discoveries in industry is often times difficult to discern because they are mainly documented in the patent literature, which can be time consuming to search, difficult to interpret, and most patents contain references to other patents, which in turn must be examined for prior art. A chronological summary of important discoveries in fatty acid, oleochemical, and triacylglycerol chemistry is shown in Table 1.1. This list is compiled from various sources including the open literature, patents, and chronological compilations from Blank (1942) and the Cyberlipid website (Anonymous, 2016). The first comprehensive book written in English that dealt with fats, oils, and waxes appeared in 1895 (Benedikt and Lewkowitsch, 1895). This book was published as an English translation of Benedikt’s German book on the subject and was subsequently revised numerous times up until 1927. The 1895 edition serves as a valuable historical reference for many of the important early discoveries in fatty acid chemistry, although, the original literature references contained therein can be difficult to locate. Eugene Blank (Blank, 1942) reported a chronological list of important dates in the history of fats and waxes through 1915. The chronology stopped at the year 1915 because events beyond this date were considered too new to have assumed an historical perspective at time the list was published. A review of fatty acid history would be incomplete without reference to Klare Markley’s extensive fivevolume set on the chemistry of fatty acids appearing in 1964 (Markley, 1964). In 1979 Everett Pryde edited a book that covered the fatty acid literature up to 1979 (Pryde, 1979). A recent review describing the contributions of Wilhelm Heintz (181780) is an excellent resource for the early events in fatty acid chemistry (Ramberg, 2013). The aim of this chapter is to provide a brief historical perspective of the events and developments in the field of fats and oils that pertain to fatty acid chemistry and their subsequent application. It is hoped that this chapter serves as a guide for the location of papers that provide further details concerning the history of fatty acids.

1.2 EARLY FATTY ACID HISTORY Fatty acids have been used by man for thousands of years and specifically in the preparation of soap. The ancient Babylonians were using soap as early as 2500 BC. The Old Testament scriptures mention soap in several passages. An excellent review of the soap industry is found in the book on soaps by Spitz (2004). By AD 800900, the soap industry was well established in

History of Fatty Acids Chemistry Chapter | 1

TABLE 1.1 Chronological Summary of Important Discoveries in Fatty Acid, Oleochemical, and Triacylglycerol Chemistry Year

Event

800

Soap produced in Germany

900

Soap produced in France

1600

Soap expands in France

1768

Candles exported to Great Britain

1779

Discovery of glycerol

1801

First oil mill in United States

1811

Bleaching with bone black

1814

Butyric acid discovered

1816

Saponification discovered

1817

Stearic acid discovered

1818

Valeric, caproic, oleic acids isolated

1819

Elaidizination of oleic acid

1823

Chevreul publishes Chemistry of Fats and Oils

1825

Candles patented in France

1825

Distillation of fatty acids

1828

Method for separation of solid and liquid fatty acids

1829

Splitting of tallow with sulfuric acid

1833

Saponification of fats with lime/pressure

1841

Myristic acid discovered

1844

First synthesis of a trigylceride

1844

Linoleic acid discovered

1848

Behenic acid discovered, ricinoleic acid discovered

1849

Lauric acid discovered

1849

Erucic acid discovered

1851

Autoclave for saponification

1852

Polymorphism discovered

1852

Interesterification discovered

1853

Term gylceride first used

1855

Correct structure of glycerol (Continued )

3

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Fatty Acids

TABLE 1.1 (Continued) Year

Event

1855

Margaric acid a mixture

1869

Margarine discovered

1879

Soxhlet extraction

1879

Saponification index

1880

Bleaching patented

1881

Hydroxy myristic acid discovered

1883

Chill roller patented

1884

Iodine value reported

1886

Diene structure of linoleic acid

1886

Tax on margarine

1887

Triene acid in hempseed oil

1889

Death of Chevreul

1892

First acetylenic acid reported

1894

Correct structure of ricinoleic acid

1895

Engine runs on peanut oil

1895

First book on the chemistry of fats and oils

1895

First English book on fats

1895

Lead salts for separation of solid/liquid triglycerides

1897

Discovery of hydrogenation/gas phase

1898

Fat-splitting patented

1898

Correct structure of oleic acid

1898

Liquid chromatography reported

1900

Benzenestearosulphonic acid as fat-splitting catalyst

1903

Hydrogenation in liquid phase

1904

First cyclopentyl acid discovered

1906

Hydrogenation of whale oil

1907

Reagent for splitting of fats

1911

Crisco shortening marketed

1915

Hydrogenation patented

1918

Continuous centrifuge patented (Continued )

History of Fatty Acids Chemistry Chapter | 1

TABLE 1.1 (Continued) Year

Event

1920

Dimer acids discovered

1921

Separation of solid and liquid fatty acids

1923

Continuous refining patented

1923

Process for lecithin

1924

Industrial interesterification patented

1927

Method for classifying fats

1929

Isomerization during hydrogenation reported

1929

Essential fatty acids discovered

1930

Votator patented

1931

X-ray diffraction used for fatty acids

1931

Continuous fat-splitting patented

1932

Spray shortening

1933

Distillation of fatty acids patented

1933

High-ratio shortenings

1934

Synthesis of oleic acid

1934

First solvent extraction plant in United States

1934

Centrifugal refining introduced

1936

Distillation of fatty acids patented

1936

Primex shortening marketed

1937

Conjugation of linoleic acid by alkali

1937

Synthesis of linoleic acid

1940

Hilditch published Chemical Constitution of Fats

1940

Fatty amines/nitriles patented

1942

DHA from fish oil

1942

Chronology of fats, oils, waxes

1944

Comparison of fat-splitting catalysts

1945

Bailey published Industrial Oil and Fat Products

1945

Displacement analysis for fatty acids reported

1945

Relative rates of fatty acid oxidation reported

1947

Swiftning shortening marketed (Continued )

5

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Fatty Acids

TABLE 1.1 (Continued) Year

Event

1948

Directed interesterification reported

1948

Bailey published cottonseed book

1949

Mechanism of hydrogenation reported

1950

Bailey published melting and solidification of fats

1950

Markley published soybeans and products, 2 volumes

1951

Second edition of Bailey’s book published

1951

Thin-layer chromatography introduced

1952

GC introduced

1953

Death of Bailey

1954

Eckey published third edition of Vegetable Fats and Oils

1954

First soft margarine—Chiffon

1955

Synthesis of mixed acid triglyceride

1955

Golden Fluffo shortening marketed

1956

Stahl advances thin-layer chromatography

1960

Theory of glyceride structures proposed

1961

Hydrogenated winterized soybean oil marketed

1963

Nobel prize for Ziegler-Natta catalyst

1964

Last volume edition of Bailey’s book

1964

Olefin disproportination reported

1964

Markley published five-volume set on fatty acids

1965

Death of Hilditch

1974

Metathesis for synthesis of mono- and dicarboxylic acids

1980

Fatty acids published AOCS Press

1993

First trait modified soybean oil commercialized

2003

Trans fat labeling law

2004

Metathesis reviewed

2004

Metathesis for long-chain dicarboxylic acids

2005

Nobel Prize for Grubb’s olefin metathesis

History of Fatty Acids Chemistry Chapter | 1

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Germany and France and by the 1600s had greatly expanded. The US soap industry is over 200 years old and has undergone much change. As early as 1714 Benjamin Franklin assisted in his father’s soap and candle business and in colonial America, candle production was growing and 500,000 pounds of candles were exported to the West Indies and Great Britain. Over the years, numerous new soap products such as bar, laundry, and deodorant soaps as well as various detergent products have been introduced. More recently, skin care products based on soaps have entered the personal care market. Although many bar soap brands have been introduced, only a few brands such as Cashmere Bouquet (1872), Ivory (1879), Lifebuoy (1887), Camay (1928), Woodbury (1899), and Palmolive (1898) have survived. Despite the long use of fatty acids in soap manufacturing, their structure, composition, and chemistry was not well understood due to the slow development of analytical and purification techniques required for their separation, purification, and identification. While inorganic chemistry had made several key advances during this time, by the early 1800s, the understanding of organic molecules and their chemistry was still in its infancy. Although glycerol was discovered in 1779 by the Swedish chemist Carl Scheele, it would be another 40 years before the nature of fats and oils would be understood. Scheele had reacted fat with lead oxide and isolated a viscous liquid, which became known as “Scheele’s sweet principle,” but the remaining fatty acid salt byproducts were not characterized. A pioneer in organic chemistry was Michel Chevreul (17861889) (Costa, 1962). Chevreul began his research by examining soap samples around the year 1811 and by 1818 he had discovered a number of fatty acids and had elucidated the chemistry of saponification by showing that fats and oils consisted of three fatty acid molecules esterified to one glycerol molecule. Nearly 50 years later, Chevreul while studying Scheele’s product isolated glycerol in the water phase for which he coined the term glycerin. However, the chemical formula for glycerin was not determined until 1855 by Charles-Adolphe Wu¨rtz (Kenar, 2007). Chevreul’s research was published in Annales di Chemie over the period of 181318. In 1823 his complete research on fats and oils was published under the title “Recherches chimiques sur les gras d orignine animale.” This work was republished in 1886 to commemorate his 100th birthday as “Chimiques sur les corps gras originine animale.” Although republished in 1886, it had never been translated from ancient French to English until Albert Dijkstra translated “A chemical study of oils and fats of animal origin” in 2009 to mark the 100th anniversary of the American Oil Chemist’s Society (Dijkstra, 2009a). Chevreul was one of the first to use a variety of new techniques such as elemental analysis, melting point determination, fractional solubility, and crystallization as a means of identifying compounds (Dijkstra, 2009a,b). Chevreul identified a fatty acid, named margaric acid (heptadecanoic acid, 17:0), which he thought to be pure substance.

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Fatty Acids

Wilhelm Heintz (181780) later showed that Chevreul’s identification of margaric acid was actually a mixture of palmitic (16:0) and stearic acids (18:0) (Ramberg, 2013). The difficulty in separating fatty acids at the time that have similar elemental composition and melting points no doubt accounted for the confusion and illustrates the difficulties organic chemists of this period faced in identifying new compounds using limited tools and crude analytical techniques. From about 184050, a number of fatty acids were discovered, the synthesis of triglycerides was accomplished, and the basic phenomena of polymorphism and interesterification of fats and oils were recognized. Wilhelm Heintz was a major contributor to fatty acid chemistry. Heintz held a position at a small university in Germany (Halle) beginning in 1851 and began to study animal fats, which he called “the fat kingdom.” He expanded upon Chevreul’s work by developing improved methods to conduct elemental analysis and determine more accurate melting points, both of which were major advances in lipid chemistry (Ramberg, 2013). Heintz was an extremely productive chemist, and authored over 200 publications on physiological chemistry, mineral analysis, and improved methods for elemental analysis and organic chemistry. As mentioned previously, Chevreul introduced some novel approaches to lipid chemistry including elemental analysis and fractional solution/crystallization, and used melting point to identify and judge the purity of fats and fatty acids. At the time, Heintz began his research on fatty acids, others had discovered a number of fats and fatty acids, all of which were defined by their melting point and chemical composition. Heintz was the first to question melting point as a measure of purity for fats and fatty acids and showed that margaric acid, described by Chevreul, was really an impure mixture of palmitic and stearic acids. He then turned his attention to butter where only four fatty acids had been identified. Heintz isolated four additional fatty acids to bring the total to eight. From this, he suggested that as a general rule, the saponification of fats contain only acids whose carbon numbers are divisible by four. His investigations on Spermaceti showed that melting point depression can occur in mixtures of fatty acids. From this discovery, Heintz concluded that the long accepted method for preparing pure compounds by repeated crystallization until the melting point no longer changed was inadequate to identify a fatty acid. From 1880 to 1900, a number of discoveries were made that had everlasting impact on fatty acid chemistry, including Soxhlet extraction, the saponification index, and iodine value. Unusual fatty acids were discovered, including fatty acids containing the hydroxyl, acetylenic, and cyclopentenyl structures. The structures of unsaturated fatty acids (ricinoleic, oleic, linoleic, linolenic) were also reported in the 1890s. Although polymorphism was discovered in 1852, the nature of the phenomenon was not fully understood until the late 1920s when Thomas Malkin

History of Fatty Acids Chemistry Chapter | 1

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introduced X-ray methods to explain why trigylcerides may exhibit multiple melting points (Malkin, 1954). Malkin’s work later came under criticism after he reported four polymorphic forms of tristearin. Both Bailey and Lutton maintained that only three polymorphic forms existed for tristearin and Lutton conclusively proved that the fourth form does not exist (Bailey et al., 1945; Lutton, 1945).

1.3 MAJOR DEVELOPMENTS IN THE OLEOCHEMICAL INDUSTRY The modern oleochemical manufacturing industry was extensively reviewed in a book by Gunstone and Hamilton (2001). Included are chapters on basic oleochemicals, amine- and anionic-based surfactants, lubricants and hydraulic fluids, biofuels, coatings and inks, analysis, new chemistry, and the environment. Applications are well covered and referenced. Normann Sonntag authored a number of review articles appearing in the Journal of the American Oil Chemists’ Society, including fat splitting, new applications for fatty acids and derivatives, and short-chain fatty acids from alcohols, olefins, and Zeigler intermediates (Sonntag, 1968). Kadesch reviewed the chemistry of fat-derived dibasic acids (Kadesch, 1954, 1979). Hastert reviewed the hydrogenation of fatty acids (Hastert, 1979). E.C. Leonard published an excellent review of polymerization and dimer acids (Leonard, 1979). Nitrogen derivatives of fatty acids were reviewed by Reck and manufacture of amides, diamides, nitriles, primary amines, and oxides as well as applications are discussed in detail (Reck, 1985). The aforementioned A.J. Stirton was a pioneer in soaps and detergents made from fats and fatty acids. He also authored five chapters for the 3rd edition of Bailey’s Industrial Oil & Fat Products (Stirton, 1964). The following sections outline some of the more important developments in various aspects of the oleochemical industry.

1.3.1 Fat Splitting Perhaps the most important discovery of the 19th century was the introduction of an industrially relevant method to split fats and oils into fatty acids and glycerin. Up until this time, fats and oils were saponified in open kettles using alkali. However, Ernst Twitchell patented a catalytic method in 1898, that became known as the Twitchell process (Twitchell, 1898). The acid catalyst was prepared by the reaction of oleic acid with sulfuric acid and naphthalene. In 1900 he reported that treatment of oleic acid and benzene with concentrated sulfuric acid yields benzene stearosulphonic acid useful as a fat-splitting reagent (Twitchell, 1900) along with additional papers on the synthesis of sulfonic acid containing stearic acid (Twitchell, 1906, 1907). In this process, melted fats with 25%50% by weight of water were mixed and then agitated while sparging with steam in an open tank for 1628 hours in

10

Fatty Acids

the presence of the catalyst. After allowing the reaction mixture to settle, the water and glycerin phase was removed and the fatty acids were recovered. Catalysts for the reaction were added at levels between 0.5% and 1.5%. Later, A.J. Stirton and colleagues showed that catalysts based on alkyl aryl sulfonates were more effective than Twitchell’s catalyst. Despite long reaction times operated in batch mode and the need for specially prepared catalysts, the Twitchell process was extensively used in the United States and England while a modified process using the improved alkylbenzene-based catalysts (Stirton et al., 1944) came to be used elsewhere in Europe. Twitchell was awarded the Perkin Medal in 1917 by the Society of Chemical Industry in recognition of his landmark achievement as well as an honorary doctorate.

1.3.1.1 Continuous Fat Splitting Although the Twitchell process represented a significant improvement over previous methods, the conditions were highly corrosive and energy intensive, and the batch method gave poor quality fatty acids having dark colors. Over time, more efficient and continuous splitting processes were developed. Several major advances in continuous fat splitting were introduced in the late 1930s. Ittner patented a countercurrent contact process with water and oil at temperatures of 200 C under pressure to yield soaps and glycerin (Ittner, 1933). Victor Mills described a continuous rapid fat-splitting method claimed to give higher yields of split fat and glycerin and a superior grade of fatty acids (Mills, 1939a,b). At various points, Procter & Gamble, Colgate, and Emery held patents similar to those of the original Mills and Ittner patents on continuous fat splitting (Ittner, 1938, 1948, 1949). Today many modern fat-splitting plants use the Colgate-Emory method, which is a continuous fat-splitting process employing a countercurrent hydrolysis reaction using steam in a pressure tower with internal heat exchange. The ColgateEmory process does not require catalysts, can be completed in 23 hours with splitting efficiencies of approximately 98%, and gives high-quality light-colored fatty acids that can subsequently be purified or separated by molecular distillation and fractionation (Barnebey and Brown, 1948).

1.3.2 Catalytic Hydrogenation Catalytic hydrogenation was a major advance in fatty acid chemistry. In 1897 Sabatier described the hydrogenation of organic compounds in the presence of finely disintegrated metals for which he was awarded the Nobel Prize in chemistry. In 1903 Normann received a patent on the hydrogenation of fatty acids and their glycerides (Normann, 1903). Normann subsequently licensed the hydrogenation technology to Joseph Crossfield, a businessman in Great Britain manufacturing soaps. Within a few years, Crossfield was

History of Fatty Acids Chemistry Chapter | 1

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convinced that hydrogenation offered the soap industry a new technology. Completely hydrogenated fats and oils would serve as a feedstock for fat splitting and a source of fatty acids for soap and candle manufacturers. Crossfield brought the hydrogenation patents to the United States and sold them to Procter & Gamble around 1907. Procter & Gamble subsequently discovered that hydrogenated fats had applications in the edible oil arena and by 1911 Procter & Gamble marketed the first all vegetable oil solid shortening made by blending partially hydrogenated and liquid cottonseed oil. In 1920 the US Supreme Court ruled that the Procter & Gamble patents were void, thereby opening up hydrogenation as a fat processing technology (List and Jackson, 2007, 2009).

1.3.3 Fatty Acid Distillation The modern US oleochemical industry developed largely on the efforts of Ralph Potts (190081), a member of American Oil Chemists’ Society (AOCS) who is often referred to as the “father of the oleochemical industry” (List, 2004b). Potts worked for the American meatpacking company, Armour and Company, and his research began with the belief that fatty acids could be distilled. With a crude-fractionating column, his belief was confirmed and by 1933 Armour built a distillation plant based on Potts’ distillation process. Armour sold fatty acids at a profit but supplies of tallow and grease exceeded the demand for fatty acids. Potts and Victor Conquest developed a plan to find new uses for fatty acids and by 1938 they had discovered a process to make fatty amines. The amine business was profitable but keeping up with the growing demand was difficult, so a new plant was built and operational by 1951. Potts designed similar plants in England, Canada, Japan, Belgium, and Brazil. Potts held numerous patents on the distillation of fatty and tall oil acids as well as patents on production of fatty acid nitriles and amines (Pool and Potts, 1944; Potts, 1948a,b, 1950, 1951, 1954, 1964, 1967; Potts and Christensen, 1943; Potts and McKee, 1936; Potts and Olson, 1953; Potts and Smith, 1957; Potts and Stalioraitis, 1971). Potts, authored numerous publications, was the second recipient of the Alton E. Bailey Award recognizing outstanding research and exceptional service in the field of lipids and associated products. Ralph Potts’ contributions to the modern oleochemical industry cannot be overemphasized, as the purification of fatty acids by distillation is a wellestablished industrial practice and still the most common and efficient means of producing high-purity fatty acids (Reck and Sonntag, 1984).

1.3.4 Fatty Alcohols Fats and oils are major sources of raw materials for soaps and detergents, with about 30% used as fatty acids and about 55% as fatty alcohols (Egan, 1968; Egan et al., 1984). Fatty alcohols can be derived from two main

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Fatty Acids

groups of natural raw materials, namely, fats and oils of plants and animals and wax esters from sources such as sperm (whale) oil or jojoba oil. Prior to 1973, fatty alcohols were produced by hydrolysis of the wax esters from sperm oil followed by fractionation of the fatty alcohols and fatty acids. However, the worldwide ban on whaling (1973) prompted the use of fats and oils for oleyl and other fatty alcohol production. The fats and oils are transesterified with methanol to give fatty acid methyl esters, which are then converted into the corresponding fatty alcohols through reduction chemistry. The fatty acid esters are reduced by either reaction with sodium and alcohol or hydrogenation in the presence of a suitable catalyst (Kastens and Peddicord, 1949). Typically, the hydrogenation reaction is carried out in the presence of hydrogen at 35004200 psi and temperatures of 300350 C using mixed catalysts that may contain chromium, zinc, copper cadmium, and aluminum. Obviously, the hydrogenation process is not suitable for preparing unsaturated fatty alcohols. For example, oleyl alcohol can be produced by sodium reduction of using tallow, olive oil, and canola oil as raw materials, which contain approximately 43%45%, 61%63%, and 60% oleic acid, respectively.

1.3.4.1 Guerbet Alcohols In 1907 Guerbet reported that linear alcohols can be converted to branched chain isomers by catalytic reactions at high temperatures (Guerbet, 1907). The product from the reaction is an alcohol with twice the molecular weight of the reactant minus one mole of water. Until recently, Guerbet alcohols have received little attention in oleochemical applications despite having many advantages over linear ones. These include good oxidative stability and lowtemperature operability as well as light colors. Guerbet alcohols can be further modified to yield Guerbet acids. For example, the Guerbet alcohol, 2-octyl dodecanol, when treated with sodium hydroxide, yields the corresponding fatty acid in high yield. A recent publication reported the synthesis of a number of Guerbet alcohols and their lubrication properties (Waykole and Bhowmick, 2014). Excellent reviews of the chemistry and properties of Guerbet alcohols are given by O’Lenick (O’Lenick, 2001, 2016).

1.3.5 Estolides Estolides are a unique class of compounds derived from fatty acids and may occur naturally as polyacylglycerides in vegetable oils or can be synthesized from unsaturated fatty acids or triglycerides (Isbell, 2011). Fatty acids with hydroxyl or epoxy groups are particularly attractive as estolide precursors. Castor and lesquerella oils are converted to estolides via successive esterifications of mid-chain hydroxyl moieties at a temperature of 250 C. The patent literature reports the synthesis of triglyceride estolides from the reaction of

History of Fatty Acids Chemistry Chapter | 1

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castor and lesquerella oils with various fatty acids using a p-toluenesulfonic acid catalyst at 150 C (Lawate, 1995). This allows the removal of water formed during the reaction as an azeotrope. Estolides generally have excellent cold flow properties and viscosity indices, thereby rendering them suitable as low-temperature lubricants. Estolides prepared from meadowfoam seed oil have poor low-temperature properties but provide good moisturizing properties for use in shampoo and conditioners (Isbell et al., 2000). Cermak and colleagues reported the synthesis of estolides from oleic and saturated fatty acids as well as their applications in lubricant formulations (Cermak et al., 2013; Cermak and Isbell, 2001). Isbell recently authored a comprehensive review of estolide studies carried out at the National Center for Agricultural Utilization Research (NCAUR) in Peoria, Illinois (Isbell, 2011).

1.3.6 Dimer and Trimer Cyclic Fatty Acids Cyclic acids were discovered in 1876 and, historically, interest in dimer acids was an outgrowth of work surrounding the thermal polymerization of vegetable oils known as heat bodying (Kappelmeier, 1933; Scheiber, 1929). When vegetable oils are heat bodied, complex oligomeric structures are formed through cyclization reactions at unsaturated sites of fatty acids. Although cyclization of unsaturated fatty acids was known and dimer acids were discovered in 1920, neither received much attention until the 1940s because the understanding of polymers and polymer chemistry was in its infancy. However, the pioneering work of Wallace Carothers, Carl Marvel, and Roger Adams at the University of Illinois gave better understanding of polymers and chemists new tools to modify fats and fatty acids. An early pioneer in the field was John Cowan, who received his PhD under Marvel at the University of Illinois. His early research at the newly opened Northern Regional Research Laboratory (now NCAUR) focused on the polymerization of fatty acids, resulting in rubber substitutes and plastics (Cowan, 1961). Fatty acids containing the trienoic structure such as linolenic acid found in linseed oil (50%55% linolenic acid), when treated with alkali at high temperatures, form a ring structure which after hydrogenation yields a saturated cyclic acid that can be esterified with alcohols (Friedrich, 1967, 1968; Friedrich et al., 1965). Friedrich synthesized a number of cyclic diesters and evaluated them for possible use as lubricants in the aviation and aerospace industries (Friedrich et al., 1965). Tetramethyl cyclobutanediol diesters from vicinally substituted cyclic acid mixtures were patented and met military specifications for aviation lubricants (Friedrich, 1968). Dimer and trimer acids are di- and polycarboxylic acids and are commercially produced by reacting unsaturated fatty acids found in vegetable oils such as tall oil, canola oil, or oleic acid in the presence of a clay catalyst. By using C18 unsaturated fatty acids, a wide variety of complicated isomeric oligomeric cyclic structures containing 36 and 54 carbon dimer and trimer

14

Fatty Acids

acids, respectively, can be obtained. These liquid oligomerized materials are unique since they never crystallize, have a molecular weight of around 560, distill with difficulty, are soluble in hydrocarbons, and are unsaturated but not conjugated. Dimer acids are reactive toward oxygen and sulfur but these reactions are easily controlled. Dimer acids are an important part of the oleochemical economy and have been commercialized and find uses in a number of applications in lubricant, pigment, cosmetic, personal care, and surfactant formulations.

1.3.7 Hydroformylation of Fatty Acids Carbon monoxide can react with unsaturated olefins (focus on unsaturated fatty acids) in a variety of ways. In the first method known as the oxo process, an unsaturated fatty acid is reacted with carbon monoxide in the presence of hydrogen and metal catalysts under high pressure to give hydroformylated products and dates back to the late 1930s (Pryde et al., 1972). The catalyst is a cobalt hydrocarbonyl complex formed in situ from a variety of cobalt compounds. Typically, extensive double bond isomerization occurs before hydroformylation thereby resulting in a mixture of isomers. Frankel reported that the use of a rhodium triphenylphosphine catalyst prevented isomerization and oleic acid gave exclusively methyl 9(10)-formylstearate (Frankel and Pryde, 1977). In the second method known as the Koch process, unsaturated fatty acids are reacted with carbon monoxide at the double bond position in the presence of sulfuric acid and water or alcohol to give a carboxylic acid or ester, respectively. The Koch process was reported in 1955 and was the first to demonstrate a feasible route to dicarboxylic acids. However, oleic acid was not reported. Subsequently, Roe and Swern reported the preparation of the corresponding diacid from oleic acid in good yields (Roe and Swern, 1960). Oleic acid was dissolved in 97% sulfuric acid and five moles of water was reacted with carbon monoxide at atmospheric pressure to give the diacid. Oleyl alcohol may also serve as the starting material. Other routes to diacids from oleic acid were reported by Reppe and Kroeper in the early 1950s (Reppe and Kroeper, 1952).

1.3.8 Ozonolysis of Fatty Acids and Triglycerides From 1960 to 1980, a considerable amount of research was reported describing the ozonolysis of fatty acids, esters, and trigylcerides (Pryde and Cowan, 1962). Ozone reacts with double bonds to form an ozonide intermediate that can be catalytically reduced to aldehydes or oxidized to shorter chain fatty acids (Kadesch, 1963). For example, oleic acid yields azelaic (9-carbon diacid) and pelargonic (nonanoic acid) acids upon oxidative ozonolysis and was commercialized in the late 1950s. Aliphatic alcohols were prepared in good yield by reductive ozonolysis of methyl oleate, followed by hydrogenation with a nickel catalyst in aprotic solvents (Pryde et al., 1968).

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1.4 CONTRIBUTIONS OF ANALYTICAL CHEMISTRY TO FATTY ACIDS A major contributor to the chemistry and composition of fatty acids was George Jamieson, who directed the Bureau of Chemistry and Soils at the US Department of Agriculture in Washington, DC (Jamieson, 1943; List, 2004a). Over the period of 191847, Jamieson and colleagues characterized the fatty acid compositions of numerous now commonplace oils including cottonseed, peanut, olive, safflower, corn, and soybean, all of which developed into important commodity oils. Jamieson also investigated a number of lesser known sources of fatty acids, some of which are of current interest as healthy oils. They include walnut, grapefruit seed, apricot, cherry seed, pecan, and avocado oils. Jamieson reviewed the literature up till 1943 (Jamieson, 1943). By the 1950s, a third revision was required and was authored by E.W. Eckey (Eckey, 1954). Some 70 years later, it remains a valuable resource for fatty acid data on plant oils from numerous species. Another pioneer in this area was Thomas Percy Hilditch (18861965). Hilditch devised or improved many procedures related to fats and oils including: ester fractionation, the use of thiocyanogen values, oxidative cleavage as a means to determine unsaturation, low-temperature crystallization, and alkali-isomerization to measure linoleic and linolenic acids (Gunstone, 2003). These contributions in analysis of fats and oils along with others relating to hydrogenation, autoxidation, and cis/trans isomerism have led many to refer to him as the “father of fats and oils chemistry” (Lie Ken Jie, 2015). His book, The Chemical Composition of Natural Oils, first published in 1940 and updated three times, is a seminal and influential contribution to chemical analysis of oils (Hilditch, 1940). By the early 1960s, gas chromatography (GC) offered fatty acid chemists, a new tool for the analysis of plant oils and fatty acids. Prior to the widespread use of GC, determination of fatty acid composition was a laborious and time-consuming endeavor, involving fractional crystallization, distillation, and subsequent chemical derivatization to determine chain length, functional group presence, and location (Lie Ken Jie, 2015). GC, along with spectroscopic methods such nuclear magnetic resonance (NMR), mass spectroscopy (MS), and Fourier transform infrared spectroscopy (FT-IR), greatly accelerated the determination of fatty acid composition from weeks to hours. Frank Gunstone, a former graduate student of Hilditch, was a prolific pioneer in the application of these methodologies to lipid chemistry (Gunstone et al., 1967; Lie Ken Jie, 2015). Chemists at NCAUR began screening germplasm for unique fatty acid compositions with potential industrial uses in the 1950s (e.g., Earle et al., 1959). A number of unusual fatty acids were found in addition to the more common ones encountered in plant oils. Among them are epoxy acids, hydroxy acids, and short-chain as well as very long-chain ( . C18) acids.

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Notable examples include meadowfoam and crambe oils (Miller et al., 1964; Miwa and Wolff, 1962). The ban on commercial whaling in 1973 brought jojoba oil (a long-chain wax ester) to commercialization as a result of research done at NCAUR (Miwa, 1984). Oil from cuphea is an excellent source of short-chain saturated fatty acids (70% caprylic acid) having a composition similar to coconut oil (Miller et al., 1964). Since the supply of coconut oil is limited, cuphea shows promise as a source of short-chain fatty acids for the oleochemical industry. Efforts are underway to bring cuphea oil to commercialization. Similarly, meadowfoam oil is a rich source of unusual very long-chain fatty acids that can be converted into estolides and other products. As mentioned previously, estolides prepared from a variety of fatty acids show great promise as lubricant additives (Isbell, 2011; Lawate, 1995). Crambe oil is a rich source of erucic acid (Miwa and Wolff, 1963). Research conducted at NCAUR showed that ozonolysis provides a route to erucamide, a monomer for nylon 13,13 production (Nieschlag and Wolff, 1971).

1.5 RECENT DEVELOPMENTS IN FATTY ACIDS A number of other important developments in the fatty acid field have occurred within the past 30 years or so. Biotechnology and the use of enzymes to modify fats, oils, and fatty acids have and are revolutionizing the entire fats and oils industry. Once thought impossible, the use of enzymes in edible oil processing has become a reality. Enzymatic degumming of crude oils and the interesterification of fat blends for trans free edible products have been commercialized in the United States and Europe (Orthoefer and List, 2015). Biotechnology led to the discovery that single cell oils offer potential for production of numerous fatty acids needed for human nutrition. These include EPA and DHA found in fish oils. Algal oils show promise as high oil yield sources for biofuel production (Chisti, 2007). The use of renewable resources represents a significant development in fatty acid chemistry and offers much potential for using green chemistry to protect the environment. Olefins can undergo a reaction known as metathesis, which has been exploited in the petroleum industry for decades. The foundations for metathesis were laid in the 1950s with the pioneering work of Anderson and Merckling (1955), who reported the first carboncarbon double bond rearrangement reaction in the titanium-catalyzed polymerization of norbenene. Banks and Bailey (1964) later discovered that olefins undergo disproportionation in the presence of catalytic tungsten and molybdenum hexachloride and tetramethyltin. In essence, olefins are cleaved and reformed to give new smaller and larger olefins. For example, propylene yields mostly ethylene and butenes, with lesser amounts of pentenes and hexenes also formed. The first successful application of metathesis chemistry to lipids was accomplished by van Dam and coworkers (van Dam et al., 1972, 1974), who reported that tungsten hexachloride/tetramethyl tin catalysts were effective at

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metathesis of fatty esters to alkenes and dicarboxylic acid dimethyl esters. Verkuijlen and coworkers (Verkuijlen et al., 1977) subsequently demonstrated that metathesis of fatty acid esters can be achieved with a heterogeneous catalyst based on rhenium oxide supported on alumina promoted by a small amount of tetramethyl tin. Grubbs and Schrock are pioneers in development of well-defined metathesis catalysts with broad functional group tolerance and high activity (Grubbs, 2004; Vougioukalakis and Grubbs, 2010). The original metathesis catalysts were ill defined, subject to poisoning and had poor functional group tolerance. In 2005 Grubbs, Schrock, and Chauvin were awarded the Nobel Prize in Chemistry for their pioneering work in metathesis catalyst development (Grubbs and Schrock) and elucidation of the reaction mechanism (Chauvin). Applications of metathesis to fatty esters were limited until the work of Grubbs and Schrock led to stable metal alkylidine complexes based on ruthenium, molybdenum, and tungsten. Several research groups in Europe and the United States have since made substantial progress in fatty ester metathesis chemistry using these new catalysts (Biermann et al., 2000, 2011; Fu¨rstner, 2000; Meier et al., 2007; Montero de Espinosa and Meier, 2012; Ngo et al., 2006; Rybak et al., 2008). Advances in plant biochemistry and traditional plant breeding have led to the development of a number of oils with modified fatty acid compositions. To date, trait-modified canola, soybean, and sunflower oils have been developed and commercialized through plant breeding and are non-genetically modified organism (GMO). Several new soybean oil varieties are nearing commercialization, including a low saturate high oleic variety and an omega3 enriched oil. Both have been developed through a combination of plant breeding and gene insertion (Wilkes, 2008; Wilkes and Bringe, 2015).

1.6 CONCLUSION In summary, fatty acids have long been important to man. Historically, from soap to candles to detergents and surfactants to biodiesel, a steady stream of new products based on fatty acids has appeared. Developments in chemistry and analytical techniques have played a major role in this progression, with fat-splitting hydrogenation, distillation, structure determination, and gas chromatographic analysis of fats and oils representing some of the most significant early developments in the chemistry and composition of fats and oils. Pioneers in these areas include Bailey, Chevreul, Heintz, Hilditch, Ittner, Jamieson, Potts, Sabatier, Twitchell, and Wurtz. Because of the important advances made by these and other researchers, renewable oleochemicals are now an important component of the worldwide chemical industry. Recent advances, including application of biotechnology and metathesis to fats and oils, demonstrate that the chemistry of fats, oils, and fatty acids is diverse and not yet fully realized, even after over 200 years of collective

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effort. Research efforts currently underway and those surely to be conducted in the future represent the next generation of important discoveries in fatty acid chemistry.

REFERENCES Anderson, A.W., Merckling, N.G., 1955. Polymeric bicycle-(2,2,1)-2-heptene, United States Patent 2,721,189. Anonymous, 2016. Chronological history of lipid science. ,http://www.cyberlipid.org/history/ history1.htm . (accessed 28.06.16). Bailey, A.E., Jefferson, M.E., Kregger, F.B., Baur, S.T., 1945. Thermal properties of fats and oils. IV. Some observations on the polymorphism and X-ray diffraction characteristics of tristearin and a highly hydrogenated cottonseed oil. Oil Soap 22, 1013. Banks, R.L., Bailey, G.C., 1964. Olefin disproportionation. A new catalytic process. Indus. Eng. Chem. Product Res. Dev. 3, 170173. Barnebey, H.L., Brown, A.C., 1948. Continuous fat splitting plants using the colgate-emery process. J. Am. Oil Chem. Soc. 25, 9599. Benedikt, R., Lewkowitsch, J., 1895. Chemical Analysis of Oils, Fats, and Waxes and of the Commercial Products Derived Therefrom. Macmillan, London. Biermann, U., Friedt, W., Lang, S., Lu¨hs, W., Machmu¨ller, G., Metzger, J.O., et al., 2000. New syntheses with oils and fats as renewable raw materials for the chemical industry. Angewandte Chem. Int. Ed. 39, 22062224. Biermann, U., Bornscheuer, U., Meier, M.A.R., Metzger, J.O., Scha¨fer, H.J., 2011. Oils and fats as renewable raw materials in chemistry. Angewandte Chem. Int. Ed. 50, 38543871. Blank, E.W., 1942. Chronological list of important dates in the history of the fats and waxes. Oil Soap 19, 110113. Cermak, S.C., Isbell, T.A., 2001. Synthesis of estolides from oleic and saturated fatty acids. J. Am. Oil Chem. Soc. 78, 557565. Cermak, S.C., Bredsguard, J.W., John, B.L., Kirk, K., Thompson, T., Isbell, K.N., et al., 2013. Physical properties of low viscosity estolide 2-ethylhexyl esters. J. Am. Oil Chem. Soc. 90, 18951902. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 23, 294306. Costa, A.B., 1962. Michel Eugene Chevreul: Pioneer of Organic Chemistry. State Historical Society of Wisconsin, University of Wisconsin Press, Madison, WI. Cowan, J.C., 1961. Twenty years of research in oils at Northern Regional Research Laboratory. J. Am. Oil Chem. Soc. 38, 1218. Dijkstra, A.J., 2009a. A chemical Study of Oils and Fats of Animal Origin by M.E. Chevreul. AOCS Press, Urbana, IL. Dijkstra, A.J., 2009b. How Chevreul (17861889) based his conclusions on his analytical results.. OCL—Oilseeds Fats Crops Lipids 16, 813. Earle, F.R., Melvin, E.H., Mason, L.H., van Etten, C.H., Wolff, I.A., Jones, Q., 1959. Search for new industrial oils. I. Selected oils from 24 plant families. J. Am. Oil Chem. Soc. 36, 304307. Eckey, E.W., 1954. Vegetable Fats and Oils. ACS Monograph Series. Reinhold Publishing Corporation, New York. Egan, R.R., 1968. The preparation and properties of amines and cationic surfactants from fatty acids. J. Am. Oil Chem. Soc. 45, 481486.

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Egan, R.R., Earl, G.W., Ackerman, J., 1984. Properties and uses of some unsaturated fatty alcohols and their derivatives. J. Am. Oil Chem. Soc. 61, 324329. Frankel, E.N., Pryde, E.H., 1977. Catalytic hydroformylation and hydrocarboxylation of unsaturated fatty compounds. J. Am. Oil Chem. Soc. 54, A873A881. Friedrich, J.P., 1967. C18-saturated cyclic acids from linseed oil: a structural study. J. Am. Oil Chem. Soc. 44, 244248. Friedrich, J.P., 1968. Tetramethyl cyclobutanediol diesters of linseed-derived C18 saturated vicinally substituted cyclic monocarboxylic acid isomer mixture, United States Patent 3,373,176. Friedrich, J.P., Bell, E.W., Gast, L.E., 1965. Potential synthetic lubricants: esters of C18saturated cyclic acids. J. Am. Oil Chem. Soc. 42, 643645. Fu¨rstner, A., 2000. Olefin metathesis and beyond. Angewandte Chem. Int. Ed. 39, 30123043. Grubbs, R.H., 2004. Olefin metathesis. Tetrahedron. 60, 71177140. Guerbet, M., 1907. Condensation and formation of methylisobutylcarbinol and dimethyl 1,2-heptanol 6 complexes. Comptes Rendus 149, 12132. Gunstone, F.D., 2003. Giants of the past: Thomas Percy Hilditch. INFORM—Int. News Fats Oils Relat. Mater. 14, 302303. Gunstone, F.D., Hamilton, R.J., 2001. Oleochemical Manufacture and Applications. John Wiley and Sons LTD, Oxford. Gunstone, F.D., Ismail, I.A., Lie Ken Jie, M.S.F., 1967. Fatty acids, part 16. Thin layer and gasliquid chromatographic properties of the cis and trans methyl octadecenoates and of some acetylenic esters. Chem. Phys. Lipids 1, 376385. Hastert, R.C., 1979. Hydrogenation of fatty acids. J. Am. Oil Chem. Soc. 56, 732A739A. Hilditch, T.P., 1940. The Chemical Constitution of Natural Fats. John Wiley and Sons, West Essex. Isbell, T.A., 2011. Chemistry and physical properties of estolides. Grasas y Aceites 62, 830. Isbell, T.A., Abbott, T.P., Dworak, J.A., 2000. Shampoos and conditioners containing estolides, United States Patent 6,051,214. Ittner, M.H., 1933. Process of making soap and glycerine, United States Patent 1,918,603. Ittner, M.H., 1938. Hydrolysis of fats and oils, United States Patent 2,139,589. Ittner, M.H., 1948. Fat hydrolysis, United States Patent 2,435,745. Ittner, M.H., 1949. Continuous fat splitting, United States Patent 2,458,170. Jamieson, G.S., 1943. Vegetable fats and oils their chemistry, production and utilization for edible, medicinal and technical purposes, second ed. Reinhold Publishing Corporation, New York. Kadesch, R.G., 1954. Dibasic acids. J. Am. Oil Chem. Soc. 31, 568573. Kadesch, R.G., 1963. Ozonolysis of fatty acids and their derivatives. In: Holman, R.T., Lundberg, W., Malkin, T. (Eds.), Progress in the Chemistry of Fats and Other Lipids, vol. VI. Pergamon Press, Oxford, pp. 291312. Kadesch, R.G., 1979. Fat-based dibasic acids. J. Am. Oil Chem. Soc. 56, A845A849. Kappelmeier, C.P.A., 1933. Chemical processes of stand-oil formation. Farben-ZTG 38, 10181020. Kastens, M.L., Peddicord, H., 1949. Alcohols by sodium reduction. Indus. Eng. Chem. 41, 438446. Kenar, J.A., 2007. Glycerol as a platform chemical: sweet opportunities on the horizon? Lipid Technol. 19, 249253. Lawate, S.S., 1995. Trigylceride oils thickened with estolides of hydroxy-containing trigylcerides, United States Patent 5,427,704. Leonard, E.C., 1979. Polymerization-dimer acids. J. Am. Oil Chem. Soc. 56, 782A785A.

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Lie Ken Jie, M.S.F., 2015. Frank D. Gunstone—teacher, researcher, and writer. Eur. J. Lipid Sci. Technol. 117, 135140. List, G.R., 2004a. Giants of the past: George Jamieson (18791959). INFORM—Int. News Fats Oils Relat. Mater. 15, 684. List, G.R., 2004b. Giants of the past: Ralph Potts (19001981). INFORM—Int. News Fats Oils Relat Mater. 15, 168. List, G.R., Jackson, M.A., 2007. The battle over hydrogenation (19031920). INFORM—Int. News Fats Oils Relat Mater 18, 403405. List, G.R., Jackson, M.A., 2009. The battle over hydrogenation (19031920). Part II. Litigation. INFORM—Int. News Fats Oils Relat. Mater. 20, 395397. Lutton, E.S., 1945. The polymorophism of tristearin and some of its homologs. J. Am. Chem. Soc. 67, 524527. Malkin, T., 1954. The polymorphism of gylcerides. In: Holman, R.T., Lundberg, W., Malkin, T. (Eds.), Progress in the Chemistry of Fats and Other Lipids, vol. II. Pergamon Press, Oxford, pp. 150. Markley, K.S., 1964. Fatty Acids, Their Chemistry, Properties, Production and Uses, second ed. Wiley Interscience, New York. Meier, M.A.R., Metzger, J.O., Schubert, U.S., 2007. Plant oil renewable resources as green alternatives in polymer science. Chem. Soc. Rev. 36, 17881802. Miller, R.W., Earle, F.R., Wolff, I.A., Jones, Q., 1964. Search for new industrial oils, IX. Cuphea, a versatile source of fatty acids. J. Am. Oil Chem. Soc. 41, 279280. Mills, V., 1939a. Continuous countercurrent hydrolysis of fat, United States Patent 2,156,863. Mills, V., 1939b. Continuous process for converting saponifiable fats into soap and glycerin, United States Patent 2,159,397. Miwa, T.K., 1984. Structural determination and uses of jojoba oil. J. Am. Oil Chem. Soc. 61, 407410. Miwa, T.K., Wolff, I.A., 1962. Fatty acids, fatty alcohols, and wax esters from Limnanthes douglassi (Meadowfoam) seed oil. J. Am. Oil Chem. Soc. 39, 320322. Miwa, T.K., Wolff, I.A., 1963. Fatty acids, fatty alcohols, and wax esters from Crambe abyssinica and Lunaria annua seed oils. J. Am. Oil Chem. Soc. 40, 742744. Montero de Espinosa, L., Meier, M.A.R., 2012. Olefin metathesis of renewable platform chemicals. Topics Organometal. Chem. 39, 144. Ngo, H.L., Jones, K., Foglia, T.A., 2006. Metathesis of unsaturated fatty acids: synthesis of long-chain unsaturated-α,ω-dicarboxylic acids. J. Am. Oil. Chem. Soc. 83, 629634. Nieschlag, H.J., Wolff, I.A., 1971. Industrial uses of high erucic oils. J. Am. Oil Chem. Soc. 48, 723727. Normann, W., 1903. Process for converting unsaturated fatty acids or their glycerides into saturated compounds, Great Britian Patent GB190301515. O’Lenick, A.J., 2001. Guerbet chemistry. J. Surfact. Deterg. 4, 311315. O’Lenick, A.J., 2016. A review of Guerbet chemistry. ,http://zenitech.com/documents/guerbet_ chemistry.pdf . (accessed 07.07.16). Orthoefer, F.T., List, G.R., 2015. Trait Modified Oils in Foods. Wiley-Blackwell, Hoboken, NJ. Pool, W.O., Potts, R.H., 1944. Preparation of amines, United States Patent 2,358,030. Potts, R.H., 1948a. Distillation of fatty acid stock by fractionation and flash steps, United States Patent 2,450,612. Potts, R.H., 1948b. Nitrile-producing method, United States Patent 2,448,275. Potts, R.H., 1950. Method of separating aliphatic nitriles, water, ammonia, and hydrogenationinhibiting impurities, United States Patent 2,504,045.

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Potts, R.H., 1951. Manufacture of nitriles and amides, United States Patent 2,546,521. Potts, R.H., 1954. Process for distilling tall oil, United States Patent 2,674,570. Potts, R.H., 1964. Nitrile hydrogenation manufacture of primary amines, United States Patent 3,163,676. Potts, R.H., 1967. Stepwise preparation of fatty acid nitriles, United States Patent 3,299,117. Potts, R.H., Christensen, C.W., 1943. Process of preparing nitriles, United States Patent 2,314,894. Potts, R.H., McKee, J.E., 1936. Fatty acid distillation, United States Patent 2,054,096. Potts, R.H., Olson, R.N., 1953. Distillation of fatty acids, tall oil, and the like, United States Patent 2,627,500. Potts, R.H., Smith, R.S., 1957. Preparation of nitriles, United States Patent 2,808,426. Potts, R.H., Stalioraitis, J.S., 1971. Hydrolysis of amides to amines, United States Patent 3,592,854. Pryde, E.H., 1979. Fatty Acids. AOCS Press, Champaign, IL. Pryde, E.H., Cowan, J.C., 1962. Aldehydic materials by the ozonization of vegetable oils. J. Am. Oil Chem. Soc. 39, 496500. Pryde, E.H., Moore, D.J., Cowan, J.C., 1968. Hydrolytic, reductive and pyrolytic decomposition of selected ozonloysis products. Water as an ozonization medium. J. Am. Oil Chem. Soc. 45, 888894. Pryde, E.H., Frankel, E.N., Cowan, J.C., 1972. Reactions of carbon monoxide with unsaturated fatty acids and derivatives: a review. J. Am. Oil Chem. Soc. 49, 451456. Ramberg, P.J., 2013. Wilhelm Heintz (18171880) and the chemistry of the fatty acids. Bullet. History Chem. 38, 1928. Reck, R.A., 1985. Industrial uses of palm, palm kernel and coconut oils: nitrogen derivatives. J. Am. Oil Chem. Soc. 62, 355365. Reck, R.A., Sonntag, N.O., 1984. A tribute to Ralph Potts. J. Am. Oil Chem. Soc. 61, 1216. Reppe, W., Kroeper, H., 1952. Verfahren zur herstellung von poly- oder oxycarbonsaeuren oder deren funktionellen abkoemmlingen, German Patent 861,243. Roe, E.T., Swern, D., 1960. Branched carboxylic acids from long-chain unsaturated compounds and carbon monoxide at atmospheric pressure. J. Am. Oil Chem. Soc. 37, 661668. Rybak, A., Fokou, P.A., Meier, M.A.R., 2008. Metathesis as a versatile tool in oleochemistry. Eur. J. Lipid Sci. Technol. 110, 797804. Scheiber, J., 1929. Reactions in the formation of stand oils. Farbe u Lack.585587. Sonntag, N.O.V., 1968. Straight-chain fatty acids from alcohols, olefins, and ziegler intermediates. J. Am. Oil Chem. Soc. 45, 1416. Spitz, L., 2004. SODEOPEC—Soaps, Detergents, Oleochemicals, and Personal Care Products. AOCS Publishing, Champaign, IL. Stirton, A.J., 1964. Fractionation of fats and fatty acids. In: Swern, D. (Ed.), Bailey’s Industrial Oil & Fat Products, third ed. Wiley Interscience, New York, pp. 10051037. Stirton, A.J., Hammaker, E.M., Herb, S.F., Roe, E.T., 1944. Comparison of fat-splitting reagents in the twitchell process. Oil Soap 21, 148151. Twitchell, E., 1898. Process of decomposing fats or oils into fatty acids and glycerine, United States Patent 601,603. Twitchell, E., 1900. Benzenestearosulphonic acid and other sulphonic acids containing the stearic radical. J. Am. Chem. Soc. 22, 2226. Twitchell, E., 1906. A reagent in the chemistry of fats. J. Am. Chem. Soc. 28, 196200. Twitchell, E., 1907. A reagent in the chemistry of fats. J. Am. Chem. Soc. 29, 566571. van Dam, P.B., Mittelmeijer, M.C., Boelhouwer, C., 1972. Metathesis of unsaturated fatty acid esters by a homogeneous tungsten hexachloride-tetramethyltin catalyst. J. Chem. Soc. Chem. Commun. 1972, 12211222.

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van Dam, P.B., Mittelmeijer, M.C., Boelhouwer, C., 1974. Homogeneous catalytic metathesis of unsaturated fatty esters: new synthetic method for preparation of unsaturated mono- and dicarboxylic acids. J. Am. Oil Chem. Soc. 51, 389392. Verkuijlen, E., Kapteijn, F., Mol, J.C., Boelhouwer, C., 1977. Heterogeneous metathesis of unsaturated fatty acid esters. J. Chem. Soc. Chem. Commun.198199. Vougioukalakis, G.C., Grubbs, R.H., 2010. Ruthenium-based heterocyclic carbene-coordinated olefin metathesis catalysts. Chem. Rev. 110, 17461787. Waykole, C., Bhowmick, D., 2014. Synthetic base stock based on Guerbet alcohols. J. Am. Oil Chem. Soc. 91, 14071416. Wilkes, R.S., 2008. Low linolenic soybeans and beyond. Lipid Technol. 20, 277279. Wilkes, R.S., Bringe, N.A., 2015. Applications of trait enhanced soybean oils. In: Orthoefer, F.T., List, G.R. (Eds.), Trait Modified Oils in Foods. Wiley-Blackwell, Hoboken, NJ, pp. 7192.

Chapter 2

Naturally Occurring Fatty Acids: Source, Chemistry, and Uses James A. Kenar1, Bryan R. Moser1 and Gary R. List2 1 2

National Center for Agricultural Utilization Research, Peoria, IL, United States, G.R. List Consulting, Washington, IL, United States

Chapter Outline 2.1 Introduction 2.2 Production of Naturally Occurring Fatty Acids 2.2.1 Chemical Splitting 2.2.2 Lipase Splitting 2.3 Purification of Fatty Acids 2.3.1 Simple Distillation 2.3.2 Fractional Distillation 2.3.3 Molecular Distillation 2.3.4 Crystallization 2.3.5 Urea Fractionation 2.4 Sources and Types of Naturally Occurring Fatty Acids 2.4.1 Saturated Fatty Acids

24 28 29 30 31 31 32 35 35 36 37 38

2.4.2 2.4.3 2.4.4 2.4.5

Unsaturated Fatty Acids Hydroxy Fatty Acids Acetylenic Fatty Acids Allenic and Cumulenic Fatty Acids 2.5 Chemistry of Naturally Occurring Fatty Acids 2.5.1 Reactions at the Carboxylic Acid Group 2.5.2 Reactions at Unsaturated Sites 2.6 Conclusion References

39 43 45 47 49 50 57 71 71

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Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00002-7 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

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2.1 INTRODUCTION There is a remarkable range of naturally occurring fatty acids (well over 1000) that are found in and constitute the major components of fats, oils (triacylglycerols), waxes, and other lipid-containing materials (Gunstone et al., 2007b). These fatty acids are structurally diverse and often have unusual and interesting structures and, although they can have from 8 to over 80 carbon atoms, they are typically comprised of an even number of carbon atoms. In addition, the alkyl chains can contain a variety of functional groups and also have from 0 to 6 carboncarbon double bonds that predominantly possess cis-geometry, although, fatty acids having trans-geometry are also known (Fig. 2.1). Because of their structural diversity, fatty acids are extensively used as feedstocks for food applications and the oleochemicals industry for the manufacture of soaps, detergents, lubricants, coatings, and cosmetics among other specialty products. Of the many fatty acids, only 2025 are widely distributed in nature, and are of commercial significance. These fatty acids range between 10 and 22 carbons in length and are obtained in large quantities from the major domesticated commodity plant oils and animal fats (Harwood and Gunstone, 2007; Zanetti et al., 2013). There are nine major commodity oils that are tracked worldwide, and include soybean, sunflower, palm, palm kernel, cottonseed, peanut, olive, rapeseed (canola), and coconut oils. These oils are derived from vegetable and tree sources, account for approximately 97% of total oil

FIGURE 2.1 General structures and functional groups that can be found in naturally occurring fatty acids.

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production, and provide lauric, myristic, palmitic, stearic, oleic, linoleic, α-linoleic, and erucic fatty acids (Gunstone, 1996; Gunstone and Hamilton, 2001). The main fatty acids from animal fats (cattle, sheep, and pigs) and fish oils are myristic, palmitic, palmitoleic, stearic, oleic, eicosenoic, arachidonic, eicosapentaenoic (EPA), docosenoic, and docosahexaenoic (DHA) acids. These fatty acids are used in food, nutrition, and oleochemical applications where they are commonly used as fatty acid mixtures rather than as individually pure fatty acids. Table 2.1 gives the fatty acid compositions found in major plant and animal sources. In the last decade (200414), annual production of vegetable oils has increased approximately 62% from 107.2 to 173.5 million metric tons (MMT) (List, 2015). In 2014, 43.8 million tonnes (25.2%) of global fats and oils production were used by the oleochemical industry for nonfood industrial purposes compared to 22.4 MMT (20.0%) in 2004. Growth was driven mainly by high petroleum prices as well as the growing demand for natural or renewable products (Anneken et al., 2000; Biermann et al., 2011; Evangelista et al., 2015). Food and nutrition continues to be the major use of natural oils and fats and approximately 129.7 MMT (74.8%) was used for this purpose in 2014 (Zanetti et al., 2013). Animal fats (pork lard and beef tallow), produced by the rendering industry, are widely used in many regions of the world for food and nonfood applications. The rendering industry is a significant part of livestock processing and uses meat industry by-products as its feedstock. After the four leading plant oils (palm, soybean, rapeseed, and sunflower), tallow and lard are the fifth and sixth largest contributors to lipid production (Gunstone, 2008; Harwood and Gunstone, 2007). In 2004 the annual production of lard and tallow was 7.52 and 8.37 MMT, respectively, while in 2006, 1.0 MMT of fish oils were produced (Harwood and Gunstone, 2007). In 2008 lard and tallow production remained steady at 8.27 and 8.66 MMT, respectively (Gunstone, 2008). The fats and oils derived from plants and animals have characteristic fatty acid profiles and the compositional distribution of fatty acids within these fats and oils is influenced not only by the botanical or animal source from which they are obtained but also to some extent by the conditions under which the plants and animals were raised (Table 2.1). Palm oil, tallow, and lard are typically rich in long-chain saturated fatty acids such as palmitic (16:0) and stearic (18:0) in addition to monounsaturated oleic acid (9c-18:1). Coconut and palm kernel oils contain high amounts of small-chain saturated fatty acids such as lauric (12:0) and myristic (14:0) acids. Olive and sunflower oils are high in oleic acid while soybean oil has more linoleic (9c,12,c-18:2) acid. Rapeseed oil is a good source of very long-chain fatty acids such as erucic acid (13c-22:1). Food grade canola is a conventional bred cultivar of rapeseed that is low in erucic acid and glucosinolates and consists predominantly of oleic acid (18:1), linoleic

TABLE 2.1 Fatty Acid Composition (Expressed as Percentage of Total Fatty Acids) of Some Common Plant and Animal Oils and Fats (Modified From Codex Alimentarius: Standard for Named Vegetable Oil CODEX STAN 210-1999, Standard for Named Animal Fats CODEX STAN 211-1999) Fatty Acid

Fat/Oil Source Plant Based

Animal Based

Coconut

Corn

Cottonseed

Olive

Palm

Palm

Peanut

Rapeseed

Kernel

Canola

Soybean

Sunflower

(Low

Pork

Beef

Lard

Tallow

Erucic Rapeseed) Oilseed content (%)

6568

5

1820

2530

4550

4550

6065

4045

4045

1820

3545





Caproic acid C6:0

ND0.7

ND

ND



ND

ND0.8

ND

ND

ND

ND

ND





Caprylic acid C8:0

4.610.0

ND

ND



ND

2.46.2

ND

ND

ND

ND

ND





Capric acid C10:0

5.08.0

ND

ND



ND

2.65.0

ND

ND

ND

ND

ND

,0.5

,0.5

Lauric acid C12:0

45.153.2

ND0.3

ND0.2

ND

ND0.5

45.055.0

ND0.1

ND

ND

ND0.1

ND0.1





Myristic acid C14:0

16.821.0

ND0.3

0.61.0

0.00.05

0.52.0

14.018.0

ND0.1

ND0.2

ND0.2

ND0.2

ND0.2

1.02.5

2.06.0

Palmitic acid C16:0

7.510.2

8.616.5

21.426.4

7.520.0

39.347.5

6.510.0

8.014.0

1.56.0

2.57.0

8.013.5

5.07.6

2030

2030

Palmitoleic C16:1

ND

ND0.5

ND1.2

0.33.5

ND0.6

ND0.2

ND0.2

ND3.0

ND0.6

ND0.2

ND0.3

2.04.0

1.05.0

C17:0

ND

ND0.1

ND0.2

0.00.3

ND0.2

ND

ND0.1

ND0.1

ND0.3

ND0.1

ND0.2

,1.0

0.52.0

Stearic acid C18:0

2.04.0

ND3.3

3.06.5

0.55.0

3.56.0

1.03.0

1.04.5

0.53.0

0.83.0

2.05.4

2.76.5

822

1530

Oleic acid C18:1

5.010.0

2.042.2

12.028.0

55.083.0

36.044.0

12.019.0

35.069.0

8.060.0

51.070.0

1730

14.039.4

3555

3045

Linoleic acid C18:2

1.02.5

34.065.6

58.078.0

3.521.0

9.012.0

1.03.5

1.46.6

11.023.0

15.030.0

48.059.0

48.374.0

412.0

16

Linolenic acid C18:3

ND0.2

ND2.0

ND1.0

,1.5

ND0.5

ND0.2

ND

5.013.0

5.014.0

4.511.0

ND-0.3

,1.5

,1.5

Arachidic acid C20:0

ND0.2

0.31.0

ND1.0

0.00.6

ND1.0

ND0.2

ND

ND3.0

0.21.2

0.10.6

0.10.5

,1.0

,0.5

Gadoleic acid C20:1

ND0.2

0.20.6

ND0.3

0.00.4

ND0.4

ND0.2

ND

3.015.0

0.14.3

ND0.5

ND0.3

,1.5

,0.5

C20:2

ND

ND0.1

ND



ND

ND

ND

ND1.0

ND0.1

ND0.1

ND

,1.0

,0.1

Behenic acid C22:0

ND

ND0.5

ND0.5

0.00.2

ND0.2

ND0.2

ND

ND2.0

ND0.6

ND0.7

0.31.5

,0.1

,0.1

Erucic acid C22:1

ND

ND0.3

ND0.3

ND

ND

ND

ND

.2.060.0

ND2.0

ND0.3

ND0.3

,0.5

ND

C22:2

ND

ND

ND



ND

ND

ND

ND2.0

ND0.1

ND

ND0.3





C24:0

ND

ND0.5

ND0.5

,1.0

ND

ND

ND

ND2.0

ND0.3

ND0.5

ND0.5





C24:1

ND

0.52.5

ND



ND

ND

ND

ND3.0

ND0.4

ND

ND





ND, non-detectable; , no value.

28

Fatty Acids

acid (9c,12,c-18:2), and α-linoleic acid (ALA; 9c,12,c,15c-18:3) acids in 51%70%, 15%30%, and 5%14%, respectively. Many industrially important oilseed crops have been domesticated for food use and have undergone many years of selection for traits amenable to agriculture practices. Long-term breeding research is ongoing for these species to develop oils having improved nutritional or agronomic properties and successes of this work include high-oleic sunflower oil and canola oil (low erucic/glucosinolate rapeseed). Development of new industrial oilseed crops that contain unique fatty acids within their oils is also of interest and is being developed through both breeding and genetic engineering approaches. Examples of these “new crop” species include crambe, pennycress, lesquerella, meadowfoam, cuphea, vernonia, and coriander (Dierig et al., 2011; Dyer et al., 2008; Isbell, 2009; Zanetti et al., 2013). These species have unusual abundances of unsaturated, short, medium, or very long-chain fatty acids or unique hydroxyl, epoxide, or acetylenic functional groups.

2.2 PRODUCTION OF NATURALLY OCCURRING FATTY ACIDS Because the majority of fatty acids occur naturally as triesters with glycerol (triacylglycerols) within plant and animal materials, processing steps must first be performed to recover the desired triacylglycerols, which are then split (hydrolyzed) to separate the fatty acids from glycerol. Traditional fat and oil processing steps can be divided into four operations, namely, recovery, refining, conversion, and stabilization of the oil and have been thoroughly reviewed elsewhere (Bockisch, 1998; Dijkstra and Segers, 2007; Gunstone, 1996; Johnson, 2002; O’Brien, 2009). Briefly, recovery, referred to as crushing or extraction for plant materials and rendering for inedible animal by-products, involves mechanical pressing and/or solvent extraction to separate the crude oil or fat from other meal components such as protein and carbohydrates. The resulting oil or fat is then refined to remove undesirable components such as pigments, phospholipids, metals, and adverse flavor and odor compounds (Dijkstra and Segers, 2007; Johnson, 2002). The oil or fat may then go through a conversion step whereby it is modified through hydrogenation, winterization, crystallization, or interesterification treatments to alter its physical properties. Finally, stabilization steps ensure that the oil or fat is in the correct crystalline form to obtain the desired functional properties and has good stability, nutrition, and safety characteristics.

Naturally Occurring Fatty Acids: Source, Chemistry Chapter | 2

29

2.2.1 Chemical Splitting To isolate fatty acids from triacylglycerols, hydrolysis (or splitting) is performed under the influence of water, temperature, and pressure. Stoichiometrically, for every triacylglycerol molecule subjected to complete hydrolysis, three molecules of water is required and liberates three fatty acids as well as one molecule of glycerol. The reaction likely occurs homgeneously in the lipophilic oil phase between triacylglycerol and a small quantity of water dissolved in the oil (Lascaray, 1952). The reaction proceeds as a series of equilibria and gives diacylglycerols and monoacylglycerols as intermediates along the way to glycerol and fatty acids. The reaction is reversible and the hydrolysis rate, equilibrium, and final product composition is influenced by both the fatty acid concentration in the oil phase and on glycerol concentration in the water phase (Lascaray, 1952). In addition, the extent of hydrolysis increases with increasing temperature and pressure since the miscibility of water in the lipid phase improves under these conditions. The equilibrium and reaction kinetics have been studied and summarized by Anneken et al. (2000). Industrially, other reagents such as methanol (methanolysis) or amines (aminolysis) can also be used in place of water to directly give the corresponding fatty acid methyl esters and fatty amines, respectively. Alkaline, high-pressure steam, and enzymatic splitting are the three major commercial processing routes used to obtain fatty acids. Early hydrolysis procedures treated fats and oils with alkali in open reaction vessels. However, this method used much energy and after the reaction was completed, the resulting fatty acid soaps required acidification to obtain the desired fatty acid products. Historically, the Twitchell and Colgate-Emery processes are the most often practiced methods for industrial-scale production of fatty acids from fats and oils. In 1898 Ernst Twitchell modernized the oil and fat industry by patenting a method (Twitchell process) to split oils and fats using an acidic naphthalene stearosulfonic acid catalyst (sulfonated long-chain alkylbenzenes) (Ackelsberg, 1958; Twitchell, 1898). Twitchell’s catalyst accelerated the hydrolysis of oils (2448 hours at B100 C) relative to past processes and had a better splitting efficiency (Sonntag, 1979). Although the Twitchell process represented a significant improvement over previous methods, the conditions were highly corrosive and energy intensive, and the batch method gave poor-quality products having dark colors. Since then more efficient and economical batch autoclave and continuous processes based on high-pressure steam such as the widely used ColgateEmery steam hydrolysis process have been developed. The Colgate-Emery process is typically conducted catalyst-free at 210330 C and 26 MPa for 23 hours under continuous counter-current conditions. Under these conditions, oil hydrolysis proceeds without the need for catalysts and provides better quality mixtures of fatty acids with splitting efficiencies on the order of

30

Fatty Acids

B95% (Anneken et al., 2000; Sonntag, 1979). However, the high temperature and pressure render this process unsuitable for splitting oils containing polyunsaturated fatty acids (PUFAs) or fatty acids containing other thermally sensitive functional groups. Other methods have also been described for fat splitting based on the use of sub- and supercritical water (Holliday et al., 1997) and solid catalysts (Satyarthi et al., 2011).

2.2.2 Lipase Splitting Because of high-energy costs, environmental concerns, and unsuitability of the aforementioned chemical-splitting methods to sensitive oils, hydrolysis of triacylglycerols into fatty acids and glycerol can be promoted by lipases and is being employed more frequently as an alternative method to produce fatty acids. Lipases (triacylglycerol hydrolases; EC 3.1.1.3) are ubiquitous enzymes found in microbes, plants, and animals, can catalyze the hydrolysis and formation of ester bonds, and are one of the most useful and wellstudied enzymes (Bornscheuer et al., 2002; Gandhi, 1997; Sharma et al., 2001). Lipases are highly selective for the carboxyl group and their natural action is to catalyze the hydrolysis of triacylglycerols to give mixtures of fatty acids, monoacylglycerols, diacylglycerols, and glycerol. Lipases are proteins having molecular weights from 9000 to 70,000 Da, require no cofactors for activity, show chemo-, regio-, and stereo-selectivity, and can also be used in organic solvents (Adlercreutz, 2013). Of the available enzymes, microbial lipases based on fungi, yeast, molds, and bacteria are used extensively. Because lipase-promoted hydrolysis can be specific and occurs under mild conditions (B620 hours at 2060 C), the amount of secondary degradation products is reduced relative to the more rigorous chemical processes. In addition, specialized reaction vessels are not required due to the mild reaction conditions. Typically, oil hydrolysis using a lipase (free or immobilized) is performed by mixing an aqueous lipase solution with oil at a temperature and pH between 30 and 50 C and 59, respectively. The reaction occurs at the interface between the oil and water phase. The reaction is influenced by many factors such as enzyme concentration, stirring, water content, and oil source. Depending on the specificity of the enzyme and reaction time, mono- or diacylglycerols may be present in the final product mixture or complete hydrolysis yielding only fatty acids and glycerol can be achieved. Despite the widespread use of enzymes in various industrial applications and the potential savings in energy costs, fat splitting using lipases is limited due to enzyme costs and inactivation/stability issues. Industrial lipase use is increasing and applications such as the enrichment of PUFAs from fish oil (Halldorsson et al., 2004; Mbatia et al., 2011) and the synthesis of structured lipids whereby certain fatty acids occupy specific positions on the glycerol backbone are being pursued (Schmid et al., 1999). The review by Biermann

Naturally Occurring Fatty Acids: Source, Chemistry Chapter | 2

31

et al. (2011) outlines other examples of lipase use in oleochemistry. Much research has been focused on engineering lipases to improve their efficiency and stability toward hydrolysis reaction conditions (Bornscheuer et al., 2002; Nagarajan, 2012). The future industrial use of lipases is promising as significant protein engineering and screening research efforts continue to improve enzyme technology and performance.

2.3 PURIFICATION OF FATTY ACIDS Once split, the resulting crude fatty acid mixtures contain water, metals, hydrocarbons, color bodies such as chlorophyll, and odor substances, in addition to high boiling phytosterols, glycerol, phosphatides, and mono- and diacylglycerol impurities that must be removed. The four fundamental approaches to purification of fatty acids include crystallization, selective adsorption, extraction, and distillation. Several excellent reviews exist on these topics (Anneken et al., 2000; Bockisch, 1998; Brown and Kolb, 1955; Cermak et al., 2012; Gunstone et al., 1994; Markley, 1964; O’Brien, 2009). Of this distillation has been practiced for over a hundred years and from an industrial viewpoint provides the most efficient and economical means to produce high-purity fatty acids (Cermak et al., 2012). Other methods such as selective adsorption, extraction, membrane filtration, and liquid chromatography are mainly utilized in laboratory settings and currently have limited or specific commercial success.

2.3.1 Simple Distillation Distillation of fatty acid mixtures may be carried out as a batch or continuous process under atmospheric or reduced pressure and is used to isolate fatty acid mixtures from contaminants (simple distillation) or as a means isolate individual fatty acids from a mixture (fractional distillation). Early commercial distillations were operated in batch mode at atmospheric pressure (Potts and Muckerheide, 1968). Crude fatty acid mixtures are obtained from splitting and refining and can vary greatly in composition and the quality of distillation is highly dependent on pretreatment processing conditions. In addition, long-chain fatty acids typically have high boiling points and are susceptible to oxidative and thermal decomposition, polymerization and dehydration reactions that may occur at elevated temperatures needed for distillation. Therefore, distillations should be conducted at the lowest permissible temperatures to achieve separation and with minimal contact time to minimize fatty acid degradation. Several excellent reviews on distillation and fractionation of fatty acids by distillation should be consulted for further details (Anneken et al., 2000; Cermak et al., 2012; Gervajio, 2005; Potts and Muckerheide, 1968).

32

Fatty Acids

Batch steam distillation at atmospheric pressure uses a direct-fired still pot fitted with a steam sparger (Markley, 1964). A distillation pot containing the crude fatty acids is heated between 260 and 316 C while sparging with steam. A 5:1 steam to fatty acid vapor ratio is typically utilized to maintain the distillation temperature and to prevent undesirable anhydride formation (Potts and Muckerheide, 1968). However, a large amount of steam is needed and a considerable percentage of fatty acids becomes entrained in the steam condensate thereby resulting in poor overall process economics. More importantly, the high temperatures and long heating times led to poor fatty acid recovery due to degradation and polymerization. With technological advances, batch distillation is currently practiced industrially on small scale, as these industrial batch units have been replaced by modern distillation apparatuses that operate continuously under high vacuum. In continuous distillation, the fatty acid feed is introduced at a rate similar to which the distillate is drawn out of the distillation unit and the high-vacuum conditions utilized allow lower distillation temperatures and residence times that result in higher quality fatty acids and economic efficiencies (Anneken et al., 2000; Lausberg et al., 2008). Although there are several designs and configurations for continuous distillation units, vacuum equipment, effective heating, good circulation for efficient mass transfer between vapor and condensate, and steam economy are paramount to achieve the lowest possible temperatures and contact times.

2.3.2 Fractional Distillation Since fatty acids are derived from natural sources, they are obtained as mixtures most commonly having chain lengths ranging between C6 and C22 that vary depending upon the type of fat or oil used and even from batch to batch within the same source (Table 2.1). To obtain more uniform and high-purity (up to 99%) fatty acid compositions, fractional distillation is commonly practiced. Fractional distillation is carried out similarly to continuous distillation. However, in fractional distillation, the main fractionating tower has bubble cap trays or column packings equipped with the ability to remove side stream fatty acid distillates and return part of these streams as reflux (Anneken et al., 2000; Cermak et al., 2012; Stage, 1984). The boiling points between C12, C14, C16, C18, and C20 acids are sufficiently different that they can be separated by distillation (Table 2.2). However, fatty acids having similar chain lengths such as stearic (18:0), oleic (18:1), linoleic (18:2), and linolenic (18:3) are more difficult to cleanly separate by distillation (Gunstone et al., 1994). Because fatty acid composition varies by feedstock, the fractionating equipment is typically customized to suit a specific feedstock and product requirements. For example, coconut and palm kernel oils contain shorter chain saturated fatty acids and up to 30 trays can be utilized to obtain high-purity fatty acid fractions due to the higher volatility and

TABLE 2.2 Nomenclature of Selected Fatty Acids and Their Respective Melting and Boiling Pointsa Symbol

Systematic Name

Melting Point ( C)

Trivial Name

Boiling Point ( C/mm Hg)

Acid

Methyl Ester

Acid

Methyl Ester

Saturated Fatty Acids 4:0

Butanoic

Butyric

2 5.3

2 95.0

164(760)

103(760)

6:0

Hexanoic

Caproic

2 3.2

2 69.6

206(760)

151(760)

8:0

Octanoic

Caprylic

15.4

2 37.4

240(760)

195(760)

10:0

Decanoic

Capric

31.0

2 13.5

150(10)

108(10)

12:0

Dodecanoic

Lauric

44.8

4.3

173(10)

133 (10)

14:0

Tetradecanoic

Myristic

54.4

18.1

193(10)

161(10)

16:0

Hexadecanoic

Palmitic

62.9

28.5

212(10)

184(10)

18:0

Octadecanoic

Stearic

70.1

37.7

227(10)

205(10)

20:0

Eicosanoic

Arachidic

76.1

46.4

204(1)

188(2)

22:0

Docosanoic

Behenic

80.0

53.2

263(10)

240(10)

24:0

Tetracosanoic

Lignoceric

84.2

58.6



198(0.2)b

9c-16:1

9-Hexadecenoic

Palmitoleic

0.5

2 34.1

180(1)

182(10)

6c-18:1

6-Octadecenoic

Petroselenic

29.1

2 1.0

9c-18:1

9-Octadecenoic

Oleic

16.3

2 20.2

223(10)

201(10)

9t-18:1

9(trans)-Octadecenoic

Elaidic

43.4

9.9

Unsaturated Fatty Acids

(Continued )

TABLE 2.2 (Continued) Symbol

Systematic Name

Melting Point ( C)

Trivial Name Acid

Methyl Ester

Boiling Point ( C/mm Hg) Acid

Methyl Ester

11c-18:1

11-Octadecenoic

cis-Vaccenic

15.4

2 24.3

11t-18:1

11-Octadecenoic

trans-Vaccenic

43.4

9.9

9c-20:1

9-Eicosenoic

Gadoleic

23.0



170(0.1)

154(0.1)b

13c-22:1

13-Docosenoic

Erucic

33.5

2 3.5

255(10)

242(10)

9c,12c-18:2

9,12-Octadecadienoic

Linoleic

2 6.5

2 43.1

224(10)

200(10)

9c,12c,15c-18:3

9,12,15-Octadecatrienoic

ALA

2 11.3

2 52.4

137(0.07)

109(0.018)

6c,9c,12c-18:3

6,9,12-Octadecatrienoic

GLA

2 11.3

2 57

125(0.05)

162(0.5)

6c,9c,12c,15c-18:4

6,9,12,15-Octadecatetraenoic

Stearidonic

Approx. 257







5c,8c,11c,14c-20:4

5,8,11,14-Eicosatetraenoic

Arachidonic

2 49.5



163(1)

194(0.7)

5c,8c,11c,14c, 17c-20:5

5,8,11,14,17-Eicosapentaenoic

EPA









4c,7c,10c,13c,16c,19c,-22:5

4,7,10,13,16,19-Docosahexaenoic

DHA









a

Taken from Budde (1968), Gunstone (1996), Gunstone et al. (2007b), Knothe and Dunn (2009). Ethyl ester.

b

Naturally Occurring Fatty Acids: Source, Chemistry Chapter | 2

35

greater stability of the shorter chain fatty acids. In contrast, fractionating stills used for rapeseed oil, containing very long-chain fatty acids such as erucic acid (22:1) that has a higher boiling point and lower vapor pressure only requires a limited number of fractionating trays to keep the reboiler below the decomposition temperature (Berger and McPherson, 1979; Cermak et al., 2012).

2.3.3 Molecular Distillation In conventional vacuum distillation, further reductions to the pressure do not further lower the boiling point of a fatty acid due to the physical hindrance of vapor flowing through the distillation equipment at low pressures. By modifying the distillation equipment to have a short vapor distillation path, very low distillation pressures can be obtained and are known as molecular distillation (or short path distillation). Molecular distillation is industrially useful for the purification of unstable or highly oxidatively unstable fatty acids under vacuum conditions below 0.01 Torr. Under these high-vacuum conditions, the distilled fatty acids have a short exposure to elevated temperatures and a small distance between the evaporator and the condenser (Anneken et al., 2000; Cermak et al., 2012; Cvengroˇs et al., 2000; Lutiˇsan et al., 2002). Two main designs are utilized to obtain short residence times (a few seconds to tenths of seconds) of the fatty acid distillate in the distillation chamber, namely, wiped film and centrifugal molecular distillation units. In both of these designs, the liquid is distributed as a uniform thin film onto a hot glass wall by wiping or by centrifugal force from a spinning rotor. Using molecular distillation, Cermak et al. (2007) enriched C8 and C10 fatty acids from a mixture of cuphea oil fatty acids. Isbell and colleagues applied this technique to field pennycress oil to enrich erucic acid for industrial applications (Isbell et al., 2015). Martins et al. (2006) reported the separation of free fatty acids from vegetable oil deodorizer distillate. Breivik and coworkers prepared highly purified concentrates of EPA (5c,8c,11c,14c, 17c20:5) and DHA (4c,7c,10c,13c, 16c,19c-22:5) acids and corresponding esters from fish oil (Breivik et al., 1997).

2.3.4 Crystallization Apart from distillation, separation and fractionation of fatty acids can be accomplished through crystallization techniques that are based on the different melting point and solubility properties of fatty acids. Crystallization of fatty acids is appealing since it is mild, the fatty acid structure is not damaged, and large quantities may be processed. Early separations were based on mechanically pressing (panning) tallow fatty acid cakes prepared by slowly cooling the tallow in aluminum pans. The fatty acid cakes were wrapped in cloth to separate liquid fatty acids (unsaturated) from solid

36

Fatty Acids

(saturated) acids (Anneken et al., 2000). However, the fractionation efficiency was poor and so this method is no longer practiced. Crystallization of fatty acids can also be accomplished on an industrial scale without solvent using melt crystallization processes (Anneken et al., 2000; Gunstone et al., 1994; Tirtiaux, 1983). In these processes, a fatty acid mixture is slowly cooled to produce a slurry of solid and liquid fatty acids that are subsequently separated through different mechanical methods to obtain various degrees of fatty acid fractionation. Careful control of temperature and processing conditions is paramount to these methods. In these processes, aqueous solutions containing wetting agents such as magnesium sulfate can be utilized to facilitate crystallization of the solid phase and separation of the liquid phase through centrifugation (Anneken et al., 2000). Purification of unsaturated fatty acids by low-temperature crystallization from solvent, most commonly acetone or methanol, was first achieved by Brown and coworkers (Brown, 1955; Brown and Shinowara, 1937; Brown and Stoner, 1937). Saturated fatty acids are typically solid at ambient temperature and can be crystallized at temperatures down to 0 C while unsaturated fatty acids are usually liquid above 0 C and crystallize at lower temperatures between 0 and 290 C. This allows fatty acids to be crudely separated into higher melting and lower melting fractions that generally consist of saturated fatty acids and unsaturated fatty acids, respectively. However, fractionation of fatty acids into clean saturated and unsaturated fractions is oversimplified, as eutectic mixtures can form and residual saturated fatty acids are intersolublized in the unsaturated liquid fraction while the solid more saturated acid fraction can trap unsaturated fatty acids. In addition, crystallation is further complicated by the varying chain lengths of the fatty acids in the fatty acid mixtures (Brown, 1955; Schlenk, 1961). Industrially, two similar solvent-based crystallization procedures have been developed based on the use of either methanol or acetone to solubilize the fatty acids and are referred to as the Emerson and the Armour-Texaco processes, respectively. Both processes are mainly used to isolate stearic and palmitic acids from oleic acids from suitable feedstock such as tallow and tall oil (Gunstone et al., 1994; Wanasundara et al., 2005).

2.3.5 Urea Fractionation The formation of crystalline ureafatty acid complexes is a well-known technique to fractionate fatty acids. This method was first described by Bengen in 1940 (Bengen, 1951) and is used to separate straight chain compounds found in milk. Because urea is readily available and cheap, it has potential to be utilized on large-scale to fractionate fatty acids. When fatty acids (among other linear compounds) are added to a solution of urea under certain conditions, urea can form hydrogen-bonded hexagonal crystalline structures that form a series of linear, parallel channels having a diameter

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ranging from 0.55 to 0.58 nm that can entrap the alkyl chains of fatty acids (Gunstone et al., 1994; Hayes, 2002a,b; Hayes, 2006; Hayes et al., 1998; Swern, 1955). The resulting crystals can then be removed from the mother liquor and the complexed fatty acids subsequently separated from the urea. Depending on the structure of the fatty acid alkyl chain, mixtures of fatty acids or esters can be selectively complexed. Long-chain saturated chains form stable urea complexes while branched and polyunsaturated chains are less stable and do not readily form. In this way, ureafatty acid complexes have been used to fractionate fatty acid mixtures containing polyunsaturated from saturated fatty acids in oils derived from fish and linseed. Recently, γ-linolenic acid (6c,9c,12c-18:3; GLA) was fractionated and purified from lipids produced by Spriulina platensis in a purity of 84% with 64% recovery (Sajilata et al., 2008). Hayes has outlined the following general observations regarding the complexation of fatty acids by urea (Hayes, 2002a,b). (1) As the number of double bonds increases the ability to form an urea complex decreases; (2) longer chain fatty acids preferentially complex relative to shorter chain fatty acids; (3) fatty acids containing trans-double bonds are complexed preferentially over those with cis-double bonds; (4) the position of the double bond in the fatty acid chain influences the ability of complexation. The main drawbacks to the separation of fatty acids using urea are related to the large amounts of solvent, chemicals, and by-products involved (Breivik et al., 1997), although Hayes has proposed a simple and ecologically responsible method to fractionate fatty acids via urea complexation (Hayes et al., 1998).

2.4 SOURCES AND TYPES OF NATURALLY OCCURRING FATTY ACIDS The following sections describe several classes of naturally occurring fatty acids that can be found in plants and animals. They include the more commercially important fatty acids that contain saturated, unsaturated, and hydroxyl, functional groups, and fatty acids that contain unusual acetylenic, allenic, and cummulenic functional groups (Fig. 2.1). The saturated and unsaturated hydroxy fatty acids are emphasized because they are common constituents in commodity oils and fats, are oleochemical precursors and nutrionally or biologically important. While not common, fatty acids containing acetylenic, allenic, and cummulenic groups are also discussed since they are unusual and not mentioned in other chapters. The following sections are not exhaustive due to space constraints but are illustrative of the main fatty acids within each functional class and their sources. Although naturally occurring fatty acids containing other functional groups such as epoxides, branching, and cyclic rings are well known, they are not discussed here because they are the subjects of other chapters.

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2.4.1 Saturated Fatty Acids Saturated fatty acids are found in both plant and animal sources. Table 2.2 lists the more common and well-known naturally occurring fatty acids of interest. These typically have an even number of carbons and fats rich in saturated fatty acids have melting points that are higher than oils more abundant in unsaturated fatty acids. Odd chain fatty acids are found in trace amounts in animal and plant lipids but are more abundant in bacterial lipids. Shortchain fatty acids such as butyric acid (4:0) are found in cow milk fat at levels around 4 wt% while coconut and palm kernel oils are the primary commercial sources for medium-chain fatty acids (8:0, 10:0, 12:0, and 14:0), with lauric (12:0) and myristic (14:0) acids as the predominant saturated fatty acids contained in these oils. Several species of families such as Lythraceae, Myristicae, and Lauraceae contain high amounts of mediumchain 10:0, 12:0, and 14:0 saturated fatty acids. Cuphea (Lythraceae) is a large genus (over 200 species) of herbs and shrubs that is unique because its seeds that contain an abundance of the medium-chain fatty acids in the oil (Dubois et al., 2007). Because of this, cuphea is being examined as a potential industrial crop in Europe and the United States. Cuphea PSR-23 is a recently developed hybrid cross between Cuphea viscosissima and Cuphea lanceolata (Knapp and Crane, 2000). The seeds from this hybrid contain roughly 35 wt% oil that is composed of 82% capric (10:0), 3% lauric (12:0), 4% myristic (14:0), and 4% palmitic (16:0) acids in addition to minor amounts of unsaturated fatty acids (Cermak et al., 2012). Other Cuphea species such as C. pulcherrima, C. koehneana and C. calophylla contain .90% 8:0, .90% 10:0, and B85% 12:0, respectively (Scrimgeour and Harwood, 2007). Currently, cuphea is not a commercial crop as agronomic challenges such as seed shattering and indeterminate growth prohibit its large-scale production. Palmitic acid (16:0) is the most abundant saturated fatty acid found in animal lipids (20%30%), and in nearly all plant seed oils (5%50%) (Table 2.2). Useful amounts of palmitic acid (upwards of 50%) are obtained from palm oil and also from cottonseed oil, lard and tallow in approximately 20%30% (Gunstone, 1996). Although well known, stearic acid (18:0) is found in much lower amounts than palmitic acid. Lard and tallow are useful sources of stearic acid (Table 2.2). Cocoa and shea butter contain approximately 30%45% stearic acid. Of course, many of the saturated fatty acids can be prepared from the corresponding unsaturated fatty acids through hydrogenation. Saturated fatty acids having alkyl chain lengths greater than 18 carbon atoms are often negligible in most seed oils and are only found at useful levels in a few unusual seed oils such as peanut (5%8%, 20:0 and 22:0) and rapeseed (2.0%60%, 22:1), and are often present in waxes.

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2.4.2 Unsaturated Fatty Acids Naturally occurring unsaturated fatty acids are those in which carboncarbon double bonds (typically in the cis-configuration) reside along the hydrocarbon backbone. Monounsaturated (monoenoic) fatty acids contain one double bond while polyunsaturated (polyenoic) acids contain multiple double bonds along the alkyl chain (Fig. 2.2). The position of the double bond along the alkyl chain is found mainly in a limited number of preferred positions (the Δ9-position is most common for 18 carbon fatty acids; Δxx designates the location of the double bond carbon in closest proximity to the carboxyl group) resulting from their biosynthesis. In PUFAs, the double bonds are nonconjugated and often separated by a methylene (CH2) group. Although these general trends are commonly observed for unsaturated fatty acids, exceptions abound and unsaturated fatty acids having trans-configurations, unusual double bond positioning, and conjugated double bonds are also be found in nature.

2.4.2.1 Monounsaturated Fatty Acids Cis-monounsaturated fatty acids (MUFAs) are common constituents of nearly all commodity oils. Over 100 MUFAs are known (Gunstone, 1996). Through biosynthetic pathways, a variety of MUFAs can be produced through desaturase and chain-elongation enzyme reactions, whereby the position of the double bond is commonly found at the Δ6-, Δ9-, or Δ12-position in the alkyl chain. Desaturase enzymes catalyze the removal of two hydrogen atoms from the alkyl chain of a saturated fatty acid to create a double bond (typically cis) while elongation enzymes add carbons in two carbon increments to the carboxyl end of the fatty acid (Harwood, 2007). For example, a Δ9-desaturase enzyme inserts a double bond into 16:0 and 18:0 saturated fatty acids to produce the corresponding 9c-16:1 (palmitoleic) and 9c-18:1 (oleic) MUFAs, respectively (Cahoon et al., 1992). Oleic acid can be subsequently elongated by two carbon atoms at the carboxyl end to give gondoic acid (11c-20:1). Similarly, palmitic acid (C16:0) can be oxidized by a desaturase enzyme to palmitoleate (9c-16:1), which can subsequently be elongated to cis-vaccenic acid (11c-18:1). The 16-carbon MUFA, palmitoleic acid (9c-16:1), is a minor acid found in animal lipids and fish oils, although it is found in 20%30% and 25%40% in macadamia nuts and the pulp of sea buckthorn, respectively. Among naturally occurring MUFAs, oleic acid (9c-18:1) is the most prevalent and is found in high amounts in many oils such as olive, peanut, palm, canola/rapeseed, and sunflower (high oleic), as well as lard and tallow (Table 2.1). Cis-MUFAs having 18 or less carbon atoms are typically liquids at room temperature while cis-MUFAs having greater than 18 carbons are low-melting solids. MUFAs having a trans-double bond configuration have higher melting points that are similar to the melting points of the

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FIGURE 2.2 Structures of some monounsaturated and PUFAs.

corresponding saturated fatty acids. In addition to geometrical configuration, double bond position also influences melting point. For example, both cisand trans-18:1 MUFAs are higher melting when the double bond is located at an even position than at an odd position; a pattern most distinct for double bonds located between carbons 4 and 14.

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Other 18 carbon MUFAs include vaccenic acid isomers (11c- and 11t18:1), which are minor components of dairy products. These are also found in sea buckthorn oil. Elaidic acid (9t-18:1), the trans isomer of oleic acid, is found in small amounts in milk. Petroselinic acid (6c-18:1) is the major component in the seed oils of Apiaceae such as coriander, which is biosynthesized from palmitic acid by a Δ4-desaturase enzyme followed by a two-carbon elongation step (Scrimgeour and Harwood, 2007). In coriander, the seeds contain 1230 wt% oil that is composed of approximately 57% 74% petroselinic acid relative to other fatty acids of the oil (Isbell, 2009). Petroselinic acid is also found in Araliaceae, Garryaceae, and Geraniaceae species. The Δ6-double bond in petroselinic acid renders it an interesting renewable feedstock for adipic acid by oxidative cleavage of the double bond. Erucic acid (13c-22:1) is an important long-chain MUFA found in useable quantities in seed oils of the Brassicaceae and Limnanthaceae families. Erucic acid is biosynthesized by elongation of oleic acid and is available on commercial scale from crambe (60%), high-erucic rapeseed (45%50%), and mustard (42%60%) seed oils. Meadowfoam seed oil from Limnanthes alba is unique in that 94 wt% of the fatty acids contain 20 or more carbon atoms. The main monounsaturated fatty is eicosenoic acid (5c-20:1) in amounts around 63% along with 17% of polyunsaturated docosadienoic acid (5c,13c-22:2) (Hayes et al., 1995).

2.4.2.2 Polyunsaturated Fatty Acids The primary natural PUFAs contain two to six double bonds, typically in the cis-configuration, that are nonconjugated and separated from one another by methylene (CH2) groups (Fig. 2.2). In plants, the number of double bonds in fatty acids rarely exceeds three, although, algae and animal (fish) fatty acids can contain up to six double bonds. These PUFAs are generally liquid at room temperature and can be further classified into two principal groups designated as n-3 and n-6 fatty acids. The n-3 and n-6 designations refer to the position of the double bond carbon in the chain closest to the terminal methyl end rather than the systematic numbering from the carboxyl end. The n-3 and n-6 fatty acids are present in most plant, animal, and commodity oils and fats. As noted for MUFAs, desaturases and chain-elongation enzymes are responsible for the production of PUFAs derived from plants, in which the double bonds are typically at the Δ9-, Δ12-, and Δ15-positions to give the corresponding n-3 and n-6 fatty acids (Scrimgeour and Harwood, 2007). Linoleic acid (9c,12c-18:2; n-6) and ALA (9c,12c,15c-18:3; n-3) are commonly found in most plant oils and abundant in several commodity oils. These fatty acids cannot be synthesized in humans and animals and must be obtained through diet. In 1929 Burr and Burr showed that linoleic and ALA are essential to proper functioning and health of human and animals (Choque

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et al., 2014). The longer chain ( . 20 carbons) PUFAs in the n-3 and n-6 families, which are also important to human health, are biosynthetically derived from linoleic acid and ALA, respectively. 2.4.2.2.1

The n-6 Polyunsaturated Fatty Acids

Linoleic acid is the shortest chain n-6 fatty acid and the most common PUFA in plant oils and can be present in commercial oils at levels .50% in cottonseed, corn, soybean, safflower, and sunflower (Table 2.1). In the human diet, linoleic acid is subsequently converted by desaturase and elongation enzymes in the liver into other longer chain n-6 PUFAs such as arachidonic (5c,8c,11c,14c-20:4) and docosapentaenoic (4c,7c,10c,13,16c-22:5) acids. Arachidonic acid is an important metabolite of linoleic acid and allegedly plays a role in obesity and in vivo enzymatic production of proinflammatory prostoglandins, thromboxanes, and leukotrienes implicated as mediators and regulators of inflammatory responses and other essential biological functions (Choque et al., 2014). Although arachidonic acid can be biosynthesized in the body, meat and fish serve as the main sources of arachidonic acid in the human diet. The yeast Mortierella alpina is a commercial source of arachidonic acid via fermentation (Harwood, 2007). Another member of the n-6 PUFA series is GLA (6c,9c,12c-18:3), and is an isomer of ALA. It is a minor component in animal fats and can be obtained in commercial quantities from the seed oils of evening primrose (Oenothera biennis), borage (Borago officinalis), and black currant (Ribes nigrum) in approximately, 9%12%, 20%25%, and 15%17%, respectively (Gunstone, 1996). In humans, linoleic acid is a precursor to GLA and is the first intermediate in the conversion of linoleic acid into arachidonic acid. Recently, GLA was examined more closely because of health and dietary claims suggesting that GLA prevents or alleviates a wide variety of human diseases (Guil-Guerrero et al., 2001). 2.4.2.2.2 The n-3 Polyunsaturated Fatty Acids The n-3 PUFA, ALA (9c,12c,15c-18:3) is found in most plant oils [soybean, canola, flax seed (or linseed), soy, perilla], nuts (walnuts), and some animal fats. As with linoleic, ALA is considered an essential fatty acid and liver desaturase and elongation enzymes convert ALA into a series of n-3 longerchain PUFAs such as stearidonic acid (6c,9c,12c,15c-18:4), eicosatetraenoic acid (8c,11c,14c,17c-20:4), EPA (5c,8c,11c,14c, 17c-20:5), and DHA (4c,7c,10c,13c, 16c,19c-22:5), although with low efficiency and in competition with n-3 fatty acid synthesis. EPA (20:5) and DHA (22:6) acids are n-3 PUFAs of great interest due to their health benefits. For instance, evidence supports their beneficial role in the prevention of various diseases, and are mainly obtained through diet. They are rarely found in plants but are commonly encountered in marine oils with fish and algal oils serving as the most

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significant commercial sources. Stearidonic acid (6c,9c,12c,15c-18:4; n-3) is a minor component of animal lipids and fish oils. It is found at low levels in blackcurrent (R. nigrum) and echium (Echium plantagineum) seed oils at approximately 5% and 7%, respectively (Scrimgeour and Harwood, 2007).

2.4.2.3 Conjugated Polyunsaturated Fatty Acids Naturally occurring fatty acids that contain conjugated double bonds are encountered in certain plants and animals. For example, calendic acid (8t,10t,12c-18:3) is a conjugated n-6 polyunsaturated trienoic fatty acid with a melting point of approximately 40 C that is found in Calendula officinalis (marigold) oil in approximately 53%62%. The conjugation in calendic acid makes it a good source of drying oils for reactive diluents in coatings such as paint (Harwood and Gunstone, 2007). Another well-known conjugated trienoic acid is 9c,11t,13t-18:3 (α-eleostearic acid, n-5), which is encountered in commercially available tung oil (Aleurites fordii) at levels of approximately 70%. Traditionally, paints were made using highly unsaturated oils like tung (or linseed oil; contains a large proportion of 9c,12c,15c-18:3) in which the highly unsaturated fatty acids undergo oxidation and subsequent polymerization. Conjugated linoleic acids (CLA) are a family of positional and geometrical isomers of linoleic acid that are found in meat and dairy sources (especially grassfed animals) as a byproduct of biohydrogenation of linoleic acid by microorganisms in ruminant animals (Pariza et al., 2001). The 9c,11t18:2, 10t,12c-18:2, 9t,11t-18:2, and 10t,12t-18:2 conjugated fatty acid isomers account for more than 90% of the CLA produced from linoleic acid via alkali isomerization. The 9c,11t-18:2 isomer is found in milk fat (approximately 1%) and is believed to be the CLA isomer responsible for interesting physiological effects (Pariza et al., 2001). CLA is being examined and marketed as a dietary supplement based on various health claims such as antiatherosclerotic and cancer effects in addition to reducing fat mass (Chin et al., 1992; Park et al., 1997).

2.4.3 Hydroxy Fatty Acids The principal naturally occurring hydroxy fatty acids are ricinoleic (12hydroxy-cis-9-octadecenoic), lesquerolic (14-hydroxy-cis-11-eicosenoic), densipolic (12-hydroxy-cis-9-cis-15-octadecadienoic), and auricolic (14hydroxy-cis-11-cis-17-eicosadienoic) acids. Fig. 2.3 depicts the structure of several hydroxy-containing fatty acids. They are found in many genera in several unrelated plant families such as Apocynaceae, Asteraceae, Brassicaceae, Coriariaceae, Euphorbiaceae, Fabaceae, Malpighiaceae, and Papaveraceae (Badami and Patil, 1980). Fatty acid-hydroxylation enzymes from animals, plants, and microorganisms typically introduce the hydroxy

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Fatty Acids

FIGURE 2.3 Structures of some hydroxyl-containing fatty acids.

group during fatty acid biosynthesis and produce a wide variety of hydroxy fatty acids (Harwood, 2007; Kim and Oh, 2013). The hydroxy moiety gives these fatty acids unique functionality and polarity that are advantageously utilized for a range of industrial products including lubricants, plastics, coatings, surfactants, cosmetics, and pharmaceuticals (Mutlu and Meier, 2010; Ogunniyi, 2006). In addition, castor oil can also be dehydrated to give a polyunsaturated oil containing a large proportion of CLA having the 9c,11t18:2 isomer (Gunstone, et al., 2007a). The best known hydroxy fatty acid is ricinoleic acid (12(R)-hydroxy-octadec-cis-9-enoic acid; 12-OH 9c-18:1), which is obtained agriculturally from castor oil (Ricinus communis L.; Euphorbiaceae). Currently, castor oil is the only oil of commercial significance that contains a hydroxy fatty acid. 12-Hydroxylase (EC 1.14.13.26) introduces the hydroxy group in ricinoleic acid stereospecifically, having an R-configuration at the carbon atom containing the hydroxy group (van de Loo et al., 1995). Ricinoleic acid is the main constituent (approximately 90%) of the total fatty acids contained in castor oil and is accompanied by small amounts of palmitic, stearic, oleic, linoleic, and 9,10-dihydroxystearic acids. Globally, the leading producers of castor are India and China, which

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collectively exceed 80% of total world production (Mutlu and Meier, 2010; Zanetti et al., 2013). Because castor seed contains ricin, a highly toxic lectin protein, alternative sources of hydroxy fatty acids are sought. Lesquerella oil produced by plants from the genus Lesquerella, such as L. fendleri and L. gordonii, is a new crop being developed for their hydroxy fatty acid content and was recently reviewed by both Isbell (Isbell, 2009) and Zanetti (Zanetti et al., 2013). These plant species produce seeds that contain approximately 30% oil that is rich in lesquerolic acid (14-OH 11c-20:1), a C20 homolog of ricinoleic acid. For example, the oil from L. fendleri contains lesquerolic and auricolic (14-OH 11c,17c-20:2) acids in 54%60% and 3%5%, respectively (Frykman and Isbell, 1997). Other Lesquerella species contain large quantities of related hydroxy fatty acids such as densipolic acid (12-hydroxy-octadec-cis-9,15-dienoic acid; 12-OH 9c,15c-18:2) (Hayes et al., 1995). Another hydroxy fatty acid similar to ricinoleic acid, isoricinoleic acid (9-hydroxy-cis-12-octadecenoic acid; 9-OH 12c-18:1), is a major component of the seed oils of Wrightia (Ahmad et al., 1986) and Strophanthus (Gunstone, 1996). The seeds of Dimorphotheca pluvialis contain 13%28% oil that is composed of up to 54% of dimorphecolic acid (9-hydroxy,-10trans,-12-trans-octadecadienoate; 9-OH 10t12c-18:2), an unusual C18 hydroxy fatty acid with the hydroxy group adjacent (allylic) to a conjugated diene system (Smith et al., 1960). This acid structure is chemically unstable and is readily dehydrated to a mixture of conjugated 18:3 acids (Δ8,10,12 and Δ9,11,13), which has characteristics similar to tung oil.

2.4.4 Acetylenic Fatty Acids Naturally occurring acetylenic fatty acids contain a carboncarbon triple bond (HCCH), are not readily available, and are often unstable compounds (Fig. 2.4). These fatty acids have a variety of biochemical and ecological functions and are found in plants, mosses, lichens, fungi, and algae. Reviews by both Minto and Kuklev have reported various aspects of naturally occurring acetylenic compounds (Dembitsky, 2006; Kuklev and Dembitsky, 2014; Kuklev et al., 2013; Minto and Blacklock, 2008). Tariric acid (6-octadecynoic acid), biosynthesized from petroselinic acid, was the first naturally occurring acetylenic fatty acid to be discovered and was identified in the seed oil of Picramnia sow (Simaroubaceae) in the 19th century (Arnaud, 1892). Probably, the most familiar acetylenic fatty acid is santalbic acid (trans-octadec-11-en-9-ynoic acid or ximenynic acid in older literature), which is found in members of Santalaceae, Olacaceae, and Opiliaceae families (Aitzetmu¨ller, 2012). Santalbic acid was first discovered in 1938 in the seeds of Santalum album (Madhuranath and Manjunath, 1938) for which the structure was elucidated in 1954 by Gunstone (Gunstone and Russell, 1955). Santalbic acid can reach up to 95% of the total seed oil fatty

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FIGURE 2.4 Structures of some acetylenic- and allenic-containing fatty acids. aThese numbers designate the total number of unsaturated centers irrespective of the bond type.

acids and is quite often found above 70% (Aitzetmu¨ller, 2012). Although santalbic acid is generally considered to be toxic to humans, it is of the interest to the chemical industry due to its reactive conjugated ene-yne groups. Stearolic acid (9-octadecynoic acid) is the acetylenic analog of oleic acid and mainly found as a minor component in nature. For example, santalbic acid and stearolic acid were isolated from Exocarpos and Santalum seeds. About 30%35% of total fatty acid composition was santalbic acid and 1%

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was stearolic acid. However, stearolic acid is found abundantly in some Pyrularia species such as P. edulis, which can contain over 50% of the fatty acid (Scrimgeour and Harwood, 2007). In the seed oil of Heisteria silvanii (Olacaceae), two conjugated ene-yne acetylenic fatty acids, santalbic acid and pyrulic acid, trans-10-heptadecen-8ynoic acid, were isolated in 3.5% and 7.4%, respectively (Spitzer, et al., 1997a). In addition, heisteric acid, cis-7, trans-11-octadecadiene-9-ynoic acid, was also isolated in 22.6%. The seed oil from Alvaradoa amorphoides (Simarubiaceae) contains 58% of tariric acid (Pearl et al., 1973). This is similar to that of Picramnia seed oils, however, in addition to tariric acid the oil also contains 15% of 17octadec-6-ynoic acid and a trace amount of 20-carbon 6-eicosynoic acid. Screening of seed oils for lipid content led to the discovery of acid (cis9-octadecen-12-ynoic acid), an acetylenic-containing fatty acid that was isolated from the seed oil of Crepis foetida (Compositae) (Mikolajczak et al., 1964). This fatty acid was the major component of the oil in approximately 60%. Gunstone and coworkers found that the seed oil of Afzelia cuanzensis (Caesalpiniaceae) contained not only crepenynic acid (44%) but also dehydrocrepenynic acid (9,14-octadecen-12-ynoic acid), which was 21% of the total fatty acids present in the oil (Gunstone et al., 1967). Vlahov later determined the acyl-composition and positional distribution of these fatty acids in the triacylglycerol mixture by nuclear magnetic resonance (Vlahov, 1996). The oil of Helichrysum bracteatum (Vent.) Andrews (Compositae) contains helenynolic acid (9-hydroxy-trans-10-octadecen-12-ynoic acid) in addition to crepenynic acid (Powell, 2009). Powell and coworkers found that 9D-hydroxy-cis-12-octadecenoic acid could be isolated from three seed oils of the family Apocynaceae: Holarrhena antidysenterica, Nerium oleander, and Nerium indicum in 73%, 11%, and 8%, respectively (Powell et al., 1969). The known occurrence of this acid was previously limited to the genus Strophanthus (9%15%), as reported by Gunstone (Gunstone, 1952).

2.4.5 Allenic and Cumulenic Fatty Acids Allenic- and cumulenic-containing fatty acids are uncommon and only limited progress toward obtaining high-level accumulation of these constituents has been made. Allenic and cumulenic fatty acids are rare in common commercially grown oilseed crops. However, they exist in less common plant species.

2.4.5.1 Allenic Fatty Acids Naturally occurring fatty acids that contain an allenic (HC 5 C 5 CH) moiety are an interesting group of fatty acids that represent a subset of naturally occurring unsaturated compounds (Fig. 2.4). The allenic group is

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comprised of two double bonds that share a common carbon atom and are rigidly held in place at right angles to one another. This arrangement of double bonds causes a twist in the molecule resulting in optical activity when asymmetrically substituted. Allenic fatty acids exhibit cytoxic, antibacterial, and antiviral activities (Dembitsky and Maoka, 2007). The most recent review on allenic fatty acids was published in 2007 by Dembitsky and Maoka (Dembitsky and Maoka, 2007) although earlier reviews have also covered various aspects of allenic and cumulenic fatty acids (HoffmannRo¨der and Krause, 2004; Smith, 1971; Taylor, 1967). Most naturally occurring allenic-containing fatty acids are found mainly as fungal metabolites in which the allenic group is part of a larger conjugated system. With regard to higher plants, laballenic [(2)-5,6-octadecadienoic or 18:2Δ5,6 allene], lamenallenic [(2)-octadeca-5,6-trans-16-trienoic or 18:3Δ5,6 allene Δ16t], and phlomic (7,8-eicosadienoic or 20:2Δ7,8 allene) acids are components of some unique seed oils. Specifically, plants from the Lamiaceae (Labiatae) species are a natural source of allenic fatty acids in up to 28% of the oil (Aitzetmu¨ller et al., 1997; Sinha et al., 1978). These seed oil-derived allenic fatty acids differ from the corresponding fungi-derived acids in that their allene group is not part of a conjugated system. The seed oil of Leonotis nepetaefolia contains laballenic acid, a C18 allenic fatty acid, in approximately 16% (Bagby et al., 1965). Synthesis of laballenic acid confirmed that it has an absolute R-configuration (Bohlmann et al., 1967; Landor and Punja, 1966). Lamium purpureum seed oil contains 16% of lamenallenic acid, whereby the acid was isolated as its methyl ester from a mixture of methyl ester fatty acids after transesterification of the oil (Mikolajczak et al., 1967). Lamenallenic acid was subsequently reported to be strongly levorotatory (Cowie et al., 1972; Mikolajczak et al., 1967). Aitzetmu¨ller and coworkers reported that the allenic fatty acid, phlomic acid, is present in small amounts in several genera of the Lamiaceae, a subfamily Lamioideae (Aitzetmu¨ller et al., 1997). The occurrence of phlomic acid was correlated with the presence of the unusual gadoleic (9c-20:1) or gondoic (11c-20:1) acids, and these are apparently produced by chain elongation of laballenic acid present in the seed oil. Chinese tallow tree seeds such as Sapium sebiferum are unusual in that they contain a desirable highly saturated oil (Chinese vegetable tallow) in the outer seed coating in approximately 2030 wt% while the kernel contains the inedible highly unsaturated oil, Stillingia oil, in approximately 1017 wt% (Jeffrey and Padley, 1991). In addition to the common oleic, linoleic, and linolenic acids, the Stillingia oil also contains a tetraester triglyceride, or estolide, in approximately 23% that is composed of 8-hydroxy5,6-octadienoic acid esterified to the glycerol backbone by the carboxylic acid moiety and also esterified at the 8-hydroxy position with 2-trans,4-cisdecadienoic acid to form the estolide linkage (Sprecher et al., 1965). Christie showed that the estolide from S. sebiferum occurs exclusively in the

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sn-3-position of the glyceride moiety and suggested that the optical activity of this molecule is mainly caused by the allenic system (Christie, 1969). Similar compounds have also been isolated and identified from Sebastiana ligustrina (Heimermann and Holman, 1972) and Sebastiana commersoniana (Euphorbiaceae) (Spitzer et al., 1997b). Naturally occurring dicarboxylic acids containing an allenic group have also been identified. Glutinic acid (2,3-pentadienedioic acid) was isolated from the leaf resin of European alder, Alnus glutinosa (Betulaceae), in 1908 (Hans, 1908). The structure of glutinic acid was subsequently confirmed in 1958 by Corsano, Capito, and Bonamico (Corsano et al., 1958) and its absolute configuration was established in 1962 (Agosta, 1962).

2.4.5.2 Cumulenic Fatty Acids The cumulenic (HC 5 C 5 C 5 CH) moiety is the one carbon atom extended homolog to the allenic group, whereby, three carboncarbon double bonds are rigidly arranged sequentially at right angles to one another. Cumulenes are generally unstable and only several cumulenic fatty acids have been identified in nature. From the root extracts of Austrian Chamomile (Anthemis austriaca), the methyl ester of 2,6,7,8-decatetraen-4ynoic acid and 9-(methylthio)- in conjunction with two isomeric δ-lactones of 5-hydroxy-9-(methylthio)-2,4,6,7,8-decapentaenoic acid was also isolated (Bohlmann and Hopf, 1973; Bohlmann and Zdero, 1971). Fatty acids containing the cumulenic functionality were isolated from scentless mayweed (Matricaria inodora L.) (Sorensen and Stavholt, 1950) in addition to two cumulene phenolics. Bohlmann and Zdero isolated and identified unstable cumelenes from Erigeron canadense (fleabane, Compositae) (Bohlmann and Zdero, 1970).

2.5 CHEMISTRY OF NATURALLY OCCURRING FATTY ACIDS Fatty acids have a long and important history as substrates for chemical synthesis. For instance, ancient civilizations such as the Sumerians and Babylonians were making soaps 40005000 years ago. Roman and Chinese candles of antiquity were produced from tallow and whale fat. By the Middle Ages, factories dedicated to soap manufacturing were found throughout Europe. More recently, biodiesel (fatty acid methyl esters) was first reported in the early 1920s. These examples illustrate the ease with which fatty acids can be modified by chemical means. This is because fatty acids contain a polar carboxylic acid and a nonpolar hydrocarbon backbone with or without one or more double bonds. Carboxylic acids and esters along with olefins are readily amenable to chemical modification. The bifunctional nature of unsaturated fatty acids thus renders them as versatile building blocks for the synthesis of a wide variety of bio-based industrial products.

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Fatty Acids

Chemical modification of the carboxylic acid and olefinic moieties will be the focus of this section, as these are the sole functional groups encountered in the five most common naturally occurring fatty acids (palmitic, stearic, oleic, linoleic, and linolenic). In addition, industrial uses of chemically modified fatty acids will be discussed. The first topic will be reactions at the carboxylic acid moiety of fatty acids.

2.5.1 Reactions at the Carboxylic Acid Group The carboxylic acid functional group readily undergoes reduction, esterification, amination, and deoxygenation, as shown in Fig. 2.5. Such transformations lead to products with useful properties, and are thus of commercial interest. Each will be discussed in greater detail in the following subsections.

2.5.1.1 Reduction Reduction of the carboxylic acid moiety results in a fatty alcohol (Fig. 2.5, reaction a). From a practical standpoint, production of fatty alcohols is more readily achieved from a methyl ester than a fatty acid. This is mostly because fatty acids create an acidic reaction medium that requires corrosion-resistant equipment and acid-resistant catalysts. Nevertheless, catalytic hydrogenolysis of fatty acids to fatty alcohols is achieved under pressure (2530 MPa) at 300 C with a copper chromite catalyst and a stoichiometric excess of hydrogen. Hydrogenolysis of methyl esters is the favored industrial route to fatty alcohols because lower temperatures (200250 C) are needed and there are no corrosion or catalyst stability issues (Voeste and Buchold, 1984). Production of unsaturated fatty alcohols is complicated by the fact that double bonds are also reduced (saturated) under the conditions of hydrogenolysis. Consequently, chemoselective catalysts such as zinc chromite must be

R

OH

a

O

d R H

R

O

b OH

R

O

R'

c

R

NH2

FIGURE 2.5 Reactions at the carboxylic acid moiety of fatty acids: a, reduction; b, esterification; c, amination; and d, deoxygenation (hydrodeoxygenation depicted).

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51

utilized to preserve unsaturation (Kreutzer, 1984). For example, New Japan Chemical Company’s (1973) selective hydrogenolysis of methyl oleate to give octadec-9-en-1-ol is accomplished commercially using a complex aluminumcadmiumchromium oxide catalyst in a continuous process at 270290 C and 19.6 MPa. Fatty alcohols and their derivatives have a wide variety of uses in consumer and industrial products either because of their surface-active properties or as a means of introducing a long-chain hydrocarbon moiety into a chemical compound (Peters, 1991). The primary use is as surfactants in detergents and cleaning products. Most fatty alcohols are used as derivatives such as poly(oxyethylene) ethers, the corresponding sulfated ethers, and alkyl sulfates, and as esters with phosphoric acid and mono- and dicarboxylic acids. Detergent-range fatty alcohols serve as building blocks for all major surfactant categories: anionic, cationic, nonionic, and zwitterionic. The alkyl sulfates derived from C12C15 fatty alcohols, such as sodium dodecyl sulfate, are incorporated into shampoos, toothpastes, dishwashing detergents, and household cleaners because of their cleaning ability, mildness, and foaming properties. The alkyl sulfates resulting from C16C18 fatty alcohols are found in powder laundry detergents and other heavy-duty cleaners. Surfactants derived from polyethoxylated alcohols are in wider use than alkyl sulfates because they are less irritating to the skin and perform better in hand dishwashing and laundry detergents. Other uses of fatty alcohols include cosmetic, pharmaceutical, lubricant, and petroleum applications (Peters, 1991). For instance, cetyl and stearyl alcohols serve as emollient additives and as bases for creams, lipsticks, ointments, and suppositories. Methacrylate esters of fatty alcohols find use as viscosity index improvers, pour point depressants, and dispersants in engine lubricants. Esters of docosanol (behenyl alcohol) are used as drag reducing agents for crude petroleum oil pipelines and a composition of octadec-9-en-1-ol and sodium lauryl sulfate is used for enhanced petroleum recovery. Fatty alcohols also serve as starting materials for fatty amine synthesis. In summary, applications of fatty alcohols and their derivatives span industrial, personal care, cosmetics, pharmaceutical, food, and petroleum industries (Egan et al., 1984).

2.5.1.2 Esterification Fatty acid alkyl esters are produced by two primary routes: direct esterification of fatty acids (Fig. 2.5, reaction b) or transesterification of triglycerides. Esterification of fatty acids takes place at elevated temperature in the presence of a stoichiometric excess of alcohol and a strong mineral acid catalyst such as sulfuric or hydrochloric acid (Miao and Shanks, 2011). Removal of water drives the equilibrium toward the desired product, as esterification is otherwise reversible if water remains in the reaction medium (Formo, 1954). Stoichiometrically, one fatty acid molecule condenses with one alcohol to yield a single fatty ester with one molecule of water eliminated as a byproduct.

52

Fatty Acids

Transesterification of triglycerides is conducted at elevated temperature in the presence of excess alcohol and an alkaline (base) catalyst. When methanol is used as the alcohol, transesterification is referred to as methanolysis and fatty acid methyl esters are produced. Methyl esters are the most common synthetic fatty esters because methanol is comparatively inexpensive relative to other monohydric alcohols and possesses the greatest volatility, thereby facilitating its removal and recovery from the reaction mixture. The classic conditions for methanolysis are 1 hour of reaction at 60 C with a 6:1 molar ratio of methanol to triglyceride and 0.5% (by weight of oil) alkaline catalyst (Freedman et al., 1984). The most common alkaline catalysts are sodium or potassium hydroxide and sodium methoxide. However, the triglyceride starting material must contain a low level of endogenous free fatty acids (,0.5 wt%) to avoid deactivation of the alkaline catalyst by reaction with free fatty acids to form soaps. In such cases, acid catalysis is preferred, as it catalyzes both esterification and transesterification to yield fatty esters free from soaps (Lotero et al., 2005). Acid catalysis is not desirable for triglycerides with low free fatty acid levels (,0.5 wt%) because it is about 4000 times slower than alkaline catalysis (Srivastava and Prasad, 2000). Numerous other catalysts are also suitable for transesterification of triglycerides, including various homogeneous and heterogeneous acids, bases, and enzymes (Di Serio et al., 2008; Lotero et al., 2005; Narasimharao et al., 2007; Ranganathan et al., 2008). The acidic catalysts are also generally suitable for direct esterification of fatty acids as alternatives to the mineral acid approach discussed previously. Industrial production of fatty esters is accomplished via transesterification of triglycerides as opposed to direct esterification of fatty acids. Because methanol is the most common alcohol employed during transesterification, fatty acid methyl esters are the most common fatty acid alkyl esters. Industrial applications of methyl esters include direct use as solvents or biodiesel, or as starting materials for fatty alcohol synthesis. Glycerol is also produced during transesterification and has numerous applications that are reviewed elsewhere (Behr et al., 2008; Behr and Gomes, 2010).

2.5.1.3 Amination Fatty amines are obtained from fatty acids (Fig. 2.5, reaction c) or esters by a multistep sequence. Initial reaction with ammonia at high temperature yields a fatty amide. Subsequent dehydration provides a fatty nitrile, which in turn is converted to a fatty amine by hydrogenation. In practice, fatty amines are prepared in two steps. First, a fatty nitrile is synthesized by concomitant amidation and dehydration. Second, the fatty nitrile is catalytically hydrogenated to yield a fatty amine. Routes to primary, secondary, tertiary, and quaternary fatty amines are depicted in Fig. 2.6.

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53

O R

OH

NH3 H2O O R

NH2

H2O

R

N R

c

R

a

N R

H2 NH3

d

b H

NH3 + N ClR

NH2

O H2

R

R

H2, NH3

g MeCl

R

N H

R

H

R

N

O e H i

R N+

R O O-

R

O HO

MeCl

f

H

N

R R

+ N Cl-

Cl MeCl

R

h

+ N ClR

FIGURE 2.6 Preparation of fatty amines from fatty acids: a, primary; b, secondary difatty; c, symmetrical tertiary; d, tertiary fatty dimethyl; e, tertiary difatty methyl; f, quaternary fatty trimethyl ammonium chloride; g, h, quaternary difatty dimethyl ammonium chloride; and i, betaine.

The principal industrial route to fatty amines is hydrogenation of nitriles. Fatty nitriles are readily prepared by batch or continuous processes at 280360 C in the presence of ammonia and a catalyst. In batch mode, zinc oxide is commonly employed as the dehydration catalyst, whereas bauxite is used for continuous processes. Other dehydration catalysts include alumina,

54

Fatty Acids

thorium, titanium oxide, manganese acetate, and cobalt salts. Removal of water generated during dehydration drives the reaction to completion. Hydrogenation of fatty nitriles to the corresponding amines is conducted at 50200 C using hydrogen at elevated pressure (3.5 MPa) in the presence of catalytic nickel or cobalt and excess ammonia to suppress secondary amine formation (Greenfield, 1967). Retention of unsaturation in the case of unsaturated fatty amines requires modification of hydrogenation conditions to minimize double bond saturation and cis/trans isomerization. Primary amines and their salts find applications as additives for fuels, bactericides, flotation agents, and as water repelling agents. They also serve as intermediates in the production of quaternary ammonium salts. The major product of nitrile hydrogenation is typically the primary amine, but production of secondary and tertiary amines is promoted by adjusting reaction conditions. For instance, high-purity secondary amines are easily prepared from nitriles at high temperature and low pressure during hydrogenation without excess ammonia. In fact, ammonia generated during secondary amine formation is continuously removed to drive the reaction to completion. This is because two primary amines condense to form a secondary amine with elimination of ammonia in a reversible reaction (equilibrium). Furthermore, copper chromite catalysts promoted with alkaline or alkaline earth compounds enhance selectivity for secondary amines (Barrault and Pouilloux, 1997). The primary industrial application of secondary amines is as intermediates in the production of difatty dimethyl quaternary ammonium salts. Symmetrical tertiary amines are prepared in an analogous manner to secondary amines. Asymmetrical tertiary amines such as methyl difatty amines and dimethyl fatty amines are prepared by reductive alkylation with formaldehyde using nickel hydrogenation catalysts (Shapiro and Frank, 1964). Tertiary amines are used as bactericides, corrosion inhibitors, cosmetics ingredients, emulsifiers, foaming and flotation agents, fuel additives, fungicides, and as intermediates for quaternary ammonium salts (Billenstein and Blaschke, 1984). Quaternary ammonium compounds are prepared by alkylation of fatty, fatty dimethyl, difatty, and difatty methyl amines with methyl chloride, dimethyl sulfate, or benzyl chloride (Billenstein and Blaschke, 1984; Reck, 1985). The most important of these are difatty dimethyl ammonium salts produced by reaction of secondary difatty amines with methyl chloride. For instance, distearyl dimethyl ammonium chloride is used as a bactericide. Tertiary amines give bactericidal quaternary ammonium salts upon reaction with benzyl or methyl chloride or dimethyl sulfate (Billenstein and Blaschke, 1984). The largest market for quaternary fatty amines is in fabric softeners. Monofatty quaternary ammonium chlorides are ingredients in liquid detergent softener formulations. Difatty dimethyl chlorides find use in the rinse cycle, and difatty dimethyl quaternary sulfates are used as laundry softeners

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during drying. Lastly, fatty amines serve as the starting points for preparation of amphoteric surfactants such as betaines that have important applications in the cosmetics and personal care industries as ingredients in shampoos, conditioners, foaming, and wetting agents. Primary fatty amines can be reacted with ethylene oxide or propylene oxide to form bis(2-hydroxyethyl) or bis(2-hydroxypropyl) tertiary amines. Analogously, secondary amines yield 2-hydroxyfatty tertiary amines upon reaction with ethylene or propylene oxide (Billenstein and Blaschke, 1984; Maag, 1984; Reck, 1985). Addition is accomplished at 170 C, and additional ethylene or propylene oxide can be added by using a basic catalyst such as sodium or potassium hydroxide. Quaternary ethoxylated and propoxylated amines can be produced in a similar fashion. Ethoxylated and propoxylated fatty amines serve as cationic surfactants. As mentioned previously, fatty amines may be produced from fatty alcohols. Fatty alcohols are reacted with ammonia or another low molecular weight primary or secondary amine to give fatty amines. Generally, primary amines are prepared from the corresponding alcohols at 50340 C under pressure (3.5 MPa) using an excess of ammonia (5:1 to 30:1) in either batch or continuous mode in the presence of a cobalt-promoted zirconium catalyst (Card and Schmitt, 1987; Koike et al., 1974; Turcotte, 1983). Secondary amines are prepared by reaction of a fatty alcohol with a primary amine at 250 C and atmospheric pressure using a selective noble metal catalyst such as platinum, palladium, or ruthenium (Yokota et al., 1988). Tertiary amines are also accessible via condensation of a fatty alcohol with ammonia or a secondary amine. For example, dimethyldodecylamine is prepared via condensation of dodecanol with dimethylamine using a nickel catalyst at 180 C and 1.1 MPa (Wattimena and Borstlap, 1979). Fatty amides are synthetic intermediates along the way to fatty amines, but amides are of interest due to their lubricity and surface-active properties. This is because fatty amides exhibit inherently strong hydrogen bonding, insolubility, incompatibility, and unreactivity. Primary amides such as erucamide are used as slip and antiblock agents for polyolefins and in other polymers such as rigid polyvinyl chloride (PVC) as external lubricants. The most widely used synthetic route to primary fatty amides is the reaction of a fatty acid with ammonia at 200 C for 1012 hours under a constant vent of excess ammonia and water byproduct at a pressure of 0.350.69 MPa (Potts and McBride, 1950). Removal of water facilitates completion of the reaction. In addition, either the fatty amide or nitrile can be produced by adjusting reaction conditions in a continuous process (Potts, 1951). Fatty diamides can be prepared by reaction of fatty acids with diamines, such as ethylenediamine, in the presence of a catalyst (Fuchizawa and Motoyoshi, 1970). The diamides are used in all of the traditional primary amide applications, but have higher commercial value because of their superior efficiency. They are also used in powder coatings, defoamers, and flow modifiers in asphalt applications.

56

Fatty Acids

2.5.1.4 Deoxygenation Deoxygenation of fatty acids and esters yields hydrocarbons containing one less carbon than the parent compound and is accomplished either by decarbonylation or decarboxylation (Fig. 2.5, reaction d). As the names imply, one results in elimination of carbon monoxide, whereas the other produces carbon dioxide. Such transformations are stoichiometric, thermal, or catalytic. Examples of the stoichiometric approach include the Barton decarboxylation, Kolbe electrolysis, and Kochi and Hunsdiecker reactions, which proceed via free radical mechanisms and thus require radical initiators and suitable hydrogen donors for completion. In most cases, reductive decarboxylation is accompanied by additional chemical functionalization. For instance, end products of the Hunsdiecker and Kochi reactions are alkyl bromides and chlorides, respectively, whereas dimerized alkanes result from Kolbe electrolysis. Lastly, terminal alkenes are produced by oxidative decarboxylation of fatty acids using catalytic lead (IV) tetraacetate in the presence of copper (II) and a base (Carlblom et al., 1973). Thermal decarboxylation is thermodynamically favorable and thus yields hydrocarbons upon exposure of fatty acids to elevated temperatures (Immer et al., 2010). However, complex mixtures are produced, including cyclic and linear alkanes and alkenes, as well as gaseous species with carbon numbers up to C5. Thermal decarboxylation also promotes cracking of unsaturated constituents into smaller hydrocarbons and fatty acids (Maher et al., 2008). Catalytic decarboxylation is an alternative to the stoichiometric and thermal approaches. However, in some cases, oxidants are required to generate the active catalyst species, such as silver (II)-catalyzed oxidative decarboxylation of unsaturated fatty acids using sodium peroxydisulfate (Na2S2O8) with Cu21 added as needed (van der Klis et al., 2011). Thus, 8(Z)-heptadecene is obtained from oleic acid. In the presence of Cu21, (Z)-heptadeca-1,8-diene is formed from oleic acid with 8(Z)-heptadecene as the major byproduct. The heptadecenes obtained from unsaturated fatty acids can be subjected to ethenolysis to give terminal alkenes (van der Klis et al., 2012). Decarbonylation using catalytic palladium or rhodium yielding alkenes requires a stoichiometric excess of acetic anhydride to form the reactive intermediate (Kraus and Riley, 2012). Palladium-catalyzed decarbonylation of unsaturated fatty acids to alkanes requires elevated pressures (Sna˚re et al., 2008). Direct decarboxylation using a palladium catalyst does not require oxidants or reactive intermediates but temperatures in excess of 300 C are needed (Immer et al., 2010). Tandem isomerization-decarboxylation of oleic acid is accomplished at 250 C using ruthenium dodecarbonyl as precatalyst to yield a mixture of heptadecene isomers (Murray et al., 2014). Hydrocarbons resulting from deoxygenation of fatty acids, esters, and triglycerides have applications as renewable fuels and, in the case of

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57

unsaturated hydrocarbons, as monomers for oligomerization to yield industrial lubricants. Regarding fuel applications, a prominent example is the hydrotreatment process in which mixtures of hydrocarbons, mostly longchain alkanes, are produced from triglycerides or fatty acid alkyl esters in the presence of hydrogen, which is summarized elsewhere (Huber and Corma, 2007). The resulting mixture thus emulates the composition of conventional petrodiesel fuel. A typical catalytic system for this process is sulfided NiMo/γ-Al2O3 or CoMo/γ-Al2O3. The product mixture can be isomerized to give chain branching to improve cold flow properties, thus rendering it suitable as renewable jet fuel. Regarding lubricant applications, unsaturated hydrocarbons can be oligomerized to yield poly(olefins) as alternatives to poly alpha olefins obtained via oligomerization of petrochemically sourced 1-decene (van der Klis et al., 2011, 2012; Wagner et al., 2001).

2.5.2 Reactions at Unsaturated Sites Double bond(s) along with the hydrocarbon backbones of unsaturated of fatty acids undergo autoxidation, photo-oxygenation, hydrogenation, epoxidation, hydroxylation, oxidative scission, metathesis, dimerization, and hydroformylation, as shown in Fig. 2.7. Such transformations lead to products with useful properties, and are thus of commercial interest. Each will be discussed in greater detail in the following subsections. It should be noted that numerous other transformations are possible at or near double bonds, but those of the greatest commercial significance are covered here.

CHO

OOH

a h

b

g

c

f

O

d

e

OH

+ O

O OH

+

OH

HO

FIGURE 2.7 Reactions at the olefinic moiety of fatty acids: a, photo-oxygenation; b, hydrogenation; c, epoxidation; d, hydroxylation; e, oxidative scission; f, metathesis (ethenolysis depicted); g, dimerization; and h, hydroformylation.

58

Fatty Acids

The interested reader is directed to section C for additional sources of information regarding this and other topics relating to fats and oils chemistry.

2.5.2.1 Autoxidation and Photo-Oxygenation Autoxidation and photo-oxygenation involve addition of oxygen to unsaturated fatty acids, esters, or triglycerides to yield a variety of oxygenated products. Autoxidation and photo-oxygenation are different from many other chemical reactions in that they may proceed whether they are desired or not and are often inevitable consequences of storage. This is because atmospheric oxygen can react with fatty compounds with or without further impetus provided by users or researchers. Such reactions result in rancidity in fats and oils and are generally considered undesirable (Lundberg, 1954). Consequently, significant effort has been invested in exploring ways to prevent or inhibit their occurrence through, for example, the use of antioxidants (Dunn, 2008). The interested reader is directed to a comprehensive review by Yin et al. (2011) along with books by Frankel (2005) and Gunstone (2004) for further information on the subject of lipid oxidation. Oxygen exists in two forms: the common ground state triplet form, 3O2, and the excited state singlet form, 1O2, which is more reactive than the triplet form by 22.5 kcal/mol. Both forms are involved in oxidation of lipids, with the triplet form leading to autoxidation and the singlet form leading to photo-oxygenation. As the name implies, photo-oxidation involves the action of light, which provides the necessary energy to excite triplet oxygen to the singlet state. Despite some similarities, there are important differences between autoxidation and photo-oxidation. For instance, photo-oxidation is an ene reaction between electrophilic singlet oxygen and an electron-rich fatty double bond, whereas autoxidation is a radical chain reaction. Photooxidation requires no induction period in contrast to autoxidation and is unaffected by antioxidants but is inhibited by singlet oxygen quenchers such as carotene. Because the singlet form is more reactive, photo-oxygenation proceeds at a much faster rate than autoxidation. In both cases, the rate of oxidation increases dramatically with increasing unsaturation in the fatty substrate. In other words, linolenates react considerably faster than linoleates, which in turn are more reactive than oleates (Cosgrove et al., 1987; Holman and Elmer, 1947). The primary oxidation products of lipid oxidation are allylic hydroperoxides (Fig. 2.7, reaction a). However, these are unstable and decompose to a variety of secondary products. Decomposition of hydroperoxides includes fission to give volatile shorter chain aldehydes and acids, dimerization and oligomerization to provide higher molecular weight constituents, and rearrangement to yield products of similar molecular weight but with different functionality. Various other types and combinations of reactions can also

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occur during decomposition, such as dehydration, cyclization, and radical substitution. Volatile unsaturated aldehydes produced during the course of hydroperoxide decomposition are responsible for the off-flavor of foods containing oxidized oils. Oxidation of lipids is promoted by elevated temperature, the presence of light, and trace amounts of extraneous materials such as metals or other oxidation initiators (prooxidants). In particular, oxidative degradation is accelerated by copper, iron, and nickel (Knothe and Dunn, 2003). Postponement of oxidation is accomplished by antioxidants and proper storage conditions, which include storage in the dark at subambient temperatures in nonmetal containers and replacement of the head space in storage vessels with an inert atmosphere to remove oxygen. Antioxidants include radical scavengers, peroxide decomposers, and metal chelators and can be natural or synthetic in origin. Examples of radical scavengers include various hindered phenols and aromatic and hindered amines. Peroxide decomposers include divalent sulfur derivatives and trivalent phosphorous compounds such as dialkyl esters of thiodipropionic acid and tris(nonylphenyl)phosphate. Metal chelators include ethylenediamine tetraacetic acid, citric acid, phosphoric acid, and amino acids. Natural antioxidants include tocopherols, tocotrienols, carotenes, flavonoids, ascorbic acid, glutathione, melatonin, and numerous polyphenolics (resveratrol, for example), among others. Among the most common antioxidants for fats and oils are butylated hydroxytoluene, n-propyl gallate, α-tocopherol, butylated hydroxyanisole, and tert-butyl hydroquinone. Just as photo-oxidation involves light-facilitated ene addition of oxygen to lipids, the thiol-ene reaction is an analogous process whereby sulfur compounds (thiols) are added in an anti-Markovnikov fashion (to the less substituted carbon) to double bonds to yield fatty sulfides. An important distinction between the two aside from oxygen versus sulfur is that photooxidation is often unintentional and/or unwanted, whereas thiol-ene reactions are deliberate acts of organic synthesis. The thiol-ene reaction is a reversible free radical addition that is initiated by light, heat, or various radical initiators such as 2,2-dimethoxy-2-phenylacetophenone (DMPA) and is useful in the production of a variety of polymer networks (Tu¨ru¨nc¸ and Meier, 2013). An example is the formation of dimer fatty esters as monomers for copolyamide synthesis via thiol-ene addition of ethane-1,2-dithiol to methyl oleate using 2.0 wt% DMPA for 16 hours at 345 nm (Unverferth and Meier, 2016). If the double bond is terminal, then the use of light and radical initiators such as DMPA is not required. For instance, various thiols such as 1-thioglycerol, 1,4-butanedithiol, 2-mercaptoethanol, and methyl thioglycolate were added to methyl 10-undecenoate at 3570 C to provide the corresponding fatty sulfides (Tu¨ru¨nc¸ and Meier, 2010).

60

Fatty Acids

2.5.2.2 Hydrogenation Hydrogenation is the process of adding hydrogen across double bonds to yield saturated analogs (Fig. 2.7, reaction b). Because saturated triglycerides have higher melting points than their unsaturated counterparts, hydrogenation is often colloquially referred to as hardening. Historically, hydrogenation played a critical role in fats and oils chemistry, as it was utilized extensively to yield plastic (deformable) fats from liquid oils, thereby rendering the margarine and shortening industries less dependent on the limited availability of fats such as tallow. Incidentally, fats are solid at room temperature, whereas oils are liquid at room temperature. In addition, hydrogenation opened markets for whale and fish oils that were otherwise too unstable for food use due to their extensive unsaturation. As discussed previously, unsaturated fatty compounds are susceptible to autoxidation and photo-oxidation. Hydrogenation reduces double bond content, which improves oxidative stability while simultaneously increasing melting point. Hydrogenation is performed in such a manner that the carboxyl moiety is retained and saturated fatty acids or esters are formed. Hydrogenation is exothermic and proceeds in stages. Complete hydrogenation of linolenic acid proceeds through linoleic and oleic acid intermediates (or the corresponding trans isomers) before finally arriving at stearic acid (Albright, 1965). The rate of hydrogenation increases with double bond content, that is, linolenic acid is hydrogenated faster than linoleic, which in turn is faster than oleic acid (Koritala et al., 1973). Both positional (double bond migration and conjugation) and geometrical (cis/trans isomerism) isomerization accompanies hydrogenation (Dijkstra, 2006). Hydrogenation proceeds at elevated pressure and temperature in the presence of hydrogen and a transition metal catalyst and can be performed in batch, fixed bed, or continuous slurry reactors. The reactors are normally constructed using 316 L stainless steel because they are resistant to corrosion by fatty acids. Catalysts for hydrogenation include platinum, palladium, rhodium, iridium, ruthenium, and copper and can be homogeneous or heterogeneous, but those most widely applied to fatty compounds are nickel-based (Koritala et al., 1973). Of the nickel-based catalysts, which include Raney nickel, the most commonly utilized is the dry reduced type protected by fat, consisting of about 25% nickel, 25% inert alumina or silicate carrier, and 50% fully hydrogenated fat. Variables affecting hydrogenation are temperature, pressure, agitation, catalyst loading and addition, and feedstock quality (Brieger and Nestrick, 1974). Agitation is important because hydrogenation produces a three-phase mixture: a liquid fatty phase, a gaseous hydrogen phase, and a solid catalyst phase. Because of this, mass transfer and diffusion limitations may inhibit completion of the reaction if insufficient agitation is applied. Proper agitation also facilitates interaction of hydrogen with the catalyst and the substrate,

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keeps the catalyst in suspension, and aids in maintaining reaction temperature. The temperature of hydrogenation is normally in the range of 150210 C because at temperatures below and above these levels, the catalyst is not sufficiently activated or may undergo degradation. Pressures of industrial hydrogenations are in the range of 2.03.5 MPa, with higher pressures yielding shorter reaction times. However, pressures above 3.5 MPa have little to no influence on kinetics. Higher catalyst loading leads to faster rates of hydrogenation, but if too much catalyst is used then a rapid decrease in hydrogen concentration may lead to undesirable dehydrogenation reactions. A typical nickel catalyst load is 100150 ppm. The optimum time to introduce the catalyst is when the mixture is at or near the reaction temperature. Prolonged exposure of the catalyst to hot fatty acids may result in deactivation. The quality of the feedstock is important because not only does it influence the quality of the product but also the rate of hydrogenation. Impurities that affect rate include oxidized fatty acids, soaps, moisture, polyethylene, and constituents that contain chemically bound sulfur, phosphorous, and halides (Irandoust and Edvardsson, 1993; Klimmek, 1984). Impurities are normally removed beforehand by clay treatment or distillation (Zschau, 1984). Often in oleochemistry, full hydrogenation is not desired. Fortunately, adjustment of reaction parameters can attenuate the level of hydrogenation, thus leading to either full or partially hydrogenated products. Typical objectives of partial hydrogenation include improving oxidative stability by decreasing polyenoic compounds while simultaneously avoiding the formation of saturated, conjugated, and trans isomers. Saturated and trans isomers are undesirable because of their relatively high melting points leading to solids at room temperature. In addition, trans isomers have been implicated as deleteriously affecting cardiovascular health when contained in food products. Conjugated products are undesirable because they are oxidatively unstable. Therefore, the trick is to selectively hydrogenate polyenoic fatty compounds such as linolenate and linoleate to monoenoic analogs without further reduction while maintaining cis-geometry. To fulfill these demanding requirements, a highly selective catalyst is needed along with careful control of reaction conditions (List et al., 2000). For example, selectivity is improved by increasing temperature and worsened by increasing pressure and catalyst load. Double bond isomerization is promoted by higher temperatures but decreases with increasing pressure and catalyst load. In addition, trans isomers are favored by use of deactivated (reused) catalyst or sulfur-poisoned catalyst. Although nickel is the preferred catalyst for hydrogenation of fats and oils, the use of other metals such as palladium, platinum, ruthenium, and rhodium leads to products with less trans compounds. Selectivity can also be enhanced by modification of catalysts with copper or lead or by addition of amines to the reaction medium (Nohair et al., 2004). Not surprisingly, copper

62

Fatty Acids

and copper chromite catalysts also display enhanced selectivity relative to nickel (Kitayama et al., 1996). In addition, catalyst activity increases in the order Pd . Pt . Rh . Ni . Ru with the activity of palladium eight times higher than nickel (Cecchi et al., 1979). However, precious metal catalysts are not used for the mass production of hydrogenated vegetable oils and derivatives because of their high-cost relative to nickel.

2.5.2.3 Epoxidation and Hydroxylation Epoxidation is the process of adding an oxygen molecule across a double bond to form a three-membered oxirane ring (epoxide) consisting of two carbons and one oxygen with loss of the double bond (Fig. 2.7, reaction c). Fatty epoxides can be synthetic or natural in origin. Natural examples include vernolic and coronoric acids, which are found in plant oils from the Compositae, Euphorbiaceae, and Malvaceae families. The classic synthetic route to epoxides from unsaturated fatty compounds is by reaction with organic peroxyacids and is known as the Prilezhaev reaction. The organic peroxyacid is generated in situ by reaction of hydrogen peroxide with a carboxylic acid. Industrial epoxidation is most often performed using peroxyformic or peroxyacetic acids. Other peroxyacids, most notably m-chloroperoxybenzoic acid (mCPBA), are commonly utilized in research settings. The first successful application of the Prilezhaev reaction to lipids was by Findley et al. (1945) in which various oxiranes were obtained in 70%90% yields from in situ generated peroxyacetic acid. An example of lipid epoxidation using stoichiometric mCPBA is that of Aerts and Jacobs (2004), who prepared oxiranes from methyl oleate, methyl linoleate, high-oleic sunflower oil, and safflower oil. In addition to peroxyacids, epoxidation can be performed using molecular oxygen, dioxiranes, methyl oxorhenium, transition metal oxides, and aldehydes, among others. Other chemical methodologies include the Shi (oxone), Sharpless (titanium tetraisopropoxide), and Jacobsen (Mn(III)salen complex) epoxidations as well as the Halcon (hydroperoxides) reaction. Besides chemical methods, epoxidation can be performed enzymatically in the presence of hydrogen peroxide using lipases such as Candida antarctica Lipase B (CALB; Novozyme 435) (Ru¨sch gen. Klaas and Warwel, 1996, 1999). In such cases, the unsaturated fatty acid is converted by reaction with hydrogen peroxide to the corresponding peroxyacid, which then acts as the epoxidizing agent during which the fatty peroxide decomposes to give back the fatty acid. Enzymatic epoxidation is of interest because it minimizes formation of by-products such as vicinal diols resulting from unwanted hydrolysis of fatty epoxides. In addition, simultaneous epoxidation and esterification of unsaturated fatty acids such as oleic acid are possible using CALB and hydrogen peroxide in the presence of an alcohol (Orellana-Coca et al., 2007).

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OH OH

CN

HN

R

RCONH2

HCN

OH

Cl

HCl

O OH O

RCOOH O

OH

H2O

R O

OH ROH

RSH RNH2

OH

SR

OH

OH

OR

NHR FIGURE 2.8 Ring-opening reactions of fatty epoxides with a variety of nucleophiles. Note that stereochemistry is not indicated. Also note that nucleophilic addition can occur at either oxirane carbon, thereby leading to a mixture of positional and geometric isomers.

However, certain alcohols such as ethanol reduce lipase activity, leading to lower product yields. Epoxidized fatty esters and plant oils have important commercial applications. A prominent example is that of epoxidized soybean oil as a plasticizer and stabilizer for PVC. Epoxidized fatty materials also find applications in powder coatings, lubricating oils, inks, adhesives, composites, and paint diluents (Tehfe et al., 2010). Another important application is as intermediates in the synthesis of other materials. Because epoxides are potent electrophiles, they readily react with a variety of nucleophiles in ring-opening reactions to yield useful products. Examples of nucleophiles include water, alcohols, amines, thiols, carboxylic acids, amides, hydrogen cyanide, and hydrochloric acid, among others, which yield vicinal diols, alkoxy alcohols, amino alcohols, hydroxy thioether, hydroxy esters, N-hydroxyalkylamides, hydroxynitriles, and chlorohydrins, respectively, upon reaction with fatty epoxides (Fig. 2.8). Polyols resulting from ring-opening of fatty epoxides with water and alcohols serve as intermediates in the synthesis of polyurethane foams (Petrovi´c, 2008). Fatty amino alcohols have important corrosion inhibition and antiwear properties as well as antioxidant behavior. Fatty acrylates from reaction of epoxides with acrylic acid yield monomers with reactive terminal double bonds that are more readily polymerizable than internal olefins, thereby providing polymers with a wide range of physical properties (Khot

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et al., 2001). Chlorohydrins resulting from capture of free hydrochloric acid by epoxidized soybean oil facilitate stabilization of PVC and thus slow its degradation. Vicinal diols are accessed either through hydrolysis of fatty epoxides or by direct hydroxylation of unsaturated fatty compounds (Fig. 2.7, reaction d). Hydrolysis of epoxides yields trans-hydroxylated products, whereas direct hydroxylation of unsaturated fatty compounds proceeds via cishydroxylation. cis-Hydroxylation of cis-double bonds gives erythro diols (same side), whereas trans-double bonds give threo diols (opposite side). Examples would be methyl 9S,10S-dihydroxystearate and methyl 9R,10Rdihydroxystearate resulting from cis-hydroxylation of methyl oleate and methyl 9S,10R-dihydroxystearate and methyl 9R,10S-dihydroxystearate resulting from cis-hydroxylation of the trans isomer (methyl elaidate). Hydrolysis of methyl 9,10-epoxystearate provides a mixture of methyl 9S,10R-dihydroxystearate and methyl 9R,10S-dihydroxystearate. Direct cis-hydroxylation of unsaturated fatty compounds is accomplished most commonly with dilute alkaline potassium permanganate or osmium tetroxide. Direct trans-hydroxylation proceeds via the Prevost reaction using iodine and silver benzoate under anhydrous conditions. Vicinal diols are useful as polyols or as intermediates in chemical synthesis. For instance, they can be cleaved to aldehydes by periodic acid or to acids by potassium permanganate.

2.5.2.4 Oxidative Scission Unsaturated fatty acids and esters are cleaved at their double bond(s) to yield smaller, more oxygenated products via a process referred to as oxidative scission (Fig. 2.7, reaction e). Oxidative scission of unsaturated fatty acids yields bifunctional and monofunctional compounds that serve as intermediates for a variety of useful products. The bifunctional compounds (dicarboxylic acids) act as monomers for the preparation of polyamides (nylons) and polyesters. For example, oxidative scission of oleic acid affords azelaic (1,9nonanedioic) and pelargonic (nonanoic) acids (Pryde et al., 1960). Azelaic acid is a precursor to plasticizers, lubricants, hydraulic fluids, and polymers such as polyesters and nylon-6,9 as well as a component in hair and skin conditioners (Ko¨ckritz and Martin, 2011). Pelargonic acid is used in herbicide formulations and in the preparation of plasticizes, lubricants, and lacquers. Oxidative cleavage of petroselinic acid gives adipic (1,6-hexanedioic) and lauric acids (Mallard and Craig, 1966). Adipic acid is widely used in the polymer industry as a monomer for nylons but is produced petrochemically via nitric acid-mediated oxidation of cyclohexanone or cyclohexanol. Lauric acid has applications in the soaps and cosmetics sectors. Lastly, oxidative scission of erucic acid provides brassylic (1,11-undecanedioic) and pelargonic acids (Nieschlag et al., 1967).

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Oxidative cleavage is most commonly accomplished via ozonolysis or with potassium permanganate. Ozonolysis is the preferred industrial route, which entails reaction of an unsaturated fatty acid with ozone and is conducted at low temperature. The resulting ozonide intermediate is decomposed to alcohols or aldehydes via reductive work-up or to acids by oxidative work-up. Azaleic, adipic, and brassylic acids are obtained from ozonolysis of oleic, petroselinic, and erucic acids employing hydrogen peroxide during an oxidative work-up procedure. Ozonolysis of methyl oleate at 24 C followed by reductive work-up with hydrogen and Raney nickel yields 9hydroxynonanoic acid and 9-nonanol upon saponification of the methyl ester with NaOH (Liu et al., 2008). Amines are formed if ammonia is substituted for hydrogen. Sodium borohydride and catalytic hydrogenation employing platinum catalysts also afford alcohols, whereas triphenylphosphine, thiourea, zinc, or dimethyl sulfide produce aldehydes upon decomposition of the ozonide intermediate. Oxidative cleavage using permanganate must be mediated to avoid chain shortening of the intended products by further oxidation. Mediation is accomplished by employing a 1:39 mixture of potassium permanganate to sodium metaperiodate according to the von Rudloff procedure (von Rudloff, 1956; Youngs, 1961). Under these conditions, permanganate is never in a concentration high enough to cause over-oxidation and is continuously regenerated by oxidation with metaperiodate. Oxidative scission can also be accomplished with catalytic amounts of transition metals such as ruthenium, osmium, palladium, manganese, tungsten, molybdenum, and rhenium in combination with oxidants such as sodium hypochlorite, sodium periodate, peracetic acid, hydrogen peroxide, and oxygen (Behr et al., 2013).

2.5.2.5 Metathesis Olefin metathesis is an equilibrium reaction in which carboncarbon double bonds are cleaved and reformed with simultaneous exchange of constituents. Metathesis has been exploited by the petrochemical industry for decades to convert inexpensive propylene into more valuable ethylene and butylene and as a component of the Shell Higher Olefins Process (SHOP) to produce higher linear alpha olefins and their derivatives from oligomerized ethylene. Application of metathesis to fats and oils was limited until the relatively recent discovery of well-defined, highly active, long-lived ruthenium and molybdenum catalysts with broad functional group tolerance (Vougioukalakis and Grubbs, 2010). Metathesis can be divided into selfmetathesis between two identically substituted alkenes and cross-metathesis between alkenes with different substitution. For example, self-metathesis of methyl oleate gives dimethyl octadec-9-ene-1,18-dioate and octadec-9-ene as products and was first reported in 1972 (van Dam et al., 1972). Selfmetathesis of fatty acids is therefore an efficient route to unsaturated diacids,

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as exemplified by the self-metathesis and hydrolysis of methyl oleate to give octadec-9-ene-1,18-dioc acid (Ngo et al., 2006). A mixture of diacids of differing chain lengths is obtained by positional isomerization of double bonds prior to self-metathesis or concurrently with the addition of isomerization catalysts compatible with ruthenium metathesis catalysts (Ohlmann et al., 2012). Metathesis itself may lead to olefin positional isomerization, especially with older generation molybdenum and tungsten-based catalysts. Unwanted isomerization complicates product mixtures and reduces yield of the intended product (Lehman Jr et al., 2003). Fortunately, olefin isomerization during metathesis can be obviated with a radical quencher such as 1,4benzoquinone (Hong et al., 2005). An example of cross-metathesis is ethenolysis (Fig. 2.7, reaction f) whereby methyl oleate is reacted with ethylene to yield methyl 9-decenoate and 1-decene and was first reported in 1981 (Bosma et al., 1981). Another example of ethenolysis is metathetical cleavage of meadowfoam seed oil methyl esters with ethylene to provide methyl 5-hexenoate along with 1hexadecene. Subsequent self-metathesis of methyl 5-hexenoate affords dimethyl 5-decenedioate as a diester suitable for polymerization. Other alkenes such as 1- and 2-butenes, 1-pentene, 1-hexene, 1-heptene, 2- and 4octenes, 1-decene, and 1-octadecene have also been cross-metathesized with plant oils and fatty esters (Patel et al., 2006). Further transformation of these cross-metathesized products at the double bond position yields various alpha, omega-bifunctional compounds suitable for polymerization, including diacids, amino acids, and cyano acids. Additional functional groups can be introduced by cross-metathesis with functionalized alkenes such as methyl acrylate, allyl chloride, acrylonitrile, and fumaronitrile (Jacobs et al., 2009; Malacea et al., 2009; Rybak and Meier, 2007). The nitrile group is then converted to an acid or amine, thus providing starting materials for the synthesis of polyesters and polyamides. Similarly, aminoesters are prepared from fatty esters via simultaneous cross-metathesis with acrylonitrile and hydrogenation (Miao et al., 2012). Cross-metathesis of methyl oleate with 1-allyl-2,3-acetonido-glycerol followed by deprotection (to remove the acetonido protecting group) and hydroxylation of the double bond affords a fatty polyol (Zerkowski and Solaiman, 2012). Other examples of metathesis include ringopening metathesis (ROM), ring-closing metathesis, ROM polymerization, and ayclic diene metathesis polymerization. Metathesis of polyunsaturated fatty esters results in complicated product mixtures consisting of a variety of products, in contrast to metathesis of monounsaturated fatty esters. For instance, self-metathesis of methyl linoleate affords a complex mixture consisting of cis- and trans-alkenes (12:1, 15:2, 18:3, 21:4, 24:5), monoesters (16:1, 19:2, 22:3, 25:4, 28:5), diesters (20:1, 23:2, 26:3, 29:4), and 1-4-cyclohexadiene (Marvey et al., 2003). Similarly, metathesis of polyunsaturated plant oils such as olive, soybean,

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palm, and linseed, leads to complicated product mixtures with excellent drying properties (Marvey et al., 2003; Refvik et al., 1999).

2.5.2.6 Dimerization Dimerization of unsaturated fatty acids yields dicarboxylic acids that are useful building blocks for the production of polyesters, polyamides, adhesives, surface coatings, lubricants, and printing inks. The most common application of dimer acids is as polyamides. The acids react with diamines to give nonreactive polyamides that are used as hot-melt adhesives for shoes and in coatings and printing inks. Reaction with triamines yields polyamides with free amino groups that can react further to act as curing agents for thermosetting epoxy resins (Vedanayagam and Kale, 1992; Vijayalakshmi et al., 1992). The resulting dicarboxylic acids can also be hydrogenated to give diols as intermediates for polyols and polyurethanes. These compounds are low vapor pressure, noncrystallizable, nonflammable liquids at room temperature that offer elasticity, flexibility, high-impact strength, hydrophobic stability, hydrophobicity, hydrolytic stability, and low glass transition temperatures. Commercial production of dimer fatty acids is accomplished in 68 hours at elevated temperature (230 C) in the presence of a cationic montmorillonite clay catalyst (Leonard, 1979). Dimerization can also occur at high temperatures (260400 C) without a catalyst. When 18 carbon unsaturated fatty acids are dimerized, 36 carbon dimers are produced containing two carboxylic acid functional groups. Trimers, higher oligomers, and isostearic acid are also produced in smaller amounts, and these can be removed or isolated via molecular distillation. Isostearic acid, arising from branching reactions along the hydrocarbon backbone of unsaturated fatty acids, typically contains one methyl branch and no unsaturation. Isostearic acid is of commercial importance due to its exceptional oxidative, thermal, and odor stability and finds applications in lubricant, pigment, cosmetic, personal care, and surfactant formulations. Reduction of isostearic acid yields isostearyl alcohol, which is used in cosmetics, deodorants, and personal care products where it provides film-forming and spreading functions. Dimerization proceeds via a variety of mechanistic pathways, thus resulting in complex product mixtures. For example, dimerization of MUFAs occurs via an ene reaction or by carbocation intermediates to yield an acyclic, unsaturated, branched structure (Fig. 2.9, reaction a). Dimerization of a MUFA with a PUFA or dimerization of two PUFAs proceeds via DielsAlder cycloaddition to yield alkyl-substituted cyclohexene rings (Fig. 2.9, reaction b). To further complicate matters, MUFAs may undergo desaturation to polyunsaturated analogs (Fig. 2.9, reaction c) to yield a mixture of Diels-Alder (cyclic) and ene-reaction (acyclic) products upon dimerization.

68

Fatty Acids

a

g c

d

b

e

+ f

b d

f

+

FIGURE 2.9 Products formed during the course of dimerization of unsaturated fatty acids: a, acyclic analogs via ene reactions; b, substituted cyclohexenes via Diels-Alder reactions; c, conjugated dienes via desaturation of MUFAs; d, substituted, unsaturated decalins via Diels-Alder reactions; e, cyclohexane and benzene derivatives via hydrogen transfer; f, saturated decalin and naphthalene derivatives via hydrogen transfer; and g, isostearic acids via branching.

In addition, dimerization of PUFAs also produces bicyclic dimers consisting of an alkyl-substituted decalin core with one double bond per ring (Fig. 2.9, reaction d). Hydrogen transfer can then convert the cylcohexene rings into cyclohexane and benzene analogs (Fig. 2.9, reaction e). Correspondingly, hydrogen transfer of the unsaturated decalin rings yields naphthalene and saturated decalin structures (Fig. 2.9, reaction f). Double bond migration, cis/ trans isomerism, conjugation, and branching (Fig. 2.9, reaction g) to yield isostearic acids can also occur during dimerization. In summary, dimerization of monoenes yields mostly acyclic (40%) and monocyclic (55%) dimers, whereas polyenes provide monocyclic (55%) and bicyclic (40%) dimers (Leonard, 1979). In commercial practice, tall oil fatty acids are most commonly used as starting materials for industrial-scale production of dimer fatty acids and isostearic acid. Because tall oil fatty acids consist primarily of oleic (30%45%) and linoleic acids (35%45%), dimerization produces a complex mixture consisting of acyclic (15%), monocyclic (55%), and bicyclic (40%) products (Leonard, 1979).

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2.5.2.7 Hydroformylation Hydroformylation is the process of adding a formyl group (CHO) and a proton across a double bond to produce an aldehyde with concurrent loss of the double bond (Fig. 2.7, reaction h). Also referred to as the oxo process, hydroformylation is an important industrial reaction and represents the most significant method for introducing a carbonyl group to a double bond. The process entails treatment of an alkene with a high pressure of carbon monoxide and hydrogen (syn gas) at elevated temperature in the presence of a transition metal catalyst. The most common catalysts for industrial hydroformylation are cobalt carbonyls and rhodium complexes of triphenylphosphine, but other transition metal compounds have been used and are mostly of academic interest. The order of catalyst activity for hydroformylation is generally accepted as: Rh .. Co . Ir, Ru . Os . Pt . Pd . Fe . Ni (Beller et al., 1995). Hydroformylation was performed initially with a dicobalt octacarbonyl catalyst at temperatures of around 150 C and syn gas pressures of 2530 MPa. A significant discovery was that rhodium chloride with trivalent ligands such as triphenylphosphine has catalytic activities, several orders of magnitude higher than those of cobalt, thus allowing the reaction to proceed at considerably lower temperatures (100 C) and pressures (12.5 MPa) (Paulik, 1972). A drawback to rhodium catalysts is that they are more expensive than cobalt catalysts, so their recovery and reuse is important from a process economics perspective. Linear terminal alkenes (alpha olefins) are the most reactive toward hydroformylation, with linear internal alkenes and especially more substituted (branched) internal alkenes exhibiting lower reactivity (Wender et al., 1956). With terminal alkenes, the aldehyde unit adds to both the primary and secondary carbons of the alkene, but proper choice of catalyst and bidentate ligand leads to selectivity for the secondary (Chan et al., 1995) or primary (Breit and Seiche, 2003) product. The linear isomer is generally more valuable than the branched isomer. Primary alcohols (so-called “oxo” alcohols) arising from hydroformylation of terminal alkenes represent the highest volume application of industrial hydroformylation. For example, the important commodity chemical 1-butanol is obtained by selective hydroformylation of propylene at the terminal carbon of the double bond to n-butryaldehyde using a phosphorous-stabilized rhodium catalyst, which is then hydrogenated to give the primary alcohol. However, aldehydes can also be subsequently oxidized to acids, reduced to amines by reductive amination or to unsaturated long-chain branched aldehydes by aldol condensation. Moreover, a reduction-elimination sequence (aldehyde-alcohol-alkene) gives access to isomerically pure alkenes elongated by one additional carbon (Franke et al., 2012). Hydroformylation is also a component of important industrial tandem catalysis reactions, such as hydroformylationhydrogenation,

70

Fatty Acids

hydroformylationisomerization, and isomerizationhydroformylation hydrogenation (Fogg and dos Santos, 2004). For example, terminal alcohols are produced from internal alkenes by tandem isomerization hydroformylationhydrogenation via the Shell Oxo Process using a chemoselective cobalt trialkylphosphine catalyst (Slaugh and Mullineaux, 1968). Selectivity in such a reaction is critical to avoid appreciable hydrogenation of the starting alkene and hydroformylation of internal alkenes. Another important example is the production of nonrenewable fatty alcohols via hydroformylationhydrogenation of C6C17 terminal alkenes arising from ethylene oligomerization employing cobalt complexes modified with phosphine ligands. Although rhodium catalysts are more active toward hydroformylation and offer milder reaction conditions, cobalt catalysts are often recruited for tandem reactions because they also catalyze olefin isomerization and hydrogenation of aldehydes to alcohols under the right conditions. The presence of bulky ligands on the cobalt catalyst restricts its access to internal carbons, thereby inhibiting secondary aldehyde formation (Reuben and Wittcoff, 1988). Hydroformylation represents an important route to primary alcohols from unsaturated fatty compounds. Primary alcohols are of interest from a practical standpoint because they are more reactive than their secondary counterparts. Accordingly, polyurethanes are more readily produced from hydroformylated fatty compounds (Fig. 2.7, reaction h) than from secondary alcohols such as those produced via hydroxylation (Fig. 2.7, reaction d) or hydrolysis of epoxides (Fig. 2.8) (Guo et al., 2006). Polyols resulting from hydroformylation of fatty materials produce softer polyurethanes than polyols by the epoxidation route because of higher molecular weights at the same functionality and an extra methylene group per double bond (Guo et al., 2006; Petrovi´c et al., 2008). Another factor influencing properties of resulting polyurethanes is catalyst choice during hydroformylation. At high conversion rates with a rhodium catalyst, a rigid polyurethane is formed, whereas under the conditions of cobalt catalysis and low conversion, a hard rubber with lower mechanical strength is produced (Guo et al., 2002). This is because cobalt catalysts afford monoaldehydes upon hydroformylation of dienes, whereas rhodium provides polyaldehydes owing to its greater catalytic activity (Frankel et al., 1973). In addition to polyurethane applications, acetalization of the resulting alcohols with glycerol or methanol forms materials with plasticizing properties for PVC (Neff et al., 1976). In the past, methyl oleate has frequently been studied as a model substrate for hydroformylation. Methyl linoleate and methyl linolenate have also been studied, but to a lesser extent. With methyl oleate, hydroformylation produces two isomeric formyl stearate esters, but other regioisomers with the formyl group between C5 and C13 are also formed as side products as a result of double bond migration (Frankel et al., 1969). Suppression of double bond migration is achieved by switching from dicobalt octacarbonyl to

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triphenylphosphine-modified rhodium during hydroformylation (Frankel, 1971). Accessing terminal aldehydes from methyl oleate is accomplished by attaching bulky diphosphate chelating ligands to the rhodium catalyst, which inhibits hydroformylation at internal carbon positions due to steric hindrance (Behr et al., 2005). With linoleates, migration of double bonds into conjugation produces mainly monoformyl esters, whereas unconjugated methyl linoleate affords diformyl esters upon hydroformylation using a rhodiumtriphenylphosphine catalyst (Frankel et al., 1973). In summary, hydroformylation of unsaturated fatty substrates represents a facile route to primary alcohols as intermediates for a number of useful materials.

2.6 CONCLUSION This chapter provides the reader with an introduction to the major structural types of naturally occurring fatty acids, and to their sources, preparation, purification, and chemistry. It is clear that the diverse array of fatty acids exhibits unusual properties and possesses interesting functionality, chemistry, biodegradability, biocompatibility, and sustainability. Fatty acids play an important role in food, nutrition, and nonfood applications and better understanding and manipulation of fatty acids will enable their continued utilization. The interested reader is directed to further sources of information regarding the purification and chemical derivatization of fatty acids, esters, and triglycerides. Comprehensive reviews on these topics include those of Baumann et al. (1988), Behr et al. (2008), Behr and Gomes (2010), Biermann et al. (2000, 2011), Corma et al. (2007), and Gunstone (2001). Books on the subject include Fats and Oils Formulating and Processing for Applications (CRC Press), Green Vegetable Oil Processing (AOCS Press), Industrial Uses of Vegetable Oils (AOCS Press), Lipid Oxidation (CRC Press), Lipid Synthesis and Manufacture (CRC Press), Oleochemical Manufacture and Applications (CRC Press), The Biodiesel Handbook (AOCS Press), The Chemistry of Oils and Fats (CRC Press), and The Lipid Handbook (CRC Press). A further comprehensive reference for information regarding all aspects of chemical technology is the multivolume KirkOthmer Encyclopedia of Chemical Technology (John Wiley & Sons).

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Chapter 3

Epoxy Fatty Acids: Chemistry and Biological Effects Arnis Kuksis and Waldemar Pruzanski University of Toronto, Toronto, ON, Canada

Chapter Outline 3.1 Introduction 83 3.2 Natural Occurrence and Structure of Epoxy Fatty Acids 84 3.2.1 Oleic and Linoleic Acid Monoepoxides and Hydroxides 84 3.2.2 Arachidonic Acid Monoepoxides 85 3.2.3 Eicosapentaenoic Acid and Docosahexaenoic Acid Monoepoxides 85 3.3 Chemical Synthesis 88 3.3.1 Direct Epoxidation 88 3.3.2 Chemo-Enzymatic Perhydrolysis 89 3.3.3 Other Chemo-Enzymatic Epoxidations 90 3.4 Biosynthesis of Epoxy Fatty Acids 90 3.4.1 Oxygenases and Lipoxygenases 91 3.4.2 Peroxygenases 91

3.4.3 Cytochrome P450-Like Oxygenases 3.5 Analysis of Epoxy Fatty Acids 3.5.1 Resolution of Regioisomers 3.5.2 Resolution of Enantiomers 3.5.3 GC/MS and LC/MS Identification of Lipid Epoxides 3.6 Biological Effects 3.6.1 Lipid Signaling 3.6.2 Cellular Effects 3.6.3 Systemic Effects 3.7 Pathological Effects 3.7.1 Toxicity 3.7.2 Inflammation and Pain 3.7.3 Angiogenesis and Cardiovascular Disease 3.7.4 Cancer 3.8 Conclusion Abbreviations References

92 94 95 97

103 104 104 105 107 108 108 108 110 111 112 112 113

3.1 INTRODUCTION There is a significant worldwide demand for replacing petroleum-derived raw materials with renewable ones (Carlsson et al., 2011; Aouf et al., 2014). Aside from polysaccharides and sugars, plant oils and animal fats are the most important renewable raw materials of the chemical industry (Metzger Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00003-9 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

83

84

Fatty Acids

and Bornscheuer, 2006; Meier et al., 2007). The latter give access to various fatty acids, which differ in carbon chain lengths, number of carboncarbon double bonds, and the presence of functional groups. Many oleochemicals are obtainable from fatty acids, including epoxidized fatty acids, which are of great interest to industry. Among the major applications of epoxidized vegetable oils and fatty acids are their use as plasticizers for polyvinyl chloride (PVC), PVC stabilizers, and diluents for paints and lubricants, and intermediate for polyurethane polyol production (Feng et al., 2014). The large-scale production of fatty acid epoxides and their incorporation into various industrial and household products should raise concern about their biological safety. There is substantial evidence that the epoxides of the long-chain polyunsaturated fatty acids (PUFAs) at least function as signaling molecules to regulate inflammation, pain, angiogenesis, and cancer (Morisseau and Hammock, 2013). This has led to extensive analytical and metabolic investigation of the formation and function of various epoxy fatty acids and their derivatives, which are discussed in the following sections.

3.2 NATURAL OCCURRENCE AND STRUCTURE OF EPOXY FATTY ACIDS Although fatty acid epoxides are known to occur naturally and can be isolated from various tissues and body fluids, only some plant seeds contain them in significant amounts.

3.2.1 Oleic and Linoleic Acid Monoepoxides and Hydroxides Plants produce a variety of epoxide containing lipids in biochemical pathways associated with host defense responses (Blee, 2002) and cutin polymer synthesis (Lequeu et al., 2003). Only a few plants produce epoxy fatty acids in significant quantities. The vernolic acid (12S,13R-epoxy-9-cis-octadecenoic acid) can be found in the seed oils from several Asteraceae genera, including Stokesia, Verninia, and Crepis (Badami and Patil, 1981; Cahoon et al., 2002) and in certain Euphorbiaceae species such as Euphorbia lagascae and Buddleja pulchella (Spitzer et al., 1996). The latter acids make up 50%90% of total fatty acids. Fig. 3.1 shows the chemical structures of fatty acid epoxides and derivatives from the C18 family (Le Quere et al., 2004). Metabolism of epoxides by human cytochrome P450 (CYP) results in the formation of several hydroxylated metabolites (ω-OH, vicinal diol, and triol). EpOME, epoxyoctadecenoic acid methyl ester, EpSTA, epoxyoctadecanoic (stearic) acid; HEpOME, hydroxyepoxyoctadecenoic acid methyl ester; HEpSTA, hydroxyepoxyoctadecanoic acid. Newman et al. (2002) have isolated the epoxyoctadecenoic acids 12 (13)- and 9(10)-EpOMe from rodent and human urine along with the corresponding dihydroxy derivatives in 314 pmol mg21 creatinine.

Epoxy Fatty Acids: Chemistry and Biological Effects Chapter | 3 Z9(10)-EpSTA

Z9(10)-EpOME

85

Z12(13)-EpOME

O COOH

OH

O

OH

OH

OH OH

OH

18.9(10)-HEpSTA 17.9(10)-HEpSTA

18.9(10)-HEpOME 17,9(10)-HEpOME

Diols HO

COOH

COOH

Diols

OH COOH

OH(Triols)

HO

18.9(10)-HEpOME

17,9(10)-HEpOME Diols

OH COOH

COOH HO

OH

OH(Triols)

OH(Triols)

FIGURE 3.1 Chemical structures of fatty epoxides and derivatives from the C18 family. Metabolism of epoxides results in the formation of several similar hydroxylated metabolites (ω-OH, vicinal diol, and triol). EpOME, epoxyoctadecenoic acid methyl ester; EpSTA, epoxyoctadecanoic acid; HEpOME, hydroxyepoxyoctadecenoic acid methyl ester; HEpSTA, hydroxyperoxyoctadecanoic acid. Reproduced with permission from Le Quere et al. (2004). Human CYP4F3s are the main catalysts in the oxidation of fatty acid epoxides. J. Lipid Res. 45, 14461458.

3.2.2 Arachidonic Acid Monoepoxides In contrast to the epoxides of oleic and linoleic acid, the natural occurrence of the epoxides of eicosatetraenoic (arachidonic) (ETE) of arachidonic acid (ARA) has been widely reported and their physiological/pathological properties have been extensively discussed (see Lagarde and Nicolaou, 2015). Olefin epoxidation via the epoxigenase reaction results in the production of four cis-epoxyeicosatrienoic acids (EETs) (14,15-, 11,12-, 8,9-, and 5,6EETs), each of which can be formed as either the R,S or S,R enantiomer. Fig. 3.2 shows the structures of regioisomers of the cis-epoxyeicosanoids and their CYP and soluble epoxy hydrolase (sEH) metabolites as originally drawn by Newman et al. (2005) and Zhang et al. (2014). Morisseau et al. (2010) have reported the distribution and quantitative levels of octadecenoic and ETE epoxide regioisomers in the central and peripheral nervous system by a number of CYP isozymes and presented their findings in tabular form (tables not shown).

3.2.3 Eicosapentaenoic Acid and Docosahexaenoic Acid Monoepoxides Arnold et al. (2010) have shown that EPA and DHA are efficient alternative substrates of ARA metabolizing enzymes in vitro. Rats given EPA/DHA supplement changed the endogenous CYP metabolite profiles: e.g. altering EET: epoxyeicosatetraenoic acid (EEQ):EDP ratio from 87:0:13 to 27:18:55 in the heart. Fig. 3.3 compares the structures of CYP and sEH metabolism of EPA and DHA as originally drawn by Arnold et al. (2010) and Zhang et al. (2014). The whole set of regioisomeric epoxides involves 5,6-, 8,9-, 11,12-,

86

Fatty Acids COOH

Arachidonic acid (ARA) Cytochrome P450 (CYP) epoxygenases O

O COOH

COOH

COOH

COOH

O

5,6-EET

8,9-EET

O

11,12-EET

14,15-EET

Soluble epoxide hydrolase (sEH) HO

OH

HO

OH

COOH

COOH

COOH HO

5,6-DiHET

8,9-DiHET

OH

11,12-DiHET

COOH HO

OH

14,15-DiHET

FIGURE 3.2 Chemical structures of metabolites of arachidonic acid (ARA) by CYP epoxygenases and sEH. 5,6-, 8,9-, 11,12-, and 14,16-EET are four regioisomers of EET; 5,6-, 8,9-, 11,12-, and 14,16-DiHET are corresponding regioisomers of DiHET, formed by sEH. Reproduced with permission from Zhang et al. (2014). Stabilized epoxygenated fatty acids regulate inflammation, pain, angiogenesis and cancer. Progr. Lipid Res. 53, 108123.

14,15-, and 17,18-EEQ from EPA, and 4,5-, 7,8-, 10,11-, 13,14-, 16,17-, and 19,20-epoxydocosapentaenoic acid (EDP) from DHA, each of which can be formed as either the R,S or S,R enantiomer (enantiomers not shown). Furthermore, each individual CYP-isoform displayed a unique regioselectivity that was dependent on the PUFA substrate. Morisseau et al. (2010) have determined the distribution of octadecenoic, ETE, and DPE epoxides of n-3 epoxy fatty acids in the central and peripheral nervous system using a number of CYP isozymes. The central nervous system (CNS) contained significantly more epoxyeicosapentaenoic acid (EpDPE) than epoxyeicosatetraenoic acid (EpETE), consistent with the observation that there is more DHA than EPA in the CNS. Except for the 7,8-EpDPE, which was present in relatively high quantity compared with the other epoxy fatty acids, the EpETEs in the CNS represented similar quantities of the EET regioisomers. Epoxidized fatty acids are currently produced by chemical oxidation of unsaturated plant oils. Despite numerous chemical methods of epoxidation of double bonds of unsaturated fatty acids, only the Prileshajev method of epoxidation has been used on an industrial scale (Aouf et al., 2014). In this method, short-chain peroxy acids are generated from the corresponding acid and hydrogen peroxide in the presence of a strong mineral acid (Swern, 1961; Aouf et al., 2014). These peroxy acids react with the CC double bonds to generate epoxidized fatty acids from the corresponding acid. Due to a potential danger of handling peroxy acids, an in situ method is generally

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87

COOH COOH

Eicosapentaenoic acid (EPA)

Docosahexaenoic acid (DHA) CYP epoxygenases

O

O

COOH

5,6-EEQ

COOH

4,5-EDP

COOH

7,8-EDP

COOH

10,11-EDP

COOH

13,14-EDP

COOH

16,17-EDP

O

O COOH

8,9-EEQ O COOH

11,12-EEQ

O COOH

14,15-EEQ

O

O COOH

17,18-EEQ

O COOH

O

19,20-EDP

O

sEH HO

OH

HO

OH

COOH

5,6-DiHETE HO

HO

OH COOH

8,9-DiHETE

HO

11,12-DiHETE OH

14,15-DiHETE

HO

7,8-DiHDPA

COOH

10,11-DiHDPA

COOH

COOH

17,18-DiHETE

13,14-DiHDPA

OH

OH

HO

COOH

COOH

COOH HO

4,5-DiHDPA

OH

COOH HO

COOH OH

HO

16,17-DiHDPA

OH COOH

OH HO

19,20-DiHDPA

OH

FIGURE 3.3 Chemical structures of metabolites of EPA and DHA by CYP epoxygenases and sEH. 5,6-, 8,9-, 11,12-, 14,14-, and 17,18-EEQ are regioisomers of EpETEs (EEQ), while 4,5-, 7,8-, 10,11-, 13,14-, 16,17-, and 19,20-EDP are corresponding regioisomers of EDPs. The 5,6-, 8,9-, 11,12-, and 17,18-DiHETE are corresponding regioisomers of DiHETE, while 4,5-, 7,8-, 10,11-, 13,14-, 16,17-, and 19,20-DiHDPA are corresponding regioisomers of DiHDPE. Reproduced with permission from Zhang et al. (2014). Stabilized epoxygenated fatty acids regulate inflammation, pain, angiogenesis and cancer. Progr. Lipid Res. 53, 108123.

preferred for large-scale epoxidation of unsaturated fatty acids, rather than a separate preparation of the peroxy acids (Rusch gen. Klaas and Warwel, 1999). To eliminate several drawbacks in these methods, enzymes such as peroxygenase and lipase were introduced in the process (Rusch gen. Klaas

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Fatty Acids

and Warwel, 1999). Lipases were shown to produce peroxy acids from hydrogen peroxide and fatty acids by a perhydrolysis reaction. Saithai et al. (2013) have presented a schematic diagram of the chemical and chemoenzymatic epoxidation process of soybean oil (Fig. 3.4).

3.3 CHEMICAL SYNTHESIS 3.3.1 Direct Epoxidation Despite the advantages of chemo-enzymatic methods of fatty acid and acylglycerol epoxidation, small-scale preparations of epoxy fatty acids have been successfully prepared by direct chemical oxidation. In an early systematic study with pure monounsaturated compounds (Swern et al., 1944), oleic acid, methyl oleate, and oleoyl alcohol were converted to the corresponding cis-9,10-epoxy derivatives, respectively, by epoxidation with perbenzoic acid. The epoxidation reaction is one of the most stereospecific reactions known and continues to be used for small-scale preparations. More recently, Van Rollins et al. (1989) and Van Rollins (1995) described the preparation of epoxides of eicosapentaenoic (EPA) and docosahexaenoic (DHA) fatty acids by reacting the methyl ester of EPA and DHA, respectively, with meta-chloroperbenzoic acid (mCPBA). The method was recently adopted by Morisseau et al. (2010) for the preparation of EpETE and EpDPE regioisomers to be used as standards for the isolation and identification of naturally occurring epoxides of C18:1 to C22:6 fatty acids in both the n-6 and n-3 series. In brief, each PUFA was treated with mCPBA, which converts cis-double bonds to ( 6 )cis-epoxides, and the regioisomer products H2C O C O HC O C O H2C O C O

O

H2O2 1. Chemical epoxidation (Prileshajev-epoxidation) H2O

HOOC

H2SO4

R

HOOC

R

2. Chemo-enzymatic epoxidation

lipase O

H2O

O

HOOC

H2O2

HOOC

R

R

O O H2 C O C O HC O C O H2C O C

O

O

H2C O C O

O

O

HC O C

O

H2C OH O

O

O

O

O

+ O

HOOC

FIGURE 3.4 Schematic diagram of the chemical and chemo-enzymatic epoxidation of soybean oil triacylglycerols. Reproduced with permission from Saithai et al. (2013). Effects of different epoxidation methods of soybean oil on the characteristics of acrylated epoxidized soybean oilco-poly (methyl methacrylate) copolymer. Express Polymer Lett. 7 (11), 910924.

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89

were isolated by normal-phase HPLC. The methyl ester of each epoxide was converted to the free acid form and freshly isolated by normal-phase HPLC. Individual regioisomers were separated by normal-phase HPLC while absorption at 192 nm was monitored. The semipreparative column [1.0 (i. d.) 3 25 cm] that was used contained 5 μm silica particles (Ultramex, Phenomenex, Torrence, CA, United States) with hexane/2-propanol, glacial acetic acid (4000:13:2, v/v/v) flowing at 7.0 mL min21 and 600 psig. The individual regioisomers were purified and used for identification and quantification of naturally occurring monoepoxides in the brain and spinal cord of rats. Likewise, Le Quere et al. (2004) synthesized racemic samples of [1-14C] Z9(10)-epoxystearic acid and [1-14C]leukotoxins [Z9(10)-EpOME and Z12 (13)-EpOME] from [1-14C]oleic acid and [1-14C]linoleic acid, respectively, using 3-chloroperoxybenzoic acid. The epoxidation of each substrate was performed in a reaction mixture (0.2 mL) containing either 0.35 mM epoxystearic acid or 0.33 mM leukotoxins together with 1.8 mM mCPBA. The reaction was initiated for 5 min at ambient temperature by the addition of mCPBA and terminated by evaporation under N2.

3.3.2 Chemo-Enzymatic Perhydrolysis Among the different lipases used in industry, the most widely employed is the one from Candida antarctica, a yeast, which actually produces two types of lipases, A and B, with B being preferred (Bjorkling et al., 1992). In such perhydrolysis reactions, hydrogen peroxide acts as the nucleophile instead of water in the deacylation step of the serine hydrolase. However, the perhydrolase activity of lipases and esterases is generally much lower than their esterase activity and some of them do not exhibit perhydrolase activity at all (Berhardt et al., 2005). When unsaturated fatty acids or their alkyl esters were treated with hydrogen peroxide in the presence of C. antarctica B lipase, their epoxidized derivatives were produced in a two-step reaction (Warvel and Rusch gen. Klaas, 1995; Aouf et al., 2014). First, the unsaturated fatty acids were converted into the corresponding unsaturated peroxy acid owing to the perhydrolysis activity of the lipase, and then the resulting unsaturated peroxy or carboxylic acids were epoxidized via an uncatalyzed Prileshajev reaction that is often referred to as “self-epoxidation reaction” in spite of the fact that it proceeds predominantly via an intermolecular process (Fig. 3.5). The chemo-enzymatic reaction can also be applied to oils and fats for the production of epoxidized plant oils. The resulting mixture contains epoxidized triacylglycerols, a small amount of epoxidized free fatty acids and some epoxidized mono- and diacylglycerols (Rusch gen. Klaas and Warwel, 1999). With addition of free fatty acids at the start, various plant oils were epoxidized with conversions and selectivities above 90%.

90

Fatty Acids O Lipase R

R

(CH2)n - COOH H2O2

(CH2)n - COOOH

R

(CH2)n - COOH

H2O2

FIGURE 3.5 Chemo-enzymatic epoxidation of unsaturated fatty acids involving a lipasecatalyzed perhydrolysis step. Reproduced with permission from Aouf et al. (2014). The use of lipases as biocatalysts for the epoxidation of fatty acids and phenolic compounds. Green Chem. 16, 17401754.

3.3.3 Other Chemo-Enzymatic Epoxidations Orellana-Coca et al. (2005a) performed the chemo-enzymatic (lipase B from C. antarctica) epoxidation of linoleic acid in toluene and observed a quantitative conversion of double bonds to give the corresponding diepoxide when operational temperature was set between 40 C and 50 C. Other organic solvents, such as butanol of dichloromethane, were also suitable, and the reaction could be carried out with plant oils. Orellana-Coca et al. (2005b) performed chemo-enzymatic epoxidation of oleic acid and methyl oleate in solvent-free medium. Epoxystearic acid and epoxystearic methyl ester were synthesized with very good yields. More recently, the use of hydrophobic and hydrophilic ionic liquids was proposed in order to improve the reaction yields in lipase-catalyzed epoxidation of methyl oleate (Silva et al., 2011). The hydrophilic ionic liquids were advantageous, resulting in the best yields and reaction kinetics. Of the nine different lipases tested, Aspergillus niger lipase in hydrophilic BMI.BF4 (1-n-butyl-3-methylimidazolium tetrafluoroborate) yielded the epoxidized compound in 89% in the first reaction hour, whereas hydrophobic BMI.PF6 (1-n-butyl-3-methylimidazolium hexafluorophosphate) yielded the same product in 67% yield. The earlier methods are not well suited for the isolation and identification of individual epoxy fatty acids and for studies of their specific physico-chemical and biological properties. For the latter purpose, enzymatic epoxidation with specific CYP and lipoxygenases (LOXs) of positional specificity have been employed.

3.4 BIOSYNTHESIS OF EPOXY FATTY ACIDS The biosynthesis of epoxy fatty acids takes place by oxygenation of preformed unsaturated fatty acids, their methyl esters or glycerophospholipids as substrates. There are three known oxidation pathways in plants: a desaturase-like oxygenase giving vernolic acid, a CYP-linked oxygenase using the same substrates, and a peroxygenase pathway. In mammals, the CYP epoxidation of fatty acids is well established.

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3.4.1 Oxygenases and Lipoxygenases The desaturase-like oxygenase was identified by Bafor et al. (1993) to synthesize vernolate (cis-12-epoxyoctadea-cis-9-enoate) in microsomal preparations from developing endosperm of E. lagascae. A P450 monooxygenase enzyme utilized linoleoyl-GroPCho and other phospholipids as the substrate producing vernoleoylGroPCho. The vernolic acid accumulated only in triacylglycerols, not in PtdCho. Lee et al. (1998) demonstrated that vernoleate biosynthesis in Crepis palaestina involves a 12 desaturase-like oxygenase epoxidizing the 12-double bond of linoleic acid linked to PtdCho, giving rise to 12,13-epoxy-18:1-9c (vernolic acid). Assays with Vernonia extracts (Lee et al., 1998) indicated some fundamental differences between the two enzymes. Thus, carbon monoxide apparently inhibited the latter activity but less so than that of the Euphorbia enzyme. Furthermore, both NADH and NADPH were necessary for activity and both supported the activity to about the same extent. The activity was inhibited by cyanide but not by anti-CYP reductase antibodies or cytochrome b5 antibodies. These results were taken to confirm that the Vernonia epoxygenase is distinctly different from the usual P450 monooxygenases and from Euphorbia epoxygenase. Recently, Radmark et al. (2015) have discussed 5-LOX as a key enzyme for leukotriene biosynthesis in health and disease.

3.4.2 Peroxygenases The peroxygenase activity was first defined by Ishimaru and Yamazaki (1977) by a labeling study using pea (Pisum sativum). Hamberg and Hamberg (1996) described peroxygenase-catalyzed fatty acid epoxidation in cereal seeds. The peroxygenase pathway is now recognized (Hanano et al., 2006) as a special branch of the LOX pathway, where oxygenation of a PUFA by LOX gives rise to corresponding fatty acid hydroperoxide, which is then used by peroxygenase as oxygen donor to oxidize an unsaturated fatty acids. Meesapyodsuk and Qiu (2011) have recently isolated the peroxygenase gene involved in the biosynthesis of epoxy fatty acids in oat (Avena sativa) and have reviewed the history of the pathway. When expressed in Escherichia coli, the AsPXG1 gene catalyzes a strictly hydroperoxidedependent epoxidation of unsaturated fatty acids. It prefers hydroperoxytrienoic acids over hydroperoxy-dienoic acids as oxygen donors to oxidize a wide range of unsaturated fatty acids with cis-double bonds. Oleic acid was the preferred substrate. The AsPXG1 could use only free fatty acid or methyl ester as substrates, not PtdCho or acyl-CoA. A second gene (AsLOX2) cloned from oat codes for 9-LOX catalyzing the synthesis of 9-hydroperoxydienoic and 9-hydroperoxy-trienoic acids, respectively, from linoleic (18:29c,12c) and linolenic (18:3, 9c,12c,15c) acids used as substrates. The

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peroxygenase pathway was reconstituted in vitro using a mixture of AsPXG1 and AsLOX2 extracts from E. coli. Incubation of methyl oleate alone with a mixture of AsPXG1 and AsLOX2 produced methyl 9,10-epoxy-stearate. Incubation of linoleic acid alone with the mixture of AsPXG1 and AsLOX2 produced two major products, 9,10-epoxy-12-cis-octadecenoic acid and 12,13-epoxy-9-cis-octadecenoic acid. The biological function of these fatty acids has not been defined, but they have been perceived as precursors of oxylipins produced in response to biotic or abiotic stress (Andreou et al., 2009) or as monomers for the biosynthesis of cutin polymers to cover aerial parts of the plant surface providing a hydrophobic structural barrier to the environment (Kato et al., 1984). Munoz-Garcia et al. (2014) have discussed the importance of the LOX-hepoxilin pathway in the mammalian epidermal barrier. Meesapyodsuk and Qiu (2011) have shown a diagram illustrating the reconstituted peroxygenase pathway in E. coli in presence of linoleic acid, which results in the formation of 9,10-epoxy-18:1-12c and 12,13epoxy-18:1-9c acids (Fig. 3.6).

3.4.3 Cytochrome P450-Like Oxygenases Cahoon et al. (2002) described a CYP-like oxygenase epoxidizing the same substrate as the desaturase-linked oxygenase (see earlier discussion), giving the same product in Euphorbia lagascae. A CYP-mediated pathway thus contrasts with the route of vernolic acid synthesis in the Asteraceae C. palaestina and Vernonia galamensis. In seeds of these plants, the 12-epoxy group of vernolic acid has instead been shown to result from the activity of a 12 -oleic acid desaturaserelated enzyme (Lee et al., 1998). However, the primary structure of the E. lagascae CYP726A1 is not related to any known fatty acidmodifying CYP enzyme, including mammalian ARA epoxygenase (e.g., CYP2J2 and CYP2J6, Cahoon et al., 2002). Capdevila et al. (1996) had earlier shown that P450BM3 catalyzed both the hydroxylation and epoxidation of ARA yielding 18-hydroxyarachidonic acid and 14,15-epoxyarachidonic in 80% and 20% yields, respectively. The absolute configuration was tentatively assigned as 15(R),16(S)-epoxyoctadeca-9,12-dienoic acid by direct comparison with published data. In contrast, with linolenic acid, Celik et al. (2005) observed exclusive epoxidation of the C-14,15-double bond with no other detectable mono-hydroxylation products. Falk et al. (2001) observed similar high levels of regioselectivity for the P450BM3-catalyzed reaction with linoleic acid, although in this case the exclusive product was C-12,13-epoxy derivative. The allylic epoxides with conjugated trienes are well known in mammalian systems, particularly the ARA-derived 5-LOX product LTA4, the immediate precursor of the proinflammatory dihydroxy LTB4 (Samuelsson et al., 1987). Niisuke et al. (2009) have compared the mechanisms of transformation in the formation of three allylic epoxides (Fig. 3.7). A chemical

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OH O

OH O

AsLOX2

OH O

9-O-OH-18:2-10t,12c

18:2-9c,12c

AsPXG1

O

OH O

12,13-epoxy-18:1-9c O OH OH

O

O

OH

9-OH-18:2-10t,12c 9,10-epoxy-18:1-12c

FIGURE 3.6 A diagram illustrating the reconstituted peroxygenase pathway in E. coli in presence of linoleic acid. Reproduced with permission from Meesapyodsuk and Qiu (2011). A peroxygenase pathway involved in the biosynthesis of epoxy fatty acids in oat[W][OA]. Plant Physiol. 157, 454463.

synthesis of an analog of the Anabaena allylic epoxide was achieved using 13-HPODE methyl ester as starting material using a method originally developed for synthesis of LTA4 and related conjugated trienoic epoxides (Fig. 3.7C). Using a method originally developed for synthesis of LTA4 and related conjugated trienoic epoxides (Corey et al., 1980; Atrache et al., 1981), Niisuke et al. (2009) obtained the equivalent results with C18:2 starting material. Transformation of the fatty acid hydroperoxide to the allylic epoxide is mechanistically quite distinct in the 5-LOX and Anabaena reactions (Fig. 3.7A,B). Conventional LTA4 synthesis by 5-LOX is initiated by abstraction of a bis-allylic hydrogen from the fatty acid hydropeoxide (Fig. 3.7A). Specifically, a “biomimetic” chemical synthesis of allylic epoxide takes place following a reaction of peroxy trifluoromesylate by basecatalyzed cleavage of the peroxide, followed by a final proton elimination to two allylic epoxides (Niisuke et al., 2009). Under the influence of the mammalian CYP enzymes (CYP-450), ARA gives rise to four regioisomers of EETs: 5,6-, 8,9-, 11,12-, and 14,15-EET (Zhang and Blair, 1994). CYP oxygenases also covert the ω-3-PUFA EPA and DHA to epoxyderivatives (Fer et al., 2006). The EPA-derived epoxides account for five EpETEs (5,6-, 8,9-, 11,12-, 14-15-, and 17,18-EETeTr), whereas the DHA-derived epoxides account for five EDP (4,5-, 7,8-, 10,11-, 13,14-, 16,17-, and 19,20-EDP) (Van Rollins, 1995). The stereoselectivity of the epoxidation reaction has been characterized by comparison with the long-chain PUFA epoxide stereoisomers obtained from the enantioselective bacterial CYP102A1 F87V (Lucas et al., 2010).

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Supplemental Material can be found at: http://www.jlr.org/content/suppl/2009/02/27/M900025-JLR20 0.DC1.html (A)

(B)

Typical lipoxygenase-catalyzed leukotriene epoxide biosynthesis

OOH

H

H

(C) Allylic epoxide biosynthesis by Anabaena minicatalase

OOH

O-SO2CF3 O H

H

O

Reaction initiated by base-catalyzed cleavage of the peroxide

H

H

H

Peroxy trifluoromesylate

Proposal: reaction initiated by cleavage of the peroxide

Reaction initiated by stereospecific hydrogen abstraction OOH

H

H

“Biomimetic” chemical synthesis of allylic epoxide

O

+

H

H

Electron transfer Peroxyl cleavage and elimination of the elements of H2O leads to LTA-type epoxide

O

+

H+

H

H

Final proton elimination leads to two allylic epoxides H+

O

H

Trans-epoxide, trans,trans,cis double bonds

O

H

Trans-epoxide, trans,trans double bonds

O

H

Isomeric trans-epoxide with trans,cis diene

FIGURE 3.7 Comparison of mechanisms of transformation to allylic epoxides. (A) LOXcatalyzed LTA-type epoxide biosynthesis. (B) Transformation of 9R-HPODE by the Anabaena enzyme. (C) Chemical biomimetic synthesis of allylic epoxide. Reproduced with permission from Niisuke et al. (2009). Biosynthesis of a linoleic acid allylic epoxide: mechanistic comparison with its chemical synthesis and leukotriene A biosynthesis. J. Lipid Res. 50, 14481455.

The stereoselectivity of the epoxidation of the last olefin of ARA (ω-6), EPA (ω-3), or DHA (ω-3) differed between the CYP isoforms, but was similar for EPA and DHA. In addition to previously reported fatty acids, Xiao and Guengerich (2012) have identified oleyl (18:1) lysophosphatidylcholine (LPC) along with other lysophosphatidylinositol (LPI), lysophosphatidylserine (LPS), lysophosphatidylethanolamine (LPE), and lysophosphatidic acid (LPA) as substrates for P450 2W1, but not diacylglycerophospholipids. The sn-1-isomer of LPC was utilized much more efficiently than the sn-2-isomer, as well, 18:1 LPC was utilized much more efficiently than the free 18:1 acid. Chiral analysis of the 18:1 epoxidation products showed an enantioselectivity for formation of (9R,10S) over (9S,10R).

3.5 ANALYSIS OF EPOXY FATTY ACIDS Depending on the epoxide synthetase, the epoxy fatty acids may be formed in situ on glycerophospholipids or in the form of free fatty acids, which may

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become subsequently incorporated into glycerophospholipids or triacylglycerols. The following brief review is confined to the analysis of free epoxy fatty acids or their methyl esters.

3.5.1 Resolution of Regioisomers The original separations of fatty acid epoxide regioisomers were obtained by thin-layer and gas-liquid chromatography (GC), and GC/MS methods, which required chemical modification of the solutes. Thus, the isomeric abundance of the linoleate-derived epoxide mixture was quantified by GC-MS following chemical hydrolysis, methylation, thiolation, and silylation as described by Newman and Hammock (2001). More recently, regioisomer separations have been obtained by normalphase and reversed phase HPLC or with LC-MS/MS. Thus, Kiss et al. (2000) used isocratic reversed phase HPLC (MeCN:MeOH:water:AcOH, 54:8:38:0.001, v/v/v/v) with photodiode array detection for the separation of all four regioisomeric cis-EETs (see Fig. 3.2), with 8,9-EET representing the predominant compound. Newman et al. (2002) quantified the epoxyeicosanoid regioisomers derived from ARA by LC-MS/MS. The solvent system was modified from Kiss et al. (2000): solvent A (acetonitrile:water:methanol, 51:40:9, v/v/v) with 0.1% glacial acetic acid; solvent B (acetonitrile:methanol, 85:15, v/v) with 0.1% glacial acetic acid. An isocratic flow of 96% solvent A and 4% solvent B at 2 mL min21 gave optimal EET isomer resolution: 14(15)-EET followed by 11(12)-EET, followed by 8(9)-EET. Newman et al. (2002) have used simultaneous multireaction monitoring to demonstrate the presence of numerous epoxy fatty acids in the urine of a hypertensive rat (Fig. 3.8). The detected EpOMEs, EETs, the epoxide extraction SSTD, and the internal standard (ISTD) are indicated with arrows. The Y-axis labels indicate the intensity of the dominant ion within each trace. Cone voltage manipulations had a dramatic effect on the intensity of the deprotonated molecular ion ([M-H]2) in MS and MS/MS, which required optimization. The authors tabulated human urinary oxylipids in pmol mg21 creatinine. In all cases, 9,10-dihydroxyoctadecenoic acid (DiHOME) was 90% of the DiHOME profile, while 8,9-DiHET was 70% of the DiHET profile. Morisseau et al. (2010) prepared the regioisomers of EpETE and EpDPE generated by reacting the methyl esters with meta-chloroperbenzoic acid. Both standards and fatty acid epoxides recovered from tissues were analyzed by LC-MS/MS. All HPLC/MS analyses were performed with a Waters ULPC separation module equipped with a 2.0 3 150 mm 5 μm Luna C18 column (Phenomenex) held at 40 C. The HPLC was interfaced with ESI probe of Quattro Ultima tandem-quadrupole mass spectrometer (Waters, Milford, MA, United States).

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Fatty Acids

4.66e6

16.43

12(13)-EpOME

295.2 > 195

17.11 16.48

10(11)-EpHep (SSTD)

5.35e5

16.99

2.93e6

283.2 > 265.2

295.2 > 171 9(10)-EpOME 17.79 14(15)-EET

16.82

2.27e4

19.71

17.67

4.31e3

18.52

11(12)-EET

4.37e4

1.69e4

19.37

18.86

8(9)-EET

5(6)-EET

319.2 > 219.1

319.2 > 208

20.16

19.71

319.2 > 155

319.2 > 191.2 20.62

3.77e6

20-HE (ISTD)

327.3 > 309.3

16.0

18.0

20.0 Time (min)

22.46

22.0

FIGURE 3.8 Simultaneous multireaction monitoring of epoxy fatty acids from a representative series of extracted ion chromatograms from the urine of a single spontaneously hypertensive rat. LC-MS/MS was performed with a Waters 2790 separation module equipped with a 2.0 3 150 mm, 5 μm Luna C18(2) column (Phenomenex) held at 20 C. The solvents were modified from Kiss et al. (2000): Solvent A 5 51:40:9 acetonitrile:water:methanol (v/v/v) with 0.1% glacial acetic acid; Solvent B 5 85:15 acetonitrile:methanol (v/v). The flow rates and times of solvent change were selected for optimum peak resolution. A 75 cm segment of 0.005 in. i.d. PEEK tubing interfaced the HPLC to the electrospray ionization probe of a Quatro Ultima tandem-quadrupole mass spectrometer (Micromass, Manchester, United Kingdom). Reproduced with permission from Newman et al. (2002). The simultaneous quantification of cytochrome P450 dependent linoleate and arachidonate metabolites in urine by HPLC-MS-MS. J. Lipid Res. 43, 15631578.

Solvent flow rates were fixed at 350 μL min21 with a cone gas flow of 125 L h21, desolvation gas flow of 650 L h21, source temperature of 125 C, and desolvation temperature of 400 C. For MS/MS argon was used as the collision gas, Morisseau et al. (2010) presented the data in a tabular form to

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show the content of fatty acid monoepoxides in rat brain and spinal cord. Although only the 17,18-EpETE regioisomer of parent EPA was detected in the brain and spinal cord, all of the EpDPE regioisomers of parent DHA were present. Except for the 7,8-EpDPE, which was present in relatively high quantity compared with the other epoxy fatty acids, the EpETEs in the CNS were in similar quantities to the EET regiosomers. Similarly, for the C18 fatty acids, the n-3 α-EpODEs were present in similar quantities in the n-6 EpOME. Levison et al. (2013) described quantification of fatty acid oxidation products using online reversed phase LC-MS/MS with synthetic standards. The use of 15(S)-HETE-d8 allowed the quantification of regioisomeric EETs, hydroxyeicosatetraenoic acids (HETEs), hydroxyoctadecadienoic acids (HODEs), oxoETEs, oxoODs, as well as linoleic acid and ARA in human plasma. Fig. 3.9 shows a representative negative ion LC-MS/MS chromatogram of unoxidized fatty acids and oxidized fatty acids extracted from human plasma (Levison et al., 2013).

3.5.2 Resolution of Enantiomers In most studies, the enantiomers have not been assessed by chiral analysis. Therefore, a correlation between the occurrence of a specific enantiomer and its putative biological action is usually impossible. The enantiomers have been resolved by chiral columns using the free acids or their methyl esters. Capdevila et al. (1991, 1996) utilized degradative ozonolysis followed by derivatization to the corresponding 3,4-epoxyhexan-1-yl benzoates for chiral analysis of 17,18-epoxy-EPA. Chiralcel OC HPLC properties of synthetic standards were then compared to those of the biologically derived samples. Chiral analysis of the EPA epoxygenase metabolite demonstrated that its biosynthesis was highly asymmetric and generated 17(S),18(R)-epoxyEPA with 97% optical purity. Le Quere et al. (2004) resolved the enantiomers of 18-hydroxy-C18epoxides as methyl ester derivatives using a mixture of solvents (hexane:isopropanol:acetic acid, 90:10:0.1, v/v/v) in an isocratic mode for 60 minutes, using a chiral column (Chiracel OB, 4.6 3 250 mm, J.J. Baker Chem. Co., Phillipsburg, NJ), as described by Pinot et al. (1992). Fig. 3.10 shows chiral-phase HPLC of radiolabeled 18,9(10)-HEpSTA generated by human recombinant CYP4F2 (A) and CYPA11 (B). Metabolites were collected from an RP-HPLC column and identified by reference to a standard (Le Quere et al., 2004). Celik et al. (2005) determined the enantioselective epoxidation of linolenic acid at the C-15,16-double catalyzed by CYPBM3 from Bacillus megaterium. The optical purity of the epoxide was obtained by an initial conversion to the methyl ester followed by HPLC analysis (Chiralcel OJ-H, Daicel Chemical Labs., Tokyo, Japan) using a racemic sample for comparison. HPLC analysis indicated that the product had an enantiomeric excess of

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Arachidonic acid

0 100

5-HETE

0 100 0 100

9-HETE

0 100

8,9-EET

11-HETE

0 100 Relative intensity (%)

8,9-EET

8-HETE

11,12-EET

12-HETE

0 100

11,12-EET

15-HETE

0 100

15-HETE-d8

0 100

PGF2

0 100

PGE

PGF2-d4

0 100

Linoleic acid

0 100

9-HODE

0 100

13-HODE

0 100

9-oxoODE

0 100

13-oxoODE

0 2

6

10 14 Retention time (min)

18

22

FIGURE 3.9 Representative LC-MS/MS chromatogram of unoxidized fatty acids and oxidized fatty acids extracted from human plasma. Separation performed on a reversed phase C18 column (2.1 3 250 mm, 5 μm) with acidified methanol/water as mobile phase at a flow rate of 0.2 mL min21 using a gradient elution. The oxidized fatty acid species had common multireaction monitoring (MRM) pairs and unique retention times; 8,9-EET exhibits the same MRM transitions as both 8-HETE and 9-HETE, and 11,12-EET shares the same MRM transitions with both 11-HETE and 12-HETE. The unique retention times of the 8,9- and 11,12-EET were established by injection of authentic standards (data not shown). Reproduced with permission from Levison et al. (2013). Quantification of fatty acid oxidation products using on-line high performance liquid chromatography tandem mass spectrometry. Free Radical Biol. Med. 59, 213.

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FIGURE 3.10 Chiral-phase HPLC of radiolabeled 18,9(10)-HEpSTA generated by human recombinant CYPF2 (A) and CYP4A11 (B). Metabolites collected from RP-HPLC column, converted to methyl ester, and analyzed using chiral-phase column (Chiracel OB, 4.6 3 250 mm) and a mixture of solvents (hexane:isopropanol:acetic acid, 90:10:0.1, v/v/v) in isocratic mode for 60 min; for residual C18-epoxides, the solvent mixture was hexane:isopropanol:acetic acid (99.7:0.2:0.1, v/v/v) at a flow of 0.8 mL min21. Radioactivity monitored by a computerized online scintillation counter (Flo-one Beta Radiometric Detector). Stereoisomers of 18,9(10)HEpSTA were generated by CYP4F2 (A) and CYP4A11 (B). Reproduced with permission from Le Quere et al. (2004). Human CYP4F3s are the main catalysts in the oxidation of fatty acid epoxides. J. Lipid Res. 45, 14461458.

60%. The absolute configuration of the epoxide was tentatively assigned as 15(R),16(S)-epoxyoctadeca-9,12-dienoic acid by direct comparison with published data for linoleic acid (Falk et al., 2001). It was assumed that the order of elution of the 15(R),16(WS)-enantiomer of linolenic acid was the same as for linoleic acid, using the identical chiral HPLC column, in which the methyl ester of 15(S),16(R)-epoxyoctadeca-9,12-dienoic acid eluted first (9.2 minutes) followed by 15(R),16(S)-epoxyoctadeca-9,12-dienoic acid (10.4 minutes). Starting with the chiral chromatographic conditions described by Zhang and Blair (1994), Kiss et al. (2008) developed a method for consecutive regional and enantiomeric separation of the four underivatized EET regioisomers within one chromatographic run employing capillary tandem column chiral-phase HPLC. It was possible to obtain a highly sensitive, direct, and simultaneous chiral analysis of all eight EET enantiomers. A chiral-phase HPLC method based on Chiralcel OD (a cellulose-3,5-dimethylphenylcarbamate impregnated silica) column was developed for direct separation of EET as free fatty acids and their methyl esters. The mobile phases contained variable proportions of either 2-propanol/hexane/AcOH(free acid) or variable proportions of 2-propanol/hexane (methyl ester). The method provided a consecutive regional and enantiomeric separation of the four underivatized EET regioisomers within one chromatographic run employing capillary tandem column chiral-phase LC/ESI-MS for identification and quantitation of

100

(A)

Fatty Acids

204 mm

198 - 202nm

IS - EPA

190

EETE 1a 1b 2a

nm 270 340

2b

3a

3b

4a

4b min

0 20 40 60 80 (B) EIC 301; 275; 257; 221; 219; 208; 205; 195; 191; 179; 175; 167; 163; 155; 151; 149; 139; 135; 129; 127; 123; 115; 113; 99 MS2 (319) 2a 2b 3a 1a 1b

100 3b 4a

4b min

0 20 (C) EIC 283; 257; 203 MS2 (301) IS - EPA

60

40

80

2

EIC 299; 281; 273; 255; 217; 235; 219; 207; 175; 163; III MS (317)

EETE

(D)

100

min

min 0

20

(E)

1a

40 1b

60

2 EIC 219; 205; 175; 113; 99 MS (319.1)

80

100

EIC 208; 195; 179; 167; 163; 149; 165 MS2 (319.1)

(F)

2a

2b min

min 40 (G)

50 3s

3b

50

60

2 EIC 221; 179; 155; 151; 139; 127; 123; MS (319.1)

(H)

60

70

EIC 219; 205; 191; 129; 115; 99 MS2 (319.1)

4a

4b

min

min 80

90

100

80

90

100

FIGURE 3.11 Simultaneous separation of the four racemic underivatized EET regioisomers into the corresponding consecutively eluting nonoverlapping pairs of enantiomers in one chromatographic run as a result of tandem capillary column coupling (a nonchiral Grom-Sil Amino4PR column for effective regioisomeric separation followed by a Chiracel OD-H chiral column for simultaneous enantiomer resolution, both with CT set to 26 C) and isocratic normal-phase elution using a 99.7:0.21:0.09:0.015 (v/v/v/v) mixture of nHex:IUPA:EtOH:AcOH as mobile phase. Reproduced with permission from Kiss et al. (2008). Direct and simultaneous profiling of epoxyeicosatrienoic acid enantiomers by capillary tandem column chiral-phase liquid chromatography with dual online photodiode array and tandem mass spectrometric detection. Anal. Bioanal. Chem. 392 (4), 717726.

eluting optical antipodes. Fig. 3.11 shows a simultaneous separation of the four racemic underivatized EET regioisomers into the corresponding consecutively eluting nonoverlapping pairs of enantiomers in one chromatographic run (Kiss et al., 2008). A tandem capillary column coupling (a nonchiral Grom-Sil Amino-4PR column, for effective regioisomeric separation, followed by a Chiralcel OD-H column for simultaneous enantiomeric resolution) was combined with an isocratic normal-phase elution using 99.7:0.21:0.09:0.015 (by volume) mixture of hexane, 2-propanol, EtOH, and AcOH as mobile phase at a flow rate of 8 μL min21. Mesaros et al. (2010) prepared [13C20]EET analog internal standard and used it to validate a high-sensitivity chiral LC/electron capture atmospheric pressure chemical ionization (ECAPCI)-MS method for the trace analysis of endogenous EETs as their pentafluorobenzyl (PFB) ester derivatives. Fig. 3.12 shows LC/MRM-MS chromatograms of EET-PFB standards and corresponding [13C]labeled internal standards. The assay was used to show

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FIGURE 3.12 LC/MRM-MS chromatograms of EET-PFB standards and corresponding [13C]labeled internal standards. Normal-phase chiral chromatography using Waters Alliance 2690 HPLC system. Gradient elution in a linear mode on a Chiralpak AD-H column (4.6 i. d. 3 250 mm; Daicel Chemical Industries, Tokyo, Japan) was used at a flow rate of 1 mL min21. Solvent A was hexane and solvent B was 2-propanol:hexane (6:4, v/v). Isocratic elution was used with 1.5% B for 12 min and then a linear gradient to 100% A for 10 min for washing the column. Mass spectrometry was done on a Thermo Finnigan TSQ Quantum Ultra AM mass spectrometer equipped with an APCI source in the EC negative ion mode. CID was performed using argon as the collision gas at 1.5 mTorr in the second (r4f-only) quadrupole. Peak are identified as shown in the figure. Reproduced with permission from Mesaros et al. (2010). Analysis of epoxyeicosatrienoic acids by chiral liquid chromatography/electron capture atmospheric pressure chemical ionization mass spectrometry using [13C]-Analog internal standards. Rapid Commun. Mass Spectrom. 24, 32373247.

the exquisite enantioselectivity of P4502C19-, P4502D6-, P4501A1-, and P4501B1-mediated conversion of ARA into EETs and to quantify the enantioselective formation of EETs produced by ARA metabolism in a mouse epithelial hepatoma cell line. Fig. 3.13 shows enantioselective biosynthesis of EETs by CYP family 2 isoforms: (1) hcCYP2C19 and (2) hCYP2D6 (Mesaros et al., 2010). There is a striking difference in the enantioselectivity of 14,15-EET formation between CYP2C19 and CYP2D6. All the

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FIGURE 3.13 Enantioselective biosynthesis of EETs by P450 family 2 isoforms: (A) hP4502C19 and (B) hP4502D6. Peaks are identified as shown in figure. LC-MS/MS conditions and column were as given in Fig. 3.12. Reproduced with permission from Mesaros and Blair (2010). Targeted chiral analysis of bioactive arachidonic acid metabolite using liquid chromatography-mass spectrometry. Metabolites 2, 337365.

regioisomers of racemic epoxy-eicosatrienoates were resolved as the PFB esters (Mesaros et al., 2010). These compounds were completely separated on columns of Chiralcel OB and OD (cellulose tris-3,5-dimethylphenylcarbamate). With most of the isomers, the columns were used in the adsorption mode, but separation of the enantiomeric 5,6-epoxy-eicosatrienotates was possible only in the reversed phase mode, that is, with a mobile phase of H2O:EtOH (30:70, v/v) (Mesaros et al., 2010). There was no problem of column collapse during the EtOH:H2O (70:30, v/v) elution of Chiralcel B (Schneider et al., 2007). An alternative method of enantioseparation of cis-EETs uses capillary electrophoresis (VanderNoot and Van Rollins, 2002a,b; Van Rollins and VanderNoot, 2003). Six EET enantiomers, 8(S)-9(R)-, 8(R)-9(S)-, 11(S)12(R)-, 11(R)-12(S)-, 14(S)-15(R)-, and 14(R)-15(S)-EETs, were successfully separated by capillary electrophoreisis (CE) using a mixture of β-CD and β-CD-sulfobutyl ether. However, analysis time was long. This can be overcome by employing another CE-based method, MEKC, used

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for the enantioseparation of 8-, 11-, 12-, and 15-HETEs (Kodama et al., 2016).

3.5.3 GC/MS and LC/MS Identification of Lipid Epoxides Lee et al. (1998) used GC/MS to identify crepianic and vernolic acids in the Arabidopsis thaliana seed fatty acids. Celik et al. (2005) used GC/MS coelution and spectral comparisons with authentic standards for the investigation of CYP epoxidation of linoleic acid methyl ester. The absolute configuration of the product was tentatively assigned as 15(R),16(S). More recently, the GC/MS methods have been replaced by LC/MS methods, including those performed with underivatized epoxy fatty acids. To efficiently conduct targeted eicosanoid analyses, HPLC separations are coupled with collision-induced dissociation (CID) and tandem mass spectrometry (MS/MS). Product ion profiles are often diagnostic for particular regioisomers. The highest sensitivity that can be achieved involves the use of selected reaction monitoring mass spectrometry (SRM/MS), whereas the highest specificity is obtained with an SRM transitions between an intense parent ion, which contains the intact molecule (M), and a structurally significant product ion (Lee and Blair, 2009; Mesaros and Blair, 2012). Newman et al. (2002) have detailed analytical methods based on LC/MS technology for highly sensitive simultaneous resolution and quantification of multiple analytes in complex samples suitable for exploring the biological activity of the epoxides. The HPLC/MS analyses were performed with a Waters ULPC separation module equipped with a 2.0 3 150 mm, 5 μm Luna C18 column (Phenomenex) held at 40 C. The sample chamber was held at 10 C. The HPLC was interfaced to ESI probe of a Quattro Ultima tandemquadrupole MS (Waters). Solvent flow rates were fixed at 350 μL min21 with a cone gas flow of 125 L h21, desolvation gas flow of 650 L h21, a source temperature of 125 C, and a desolvation temperature of 400 C. For MS/MS experiments, argon was used as a collision gas at a pressure of 2.3 3 1023 Torr. The optimized declustering potential needed to produce the molecular ion and the collision energy was adjusted as suggested by Newman et al. (2002). As observed for linoleate and arachidonate epoxides and diols (Newman et al., 2002), the most characteristic daughter ion for each analyte was the one resulting from breakage of epoxide ring or the bond between the two alcohol groups. Surrogate standards [11,12-EET-d8, 10,11-dihydroxyundecanoic acid (10,11), and 10(11)-epoxyheptadecanoic acid (10,11-EpHep)] were added to samples before extraction. Xiao and Guengerich (2012) used the LC/MS metabolomic and isotope labeling approach to conduct untargeted substrate searches in human colorectal cancer samples. A series of lysophospholipids and FFAs were identified as novel substrates for P450 2W1 and the isomer and enantiomer selectivity determined. Enantiomeric 9,10-epoxystearic acids obtained by hydrogenation

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of the epoxides and released by chemical hydrolysis were identified by normal-phase HPLC with Waters Alliance 2695 HPLC pump (Waters) and a Chiralpak AD column (5 μm, 4.6 mm 3 25 cm). An isocratic solvent of 100:2:0.05 (v/v/v) hexanes:CH3OH:CH3CO2H mixture was used to resolve the enantiomers at a flow rate of 1 mL min21 at room temperature. The retention times of the (9S,10R)- and (9R,10S)-epoxystearic acids were 16.9 and 18.7 minutes, respectively. Optically pure (9S,10R) and (9R,10S)-epoxystearic acids were prepared by hydrogenating pure (9S,10R)-epoxy-12Z-octadecenoic acid and (9R,10S)-epoxy-12Z-octadecenoic acid (Gao et al., 2009) with Pd powder under a H stream for 3 minutes (Tang et al., 2009). Levison et al. (2013) have described quantification of fatty acid oxidation products using online reversed phase HPLC with tandem mass spectrometry. In the protocol, addition of synthetic internal standard to the sample, followed by base hydrolysis at elevated temperature, and liquidliquid sample extraction with lighter than water solvents led to isolation of the oxidized fatty acids. Using 15(S)-HETE-d8 allowed the quantification of regioisomeric EETs, HETEs, HODEs, oxoETEs, oxoODs, as well as linoleic acid and ARA in human plasma.

3.6 BIOLOGICAL EFFECTS The biological effects of epoxy fatty acids include signaling and lipid mediator activity. Spector and Kim (2015) have recently summarized the biological mechanisms and functions of epoxy fatty acids using 11,12-EET as an example (Fig. 3.14). As shown in the outline, many EET functions occur through a membrane receptor-dependent mechanisms, which activate signal transduction pathways that modulate ion channels and transcription factors in the target cell. In most studies, the enantiomers have not been assessed by chiral analysis. Thus no correlation between a specific enantiomer and its putative biological action was possible. Herein is a brief summary of biological activity of epoxy fatty acids limited to specific regioisomers and enantiomers. References to general effects related to changes in diets and lipid classes, or changes in enzyme levels and enzyme activity, have been limited or excluded.

3.6.1 Lipid Signaling Both ω-3 and ω-6 fatty acid series are substrates of CYP epoxygenases, which convert them to epoxy signaling lipids including EETs derived from the ω-6 ARA and EDP derived from ω 3-DHA (Capdevila et al., 1992; Zeldin et al., 2001). It has been shown that EDPs are more potent than EEts in vasodilation, and vascular tone and antiinflammation (Morriseau et al., 2010; Node et al., 1999).

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FIGURE 3.14 Mechanism of action of EETs using 11,12-EET as an example (solid arrows). Some EET responses may occur through intracellular effects or direct interactions with ion channels (dashed arrows). Listed are the signal transduction pathways, ion channels, and transcription factors that are targeted by EETs in various cells, and the functional responses that occur in the cardiovascular, renal, and nervous system. Reproduced with permission from Spector and Kim (2015). Cytochrome P450 epoxygenase pathway of polyunsaturated fatty acid metabolism. Biochim. Biophys. Acta 1851, 356365.

EETs have been shown to inhibit inflammation via blocking NF-κB, an important signal regulating inflammation. The NF-κB complex is sequestered in the cytoplasm through binding to the inhibitory protein 1kBα. Node et al. (1999) showed that 11,12-EET, but not 14,15-EET, inhibited tumor necrosis factor (TNF)-αinduced 1kBα degradation and nuclear translocation of NFκB in endothelial cells, suggesting that EETs inhibit inflammation via blocking NF-κB signal activation. Panigrahy et al. (2010a) have discussed EET signaling in cancer.

3.6.2 Cellular Effects Unesterified EETs are taken up from extracellular fluid by various cells to be incorporated into cellular glycerolipids (Spector et al., 2004). Lee and Blair (2009) developed a targeted lipidomics approach that makes it possible to directly analyze chiral epoxides generated in cellular systems by LC/MS/ MS. EETs, ω-3-PUFA epoxides, and the corresponding diols are present in the glycerophospholipids of plasma lipoproteins (Shearer et al., 2010; Kuksis and Pruzanski, 2013). EETs are incorporated into the sn-2-position of cell glycerophospholipids, mostly in PtdCho, but 14,15-EET is also found in

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PtdIns (Spector et al., 2004). These compounds are released when plasma lipoproteins are hydrolyzed by lipoprotein lipase from very low-density lipoproteins (LDLs) (Shearer and Newman, 2008). The epoxy and hydroxy fatty acids of plasma LDL and HDL PtdCho have been shown to be released by secretory phospholipases, groups IIA, V, and X in vitro (Kuksis and Pruzanski, 2013). The biological actions of 11,12-EET in endothelial cells are specific to the R/S-enantiomer and require the Gs protein (Ding et al., 2014). The results suggest that a Gs-coupled receptor in the endothelial cell membrane responds to 11(R),12(S)-EET and mediates the protein kinase A (PKA)-dependent translocation and activation of TRPC6 channels. This process affects angiogenesis (Ding et al., 2014). The EETs have potent vasodilator and antiinflammatory activities, and EETs can inhibit platelet aggregation depending on their chirality and regiochemistry. The antiinflammatory effects of EETs in vitro are regioselective: 11,12-EET has the most potent effect, followed by 8,9- and 5,6-EET, while 14,15-EET is inactive (Node et al., 1999). 11,12- and 8,9-EET inhibit basal TNF-α production in THP-1 cells. 11,12-EET dose-dependently suppress LPS-induced PGE2 formation by inhibiting the enzyme activity (Kozak et al., 2003). They are lipid mediators that regulate inflammation and vascular tone and drugs that raise EET levels are in clinical trials for the treatment of hypertension. The epoxide hydrolases (EHs) are present in all living organisms, and transform epoxide containing lipids by the addition of water. In plants and animals, many of these lipid substrates have potent biological activities, such as host defenses, control of development, regulation of inflammation and blood pressure, and tumor growth and metastasis (Newman et al., 2005). It was generally believed that epoxide hydrolysis eliminated the biological activity of these lipids. However, the dihydroxyeicosatrienoic acid (DiHET) products are active in some systems, for example, vasodilation, sodium channel activation. Newman et al. (2005) have reviewed the EHs and their roles and interactions with lipid metabolism. Seven distinct EH subtypes were recognized in higher organisms, including the plant soluble EHs, the mammalian soluble EHs, the hepoxilin hydrolase, leukotriene A4 hydrolase, the microsomal EH, and the insect juvenile hormone EH (Newman et al., 2005). For structures of the dihydroxy products of hydrolysis of the epoxides of ARA (EETs), EPA (EEQs), and DHA (EPD), see Figs. 3.2 and 3.3. The 14,15-, 11,12-, and 8,9-EETs are excellent substrates for the soluble EHs (Roman, 2001). A number of structurally different but potent small molecule inhibitors (soluble epoxide hydrolase inhibitor, sEHIs) have been demonstrated to stabilize the EpFAs in vivo, strongly indicating that the mechanism of action of sEHI is through stabilization and prolonging the activity of EpFAs (Morisseau and Hammock, 2013; Inceoglu et al., 2013).

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Shen and Hammock (2012) have discussed the chemistry of inhibition of sEHs, which may serve as a target for therapeutic interference, but have cautioned whether sEH inhibition is a robust mechanism to treat hypertension and/or diabetes.

3.6.3 Systemic Effects At the systemic level, the EETs have significant roles in the regulation of vascular, cardiac, pulmonary, and renal physiology (Spector et al., 2004; Spector and Kim, 2015). They are potent regulators of smooth muscle tone, cell proliferation, and migration. EETs are hydrolyzed to their vicinal diols or DiHETs. Spector and Kim (2015) have visualized the mechanism of action of EETs via putative EET receptor located in the cell membrane (Fig. 3.14). The actions of EETs have been shown to be elicited by specific optical isomers. Thus, 14(R),15(S)-EET is a stereospecific inhibitor of cyclooxygenase (COX), 11(R),12(S)-EET is a potent renal vasodilator, while 8(S),9(R)EET is adrenal vasoconstrictor. Their optical antipodes, 14(S),15(R)-, 11 (S),12(R)-, and 8(R),9(S)-EET, respectively, are inactive (Zhang and Blair, 1994). In the brain, EETs are involved in controlling the cerebral blood flow (Puppolo et al., 2014). A deletion of sEH, the enzyme that metabolizes EETs to dihydroxyeicosatetraenoic acids (DiHETEs), was found to be protective against ischemic brain injury. CYP-derived EETs (and their hydration products, the DiHETs) are vasodilators (Natarajan and Reddy 2003), whereas P450-derived 20-HETE is a vasoconstrictor (Miyata and Roman, 2005). The enzymic oxidation of ARA is a key factor in the blood pressure regulatory cascade (Roman, 2001). Hydroxylation of the ω-carbon of the ARA chain yields 20-HETE, a potent vasoconstrictor (Roman, 2001). The action of 20-HETE is opposed by the epoxides of ARA, that is, EETs, which increase the open state probability of K 1 Ca channels, in particular, the 11(12)-EET. The EET regioisomers also affect mitogenesis and sex or developmental hormone secretion (Newman et al., 2002). EETs (from ARA) produce vascular relaxation, have antiinflammatory effects on blood vessels and in the kidney promote angiogenesis, and protect ischemic myocardium and brain (Capdevila et al., 2000; Spector et al., 2004; Spector and Kim, 2015). A high regio- and stereo-specificity has been observed when testing the effects of chemically synthesized epoxides from ARA and EPA. For example, renal rat arteries were dilated by 11(R),12(S)-EET but not by 11(S),12 (R)-EET or 14,15-EET enantiomers (Zou et al., 1996). Also in rats, among all EETTeTr enantiomers, only the 17(R),18(S)-enantiomer, not 17(S),18(R), was effective on calcium-activated potassium (BK) channels in cerebral arteries (Lauterbach et al., 2002). However, in porcine coronary microvessels, all regioisomeric EETTeTr had vasodilatory potencies (Zhang et al.,

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2001). EPA- and DHA-derived epoxides are potent dilators of coronary arterioles (Hercule et al., 2007; Lauterbach et al., 2002; Ye et al., 2002), pulmonary artery (Morin et al., 2009), and inhibit platelet aggregation (Van Rollins, 1995). The biological activity of DHA-derived epoxide regioisomers and enantiomers is still largely unknown as are the identities involved in EPA and DHA metabolism (Serhan and Chang, 2008; Serhan and Petassis, 2011; Serhan et al., 2015).

3.7 PATHOLOGICAL EFFECTS 3.7.1 Toxicity The P450s in the CYP1 and CYP2 gene families metabolize linoleic acid at rates comparable to ARA and produce linoleic acid monoepoxides as major products. Moran et al. (2000) tested the cytotoxic properties of linoleic acid, linoleic acid monoepoxides, and corresponding diols in a rabbit renal proximal tubule model. They were found to be toxic at concentrations of 100500 μM and disrupted mitochondrial function with subsequent loss of ion transport and cell death. Moran et al. (2000) found no evidence that oxidative stress plays a significant role in the toxicity of these compounds. Le Quere et al. (2004) have suggested that epoxidized fatty acids from the C18 family (C18-epoxides) such as a Z9(10)-epoxyoctadecanoic acid [Z9 (10)-EpSTA], Z9(10)-epoxyoctadec-Z12-enoic acid [Z9(10)-EpOME, leukotoxin], and Z12(13)-epoxyoctadec-Z9-enoic acid [Z12(13)-EpOME, isoleukotoxin] should be regarded as toxic and or defensive substances in vivo because they have been described as toxic metabolites in mammals (Moran et al., 2000) and as defense compounds in infected plants (Kato et al., 1984). Greene et al. (2000) have reported toxicity of epoxy fatty acids and related compounds to cells expressing human sEH. Le Quere et al. (2004) have shown that the hydroxylation of Z9(10)EpSTA, Z9(10)-EpOME (leukotoxin), and Z12(13)-EpOME (isoleukotoxin) and that of monoepoxides from ARA EET are important in the regulation of leukotoxin and EET activity.

3.7.2 Inflammation and Pain Inflammation is a common pathological process, therefore modulation or inhibition of inflammation is important in therapeutic strategy. Lipid mediators play a central role in regulation of inflammation (Hansson, 2005), including PGE2, which is a COX-2 metabolite of ARA. Pruzanski and Kuksis (2014) have suggested that secretory phospholipases A2 may mediate inflammatory and proatheromatous changes in rheumatic diseases.

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Recently, EETs have been demonstrated to have potent antiinflammatory effects in vitro and in vivo (Node et al., 1999), implying that stabilization of EpFAs is a promising strategy to treat inflammatory disorders. A potential therapeutic application for EpFAs or sEHI is for alleviation of inflammatory and neuropathic pain. The use of sEHI and EpFAs for therapy of pain has been reviewed by Inceoglu et al. (2007) and Wagner et al. (2011). The basis for testing sEHIs stemmed from observations that sEHIs reduce inflammation (Schmelzer et al., 2005). Observation that direct administration of EETs in absence of sEHI produced the same effect and the structural diversity of sEHI tested indicates that EpFAs are the primary regulators of pain relief (Inceoglu et al., 2006). Other EpFAs including metabolites of DHA and EPA have similar effect on reducing inflammatory reaction; however, the effect is less potent for EPA metabolites (Morisseau et al., 2010). The EDPs have been shown to be the most potent EpFA vasodilators (Ye et al., 2002). EDP regioisomers (except 5,6EDP, which is chemically unstable) were all active, while the corresponding diol 13,14-DiHDPA was .1000-fold less active. Animal experiments supported the antihypertensive effects of ω-3 EpFAs. Stabilized 19,20 EDPs also suppressed hypertension, suggesting that the antihypertensive effect was at least partially mediated by the formation of EDPs (Ulu et al., 2013). In carrageenan-induced inflammatory pain model in rats, EDPs and EETs had similar efficacy on reduction of pain, while the effects of EEQs were less evident (Morisseau et al., 2010). The parent fatty acids and the corresponding fatty acid diols lacked such effects. The effects of EDPs on pain were regio-specific. 13,14-EDP was the most potent regioisomer, followed with 16,17- and 19,20-EDP. These studies demonstrated that similar to EETs, the ω-3 EpFAs also have potent effects to reduce inflammation and pain. In vivo studies of EETs on inflammation provide strong evidence to support the antiinflammatory effects of EETs in various inflammatory models, suggesting that inhibiting sEH to stabilize EETs is a promising therapeutic strategy to treat inflammatory disorders, although inconsistencies exist (Davis et al., 2011; Fife et al., 2008). The antiinflammatory effects of EETs are regioselective: 11,12-EET showing the most potent effect, followed with 8,9- and 5,6-EET, while 14,15-EET was inactive (Zhang and Blair, 1994, and references cited therein). A recent study indicated that 14,15-EET inhibited TNFα-stimulated inflammation in human bronchi, suggesting this EET regioisomer is also biologically active to suppress inflammation in some systems (Morin et al., 2008). Other studies (Kundu et al., 2013) indicate that pharmacological inhibition of sEH not only stabilizes but increases level of antiinflammatory EETs. Serhan and Petassis (2011) and Serhan et al. (2015) have discussed the role of resolvins and protectins in inflammation resolution, while Serhan

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et al. (2015) have described new proresolving families of mediators in acute inflammation and resolution bioactive metabolome. Miyata and Arita (2015) have pointed out that several reports have indicated that biosynthesis of antiinflammatory and proresolving lipid mediators is dysregulated in severe asthma, suggesting that an imbalance between pro- and antiinflammatory molecules causes exacerbation of inflammation observed in asthmatic patients.

3.7.3 Angiogenesis and Cardiovascular Disease Imig (2006) has pointed out that sEH has become a potential therapeutic target in the management of cardiovascular disease due to biological effects of its epoxide substrates on vasodilation and inflammation. In contrast, Enayetallah et al. (2012) have suggested that inhibition of EH domain could abolish its cholesterol lowering effect thus resulting in an undesirable effect in cardiovascular disease. As opposed to EETs, ω-3-PUFA epoxides suppress angiogenesis, 19,20EpDPE, and other EpDPE regioisomers decreased vascular endothelial growth factor (VEGF)induced angiogenesis in mice and suppressed fibroblast growth factor 2induced migration and protease production in human umbilical vein endothelial cells (Spector and Kim, 2015). Ulu et al. (2013) demonstrated that a diet rich in ω-3 PUFAs (EPA and DHA) lowers systolic BP in angiotensin-IIdependent hypertension when compared with animals on a diet rich in ω-6 FUFAs. This reduction in BP was enhanced by treatment with a sEH inhibitor (TPPU, 1-trifluoromethoxyphenyl-3-[1-propionyl-4-yl] urea). Among the regioisomers of DHA, epoxides that were investigated increased tissue levels of 19,20-EDP correlated well with the reduction in BP. Ulu et al. (2014) have also shown that treatment with 19,20-EDP and an sEH inhibitor had a larger effect as compared to either treatment alone. The effect of 19,20-EDP and TPPU was more efficacious than the combination of 14,15-EET and TPPU. The CYP epoxygenases and the metabolites the EETs generate, clearly have cardiovascular protective effects, but the findings of Panigrahy et al. (2012) indicate that EETs also promote tumor growth and metastasis under the same conditions. EDPs derived by CYP from the ω-3 fatty acid DHA inhibit VEGF and fibroblast growth factor 2induced angiogenesis in vivo, and suppress endothelial cell migration and protease production in vitro via the VEGF receptor 2dependent mechanism (Zhang et al., 2013). Wang et al. (2014) have shown that CYP2J2-derived EETs suppress ER stress response in the heart and protect against cardiac failure by maintaining intracellular Ca21 homeostasis and sarcoplasmic/endoplasmic reticulum calcium ATPase expression and activity.

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3.7.4 Cancer Little has been known about the role of epoxyeicosanoids in cancer (Panigrahy et al., 2010a,b). Using genetic and pharmacological manipulation of endogenous EET levels, Panigrahy et al. (2012) have demonstrated that EETs are important in primary tumor growth and metastasis in various mouse models of cancer. Using genetically altered mice that exhibit high endothelial EET levels, a dramatic increase was seen in the growth of B16F10 melanoma, T241 fibrosarcoma, and Lewis lung carcinoma in Tie2CYP2c8-Tr, Tie2-CYP2J2-Tr, and sEH-null mice compared with wild mice, suggesting that endothelial EET promotes primary tumor growth. Furthermore, plasma 11,12- and 14,15-EET levels were elevated 15-fold in sEH-null tumor-bearing mice when measured on Day 22 after injection of T241 fibrosarcoma cells. Furthermore, using a model in which resection of a primary tumor stimulated development of distant metastases 1417 days after resection, Panigrahy et al. (2012) showed that EETs promote spontaneous metastatic growth (as opposed to inducing metastases by intravenous injection of tumor cells). Importantly, systemic administration of 14,15-EET via osmotic minipumps in wild-type mice at the time of liquid-liquid chromatography (LLC) tumor resection stimulated threefold increase in the number of surface lung metastases compared with vehicle-treated controls and led to development of liver, kidney, and distant lymph node metastasis 12 days after resection of the primary LLC tumor. Panigrahy et al. (2012, 2013) have shown that elevated EETs trigger massive metastatic spread and escape from tumor dormancy in several tumor models (Ding et al., 2014). Zhang et al. (2013) have shown that the corresponding EDTs produced by CYP epoxyoxygenases from DHA (4,5-, 7,8-, 10,11-, 13,14-, 16,17-, and 19,20-EDP) have an effect opposite to that of EETs on tumor growth and metastasis. Zhang et al. (2014) have proposed to use pharmacological inhibitors of sEH to stabilize endogenous EpFAs. When EPDs are administered with a low dose of sEH inhibitor, EDPs are stabilized in circulation, causing approximately 70% inhibition of primary tumor growth and metastasis, but the mechanism by which these ω-3-lipids inhibit angiogenesis and tumorigenesis is poorly understood. A widely accepted theory to explain the health promoting effects of ω -3 fatty acids is that they suppress the metabolism of ω-6 ARA to form proangiogenic proinflammatory eicosanoids or serve as alternative substrates to generate ω-3 lipid mediators with beneficial actions (Rose and Connelly, 1999). Since fatty acid epoxides are highly unstable in vivo (Catella et al., 1990), presumably due to sEH abundantly expressed in numerous tissues for which EDPs are highly efficient substrates (Morisseau et al., 2010), coadministration of a low dose of sEHI was required to stabilize EDPs in circulation, leading to a dramatic inhibition of tumor growth and metastases. Xiao

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and Guengerich (2012) have characterized the orphan human CYP 2W1 in selective epoxidation of lysophospholipids in human colorectal cancer (Tang et al., 2009). Zhang et al. (2014) have followed up their earlier reports (Panigrahy et al., 2012; Zhang et al., 2013) with an extensive review of pharmacological inhibitors that stabilize endogenous EpFAs in view of the consideration of these inhibitors for human clinical uses. Zhang et al. (2014) concluded that the biological effects of sEHIs (or EETs) on tumorigenesis have a high threshold. This raises the question if the therapeutic index of sEHI is sufficiently high to justify their long-term use as pharmaceuticals. According to Zhang et al. (2014), the variable expression pattern of CYP epoxygenases and sEH makes it difficult to investigate the significance of CYP/sEH pathway in tumorigenesis.

3.8 CONCLUSION Since only a few plants produce significant amounts of vernolic and related epoxy fatty acids, industrial extraction and production of epoxidized acids are not practical. Therefore, epoxy fatty acids are currently produced by chemical oxidation of unsaturated plant oils. Except for increased safety of operation, the methodology of chemical epoxidation of fatty acids has remained largely as originally developed. In contrast, the biochemistry and physiology of the epoxy acids have greatly advanced. The epoxy acids have become recognized as lipid mediators, although the C18 series of epoxides has been less extensively investigated. The C20 series of epoxides regulate inflammation and vascular tone, with high endothelial levels promoting primary tumor growth. The corresponding C22 series of epoxides has the opposite effect. A large-scale production of fatty acid epoxides and their incorporation into various industrial and household products raises concern about biological safety of epoxy fatty acids and their derivatives. In any event, the physiological effects need to be further investigated, especially if pharmacological inhibitors are to be used as stabilizers of EH.

ABBREVIATIONS ARA arachidonic acid CID collision-induced dissociation CNS central nervous system COX cyclooxygenase CYP cytochrome P450 DHA docosahexaenoic acid DiHDPE dihydroxydocosapentaenoic acid DiHET dihydroxyeicosatrienoic acid DiHETE dihydroxyeicosatetraenoic acid DiHETrEs dihydroxyeicosatrienoic acids

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DiHOME dihydroxyoctadecenoic acid EDP epoxydocosapentaenoic acid EET epoxyeicosatrienoic acid EPA eicosapentaenoic acid EpDPE epoxyeicosapentaenoic acid EpETE epoxyeicosatetraenoic acid EpOME epoxyoctadecenoic acid EpSTA epoxyoctadecanoic acid HEpOME hydroepoxyoctadecanoic acid HEpSTA hydroxyepoxyoctadecanoic acid HETE hydroxyeicosatetraenoic acid HODE hydroxyoctadecadienoic acid LOX lipoxygenase sHE soluble epoxide hydrolase sEHI soluble epoxide hydrolase inhibitor TNF-α tumor necrosis factor-α

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Chapter 4

Acetylenic Epoxy Fatty Acids: Chemistry, Synthesis, and Their Pharmaceutical Applications Valery M. Dembitsky1 and Dmitry V. Kuklev2 1

National Scientific Center of Marine Biology, Vladivostok, Russia, 2University of Michigan Medical School, Ann Arbor, MI, United States

Chapter Outline 4.1 Introduction 121 4.2 Occurrence Epoxy Acetylenic Fatty Acids in Nature 122 4.3 Lipids Containing Epoxy Acetylenic Fatty Acids 125 4.4 Epoxy Acetylenic Furanoid and Thiophene Fatty Acid and Derivatives 128 4.5 Pyranone and Macrocyclic Epoxides 129

4.6 Acetylenic Cyclohexanoid Epoxy Fatty Acids 130 4.7 Determination or Epoxy Acetylenic Lipids 131 4.8 Synthesis of Epoxy Acetylenic Lipids 136 4.9 Concluding Remarks 141 References 142 Further Reading 146

4.1 INTRODUCTION Natural acetylenic epoxides and related compounds display important biological activities, including antitumor, antibacterial, antimicrobial, antifungal, phototoxic, and other chemical and medicinal properties (Dembitsky, 2006; Dembitsky and Levitsky, 2006; Dembitsky et al., 2006; Minto and Blacklock, 2008; Carballeira, 2008; Bador and Paris, 1990; Siddiqi and Dembitsky, 2008). Compounds with acetylene, vinylacetylene, and acetylene-allenetype systems of bond(s) were first found in the late 19th century in some mushrooms. Because molecules containing these fragments are most often unstable, their presence in natural objects appeared unusual. However, as experimental findings have been accumulated, it turned out that compounds of this type are characteristic of natural life, and are widely

Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00011-8 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

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O

O

O

O

R1

R1

n

n R

R

R1

n = 1 or 2

O

n = 1, 2, 3, 4, 5 R

FIGURE 4.1 Graphical display of chemical structures of natural acetylenic oxiranes: R, R1 5 H, alkyl, phenolic, heterocyclic, etc.

represented and perform important functions—in particular: act as antibiotics, anticancer, antibacterial, and other agents. Acetylenic epoxides, and related liphophilic metabolites that contain the [CC] bond(s) and ethylene oxide group (ethylene oxide, also called oxirane), are rare in nature (Siddiqi and Dembitsky, 2008; Kuklev et al., 2013). Graphic chemical structures are shown in Fig. 4.1. Intensive chemical and pharmacological studies during the last five decades have led in many cases to validation of traditional claims and facilitated identification of the traditional medicinal plants and of their active principles (Minto and Blacklock, 2008; Siddiqi and Dembitsky, 2008). More than 1000 acetylenic metabolites have been isolated and identified from plant and animal species (Dembitsky, 2006; Dembitsky and Levitsky, 2006; Dembitsky et al., 2006; Siddiqi and Dembitsky, 2008; Kuklev et al., 2013; Christensen, 1992; Christensen and Jakobsen, 2008; Christensen and Lam, 1990, 1991a,b; Pan et al., 2009). Thousands of plant and marine acetylenic oxiranes are being screened worldwide to validate their use as anticancer drugs, but terrestrial acetylenic compounds comprise an especially interesting group of anticancer agents and other biologically active compounds (Dembitsky, 2006; Dembitsky and Levitsky, 2006; Dembitsky et al., 2006; Minto and Blacklock, 2008; Siddiqi and Dembitsky, 2008). This chapter is the first article devoted to natural acetylenic oxiranes. It focuses on origin, structures, and biological activities of natural acetylenic oxiranes and selected semi- and/or synthetic-related compounds. Their structure and biological activities, modes of action, and future prospects are discussed.

4.2 OCCURRENCE EPOXY ACETYLENIC FATTY ACIDS IN NATURE The aerial parts of Erigeron philadelphicus afforded the isomeric acetylenic epoxides, (Z)-methyl 8-((2S,3R)-3-methyloxiran-2-yl)octa-2-en-5,7-diynoate (1) and (Z)-methyl 8-((2R,3R)-3-methyloxiran-2-yl)octa-2-en-5,7-diynoate (2) (Jakupovic et al., 1986). The same fatty acids (FAs) (1 and 2) were detected along polyacetylenes of Chrysoma pauciflosculosa (Menelaou et al.,

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1992). Two derivatives of matricaria esters, (Z)-methyl 8-(3-(hydroxymethyl) oxiran-2-yl)octa-2-en-5,7-diynoate (3) and (Z)-methyl 8-(3-(acetoxymethyl) oxiran-2-yl)octa-2-en-5,7-diynoate (4), have been detected in extract of the rabbitbrush Chrysothamnus nauseosus (Rose et al., 1980a), and they were found to inhibit feeding of third-instar Colarado potato beetle larvae. The (Z)-methyl 8-(3-(acetoxymethyl)-oxiran-2-yl)octa-2-en-5,7-diynoate (4) was isolated from Chrysothamnus parryi (Bohlmann et al., 1979). The seed oil of Crepis foetida (family Compositae) contains 60% of a FA, which has been identified as 8-((2S,3R)-3-(hept-2-yn-1-yl)oxiran-2-yl) octanoic acid (also as crepenynic acid, 5) (Mikolajczak et al., 1964). Acetylenic acid, 4-(3-(trideca-2,4-diyn-1-yl)oxiran-2-yl) butanoic acid (6) and methyl ester, methyl 4-(3-(trideca-2,4-diyn-1-yl)oxiran-2-yl) butanoate (7) as inhibitors of 3-hydroxy-3-methylglutaryl coenzyme A reductase were found in the root bark of Paramacrolobium caeruleum (Patil et al., 1989). Two acetylenic acid, methyl esters, (Z)-methyl 8-(3-(acetoxymethyl)oxiran-2-yl)octa-2-en4,6-diynoate (8) and (Z)-methyl 8-(3-(hydroxyl-methyl)oxiran-2-yl) octa-2-en-4,6-diynoate (9) as antifeedants, were isolated from rabbitbrush C. nauseosus (Asteraceae). Both metabolites inhibited feeding of third-instar Colarado potato beetle larvae (Leptinotarsa decemlineata) (Rose et al., 1980b). O

MeO

HO

O

1

MeO O MeO

OH

3

O

6

O

O

7

O

O

O

O O

5

MeO

O 2

O HO

O

OAc

MeO

O

OAc 8

MeO

O O

4

MeO

O 9

OH

Acetylenic acid, 4-((2S,3S)-3-(pent-1-yn-1-yl)oxiran-2-yl) butanoic acid (10), was prepared as an inhibitor of human neutrophil LTA4 hydrolase (Evans et al., 1986). An unusual acetylenic amide, (E)-3-(hexa-3,5-diyn-1-yl)-N-styryloxirane2-carboxamide (11), has been isolated and its structure elucidated from extract of Spilanthes alba (Bohlmann et al., 1980). Acetylenic N-alkylamide, (2R,3R)-3(hexa-3,5-diyn-1-yl)-N-phenethyloxirane-2-carboxamide (12), a compound having evidence of immune stimulating properties, was isolated from extract of Spilanthes acmella (Boonen et al., 2010), in Spilanthes acmella flowers (Nagashima and Nakatani, 1992), in the roots of Acmella ciliata (Martin and Becker, 1985), and in the aerial parts of Salmea scandens (Bohlmann et al., 1985). (2S,3S)-3-(hexa-3,5-diyn-1-yl)-N-phenethyloxirane-2-carboxamide (13) was

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Fatty Acids

isolated from the leaves and the flower heads of Acmella radicans var. radicans (Asteraceae) (Rios-Chavez et al., 2003). The unusual (S,E)-methyl 11-((2S,3S)-3-(6-bromohex-5-yn-1-yl)oxiran-2yl)-9-hydroxyundec-10-enoate (14) was isolated from N-fixing lichen Leptogium saturninum and Peltigera canina (Rezanka and Dembitsky, 1999). Leptogium saturninum (order Peltigerales) displayed strong activity of multicopper oxidases (e.g., tyrosinase) as well as heme-containing peroxidases (Liers et al., 2011; Dembitsky, 2003). Peltigera sp., a cyanolichencontaining Nostoc as cyanobiont, produced arginase and arginine (Diaz et al., 2009; Dembitsky and Rezanka, 2005), also produced of phycobiliprotein pigments (Czeczuga et al., 2011), and displayed laccase activity (Laufer et al., 2006). Both lichen species contain unusual lipids and FAs (Dembitsky, 1992, 1996; Dembitsky et al., 1991). The succinate ester of panaxydol, 4-((8-((2R,3S)-3-heptyloxiran-2-yl)octa-1en-4,6-diyn-3-yl)oxy)-4-oxobutanoic acid (15), was obtained from Panax ginseng and esterification with succinic anhydride (Hirakura et al., 1992). Compound (15) and methyl succinate (16) showed IC50 values of 0.06 and 12.7 μg mL21 for inhibiting the proliferation of L2110 and Hela cells, respectively. Panaxydol linoleate, (9Z,12Z)-8-((2S,3R)-3-heptyloxiran-2-yl)octa-1-en-4, 6-diyn-3-yl octadeca-9,12-dienoate (17) and ginsenoyne A linoleate, (9Z,12Z)8-((2S,3R)-3-(hex-5-en-1-yl)oxiran-2-yl)octa-1-en-4,6-diyn-3-yl octadeca-9,12dienoate (18) were found in extract of the root of P. ginseng and they showed cytotoxic activities against murine and human malignant cells (DT, NIH/3T3, L-1210, HeLa, T24, and MCF7 cells) in vitro (Hirakura et al., 2000). OH HO O

10

H

O

H N

14

Br O

OMe

11 O

H N

H

O

O

O

H

H O

O

O

H N

O

12

H

OH O

H

H O

O 15

O

O

13

O

H

OMe O

O 17 O O O 18 O O

16

H

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125

The myxomycetes (plasmodial slime molds) are a group of fungus-like organisms usually present and sometimes abundant in terrestrial ecosystems. The myxomycete life cycle involves two very different trophic (feeding) stages, one consisting of uninucleate amoebae with or without flagella and the other consisting of a distinctive multinucleate structure, the plasmodium (Speijer, 2008). Their chemical constituents—more than 100 natural compounds from 26 species of 4 orders—from myxomycetes were reported in some review articles (Dembitsky et al., 2005; Ishibashi, 2005; Ishibashi and Arai, 2012). The slime mold Lycogala epidendrum, commonly known as wolf’s milk or groening’s slime, is a cosmopolitan species. Recently, some interesting lipids were isolated from this myxomycete. Specifically, rare FAs are 7((2S,3S)-3-((4Z,6Z)-nona-4,6-dien-1-yl)oxiran-2-yl)hepta-4,6-diynoic acid (19a), methyl 7-((2S,3S)-3-((4Z,6Z)-nona-4,6-dien-1-yl)oxiran-2-yl)hepta4,6-diynoate (19b), 7-((2S,3S)-3-(non-8-en-1-yl)oxiran-2-yl)hepta-4,6diynoic acid (20), and 7-((2S,3S)-3-(oct-7-yn-1-yl)oxiran-2-yl)hepta-4,6diynoic acid (21). O HO O

19a

MeO O

O

H

H

O

H

19b

HO O

H

20

O

HO

H

21

O

4.3 LIPIDS CONTAINING EPOXY ACETYLENIC FATTY ACIDS Three triacylglycerols, named lycogarides A (22), B (23), and C (24), have been isolated from the myxomycete L. epidendrum (Hashimoto et al., 1990, 1994). More recently, two unusual triacylglycerols, lycogarides D (25) and E (26), and two diacylglycerols, lycogarides F (27) and G (28), along with the

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Fatty Acids

known lycogalic acid dimethyl esters A and B were reported (Buchanan et al., 1996). Discoveries of di- (28 and 29) and triglycerides (2227) containing acetylenic epoxy FAs are uncommon in nature. Some other examples are described in the scientific literature. Gunstone and Sealy (1965) have reported that the FAs of the seed oil of the tree Ongokea gore, which is also known as isano or boleko oil, contain several acetylenic acids. The triacylglycerides (TAG) of boleko oil contain the following FAs: 6% saturated (C14, C16, C18); 19% oleic and linoleic; 51% of five acetylenic acids (17-octadecene-9, 11-diynoic, 13-octadecene-9,11-diynoic, 11-octadecen-9-ynoic, 9,12-octadecadiynoic, and 13,17-octadecadiene-9,11-diynoic acids), 22% of four hydroxy derivatives of acetylenic acids (8-hydroxy-17-octadecene-9,11-diynoic, 8-hydroxy-13,17-octadecadiene-9,11-diynoic, 8-hydroxy-13-octadecene-9,11diynoic, and 8-hydroxy-9,12-octadecadiynoic acids), and 2% of a dihydroxystearic acid. Two acetylenic FAs (20-heneicosen-6-ynoic and 18-nonadecen-4-ynoic acids) and a triglyceride containing these acetylenic acids have been isolated from the leaves of Hymenodictyon excelsum, family Rubiaceae (Nareeboon et al., 2009). Extract of H. excelsum showed anticoagulant, antiinflammatory, and sunscreening effects (Jagdishprasad and Rao, 1998). Acetylenic FAs in TAG varied from 6.6% in the moss Calliergon cordifolium to 80.2% in the liverwort Riccia antipyretica. Three acetylenic acids were identified among the monoenoics (6a-18:1, 9a-18:1, and 12a-18:1) and dienoics (6a,9c-18:2, 9a,12c-18:2, and 9c,12a-18:2). Four acetylenic acids were identified among the polyenoics 6a,9c,12c-18:3, 8a,11c, 14c-20:3, 6a,9c,12c,15c-18:4, and 5a,8c,11c,14c-20:4 (Dembitsky and Rezanka, 1995). Some usual conjugated ene-yne acetylenic FA, trans-10-heptadecen-8ynoic (pyrulic, 7.4%) acid, trans-11-octadecen-9-ynoic (ximenynic, 3.5%) acid, cis-7,trans-11-octadecadiene-9-ynoic (heisteric, 22.6%) acid, and 9,10epoxystearic acid, could be identified in the seed oil of Heisteria silvanii (Olacaceae). Two further conjugated acetylenic FA, 9,11-octadecadiynoic (0.1%) and 13-octadecene-9,11-diynoic (0.4%) acids, were identified tentatively by their mass spectra. Twenty six species of the separated TAG were identified by means of their abundant quasi-molecular ion [M 2 H]2 and their corresponding carboxylate anions [RCOO]2 of the FAs, respectively. The major molecular species of the TAG were found to be 16:0/18:1/18:1, 16:0/18:1/18:3 (heisteric acid), 17:2 (pyrulic acid)/18:1/18:1, and 18:1/18:1/ 18:3 (heisteric acid). The TAG containing acetylenic FA also was found (Spitzer et al., 1997).

127

Acetylenic Epoxy Fatty Acids Chapter | 4 O O O

5

O

O

O

O

O

3

4

22 Lycogaride A

O O

O

5

O

O

O

O

O

12

4

23 Lycogaride B

O

O

O O

5

O

O

O O

O

4

O

H

24 Lycogaride C

The triacylglycerols of type AAA, ABA, ABA, AAB, and AAB (containing positional isomers of acetylenic FA) were prepared and their 1H and 13C NMR spectroscopic properties were studied (Gehrt et al., 1998; Bellina et al., 2004). Gehrt et al. (1998), using the lipase from Candida cylindracea and Candida rugosa, were able to catalyze the release of 10-undecynoic acid and 9-octadecynoic acid from the corresponding TG, but less readily the 13-docosynoic acid in the case of glycerol tri-(13-docosynoate).

128

Fatty Acids O O O

Et

O

25 Lycogaride D

O

O O

Et O O

Et O O O

R O

O

26 Lycogaride E, R = saturated FA 27 R = unsaturated FA

O

Et

O

O

Et O O

Et

O

28 Lycogaride F

OH

O O

O

Et O

O O O

Et O

O OH

Et 29 Lycogaride G

4.4 EPOXY ACETYLENIC FURANOID AND THIOPHENE FATTY ACID AND DERIVATIVES Two antifungal acetylenic epoxides, wyerone epoxide (30) and wyerol epoxide (31), were identified in Vicia faba (Hargreaves et al., 1976). Wyerone epoxide (30) accumulated in limited lesions formed by both Botrytis cinerea and Botrytis fabae. Products of the metabolism of (31) by B. cinerea and B. fabae were identified as wyerol epoxide (31) and dihydrodihydroxy-wyerol, respectively. Two acetylenic antibiotics, cepacin A (32) and B (33), have been isolated from the fermentation broth of Pseudomonas cepacia SC 11,783 (Parker et al., 1984). Cepacin A has good activity against Staphylococci (MIC

129

Acetylenic Epoxy Fatty Acids Chapter | 4

0.2 μg mL21), but weak activity against Streptococci (MIC 50 μg mL21) and the majority of Gram-negative organisms (MIC values 6.3 approximately greater than 50 μg mL21). Cepacin B (33) has excellent activity against staphylococci (MIC less than 0.05 μg mL21) and some Gram-negative organisms (MIC values 0.1 approximately greater than 50 μg mL21). Aporpinone B (34) and 10 -acetylaporpinone B (35) with an unusual skeleton containing an acetylene unit were isolated from the culture of the wood inhabiting fungus Aporpium caryae (Basidiomycete). Both acetylenic oxiranes (34, 35) showed weak to moderate antibacterial activity against Bacillus subtilis, Staphylococcus aureus, and Escherichia coli (Levy et al., 2003). Foeniculacin (aromatic acetylenic epoxide, 36) was detected in stem extract of endemic to the Canary Islands, Argyranthemum foeniculaceum (Gonzalez et al., 1987). Extracts from Argyranthemum adauctum, A. foeniculaceum, and Argyranthemum frutescens showed antimicrobial activity against Gram-positive and Gram-negative bacteria and cytotoxic activity against HeLa and Hep-2 cell lines (Gonzalez et al., 1997). O O

HO

OH

O

O

O . O

32 Cepacin A 30 Wyerone epoxide

Et

O

O O

HO

H

O O

OH

O . OH 33 Cepacin B

31 Wyerol epoxide O

O

OH 34 Aporpinone B

H

Et

O

O

H

O

O

O

O

H

O OAc

S

MeO

36

35 1'-Ac Aporpinone B O

OMe

4.5 PYRANONE AND MACROCYCLIC EPOXIDES Bioactive acetylenic oxiranes (3740) have been isolated from leaves or roots of some plants, fungi, and lichens. Nitidon (37), a highly oxidized pyranone derivative produced by the corticoid fungus Junghuhnia nitida (Meruliaceae), was isolated and its several biological activities were

130

Fatty Acids

evaluated. Compound (38) exhibited antibiotic and cytotoxic activities and induced morphological and physiological differentiation of tumor cells at nanomolar concentrations (Gehrt et al., 1998). The first total synthesis of naturally occurring (2)-nitidon (37) and its enantiomer (38) was reported. Both enantiomers of nitidon have been found to exhibit significant cytotoxic activity against human cancer cell lines in vitro (Bellina et al., 2004). Junghuhnia nitida is a fungus that breaks down wood deciduous trunks by a white rot (Westphalen et al., 2011). Ivorenolide A (39), a novel 18-membered macrolide featuring conjugated acetylenic bonds and five chiral centers, was isolated from Khaya ivorensis. Aqueous extracts from the K. ivorensis stem-bark of the showed antiplasmodial activity. Both compound (40) and its synthetic enantiomer (40) showed potent and selective immunosuppressive activity (Zhang et al., 2012). HO

O

O

OH

O H

37 trans-(–)-Nitidon

O

O O

OH

H

39 Ivorenolide A HO

O

OH

O H

38 cis-(+)-Nitidon

O

O OH

O

O

H 40 Enantiomer

4.6 ACETYLENIC CYCLOHEXANOID EPOXY FATTY ACIDS Tricholomenyns C (41) and D (42), which found in extract of the fruiting bodies of T. acerbum and other species of the genus Tricholoma, are the first naturally occurring dimeric dienyne geranyl cyclohexenones (Garlaschelli et al., 1996). Tricholoma acerbum is a fairly large genus of mycorrhizal-gilled mushrooms. The tricholomenyns efficiently inhibit mitosis of T-lymphocyte cultures and are potent as anticancer agents.

Acetylenic Epoxy Fatty Acids Chapter | 4

131

OAc

O O

O COOH

O

O

41 HO

OAc

OAc O O OH

O COOH

O 42 HO OAc

4.7 DETERMINATION OR EPOXY ACETYLENIC LIPIDS The determination of epoxy acetylenic lipids is a complex problem that is solved by using a combination of modern analytical methods. The most important methods for determination of triple bonds are carbon-13 NMR and FT-IR (Fourier transform infrared spectroscopy). In 13 C NMR spectra, acetylenic-bonded carbon atoms appear as singlets with a chemical shift of 6080 ppm. In FT-IR spectra, triple bonds appear as a weak signal around 21202250 cm21, and if the triple bond is terminal, the terminal proton signal presents at 3320 cm21 (stretch) and 660 cm21 (bend). The position of the triple bond within the molecule is usually determined by combination of the results of mass spectrometry, 1H and 13C NMR and, sometimes, by UV spectroscopy if triple bond(s) is a part of a chromophore. In proton-NMR spectra, triple bonds create hydrogen-free zones, and if the triple bond is conjugated with double bond(s), it creates a chromophore with a specific absorbance in the UV spectrum. Diagnostic peaks resulting from mass-spectrometric fragmentation of these alkyne molecules help in deducing the specific positions of triple bonds in the molecule. Epoxide groups are primarily determined by 13C NMR and mass spectrometry (sometimes, by mass spectrometry of derivatives). In 1H NMR

132

Fatty Acids

spectra, protons in an epoxy group produce signals at chemical shifts of 23 ppm, with the usual coupling constants of B4 Hz for cis-epoxides and 2 Hz for trans ones. In 13C NMR spectra, carbon atoms of epoxy groups produce signals at chemical shifts of 5060 ppm. The usual method in epoxide mass spectrometry is to derivatize the group by opening it using trimethylsilyl chloride, acetyl chloride, or similar compounds in order to increase the intensity of characteristic fragments pointing on the position of an epoxy ring in the given molecular structure. For example, in determining the structure of an unusual sesquiterpenoid (43) from the green alga Caulerpa prolifera (Amico et al., 1978), the presence of a triple bond in its structure was confirmed by the presence of a weak stretch band at 2200 cm21 in the IR spectrum, and by 13C NMR demonstrating two lines characteristic of a nonterminal triple bond as singlets at 93.8 and 84.9 ppm. In the structure of 9,12,15-octadiene-6-ynoic acid (44), an acetylenic acid obtained from mosses (Anderson et al., 1974), the presence of a triple bond can be confirmed by a small peak at 1335 cm21, while the absence of absorption at 2150 cm21 indicated that unsaturated triple bonds would be neither terminal nor near the carboxyl group. In addition, the Raman spectrum of the ester showed absorption at 2250 cm21, which is characteristic of triple bonds in such molecules. Its methyl ester was also shown to have a triple bond at the sixth position by mass spectrometry. The presence of alkyne groups in the structure of FAs (45) isolated from the leaves of H. excelsum (Nareeboon et al., 2009) has been proven by the presence of signals for two quaternary carbons associated with alkyne functionality at 81.3 and 77.9 ppm, and for the vinyl ABX, 1H NMR signals at 5.79 and 4.974.91 ppm. The 1H1H COSY spectrum located the position of the triple bond in the carbon chain. Additional confirmation of the position of the alkyne group came from analysis of massspectroscopic fragmentation—this confirmed the presence of two acetylenic FAs with delta-4 and delta-6 positions of a triple bond. The authors describe the IR peaks, but do not propose any structurefunction relationship. In the study of molecular structures of cytotoxic components (46, 47) of American ginseng (Panax quinquefolius) (Fujimoto et al., 1991), the triple bonds were determined by the presence of signals with chemical shift of 7078 ppm in 13C NMR, and the presence of an epoxide group was determined by signals with shift of 3.153.06 ppm, with a 4.4 Hz (cis-stereoisomer) constant in 1H NMR spectrum.

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133

In the structure of the epoxy acetylenic carotenoid halocynthiaxanthin (48) (Konishi et al., 2006) found in the sea squirt Halocynthia roretzi, the triple bond at the 70 -80 position is determined by the presence of signals at 89.2 (70 ) and 98.4 (80 ) ppm in the 13C NMR spectrum, and the epoxide group by signals at 66.2 (5) and 67.0 (6) ppm. Angelicol B, (Z)-2-(3-hydroxypent-1-ynyl)-3-(non-1-enyl)oxiran-2-ol (49), a compound isolated from Angelica keiskei (Luo et al., 2012), produced a characteristic acetylenic signal at 2134 cm21 in the IR spectrum, and signals associated with the presence of two acetylenic carbons at 80.1 (C) ppm and 68.5 (CH) in the 13C NMR spectra. Its selective UV absorption with maxima at 258 and 284 nm points to conjugation of the groups described earlier. In the structures of six acetylenic compounds, which were isolated from the leaves of Artemisia lactiflora (Compositae), an edible plant of Thailand (Nakamura et al., 1999), the presence of two triple bonds conjugated with one double bond was proven by the specific absorption in the UV spectrum with maxima at 225, 265, 278, and 293 nm. For example, in one of the compounds, authors reported a signal at 2140 cm21 in the IR spectrum and six singlets in 13 C NMR spectra at 68.9, 80.1, 80.8, 82.4, 85.4, and 166.0 ppm, which identified its structure as (2R,E)-4-(hexa-2,4-diyn-1-ylidene)tetrahydro-3,6-dioxaspiro [bicyclo[3.1.0]hexane-2,20 -pyran] (50), structure 4 in the cited paper.

134

Fatty Acids

In the structures of the polycyclic epoxy acetylenic antibiotics deoxydynemicin A and dynemicin A1 (51), found in the culture broth of a strain of Actinomycetes (Shiomi et al., 1990), two triple bonds in combination with a double bond forming an a,d,a-structure have been identified using 13C NMR by the presence of singlet peaks with chemical shifts of 99.389.5 and 88.898.0 ppm in deoxydynemicin A (at atoms 2324 and 2728, respectively), and shifts of 99.090.5 and 90.798.2 ppm in dynemicin A1 (at atoms 2324 and 2728, respectively). The epoxide group was identified by the presence of singlet peaks at 63.3 and 70.2 ppm in the case of deoxydynemicin A (at atoms 522, respectively) and 64.2 and 72.7 ppm in dynemicin A1 (at atoms 522, respectively). A new polyacetylenic antibiotic, oploxyne A (52), was isolated from the stem of Oplopanax elatus. The structure of the compound was determined to be 9,10-epoxyheptadeca-4,6-diyne-3,8-diol, on the basis of its UV, MS, and NMR data (Yang et al., 2010). Two conjugated double bonds were revealed by the signals in 13C NMR at 81.0, 68.7, 70.3, and 77.4 ppm, and selective absorption in the UV spectrum at 243 nm. In the IR spectrum, two bands of absorption at 2253 and 2146 cm21 were ascribed as selective absorption by triple bonds. The epoxide group was identified on the basis of 1H NMR signals at 3.16 and 3.07 ppm with a coupling constant J 5 4 Hz (cis-epoxy group)—as well as the presence of two singlets in the 13C NMR spectrum at 58.058.1 ppm.

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135

Gymnasterkoreayne B is a polyacetylenic compound (53) isolated from the roots of Gymnaster koraiensis, which exhibits significant cytotoxicity against L-1210 tumor cells with an ED50 value of 3.3 μg mL21 (Jung et al., 2002). The structure of the oily compound was established spectroscopically, including 2D NMR experiments. Two conjugated triple bonds absorbed UV with a maximum at 250 nm, and featured a diagnostic absorption band in the IR spectrum at 2154 cm21. Four carbons in the conjugated system of two triple bonds were determined by chemical shifts of four singlets at 78.7, 67.5, 68.8, and 76.6 in 13C NMR. The epoxide group was determined by 13C NMR by the presence of two signals at 56.9 and 46.1, and by two complimentary multiplets in 1H NMR at 3.13 and 3.19 ppm. Gummiferol is a new cytotoxic acetylenic diepoxide compound (54) isolated from the leaves of Adenia gummifera (Fullas et al., 1995). Its three conjugated triple bonds absorbed UV with maxima at 287 nm. However, in the IR spectrum, no absorption at 2140 cm21 was reported; four signals of the protons of two epoxide groups presented at 3.46, 3.39, 3.35, and 3.04 ppm with coupling constants J 5 2.0 Hz (trans-orientation of oxirane rings), six acetylenic carbons making three conjugated triple bonds were determined by signals at 77.2, 70.1, 62.4, 62.8, 69.0, and 73.9 ppm in the 13C NMR spectrum. Ivorenolide A (39) (Zhang et al., 2012) is a novel 18-membered macrolide featuring two conjugated triple bonds, one oxirane ring, and five chiral centers, was isolated from K. ivorensis. The structure of ivorenolide A was fully determined by spectroscopic analysis including 1H, 13C NMR, FT-IR, and high-resolution mass spectrometry. Four carbons in the conjugated system of two triple bonds were determined by four singlets with chemical shifts of 78.5, 68.7, 70.3, and 81.3 ppm in 13C NMR, and the triple bonds produced a weak and sharp absorption band at 2150 cm21 in the IR spectrum. The epoxy group was determined by 13C NMR by the presence of two signals at 57.0 and 61.1 ppm, and by 1H NMR showing two multiplets at 3.12 and 3.55 ppm with a coupling constant J 5 4.1 Hz (cis-oxirane ring). An aliphatic acetylenic alcohol (55) isolated from Saussurea katochaete (family Asteraceae) collected in China (Saito et al., 2012) has two conjugated triple bonds, one hydroxyl group, one terminal double bond, and one oxirane cycle. Four carbons in the conjugated system of two triple bonds were determined by four singlets with chemical shifts of 81.8, 71.5, 71.4, and 65.6 ppm in 13C NMR, and the triple bonds in this molecule show an absorption band at 2253 cm21 in the IR spectrum. The epoxy group was determined by 13C NMR showing two signals at 60.9 and 61.1 ppm, and by 1 H NMR showing two multiplets at 3.06 and 2.59 ppm with a coupling constant J 5 3.8 Hz (cis-epoxide ring).

136

Fatty Acids

4.8 SYNTHESIS OF EPOXY ACETYLENIC LIPIDS The complexity of the synthesis of epoxy acetylenic compounds and their low stability under normal conditions are the primary reason why in the current literature only a few examples of practical syntheses of these compounds are described. However, a set of modern methods of organic synthesis allows obtaining the epoxy acetylenic compounds in various ways. The most common method of introduction of a triple bond into a complex molecule is a coupling of an acetylenic synthon with another fragment of the molecule. The epoxy group is prepared in several ways—the most common methods are the epoxidation of a double bond by Prilezhaev reaction [e.g., by mCPBA (meta-chloro-perbenzoic acid) in CH2Cl2], Sharpless epoxidation, and cyclization of halohydrins and similar compounds (e.g., tosylates). The order of introduction of various functions depends mostly on the target compound—there are examples of the construction of the goal structure around a triple bond, around the system of multiple bonds, and around an epoxy group. A very common method in the syntheses of such type of structures is the epoxidation of a double bond at the very last stage of the synthesis (the double bond can be considered as pro-epoxy double bond in these cases).

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137

Introduction of acetylenic parts to an epoxy compound was performed during the preparation of terminal alkynyl derivatives of 19-acetylgnaphalin (56) (Montenegro et al., 2010). Prior to the use of a bis(alkynyl)lithium reagent, 19-acetylgnaphalin (56) was reacted with one equivalent of Lithium (trimethylsilyl)acetylide (TMS)-acetylide in tetrahydrofuran (THF) at 278 C. Treatment of the reaction product with tetrabutylammonium fluoride produced the alkynyl derivative (57) in 65% isolated yield (two steps). Analogously, reaction of 19-acetylgnaphalin (56) with the lithium reagent prepared “in situ” by deprotonation of 1,3-diethynylbenzene with lithium hexamethyldisilazanide in THF at 278 C produced the goal epoxy diacetylene (58) as the sole reaction product in 65% yield. Analytical and spectroscopic data for the reaction products were in an agreement with the proposed structures. In the process of the development of the synthesis of the antibiotic dynemicin A (Shair et al., 1996), it was described an illustrative synthesis of epoxy acetylene compound (62) where the first step was the introduction into the molecule an acetylenic bond to form compound (59), then the resulted compound was epoxidized by Prilezhaev reaction to yield compound (60), and the third step added another acetylenic and a double bond (61) in a single unit, which then was cyclized by a condensation with a keto group to form a ring (62).

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Fatty Acids

In a communication (Zhang et al., 2012), ivorenolide A (65), a synthesis of an enantiomer of unprecedented immunosuppressive 18-membered macrolide previously isolated from K. ivorensis, which was featured by two conjugated acetylenic bonds, an epoxy group, and five chiral centers. The bioinspired asymmetric total synthesis of the enantiomer was achieved in 12 steps with 22% overall yield (63-65). In the described synthetic approach, the epoxy group was created by epoxidation of a double bond by Prilezhaev reaction with m-CPBA in CH2Cl2 as a reagent (64-65), while the conjugated system of two acetylenic bonds was The similar approach was used (Franck-Neumann et al., 1990) in the synthesis of the sesquicarene series using a C7-vinylalkynylcarbene (66-69). The introduction of an epoxy group was performed on the very last step of the synthesis by Prilezhaev reaction with m-CPBA in CH2Cl2 as a reagent.

Sometimes, building of the goal structure is based around the epoxide ring. One of these rare examples of such approach is shown during a synthesis of a naturally occurring antifeedant (73) (Grandjean et al., 1992). Where, the optically pure (2S,3R)-4-butyryloxy-2,3-epoxybutan-1-ol (70) was first oxidized under Swern conditions to yield the aldehyde. The resulting unstable epoxyaldehyde was treated with a mixture of triphenylphosphine and carbon tetrabromide in presence of trimethylamine to give dibromovinylepoxide (71) in 61% yield. After ester cleavage and silylation of the resulting alcohol, elimination using sodium hexamethyldisilazide cleanly provided the bromoepoxyacetylene (72) in

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high yield (85%). The last was then coupled with enyne yielding the epoxy diacetylenic ester with 92% yield, and after a quantitative acylation, the goal acetate (73) was synthesized. In the first total synthesis of naturally occurring (2)-nitidon (77) (Bellina et al., 2004) and its enantiomer, a system of three conjugated bonds (two acetylenic and one double) was created employing a modification of the CadiotChodkiewicz reaction (74-76). Two enantiomerically pure compounds were synthesized by the Sharpless asymmetric epoxidation of an (E)-2-ene-4,6-diyn-1-ol on the final step. The synthesis was made in five steps and 18% overall yield. A rare example of creating the goal epoxide by a cyclization of the corresponding acetylenic alpha-chlorohydrins by t-Bu-OK in ether is ascribed by Bernard et al. (1989). The authors coupled the commercially available 2methyl, 2-amino, but-3-yne (78) with various alpha-chloro carbonyls [three ketones and one aldehyde, with good preparative yields for ketones (78%45%) and the detectable one (5%) for the aldehyde] to yield alfachlorohydrines (79), the lasts were cyclized to produce the goal gammaamino alpha-acetylenic epoxides (80).

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Synthesis of the panaxydol analogs (83) (Kim et al., 1999) was carried out using Takahashi’s method by treatment of the Grignard reagent of diacetylene (81), then the last was condensed with allyl aldehyde to give hept-1-ene-4,6diyn-3-ol, which further underwent reactions with ethyl magnesium bromide and subsequently with various alkyl bromides to yield 7-alkylallyl-hept-1-ene-4,6diyn-3-ol derivatives (82). Regiospecific epoxidation of the double bond at C-9 of heptadeca-1,9-cis-diene-4,6-diyn-3-ol was performed under Prilezhaev reaction conditions by m-CPBA in CH2Cl2 to yield the goal epoxy acetylenes (83). A highly stereoselective and stereodivergent synthesis of two possible diastereomers of (2)-gummiferol was ascribed in Takamura et al. (2011), the synthesis (84-88) is representing a state of the art in the modern chemical synthesis. The epoxy alcohol (84) was prepared by Sharpless epoxidation followed by

ParikhDoering oxidation of a commercially available (2E,4E)-6-((tert-butyldimethylsilyl)oxy)hexa-2,4-dien-1-ol and subsequent two-carbon elongation of the corresponding aldehyde obtained under the MasamuneRoush conditions afforded α,β-unsaturated ester, which was carefully reduced by DIBAL-H. The alcohol (84) was epoxidized by second stereo-controlled Sharpless epoxidation with (1)-DIPT when the present epoxy ring and the hydroxyl group were used for epoxidation asymmetric induction. The ParikhDoering oxidation of (85), dibromo-olefination utilizing CoreyFuchs protocol in the presence of Et3N, dehydrobromination afforded TBS protected allylic alcohol (86) in 57% yield in three steps. The bromoacetylene (86) was reacted with a corresponding diacetylene under the optimized conditions of CadiotChodkiewicz reaction to form the desired coupling product (87). TBS-deprotection and acetylation of the resulting allylic alcohol provided acetate (88) in 48% yield from (86).

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The first total synthesis of recently isolated diacetylene alcohols oploxyne A (93), oploxyne B, and their C-10 epimers was recently ascribed (Yadav et al., 2011). The key steps involved are base-induced double elimination of a carbohydrate-derived β-alkoxy chloride (89) to generate the chiral acetylenic alcohol (90) and CadiotChodkiewicz cross-coupling reaction to attach a second acetylenic bond to the structure (90) to form the diacetylene (91). This synthesis is representing an unusual way in creating epoxy cycle where tosylation of alcohol (91) is used to create a protected triol (92), which was after selective deprotection with trifluoroacetic acid was converted to epoxy acetylenic alcohol (93) by treatment with diisopropylethylamine in 75% yield.

4.9 CONCLUDING REMARKS Terrestrial and marine secondary metabolites are unique sources for pharmaceuticals, food additives, flavors, and other industrial materials. Accumulation of such metabolites often occurs in plants subjected to stresses including various elicitors or signal molecules. At the present time, more than 50 acetylenic oxiranes and related compounds have been isolated from living organisms. Natural, semisynthetic, and synthetic acetylenic oxiranes, and their analogs and derivatives have been discovered and/or synthesized and evaluated for their biological activity. Inspired by the intriguing biological activities of many acetylenic natural products, polyyne moieties are now introduced in compounds that have pharmacological activity. The many functionalized acetylenic oxiranes thus obtained exhibit an impressive array of activities, such as enzyme inhibitor activities, cytotoxic, or antiviral activities. Without doubt, other important new acetylenic oxiranes possessing important biological activities will be discovered in the near future.

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Hashimoto, T., Akazawa, K., Tori, M., Kan, Y., Kusumi, T., Takahashi, H., et al., 1994. Three novel polyacetylene triglycerides, lycogarides A-C, from the myxomycete Lycogala epidendrum. Chem. Pharm. Bull. 42, 15311533. Hirakura, K., Mihashi, H., Fujihashi, T., Okuma, T., 1992. Isolation of acetylene compounds from Panax ginseng C.A. Meyer and preparation of their derivatives as antitumor agents. Japanese Patent: Jpn. Kokai Tokkyo Koho, 13 pp., JP 04264044 A. Hirakura, K., Takagi, H., Morita, M., Nakajima, K., Niitsu, K., Sasaki, H., et al., 2000. Cytotoxic activity of acetylenic compounds from Panax ginseng. Nat. Med. (Tokyo) 54, 342345. Ishibashi, M., 2005. Isolation of bioactive natural products from Myxomycetes. Med. Chem. 1, 575590. Ishibashi, M., Arai, M.A., 2012. Bioactive natural products from Myxomycetes having effects on signaling pathways. Heterocycles 85, 12991332. Jagdishprasad, P., Rao, N.S., 1998. Anticoagulant and anti-inflammatory and sunscreening effects of Hymenodictyon excelsum. Indian J. Pharm. 20, 221222. Jakupovic, J., Chau-Thi, T.,N., Fischer, N.,H., 1986. Isomeric epoxides of matricaria ester from Erigeron philadelphicus. Phytochemistry 25, 12231224. Jung, H.J., Min, B.S., Park, J.Y., Kim, Y.H., Lee, H.K., Bae, K.H., 2002. Gymnasterkoreaynes AF, cytotoxic polyacetylenes from Gymnaster koraiensis. J. Nat. Prod. 65, 897901. Kim, S.-I., Lee, Y.-H., Ahn, B.-Z., 1999. Synthesis of Ginseng diyne analogs and their antiproliferative activity against L1210 cells. Arch. Pharm. (Weinheim) 332, 133136. Konishi, I., Hosokawa, M., Sashima, T., Kobayashi, H., Miyashita, K., 2006. Halocynthiaxanthin and fucoxanthinol isolated from Halocynthia roretzi induce apoptosis in human leukemia, breast and colon cancer cells. Comp. Biochem. Phys. 142C, 5359. Kuklev, D.V., Domb, A.J., Dembitsky, V.M., 2013. Bioactive acetylenic metabolites. Phytomedicine 20, 100115. Laufer, Z., Beckett, R.P., Minibayeva, F.V., Luthje, S., Bottger, M., 2006. Occurrence of laccases in lichenized ascomycetes of the Peltigerineae. Mycol. Res. 110, 846853. Levy, L.M., Cabrera, G.M., Wright, J.E., Seldes, A.M., 2003. 5H-Furan-2-ones from fungal cultures of Aporpium caryae. Phytochemistry 62, 239243. Liers, C., Ullrich, R., Hofrichter, M., Minibayeva, F.V., Beckett, R.P., 2011. A heme peroxidase of the ascomyceteous lichen Leptogium saturminum oxidizes high-redox potential substrates. Fungal Gen. Biol. 48, 11391145. Luo, L., Wang, R., Wang, X., Ma, Z., Li, N., 2012. Compounds from Angelica keiskei with NQO1 induction, DPPH scavenging and a-glucosidase inhibitory activities. Food Chem. 131, 992998. Martin, R., Becker, H., 1985. Amides and other constituents from Acmella ciliata. Phytochemistry 24, 22952300. Menelaou, M.A., Foroozesh, M., Williamson, G.B., Fronczek, F.R., Fischer, Helga D., Fischer, N.H., 1992. Polyacetylenes from Chrysoma pauciflosculosa: effects on Florida sandhill species. Phytochemistry 31, 37693771. Mikolajczak, K.I., Smith Jr, C.R., Bagby, M.O., Wolff, I.A., 1964. A new type of naturally occurring polyunsaturated fatty acid. J. Org. Chem. 29, 318322. Minto, R.E., Blacklock, B.J., 2008. Biosynthesis and function of polyacetylenes and allied natural products. Prog. Lipid Res. 47, 233306. Montenegro, H.E., Ramirez-Lopez, P., De la Torre, M.C., Asenjo, M., Sierra, M.A., 2010. Two versatile and parallel approaches to highly symmetrical open and closed natural productbased structures. Chem. Eur. J. 16, 37983814.

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Yadav, J.S., Boyapelly, K., Alugubelli, S.R., Pabbaraja, S., Vangala, J.R., Kalivendi, S.V., 2011. Stereoselective total synthesis of (1)-oploxyne A, (-)-oploxyne B, and their C-10 epimers and structure revision of natural oploxyne B. J. Org. Chem. 76, 25682576. Yang, M.C., Kwon, H.C., Kim, Y.-J., Lee, K.R., Yang, H.O., 2010. Oploxynes A and B, polyacetylenes from the stems of Oplopanax elatus. J. Nat. Prod. 73, 801805. Zhang, B., Wang, Y., Yang, S.-P., Zhou, Y., Wu, W.-B., Tang, W., et al., 2012. An unprecedented immunosuppressive macrolide from Khaya ivorensis: structural elucidation and bioinspired total synthesis. J. Am. Chem. Soc. 134, 2060520608.

FURTHER READING Girard, Y., Rokach, J., 1985. Leukotriene antagonists and acceptable salts. Patent: EP 1984-309045 Eur. Pat. Appl., 102 pp.

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Chapter 5

Carbocyclic Fatty Acids: Chemistry and Biological Properties Moghis U. Ahmad, Shoukath M. Ali, Ateeq Ahmad, Saifuddin Sheikh and Imran Ahmad Jina Pharmaceuticals, Inc., Libertyville, IL, United States

Chapter Outline 5.1 Introduction 5.2 Naturally Occurring Cyclopropene Fatty Acids 5.2.1 The Halphen Test 5.2.2 Isolation of Cyclopropene Fatty Acids From Seed Oils 5.2.3 Chemical Characterization 5.3 Synthesis and Characterization of Sterculic Acid 5.3.1 Characterization of Dihydrosterculic Acid 5.3.2 Total Synthesis of cisCyclopropane Fatty Acids 5.3.3 Deuterated Cyclopropene Fatty Acids 5.4 Biosynthesis of Cyclopropane and Cyclopropene Fatty Acids 5.5 Mass Spectrometry of Cyclopropene Fatty Acids

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5.5.1 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropene Fatty Acids 5.5.2 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropane Fatty Acids 5.6 Physiological Properties of Cyclopropene Fatty Acids 5.7 Cyclopropaneoctanoic Acid 2-Hexyl in Human Adipose Tissue and Serum 5.7.1 Cyclopropaneoctanoic Acid 2-Hexyl in Patients With Hypertriglyceridemia 5.8 Leishmania Cyclopropane Fatty Acid Synthetase 5.8.1 Leishmania: A Fungal Infection 5.9 Conclusion References Further Reading

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5.1 INTRODUCTION A wide array of unusual fatty acids is found in seed oils (Badami and Patil, 1980; Van de Loo et al., 1993). These fatty acids contain three-member carbocyclic rings, namely cyclopropene fatty acids (CPE-FAs) and cyclopropane fatty acids (CPA-FAs). The CPE-FAs have been reported in the seed oils from plant species of the Malvaceae, Sterculiaceae, Tiliaceae, and Bombacaceae families (Smith, 1970; Christie, 1970). Sterculic acid [8-(2octyl-1-cyclopropenyl)octanoic acid] is often the prevalent CPE-FA, but malvalic acid [7-(2-octyl-1-cyclopropenyl)heptanoic acid], one-carbon shorter in chain length than sterculic acid (Fig. 5.1A) is also a significant component. They are present in different amounts in different plant species, the highest content being reported for Sterculia foetida (Sterculiaceae) seed oil, where the sterculic acid content is approximately 55% (Corl et al., 2001). Seed oil of Eriolaena hookeriana (Sterculiaceae) contains malvalic (25.8%) and sterculic (6.0%) acids (Ahmad et al., 1979). Eriolaena hookeriana seed oil is the second known species of the Sterculiaceae plant family in which the content of malvalic acid is greater than sterculic acid. It is a major source of malvalic acid, similar to Pterospermum acerifolium (Sterculiaceae), which has 16% malvalic acid and small amount of sterculic acid (Roomi and Hopkins, 1970). In Litchi chinensis seed oil, dihydrosterculic acid is the major carbocyclic fatty acid (Lie Ken Jie and Chan, 1977; Gaydou et al., 1993). Long-chain CPA-FAs are reported in various polar lipid classes of leaves (Kuiper and Stuiver, 1972), whereas both CPA-FAs and CPE-FAs are found H2 C

(A) C

C

CO2 H

A H2 C C HO2C

C

B

H2 C

(B) HC

CH

CO2 H

C H2 C HC

CH

CO2 H

D

FIGURE 5.1 (A) Chemical structures of CPE-FAs. (B) Chemical structures of the main CPA-FAs.

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in root, leaf, stem, and callus tissues in plants of the Malvaceae (Yano et al., 1972a,b; Schmid and Patterson, 1988a,b). A number of seed oils from different plant families have been investigated and found to contain both malvalic and sterculic acids, sometime accompanied by smaller proportions of one or both analogs of CPA-FAs (Smith et al., 1961; Wilson et al., 1961; Ahmad et al., 1976, 1978, 1979, 1981; Bohannon and Kleiman, 1978; Husain et al., 1980; Babu et al., 1980; Berry, 1980; Mustafa et al., 1986; Schmid and Patterson, 1988a,b; Daulatabad et al., 1998; Alitzetmuller and Vosmann, 1998; Herrera-Meza et al., 2014). In some cases, seed oils containing malvalic acid usually contain small amounts of epoxy fatty acids. Recently, CPE-FAs and CPA-FAs have been reported in Mediterranean nuts (Hanus et al., 2008). CPE-FAs have been the subject of many investigations due to their profound biological effects (Roehm et al., 1970; Phelps et al., 1965; AbouAshour et al., 1970), medical and mitogenic effects on animals (Tumbelaka et al., 1994; Andrianaivo-Rafehivola et al., 1995; Matlock et al., 1985), carcinogenic (Hendrick et al., 1980; Sinnhuber et al., 1974), and cocarcinogenic (Sinnhuber et al., 1968a,b; Lee et al., 1969, 1971) properties. Cottonseed and kapok oils are used for human consumption and contain sterculic and malvalic acid. These two fatty acids have been shown to cause numerous physiological disorders in farm and laboratory animals, such as retarded growth, postnatal mortality, altered lipid metabolism, inhibition of acyl desaturase, reduced microsomal enzyme activity, atherosclerosis in rabbits, and cancer in trout. Metabolism of sterculic acid was studied by Pawlowski et al. (1983), and it was reported that neither rabbits, trout, nor rats are able to metabolize the cyclopropene ring of 14C-sterculic acid to carbon dioxide. The primary route of excretion in rat is the urine, whereas it is excreted primarily through the bile in the rabbit and trout. The toxic responses, the products, and the patterns of metabolism of sterculic acid are similar between trout and rabbits, and different from rats. Due to the biological adverse effect, the presence of CPE-FA in food is dangerous to human health (Artman, 1969; Nolen et al., 1967; Potteau et al., 1970). CPA-FAs (Fig. 5.1B) are known to be components of bacterial membranes. The cis-11,12-methyleneoctadecanoic acid was first isolated from the phospholipids of Lactobacillus arabinosus and given the trivial name of Lactobacillic acid. It has also been found in a wide range of bacterial species, both Gram-negative and Gram-positive, and it is often accompanied by cis-9,10-methylenehexadecanoic acid and other homologs. Some organisms contain cis-9,10-methyleneoctadecanoic acid (dihydrosterculic acid) derived from oleic acid, together with homologs of fatty acids (C16 and/or C20 in chain length). Marseglia et al. (2013) reported the presence of CPA-FAs in milk and cheese in an amount of 0.12% of the total fatty acids. The same research group (Caligiani et al., 2014) reported that cow milks were generally positive to CPA-FAs (0.014%0.015% of total fatty acids), while goat, yak, and sheep milk were negative. Lactobacillic acid and dihydrosterculic acid are components of bacterial membranes and have been recently detected

150

Fatty Acids

in milk from cows fed with maize silage. This suggests that the presence of CPA-FAs in milk and dairy products is derived from their presence in silage forges, where CPA-FAs can be released by bacteria during fermentation conditions. On the contrary, lactic acid bacteria, ubiquitous in fermented milk and cheeses, seem not to be able to release CPA-FAs in the conditions of milk fermentation. Therefore, CPA-FA can serve as molecular markers, to distinguish milk from dairy cows fed with silage-based diet from milk from cows fed with hay-based diets. The presence of CPA-FA in dairy products could be used as a marker of silage feeding. Recently, CPA-FAs are also identified in adipose tissue and serum of humans and rats (Sledzinski et al., 2013). Fatty acids of adipose tissue and serum extracted from obese women were identified as cyclopropaneoctanoic acid 2-hexyl (also called cis-9,10-methylenehexadecanoic acid), cyclopropaneoctanoic acid 2-octyl, cyclopropanenonanoic acid, and 2-[[2-[(2-ethylcyclopropyl)methyl]cyclopropyl]methyl] acid. The cyclopropaneoctanoic acid 2-hexyl was the main CPA-FA (approximately 0.4% of total fatty acids in human adipose tissue and approximately 0.2% of total fatty acids in the serum). The cyclopropaneoctanoic acid 2-hexyl has also been found in serum and adipose tissue of rats in amounts comparable to humans. The content of cyclopropaneoctanoic acid 2-hexyl decreased in adipose tissue of rats on restricted diet. Adipose tissue cycloproaneoctanoic acid 2-hexyl is mainly stored in triacylglycerols and storage of this CPA-FA is affected by restriction in diet. In further study, the increased level of cyclopropaneoctanoic acid 2-hexyl was observed in patients with hypertriglyceridemia-related disorders and was associated with chronic kidney disease (CKD) and obesity (Mika et al., 2016). This chapter describes the chemistry of CPE-FAs and CPA-FAs, biosynthesis, biological properties, and their industrial application. A review on the chemistry of cyclopropene compounds was published earlier (Carter and Frampton, 1964).

5.2 NATURALLY OCCURRING CYCLOPROPENE FATTY ACIDS The application of modern methods has shown that the occurrence of CPEFAs in natural oils is not as uncommon as was once believed. CPE-FAs are reported in seed oils of plant families, Malvaceae, Sterculiaceae, Tiliaceae, Bombacaceae, Leguminosae, Ranunculaceae, and references cited in the introduction section of this review. Fatty acids in naturally occurring glycerides, plant, or animal are almost always found in homologs series with even numbered chains differing in length by multiple of two carbons. In contrast, sterculic and malvalic acids differ in chain length by a single carbon; malvalic acid has an odd-numbered (C17) chain length. These two homologs acids, sterculic and malvalic, have often been found to occur together, and give positive Halphen test (Halphen, 1897, 1898).

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151

5.2.1 The Halphen Test Halphen test is over hundred years old analytical method that was used to check adulteration or replacement of one vegetable oil with another. Many early studies were erroneous as a result of inadequate equipment and procedure. In the original method described by Halphen (1897, 1898), equal volume of the oil under examination (13 mL), amyl alcohol, and carbon disulfide containing 1% free sulfur were placed in tube and heated on boiling water bath for 1015 minutes. A red or orange color develops in the presence of CPE-FAs containing seed oils, for example, cottonseed oil. Several investigators modified the procedure to change heating time, temperature of the water bath, excess of sulfur, use of amyl alcohol, addition or substitution of an additional reagent like pyridine, and use of sealed tube or pressure flasks in place of condensing tubes (Carter and Frampton, 1964). Deutschman and Klaus (1960) studied the Halphen test with modifications in reaction conditions and proposed a reproducible color development. A very small amount of fatty acid with a cyclopropene ring was suggested as the material in cottonseed oil responsible for a positive Halphen reaction (Dijkstra and Duin, 1955; Gunstone, 1955). The reaction with sterculic acid indicated that the reaction involved an opening of the ring across the single bonds (Faure, 1956; Nunn, 1952). With the development of Halphen color (orange or red color), the absorption bands at 1869 and 1010 cm21 attributed to cyclopropene ring disappears. A strong band at 1900 cm21 appears and subsequently disappearance was attributed to the double bond in the grouping SC 5 S, which was first formed by the reaction between carbon disulfide (CS2) and cyclopropene ring, and then polymerized across the C 5 S double bond. The addition of sulfur and amyl alcohol was not required. The structures of two-colored products produced in Halphen test were studied with the reaction of 1,2-diethylcyclopropene as shown in Fig. 5.2. H2 C

CH3 S

S

S

C

CH3 SH

S C

C

H3 C

C

H3 C

H2 C

S H3 C

S

H 3C

FIGURE 5.2 Structures of two-colored products formed in the Halphen test with 1,2-diethylcyclopropene. Adapted from Carter and Frampton, 1964. Review of the chemistry of cyclopropene compounds. Chem. Rev. 64, 497525.

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Fatty Acids

The structures were established by a combination of IR, UV, nuclear magnetic resonance (NMR), and mass spectral methods.

5.2.2 Isolation of Cyclopropene Fatty Acids From Seed Oils Sterculic acid was first isolated from the seed oil of S. foetida (Sterculiaceae) by urea clathrate (Nunn, 1952) and its structure was proposed to be ω-(2-n-octylcycloprop-1-enyl) octanoic acid. The acid was isolated by low-temperature saponification of the oil followed by fractional crystallization of the urea clathrates of the acids from methanol. Final purification of sterculic acid was done by low-temperature crystallization from acetone. In addition, the presence of CPE-FAs in the plant family Malvaceae was studied on the oils extracted from the seeds and leaves of Malva verticillata, Malva parviflora, and Gossypium hirsutum (cottonseed). The solvent extracted oil was saponified under mild reaction condition and the total fatty acids were subjected to low-temperature crystallization from acetone. The Halphen positive acids collected in the filtrate and further purified by reversed-phase partition chromatography followed by low-temperature crystallization in acetone and petroleum ether. The isolated biologically active fatty acid was characterized as a homolog of sterculic acid and was assigned the name malvalic acid (Shenstone and Vickery, 1956; Macfarlane et al., 1957). In general, the isolation procedures adopted by various group of researchers used either urea-complex fractionation or the low-temperature crystallization method (Nordbay et al., 1962; Kircher, 1964). Sterculia foetida and Bombax olagineum are the richest source of sterculic acid (Shenstone and Vickery, 1961; Cornelius and Shone, 1963) and malvalic acid in lower concentration. The seed oil of E. hookeriana (Sterculiaceae) contains malvalic (25.8%) and sterculic (6.0%) acids (Ahmad et al., 1979). Eriolaena hookeriana is the second known species of the Sterculiaceae plant family whose seed oil contains more malvalic acid than sterculic acid. Ahmad et al. (1979) used gas chromatography-mass spectrometry (GC-MS) analysis of the silver nitratemethanol treated methyl esters for unequivocal characterization of the individual CPE-FAs.

5.2.3 Chemical Characterization Nunn (1952) assigned the structure of sterculic acid and proposed the nomenclature ω-(2-n-octylcycloprop-1-enyl) octanoic acid. The structure was assigned on the basis of reactions like hydrogenation, oxidation, reduction, halogenation, and polymerization as described later.

5.2.3.1 Hydrogenation Sterculic acid on hydrogenation with palladiumcalcium carbonate gave dihydrosterculic acid and showed the consumption of hydrogen equivalent to

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5 H2 C H3 C

HC

O

O CH (CH 2)7

(CH2) 7

C

H3 C

OH

(CH 2)7

(CH2) 7

Dihydrosterculic acid

+

(CH2) 7

O

CH3

H2 C C

OH

n-Nonadecanoic acid

H2/Pd

H3 C

153

O C (CH 2)7

H2/PtO 2

H3 C

C

CH (CH2) 7

OH

(H2 C)8

OH

10-Methyloctadecanoic acid

Sterculic acid

+

H3 C (CH2) 8

CH3

O

CH

C (CH 2)7

OH

9-methyloctadecanoic acid FIGURE 5.3 Hydrogenation products of sterculic acid.

one double bond. Similarly, dihydromalvalic acid was obtained from malvalic acid by hydrogenation using palladiumcalcium carbonate catalyst (Macfarlane et al., 1957). However, hydrogenation in the presence of platinum catalyst (PtO2) absorbs one additional hydrogen atom and resulted in a mixture of n-nonadecanoic acid and two methyl-substituted octadecanoic acids (Fig. 5.3). Oils containing both sterculic and malvalic acids on prolong hydrogenation produced dihydro derivatives and branched chain products instead of straight chain products (Wilson et al., 1961).

5.2.3.2 Oxidation Ozonolysis of sterculic acid in acetic acid at low temperature provides an ozonide, which on oxidation with hydrogen peroxideacetic acid gives pelargonic acid and azelaic acid as the main product (Nunn, 1952). Alkaline hydrolysis of the dioxo compound gives four products, of which three products, methyl n-octyl ketone, azelaic acid, and 9-oxodecanoic acid, were isolated and identified (Faure and Smith, 1956). Oxidation of sterculic acid with potassium permanganate (KMnO4) in acetone gives pelargonic acid and azelaic acid as the main products (Fig. 5.4A). 5.2.3.3 Reduction Sterculic acid on reduction with lithium aluminum hydride (LiAlH4) produces a stable sterculyl alcohol (Nunn, 1952) (Fig. 5.4B). The cyclopropene

154

Fatty Acids O

O

O

(A)

+

C OH

H 3C(H2 C)7

Pelargonic acid

C

C

(CH2) 7

HO

OH

Azelaic acid H2O2 AcOH

KMnO4 Acetone H2 C H3 C

C

C H2

H3 C(H 2C) 7

OH

(CH 2)7

C

C

1. O 3

C

(CH2) 7

O

O

O

2. H 2/Pd

(CH 2)7 CO2H

9,11-Diketononadecanoic acid

Sterculic acid KOH O C CH3

H 3C(H2 C)7

O

O

Methyl n-octyl ketone

+

C

C

(CH2) 7

H 3C

9-Oxodecanoic acid

+

+ Pelargonic acid

(B) H3 C

H2 C C (CH2) 7

Azelaic acid

H2 C

O

LiAlH4 C (CH 2)7

Sterculic acid

OH

OH

H3 C

C (CH2) 7

H2 C

C

(CH 2)7

OH

Sterculyl alcohol

FIGURE 5.4 (A) Oxidation products of sterculic acid. (B) Reduction products of sterculic acid.

ring stays intact on reduction. The sterculyl alcohol was methylated to form the methyl sterculyl ether followed by reduction to form 1,2-dioctylcyclopropene, also known as sterculene (Nordbay et al., 1962). The infrared spectra of these preparations were compared with sterculic acid and its methyl ester to confirm the presence of cyclopropene ring.

5.2.3.4 Halogenation Methyl sterculate on heating with sodium iodide in acetone gives diiodide. The diiodo ester on heating at various temperatures up to 190 C, heating under reflux with zinc dust in acetone, heating with pyridine, quinolone, or

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5 H2 C

(A) H 3C(H2 C)7

C X

H 3C(H2 C)7

H2 C CH2CH2 (CH2 )5 CO2Me

C H2 C

C H

H 3C(H2 C)7

C

Y

C

C

CH2CH2 (CH2 )5 CO2Me

C

CHCH 2(CH2) 5CO 2Me

H2 C

C H

CH

CH(CH2 )5CO2Me

C H

H 3C(H2 C)7

CH2 H 3C(H2 C)7

155

CH2 H2 C

CH

CH(CH2 )5CO2Me

H 3C(H2 C)7

H C

C

CHCH 2(CH2) 5CO 2Me

X, Y = I, I; H, Br; H , I

O

CH2

(B) R

H C

C

O

(CH 2)7 X

+ CH2 R

H C

O

C (CH 2)7

O

X H2 C

+

O CH2X

H3 C(H 2C) 7

(CH 2)7

O

R

C

O H C (CH 2)7

X = Br or Cl

O

+ CH2X

R = CH 3(CH2)7HC H3 C(H 2C) 7

O

C (CH 2)7

O

FIGURE 5.5 (A) Dehalogenation products of sterculic acid. (B) Halogenation products of sterculic acid.

potassium hydroxide gives a series of products. The decomposition of hydrobromides and hydroiodides of methyl sterculate gives similar products (Fawcett and Smith, 1960). The reaction products (Fig. 5.5A) were expected to produce on decomposition depending on the severity of the decomposition method. The reaction of hydrogen halides with CPE-FAs produces four isomeric monounsaturated monohalo moieties (Fig. 5.5B). The mechanism was analogous to that postulated for the polymerization of sterculic acid (Rinehart et al., 1959).

156

Fatty Acids

C H3 C(H 2C) 7

O

O

CH2

CH2 H C

C (CH 2)7

H C

OR' H3 C(H 2C) 7

OR''

OR'

(B)

CH2OR''

O

H C

C (CH 2)7

H3 C(H 2C) 7

(CH 2)7

OR''

(A)

C

C C

CH2OR'' O HC OR'

C

H3 C(H 2C) 7

(C)

C (CH 2)7

OR'

(D)

FIGURE 5.6 Polyesters (AD) of sterculic acid where R and Rv are sterculic acid residues.

5.2.3.5 Polymerization Sterculic acid is unstable and polymerization occurs at room temperature and even starts slowly at 0 C, as indicated by the increase of the equivalent weight with time. It was suggested that sterculic acid polymerized by the addition of carboxyl group across the double bond of cyclopropene ring (Nunn, 1952; Faure and Smith, 1956). The polymerization proceeds by isomerization of the cyclopropene ring with carboxylic acid addition to give polymeric mixtures of compounds (Fig. 5.6) (Rinehart et al., 1959). The polymerized products were found negative to Halphen test and insoluble in hot methanol. Polymerization and acetolysis reactions of sterculic acid have been shown to proceed via opening of the cyclopropene ring by carboxylate group to yield four isomeric products (Rinehart et al., 1961). The polymer was saponified at room temperature to the corresponding unsaturated hydroxy acids followed by acetylation to give the unsaturated acetoxy acids. The acetoxy acids were also prepared by heating sterculic acid in excess glacial acetic acid, and the isomeric products were identified by oxidative degradation and infrared spectra. The infrared spectrum of the purified polymer was in agreement with that described by Faure and Smith (1956) indicated the absence of cyclopropene ring (1869 and 1010 cm21). Bands were reported at 1737 and 1169 cm21 and suggested the presence of ester, while bands at 1648 and 901 cm21 and at 1712 and 960 cm21 were indicative of unsym-disubstituted olefin and terminal carboxyl groups, respectively.

5.3 SYNTHESIS AND CHARACTERIZATION OF STERCULIC ACID The assignment of ω-(2-n-octylcycloprop-1-enyl) octanoic acid by Nunn (1952) as the structure of sterculic acid (I, Fig. 5.7) was subsequently confirmed by synthesis (Castellucci and Griffin, 1960). Chemical synthesis was

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5

H2 C C

H3 C

H2 C

O C

C

(CH 2)7

(CH2) 7

OH

H3 C

HC

O C

CH C H

(CH2) 5

CH(CH2) 7

I

OH

II H2 C

R1 HC

157

C

H C R2

C H

R1

C H

IIIa,b

C

R2

IVa,b

IIIa, R1 = IIIb, R2 = IVa, R1 = IVb, R2 =

n-C7H10 ; R2 = – (CH2)7CO2H n-C8H10; R2 = – (CH2)6CO2H n-C8H10; R2 = – (CH2)7CO 2H n-C8H10; R2 = –(CH2)7CO2H

H C H2 C

CO2H

C

H3 C C

H

H C

CO2H

C

CO2 H

C

CO2 H

V

VI

FIGURE 5.7 Proposed structures of sterculic acid.

done using acetylenic precursor, stearolic acid, which was treated with methylene iodide in the presence of zinccopper couple catalyst for 9 hours under reflux, following the general procedure of Simmons and Smith (1959). Fractional crystallization of the urea adduct led to the isolation of the sterculic acid (I), m.p. 18.919.9 C (lit. m.p. 19.319.9 C). The properties of the synthetic sterculic acid were compared with those of the isolated cyclopropene acid from seed oils and were found to be identical. The acid gave red color with Halphen reagent. The infrared spectra of malvalic acid, dihydromalvalic acid compared with the synthetic sterculic acid, and dihydrosterculic acids found that the acids were analogous (Macfarlane et al., 1957). Both CPE-FAs gave bands at 1872 and 1008 cm21. Hydrogenation of the acid using palladiumcharcoal catalyst gave dihydrosterculic acid resulted in the disappearance of the band at 1872 cm21 and the shifting of the 10081020 cm21 (cyclopropane). In the high-frequency region of the spectra, no absorption was observed for malvalic or sterculic acid or their methyl ester. Absorption for the stretching frequencies of carbonhydrogen groups in the cyclopropane ring was noted at 30563058 and 29882990 cm21 for the dihydro acids and for the lactobacillic acid, a naturally occurring CPA-FA.

158

Fatty Acids

NMR data (Rinehart et al., 1958) further supported the assigned structure for sterculic acid. The NMR spectrum of the sterculic acid showed signals at δ 6.88 (COOH), δ 3.50 (chain CH2), δ 3.95, δ 3.85 (split) (CH3), δ 2.53 (CH2 adjacent to carboxyl or olefin, unresolved, integrated intensity of 67 protons); δ 4.03 (CH2 of a cyclopropene group). Most significant, no signal was observed in the region expected for absorption by olefinic proton. The absence of signals in this region excludes the alternate formulas II, IIIa, IIIb, IVa, and IVb (Fig. 5.7). In the structure IIIa, or IIIb, the double bond is exocyclic to the cyclopropane ring, and in the structure IVa or IVb, the double bond is in the ring but trisubstituted. These formulas place the cyclopropane ring at the same position as I but differ from I in the double bond location. Formula IIIa or IIIb are the analogy in that ozonolysis of Feist’s acid (V), with double bond exocyclic to a cyclopropane ring gives no formaldehyde but the main products are expected from the endocyclic olefinic structure (VI). All alternate formulas (II, IIIa, IIIb, IVa, and IVb) differ from structure I in that each has at least one olefinic hydrogen atom, while I has none. The NMR showed that the acid lacked olefinic hydrogens, and therefore eliminates the possibility of alternate structure ω-(2-n-hexylcyclopropyl)dec-9-enoic acid (II) (Fig. 5.7) proposed by Varma et al. (1955a, 1955b, 1957). This conclusion was further supported by the comparison of spectra of S. foetida oil, sterculic acid, dihydrosterculic acid, methyl dihydrosterculate, 9,11-diketononadecanoic acid (Hopkins and Bernstein, 1959; Hopkins, 1961), and formula I (Fig. 5.7) unequivocally assigned to sterculic acid.

5.3.1 Characterization of Dihydrosterculic Acid 2-Octyl cyclopropaneoctanoic acid commonly referred as CPA-FA occurs naturally in the phospholipids of bacterial membranes, in seed oils containing CPE-FAs, and in Litchi sinensis oil. Several research groups reported the synthesis and analytical data of CPA-FAs (Stuart and Buist, 2004; Jing et al., 2004; Cryle et al., 2005; Coxon et al., 2003; Tashiro et al., 2002; Hartmann et al., 1994; Gunstone and Perera, 1973; Longone and Miller, 1967; Minnikin, 1966), and signals were assigned to methylene protons in the 1H NMR spectra, including distinguishing the cis and trans protons (Minnikin, 1966; Longone and Miller, 1967). It is observed that 1 H NMR data for the salient signals do not always agree or assignments in some cases are incomplete. Konthe (2006) used dihydrosterculic acid and its methyl ester as compounds of representative of CPA-FAs. The 1H NMR (500 MHz) and 13C NMR (125 MHz) data are presented along with assignments of key peaks, and 2D homo- and heteronuclear correlations were acquired with CDCl3 as solvent. The assignments relating to the cyclopropane ring are shown in Fig. 5.8.

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5 –0.30 H

H

159

0.60

10.93 0.68 1.17

0.68

H

H

H

1.17

H

R1

28.68 H

1.40

15.78

28.73

R2

H

1.40

FIGURE 5.8 NMR chemical shifts (ppm) assigned to the cyclopropane moiety in methyl dihydrosterculate (13C shifts are italicized) R1 5 (CH2)6COOMe and R2 5 (CH2)6CH3. Adapted from Konthe, 2006. NMR characterization of dihydrosterculic acid and its methyl ester. Lipids 41, 393396.

Peaks at 20.30, 0.60, and 0.68 ppm were observed in agreement with earlier literature data (Tashiro et al., 2002; Stuart and Buist, 2004; Cryle et al., 2005), and correlated with 13C NMR shifts at 10.93 ppm for the methylene (CH2) moiety of the cyclopropane ring; 15.75 and 15.78 ppm for the methine (CH) carbons. The 2D homonuclear correlation showed that the peaks at 20.30 and 0.60 ppm were assigned to the two C3 protons (cis and trans) of the cyclopropane ring and the two methine protons resonate slightly downfield at 0.68 ppm. The peak at 1.17 ppm, which was not resolved or listed in the earlier literature, correlates with the signal of the methine protons at 0.68 ppm and is therefore assigned to methylene protons to the cyclopropane ring. However, the peak at 1.17 ppm is caused by two protons. In heteronuclear correlation, it correlates with 13C NMR signals at 28.68 and 28.73 ppm. These two 13C NMR signals also correlate with the downfield region at 1.40 ppm, which appears as broad methylene peak and is likely due to shielding effects of the cyclopropane ring and similar to the differing shifts of the methylene protons in the cyclopropane ring; one proton each of the two methylene units, to the cyclopropane ring at C8 and C11 of the fatty acid chain, is responsible for the peak at 1.17 ppm. The downfield correlation is caused by the two other protons of these methylenes. According to the previous findings, the upfield signal is assigned to the cis proton and the downfield signal to the trans proton. This signal of the trans proton is resolved from the peak of the two methine protons of the cyclopropane ring, which is located at 0.68 ppm. The four protons attached to the two methylene carbons α 2 to the cyclopropane ring also show a split signal. Two of these protons, one from each methylene moiety, display distinct shift at 1.17 ppm and the signal of the other two protons is observed at 1.40 ppm, within the broad methylene peak.

160

Fatty Acids

5.3.2 Total Synthesis of cis-Cyclopropane Fatty Acids The synthesis of enantiomeric pairs of four cis-CPA-FAs, dihydromalvalic acid, dihydrosterculic acid, lactobacillic acid, and 9,10-methylenehexadecanoic acid, is reported (Shah et al., 2014). The synthetic approach commences with Rh2(OAC)4-catalyzed cyclopropenation of 1-octyne and 1-decyne, followed by chromatographic resolution of racemic 2alkylcycloprop-2-ene-1-carboxylic acids. Saturation of the individual diastereomeric N-cycloprop-2-ene-1-carbonylacyloxazolidines, followed by elaboration to alkylcyclopropylmethylsulfones, allowed Julia-Kocienski olefination with various ω-aldehyde esters. Finally, saponification and diimide reduction afforded the individual cis-CPA-FA enantiomers. The four most prominent CPA-FAs are those derived from: cis-palmitoleic acid (cis-Δ9 C16:1), namely cis-9,10-methylenehexadecanoic acid; 8Zheptadecenoic acid (cis-Δ8 C17:1), namely dihydromalvalic acid; oleic acid (cis-Δ9 C18:1), namely dihydrosterculic acid; and cis-vaccenic acid (cis-Δ11 C18:1), namely lactobacillic acid (Fig. 5.9).

(CH 2)7

z

(CH 2)5 10

(CH 2)5

CO2 H

9

cis-Palmitoleic acid

9

z

8

(CH 2)7

z

10

CO2 H (CH 2)6

CO2 H

Oleic acid

(CH 2)5 12

8

cis

CO2 H (CH 2)6

(CH 2)7

(CH 2)7

cis

10

9

CO2 H

Dihydrosterculic acid (3)

(CH 2)9

z

9

(CH 2)7

Dihydromalvalic acid (2)

(CH 2)7 9

CO2 H

9

10

cis-9,10-Methylenehexadecanoic acid (1)

8Z-Heptadecenoic acid

(CH 2)7

(CH 2)7

cis

11

cis-Vaccenic acid

CO2 H

(CH 2)5 12

(CH 2)9

cis 11

CO2 H

Lactobacillic acid (4)

FIGURE 5.9 Structures of cis-CPA-FAs 14 from unsaturated fatty acids. Adapted from Shah et al., 2014. Total synthesis of cis-cyclopropane fatty acids: dihydromalvalic acid, dihydrosterculic acid, lactobacillic acid, and 9,10-methylenehexadecanoic acid. Org. Biomol. Chem. 12, 94279438.

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5

161

Shah et al. (2014) reported a general route to both enantiomers of four common cis-CPA-FAs in enantiopure forms using a diastereomeric resolution of cyclopropanecarboxamides, originally developed by Liao et al. (2004) as a key enabling step, allowing the synthesis of both enantiomers of the four CPA-FAs depicted in Fig. 5.9.

5.3.3 Deuterated Cyclopropene Fatty Acids Certain CPE-FAs are potent and could be useful for affinity labeling of desaturases in insect pheromone biosynthetic studies. A series of novel, selectively deuterated CPE-FAs analogs, have been synthesized and characterized by Gosalbo et al. (1993). Selective incorporation of one to five deuterium atoms into CPE-FAs at different sites and from moderate to high yields has been developed by this group of chemists. The developed methods should easily be applicable to the preparation of the corresponding tritiated analogs. In most cases, deuterium was introduced in the last step of the synthesis to optimize the procedure for the preparation of the corresponding radioactive materials. For example, deuterated cyclopropene fatty ester (1a) (Fig. 5.10) in which deuterium is placed at the ω-position was synthesized following the reaction sequence depicted in Scheme 5.1.

(CH 2)3 R1

C

C

R1

(CH2) nCOR2

C

C

(CH 2)nCOR2

CH

CH

D

R3

3a: R1 = C3H7, R2 = OCH3, n = 10 4b: R1 = C3H 7, R2 = OH, n = 10 4a: R1 = C4H9, R2 = OCH3, n = 9 4b: R1 = C4H 9, R2 = OH, n = 9 5a: R1 = C5H11, R2 = OCH 3, n = 8 5b: R1 = C5H11, R2 = OH, n = 8

1a: R1 = D, R2 = OCH3, R3 = H, n = 10 1b: R1 = D, R2 = OH, R3 = H, n = 10 2a: R1 = H, R2 = OCH3, R3 = D, n = 8 2b: R1 = H, R2 = OH, R3 = D, n = 8 (CH 2)3

(CH2) 7CD 2CD 2CO

R

H3 C

D

6a: R = OCH3 6b: R = OH FIGURE 5.10 Synthetic deuterated cyclopropane fatty esters. Adapted from Gosalbo et al., 1993. Synthesis of deuterated cyclopropene fatty esters structurally related to palmitic and myristic acids. Lipids 28, 11251130.

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Fatty Acids

A chlorine atom was selected as precursor of deuterium. Alkylation of acetylene (7a) with 1-chloro-3-iodopropane followed by jones oxidation and methylation of the resulting fatty acid (8b) gives intermediate compound (8a). The cyclopropene ring was introduced by the reaction of alkyne (8a) with ethyl diazoacetate in the presence of activated Cu-bronze as a catalyst, and the resulting diester (10a) was hydrolyzed at room temperature. Decarbonylation of diacylchloride (10b) was accomplished with diethyl ether solution of ZnCl2. Reduction of cyclopropenium ions thus formed was accomplished with NaBH4 in sodium hydroxide/methanol at low temperature. The chlorine atom remains unaffected under this reduction condition and successfully reduced by NaBD4 in dimethysulfoxide at 70 C.

HC

C

(CH2 )n

1. MeLi or LDA/THF 2. RX/HMPA OTHP

(CH 2)3 R1

3. CrO3/H 2SO 4/acetone

7a: n = 10 7b: n = 8

K 2CO 3/CH3I/DMF

C

C

(CH2 )nCOR 2

8b: R1 = Cl, R2 = OH, n = 10 9b: R1 = H, R2 = OH, n = 8 8a: R1 = Cl, R2 = OCH 3, n = 10 9a: R1 = H, R 2 = OCH 3, n = 8

N 2CHCO2Et/Cu-bronze (CH 2)3 R1

C

C CH

(CH2 )nCO2 CH3

1. (COCl)2/r.t. 2. ZnCl2/Et2O/CH 2Cl2 3. NaBH4 or NaBD 4 MeOH

(CH 2)3 R1

C

C

(CH2 )nCOR 2

CH

R2 COR3 NaBD 4/DMSO/70ºC

12: R1 = Cl, R2 = H, n = 10 1a: R1 = D, R2 = H, n = 10 2a: R1 = H, R2 = D, n = 8 KOH/EtOH/r.t.

10a: R1 = Cl, R2 = OCH3, R3 = OEt, n = 10 11a: R1 = H, R2 = OCH3, R3 = OEt, n = 8 10b: R 1 = Cl, R2 = R3 = OH, n = 10 11b: R 1 = H, R2 = R3 = OH, n = 8

SCHEME 5.1 Synthesis of deuterium labeled cyclopropene fatty esters (1a, 2a). Adapted from Gosalbo et al., 1993. Synthesis of deuterated cyclopropene fatty esters structurally related to palmitic and myristic acids. Lipids 28, 11251130.

The synthesis of other deuterated cyclopropene fatty ester (2a) is also outlined in Scheme 5.1. Alkylation of acetylene (7b) with 1-bromopropane followed by jones oxidation and methylation of resulting acid (9b) produces 9a. Transformation of 9a to final product (2a) was accomplished as described earlier for the compound 1a. Ring-labeled cyclopropene fatty esters (3a5a) were synthesized by decarbonylation reaction followed by reduction of diacyl chlorides (13a13c) following the procedure of Arsequell et al. (1992). Both decarbonylation and cyclopropenium ion reduction was carried out as described earlier and NaBD4 was used as reducing agent (Scheme 5.2). Details of synthesis and experimental procedures are given in the original publication (Gosalbo et al., 1993).

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5 R HO2C

H C

C

R

O

C (CH 2)n

13a: R = C3H7, n = 10 13b: R = C4H9, n = 9 13c: R = C5H11, n = 8

163

1. (COCl)2/r.t.

C OH

2. ZnCl2/Et2O/CH2Cl 2

D

C

H C

O

C (CH 2)n

3. NaBD4/MeOH

C OCH 3

3a: R = C3H7, n = 10 4b: R = C4H9, n = 9 5a: R = C5H11, n = 8

SCHEME 5.2 Synthesis of deuterium labeled cyclopropene fatty esters (3a, 5a). Adapted from Gosalbo et al., 1993. Synthesis of deuterated cyclopropene fatty esters structurally related to palmitic and myristic acids. Lipids 28, 11251130.

5.4 BIOSYNTHESIS OF CYCLOPROPANE AND CYCLOPROPENE FATTY ACIDS CPA-FAs containing three-member carbocyclic rings are found in both bacteria and plants. The bacterial CPA-FA, lactobacillic acid, is produced by the addition of methylene group derived from the methyl group of S-adenosyl methionine to the double bond of cis-vaccenic acid attached to phosphatidylethanolamine (Chung and Law, 1964; Zalkin et al., 1963). Although CPAFAs are widely distributed in bacterial lipids, no CPE-FAs have been reported in bacteria. One striking difference between the bacteria CPA-FAs and the plant CPA-FAs is that the bacteria CPA-FAs are normally esterified at the sn-2-position of phosphatidylethanolamine, whereas the plant CPAFAs are preferentially esterified at the sn-1 position of phosphatidylcholine. The desaturated products of CPA-FAs, CPE-FAs, are only reported in plants. CPA-FAs are usually minor components with CPE-FAs more abundant in plants. Both CPA-FAs and CPE-FAs are distributed across several plant orders, mostly Malvales. The sterculic acid is often the major CPE-FAs, but malvalic acid, one-carbon shorter in chain length than sterculic acid, can be a significant component. Seed oil of S. foetida contains up to 78% CPE-FAs, mainly sterculic acid (Badami and Patil, 1980; Pasha and Ahmad, 1992). CPA- and CPE-FAs are possibly functioning as antifungal agents in plants. The biosynthetic pathway of CPA-FAs in bacteria is studied in great detail and well understood (Grogan and Cronan, 1997). The first CPA-FA synthase gene was cloned from Escherichia coli on the basis of its ability to complement a CPA-FA-deficient mutant (Grogan and Cronan, 1984). It was shown that bacterial CPA-FAs were synthesized from monounsaturated fatty acids by the addition of a methylene group, derived from S-adenosylmethionine, across the double bond. The monoenoic fatty acyl substrates are esterified to phospholipids, mostly phosphatidylethanolamine (Grogan and Cronan, 1997). However, less attention has been paid to the biosynthesis of CPE-FAs in plants. Yano et al. (1972a,b) explored the biosynthesis of CPAFAs and CPE-FAs in immature seeds, leaves, and callus tissue cultures of

164

Fatty Acids H2 C

O H3 C

H C (CH 2)7

H C

(CH 2)7C

Oleic acid

CH2 addition OH

H3 C

HC

CH

(CH2 )7

Cyclopropane synthase

H2 C

O C (CH2) 7

Desaturation

H3 C

OH

α-Oxidation H2 C

(CH2 )7

C (CH2) 7

Sterculic acid

α-oxidation

HC

O C

(CH2 )7

OH

Dihydrosterculic acid

H3 C

C

H2 C

O CH

H3 C

C (CH2) 6

Dihydromalvalic acid

OH

C (CH2 )7

O C

C (CH2) 6

OH

Malvalic acid

FIGURE 5.11 Proposed pathway for the biosynthesis of sterculic acid from oleic acid. Adapted from Yano et al., 1972. The biosynthesis of cyclopropane and cyclopropene fatty acids in higher plants (Malvaceae). Lipids 7, 3545.

several species of Malvaceae, and proposed that the pathway involved initial formation of dihydrosterculic acid from oleic acid with subsequent desaturation to sterculic acid (Fig. 5.11). The chemical characterization of labeled CPA-FAs and CPE-FAs obtained from incubation with L-[14CH3] methionine confirmed that the ring methylene group was derived from the methyl group of methionine. The time variation in the distribution of radioactivity in the products of incubation with [14CH3] methionine and [2-14C] acetate demonstrated that the pathway involved initial formation of dihydrosterculic acid from oleic acid with subsequent desaturation to sterculic acid and α-oxidation to malvalic and dihydromalvalic acids. It is confirmed that methionine, presumably as S-adenosyl methionine, is the methylene donor. The conversion of oleic acid to dihydrosterculic and sterculic acids has been demonstrated confirming the suggestion of Hooper and Law (1965) and Johnson et al. (1967). Wilson et al. (1961) demonstrated the cooccurrence of sterculic and malvalic acids and the corresponding CPA-FAs in various seeds and suggested that methylene addition to oleic acid gives rise to dihydrosterculic acid, which was desaturated to sterculic acid. 8-Heptadecenoic acid was similarly the precursor of dihydromalvalic acid and malvalic acid. The route of synthesis of malvalic acid is of interest because it forms the major cyclopropene component in some seeds due to the presence of ring in the 8,9-position of fatty acid chain. From the observed activities of dihydromalvalic and malvalic acids, it is likely that dihydromalvalic acid is the immediate precursor of malvalic acid but it is not evident whether the dihydromalvalic acid produces by the α-oxidation of dihydrosterculic acid or from the addition of a methylene group to a C17-monounsaturated fatty acid (Martin and Stump, 1959). The C17-monounsaturated fatty acid is present in small amounts in seeds and could be formed from α-oxidation of oleate (Smith and Bu’Lock, 1964). The labeled C17-CPA-FA could have been formed either from palmitoleic acid or from dihydrosterculic acid by chain

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shortening by β-oxidation. In is interesting to note that the results have shown similar biosynthetic pathways for the formation of CPE-FAs for all three types of seeds and seed oils examined, namely those rich in sterculic acid or malvalic acid, and those with a low CPE-FA content.

5.5 MASS SPECTROMETRY OF CYCLOPROPENE FATTY ACIDS The bond strain in three-membered rings of CPE-FAs makes the purification and derivatization difficult. Derivatization is done to improve the stability and other useful characteristics, and that helps in studies of the structure determination (Hooper and Law, 1968). Different types of derivatives were prepared for MS of CPE-FAs, such as the conversion to the diketo acid by ozonolysis, addition of thiols, and catalytic reduction to the saturated cyclopropane compounds (Fig. 5.12). The 1,3-diketo fatty acids prepared by ozonization of CPE-FAs and were purified by silicic acid chromatography. The diketo acids are crystalline solids and stable compounds. The esters of various chain lengths can be separated by gas-liquid chromatography (GLC) or reverse-phase HO C

(CH 2)n

H C

H C

(CH 2)7

CH 3

C

C H2

O

O

O

C

C

HO (CH 2)n

(CH 2)7

CH 3

C H2

O

Pd/CaCO3 H2

1. O3 2. Pd/CaCO3

HO C

(CH2 )n

C

C

(CH 2)7

CH 3

C H2

O

CH3SH

H

SCH 3 H

SCH3

HO C O

(CH 2)n

C

C C H2

(CH 2)7

CH 3

+

HO C O

(CH 2)n

C

C

(CH 2)7

CH 3

C H2

FIGURE 5.12 Derivatives of CPE-FAs for MS: malvalic acid (n 5 6); sterculic acid (n 5 7). Adapted from Hooper and Law, 1968. Mass spectrometry of derivatives of cyclopropane fatty acids. J. Lipid Res. 9, 270275.

166

Fatty Acids

chromatography, and mass spectra of these esters are excellent method for locating the position of cyclopropene ring in fatty acid chain (Morris and Hall, 1967). The addition of methane thiol across the double bond of the ring gives stable derivatives and can be recovered from gas chromatographic effluents and useful in gas chromatographic separations (Raju and Raiser, 1966). The addition of thiol to the double bond gives an unresolved mixture of isomers in which the sulfur atom attached at one or the other end of the original double bond. A mass spectrum of the mixture of methane thiol adducts of cyclopropane acid esters makes a definitive assignment of the ring position. The technique is simple and quantitative. The CPE-FAs can also be reduced to less reactive cyclopropane acids by catalytic hydrogenation in the presence of palladium catalyst in alcohol. Further reduction to the ring-opened compounds helps in the assignment of cyclopropene ring by MS (Polacheck et al., 1966; McCloskey and Law, 1967). However, this process is complicated than conversion of CPE-FAs to diketones and thiol addition products. Location of the cyclopropene ring from the mass spectrometric studies of diketo and thiol derivatives of sterculic and malvalic acids appears to be a convenient method. Mass fragmentation pattern of diketo esters and methane thiol adduct of sterculic and malvalic acids is reported by Hooper and Law (1968). The simple cleavage at the carbonyl group of the derivatized ester, McLafferty rearrangement (McLafferty, 1959) of the major ion, and loss of methanol from parent ion helps in the location of the ring and chemists should use this as reference spectra.

5.5.1 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropene Fatty Acids It is evident from earlier investigations that seed oils containing CPE-FAs and CPA-FAs are difficult to quantitatively analyze by GLC and hydrogen bromide titration methods. Because of high column temperature, CPE-FAs methyl esters tend to isomerize and decompose as they pass through GLC column (Recourt et al., 1967). In addition, GLC data show that malvalic acid peak is masked by the linoleic (C18:2) acid peak (Wilson et al., 1961) and that the corresponding CPA-FA may also be obscured by the presence of oleic acid (Miwa, 1963). GC-MS studies of the silver nitratemethanol treated methyl esters of naturally occurring CPE-FAs present in the seed glycerides helps in the unequivocal characterization of individual cyclopropene, sterculic, and malvalic acids (Ahmad et al., 1979). A detailed mass spectral study of the seed glyceride of E. hookeriana was reported to characterize the component CPE-FAs. Glycerides were converted to methyl esters by transesterification with 1% methanolic sodium methoxide. The cyclopropene esters were converted into their silver nitrate derivatives by treating methyl esters with an excess of anhydrous methanol saturated with silver nitrate at room temperature overnight

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167

(Schneider et al., 1968). The mass spectra of silver nitrate derivatives of methyl sterculate and methyl malvalate (Scheme 5.3) confirmed the GC identification. In addition, the spectra of the normal esters showed the appropriate molecular ions. The GC-MS study presented here is of a binary mixture. H 3C(H2C)7

C

C

(CH3) nCOOCH 3

C H2

n = 6 (Malvalic) n = 7 (Sterculic)

AgNO3 MeOH

CH2

CH2 OCH3 H 3C(H2C)7

C H

C

(CH2) 6COOCH3

H 3C(H2C)7

( 1)

( 5)

+

H C

O

O

C

(CH2) 6COOCH3

H 3C(H2C)7

( 2)

C

C

( 6)

+ CH2

CH2 OCH3 C

C H

(CH2) 7COOCH3

H 3C(H2C)7

C

C

( 3)

( 7) O

+

+

H C

( 4)

C

(CH2) 7COOCH3

(CH2) 7COOCH3

O

CH2 OCH3 H 3C(H2C)7

(CH2) 6COOCH3

CH2

+

H 3C(H2C)7

(CH2) 6COOCH3

+ CH2 OCH3

H 3C(H2C)7

C

C

H 3C(H2C)7

C

( 8)

C

(CH2) 7COOCH3

CH2

SCHEME 5.3 Silver nitrate derivatives of cyclopropene acids for MS. Adapted from Ahmad et al., 1979. Eriolaena hookeriana seed oil: a rich source of malvalic acid. Chem. Phys. Lipids 25, 2938.

Mass spectra of ether derivatives (1, 2 and 3, 4) showed the molecular ion peaks at m/e 326 (for malvalic) and m/e 340 (for sterculic), respectively. The diagnostic ions (IIV), which indicate the position of cyclopropene ring, are important components in each spectrum (Scheme 5.4). A characteristic peak at m/e 152 (V) was also observed, which is believed to arise from cleavage of ether derivative with respect to side-chain ether group (Scheme 5.5).

CH2OCH3 H3C(H2 C)7

C

H C

M n = 6 (m/e = 326) n = 7 (m/e = 340)

(CH 2)nCOOCH3

(1 and 3)

CH2OCH3

CH2OCH3

H C

C

I

(CH 2)nCOOCH3

H3C(H2 C)7

n = 6 (m/e = 213) n = 7 (m/e = 227)

CH

C

II (m/e = 183)

CH2 H3C(H2 C)7

CH2OCH3 H C

C

CH2CH=CH2

H3C(H2 C)7

III (m/e = 193)

H C

C

CH2

IV (m/ e = 197)

SCHEME 5.4 Mass fragment ions of ether derivatives (I and III) used for diagnostic purpose. Adapted from Ahmad et al., 1979. Eriolaena hookeriana seed oil: a rich source of malvalic acid. Chem. Phys. Lipids 25, 2938.

CH2OCH3 H3C(H2 C)7

C

H C

H2C (CH2) nCOOCH 3

H3C(H2 C)7

C

OCH 3 (CH2) nCOOCH 3

C H

(M)

CH2 H3C(H2 C)7

C

H2C H C

OCH3

C

H3C(H2 C)7

OCH 3 CH

(m/ e = 183) CH2 CH2 H3C(H2 C)7

C

—OCH3 H C

OCH3

H3 C(H 2C) 7

C

CH

V (m/ e = 152) SCHEME 5.5 A characteristic fragment ion (m/e 5 152) of ether derivative. Adapted from Ahmad et al., 1979. Eriolaena hookeriana seed oil: a rich source of malvalic acid. Chem. Phys. Lipids 25, 2938.

169

Carbocyclic Fatty Acids: Chemistry and Biological Properties Chapter | 5

The molecular ion and fragmentation pattern of two ether isomers (2 and 4) of sterculic and malvalic acids are also very similar (Scheme 5.6). A characteristic peak at m/e 152 (V) arises from these two isomers.

CH2 OCH3 H3C(H2 C)7

C H

C

M n = 6 (m/e = 326) n = 7 (m/e = 340)

(CH2)nCOOCH 3

(2 and 4)

CH2 OCH3

CH2 OCH3 HC

C

(I')

H3C(H2 C)7

(CH 2)nCOOCH3

n = 6 (m/e = 213) n = 7 (m/e = 227)

C

C H

(II') (m/e = 183)

CH2 H3C(H2 C)7

C H

C

CH2 OCH3 CH2 CH=CH2

(III') (m/ e = 193)

H3C(H2 C)7

C H

C H

C

(II') (m/ e = 183)

CH2

(IV') (m/ e = 197)

CH2 OCH3 H3C(H2 C)7

C

CH 2

—OCH3

H3 C(H 2C) 7

C H

C

(V') (m/e = 152)

SCHEME 5.6 Mass fragment ions of ether derivatives (II and IV) used for diagnostic purpose. Adapted from Ahmad et al., 1979. Eriolaena hookeriana seed oil: a rich source of malvalic acid. Chem. Phys. Lipids 25, 2938.

170

Fatty Acids

Similarly, the cleavage on either side of the keto group in ketone derivatives (58) (Scheme 5.3) further supports the position of cyclopropene ring in the fatty ester chain. The fragment pattern of the 8-keto derivative (5) showed a weak molecular ion peak of m/e 310. The diagnostic ion peaks at m/e 143 (VI) (Scheme 5.7) arise from m/e 310, by the preferred cleavage α-to-keto group, are alone sufficient to locate the cyclopropene ring at 8,9-position on the chain. The other fragment ion peak at m/e 139 (VII) corresponds to cleavage on the other side of the keto group and supports the position of cyclopropene ring at 8,9-position. The isomeric ketone (6) also supports the cyclopropene ring between C8 and C9 position. The diagnostic ion peak at m/e 141 (VIII) corresponds to the cleavage between α,β-unsaturated ketone.

CH2 H 3C(H2 C)7

C

O (CH 2)6 COOCH3

C

H2C

(CH 2)5 C

OCH 3

+

H 3C(H2 C)7

C

CH2

O

(V) M (m/ e = 310)

(VII) (m/ e = 139)

(VI) (m/ e = 143)

O H 3C(H2 C)7

C

C

H 3C(H2 C)7

(CH 2)6 COOCH3

C

O

CH2

(VIII) (m/e = 141)

M (m/e = 310)

CH2 H 3C(H2 C)7

C

CH2 C

(CH 3)7 COOCH3

H 3C(H2 C)7

C

CH2

+

H 3C(H2 C)7

C

C

O

O

M (m/e = 324)

(VII) (m/ e = 139)

(IX) (m/e = 167)

SCHEME 5.7 Mass fragment ions of keto derivatives (V and VIII) used for diagnostic purpose. Adapted from Ahmad et al., 1979. Eriolaena hookeriana seed oil: a rich source of malvalic acid. Chem. Phys. Lipids 25, 2938.

The mass spectrum of 9-keto derivative (7) of sterculic acid, showed an intense ion peak at m/e 167 (IX), corresponds to one of the fragments of α-keto cleavage, correctly positions the cyclopropene ring at 9,10-position. Similarly, the diagnostic ion peak at m/e 141 (VIII), corresponding to the cleavage between α,β-unsaturated ketone from the isomeric ketone

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171

(8), further supports the cyclopropene ring between C9 and C10 position. The mass spectral data of the two binary mixtures of the ether derivatives (14) and mono-ketone derivatives (58) of cyclopropene esters (Scheme 5.3) convincingly position the ring at the 8,9-position (methyl malvalate) and 9,10-position (methyl sterculate). The chemists working in this area should use this method for the characterization of cyclopropene ring in seed glyceride or synthetic mixtures in milligram quantity. The GC-MS has been so far the method of choice for identifying the CPE-FAs and CPA-FAs in fats and oils present naturally or produced during heating or similar treatment.

5.5.2 Gas Chromatography-Mass Spectrometry Analysis of Cyclopropane Fatty Acids Food authentication represents an important issue for the food industry because consumers are becoming interested in the quality and origin of food. Consumers are more interested in the quality and origin of foods, such as organic, protected denomination of origin (PDO), protected geographical indication (PGI) products. The higher prices of PDO products encourage bringing more counterfeiting products in the market especially in the dairy sector. The most important issue in the authentication is related to PDO cheeses, which are high commercial-value products. The cheese production can differ according to the feeding system of the animals providing the milk, the starters used, the heating temperature, the salting, the use or lack of use of preservatives, and the ripening time. All these parameters generate defined characteristics and can be detected by analytical techniques. CPA-Fas, such as lactobacillic acid and dihydrosterculic acid, are components of bacterial membranes and have been recently detected in milk and dairy products from cows fed with corn silage. GC-MS method for the detection of CPA-FAs in cheese, as new molecular markers, is recently developed (Caligiani et al., 2016). Limit of detection and quantitation of CPA-FAs were, respectively, 60 and 200 mg kg21 of cheese fat. The analysis of CPAFAs can be a useful tool for the quality control of PDO cheeses, such as Parmigiano Reggiano, whose specifications of production do not allow the use of silages. The presence of CPA-FAs could be used as one of the markers of Parmigiano Reggiano authenticity.

5.6 PHYSIOLOGICAL PROPERTIES OF CYCLOPROPENE FATTY ACIDS A variety of biological effects have been observed in several species of animals upon ingestion of small quantities of CPE-FAs. Two storage disorders “pink white” and “pasty yolk” are known to develop in stored eggs associated with the feeding of CPE-FAs containing substances like cottonseed oil, cottonseed meal, S. foetida oil (containing both sterculic and malvalic acids).

172

Fatty Acids

The pink-white condition is related to increased diffusion processes in the egg during storage, and the pasty yolk condition is related to an increase in the proportion of saturated to unsaturated fatty acids in yolk (Shenstone and Vickery, 1956, 1959; Masson et al., 1957). The pink-white defect develops during storage and is associated with increased diffusion from the albumen and the yolk. The yolk enlarges at the expense of the albumen and becomes a pink-orange color, proteins are absorbed into the yolk from the albumen, iron migrates from the yolk and reacts with conalbumin of the albumen to form a pink chelate, and its pH also changes. The pasty yolk condition in the affected eggs gradually develops during storage at normal temperature, but it can be induced quickly in any affected egg by the exposure to low temperature. The texture of the yolk is hard to the touch and this is attributed to an increase in the ratio of saturated fatty acids to unsaturated fatty acids (stearic acid vs oleic acid). However, when the cyclopropene ring was hydrogenated, the biological effects were destroyed. A significant positive correlation was also found to exist between the intensity of pink-white discoloration and the concentration of iron (59Fe) in egg albumen. The findings of Abou-Ashour and Edwards (1970) provide evidence that a diffusion of 59Fe from the yolk into the albumen happens and is responsible in pink-white discolored eggs. Bain and Hall (1970) compared the structures of the albumen, the yolk, and the vitelline membrane in new-laid and stored eggs from hens fed a CPEFAs (e.g., methyl sterculate) with those from eggs of hens fed a normal diet and related differences in them to the cause of the pink-white or pasty yolk condition. The fine structure of the inner layer of the vitelline membrane developing around the ova in the ovaries of hens fed methyl sterculate was also compared with that in normal hens, to see any structural differences were present in the vitelline membranes when the yolk was being formed. No significant structural changes were reported in the vitelline membrane to account for the increased diffusion from the albumen and the yolk. This observation supports existing opinion that normal diffusion in the egg is controlled by the physicochemical properties of the yolk rather than by resolvable structure in the egg. It is likely that these barriers to diffusion are altered when CPE-FAs are included in the diet. It has also been suggested that the diffusion process in such eggs may be influenced by the increase in stearic acid as mentioned earlier. A comparison of the fatty acid distribution in the tissues and egg yolk lipids of normal chickens with that of hens fed cottonseed oil or S. foetida seeds indicated that fatty acid metabolism of the hen was disturbed by the CPE-FAs containing materials. It is observed that hens fed the test rations had higher levels of stearic acid in the egg yolk lipids, their livers, blood plasma, and ovaries than hens fed the normal diet. The mechanism regulating the equilibrium that normally existed between stearic and oleic acids was upset so that a greater proportion of stearic acid was produced at the expense of oleic acid.

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A mechanism postulated to account for the biological activity of the cyclopropene compounds was the reaction of cyclopropene ring with sulfhydryl groups in protein, which are closely associated with the lipids. The physical and biochemical properties of the protein molecule are altered by the irreversible addition of protein sulfhydryl groups to the cyclopropene ring (Kircher, 1964). Kircher studied the reaction of methyl mercaptain and 2-mercaptopropanoic acid with cyclopropene compounds and suggested that this phenomenon may have its counterpart in the animal body and could cause the physiological effects when CPE-FAs are ingested. The finding supports the hypothesis that the increase in saturated fatty acids at the expense of the corresponding monoenes observed when animals ingest small amount of CPE-FAs is due to binding of the thiol groups of acyl desaturase by the cyclopropene groups. It is quite apparent that cyclopropene ring can react with active sulfhydryl groups in enzymes. The addition products of sulfhydryl groups with cyclopropenes are stable to acid and alkaline conditions. This suggest that it is possible to isolate the addition products of sulfhydryl enzymes with CPE-FAs as peptides and the analysis of the complex will give valuable information concerning the active center of the enzyme. Raju and Raiser (1967) further investigated to test this hypothesis. They studied the effect of CPE-FAs on the aerobic desaturation of stearic to oleic acid in rats and rat liver extract, and demonstrated inhibition activity of fatty acyl desaturase by CPE-FAs. Further evidence was obtained that the enzyme contains sensitive sulfhydryl groups. There are a number of enzymes known to contain sulfhydryl groups, which are essential for their activity. Some of these enzymes are more sensitive to CPE-FAs than others because some of these sulfhydryl groups are more reactive than others. At lower level CPEFA may attack only the highly reactive thiol groups. At higher concentrations, a number of less reactive thiol groups may be attacked resulting in the inhibition of many vital enzymes and leading to animals’ death. Evidence was obtained that the mechanism of inhibition is the irreversible binding of enzyme sulfhydryl groups with the cyclopropene group. Some of the biological effects of cyclopropene compounds have been reviewed earlier by Phelps et al. (1965).

5.7 CYCLOPROPANEOCTANOIC ACID 2-HEXYL IN HUMAN ADIPOSE TISSUE AND SERUM Sakurada et al. (1999) identified cis-9,10-methylenehexadecanoic acid (also called cyclopropaneoctanoic acid 2-hexyl) in phospholipids of human, rat, bovine heart, and rat liver. Sledzinski et al. (2013) reported: (1) cyclopropaneoctanoic acid 2-hexyl in human and rat adipose tissue and serum, (2) mainly stored as a component of triacylglycerols in human adipose tissue. Other CPA-Fas namely cyclopropaneoctanoic acid 2-octyl, cyclopropanenonanoic acid, and 2-[[2-[2-ethylcyclopropyl)methyl]cyclopropyl]methyl]

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CO2 H

(A)

CO2 H

(B)

CO2 H

(C)

CO2 H

(D) FIGURE 5.13 Chemical structure of CPA-FAs identified in human adipose tissue: cyclopropaneoctanoic acid 2-hexyl (A), cyclopropaneoctanoic acid 2-octyl (B), 2-[[2-[(2-ethylcyclopropyl) methyl]cycylopropyl]methyl] acid (C), and cyclopropanenonanoic acid (D). Adapted from Sledzinski et al., 2013. Identification of cyclopropaneoctanoic acid 2-hexyl in human adipose tissue and serum. Lipids 48, 839848.

(Fig. 5.13) were detected in small amounts (up to 0.05% of total fatty acids) in adipose tissue of some patients and were undetectable in human serum. The presence of cyclopropaneoctanoic acid 2-hexyl in adipose tissue and serum suggests that adipose tissue can take up and release CPA-FAs into circulation. The storage of CPA-FAs in adipose tissue may protect other organs from exposure to excessive CPA-FAs. The question arise about the source of cyclopropaneoctanoic acid 2-hexyl (and other CPA-FAs) (Fig. 5.13) in human adipose tissue and serum. In theory, there are three possibilities. First, CPA-FAs may originate from food. However, cyclopropaneoctanoic acid 2-hexyl was not detected in lipids extracted from laboratory food fed to rats. Second, cyclopropaneoctanoic acid 2-hexyl could originate from intestinal bacteria (Wood and Reiser, 1965). This CPA-FA was not detected in the rat intestinal content. It cannot be excluded that CPA-FA is present in food and is produced by intestinal bacteria at low level, below the limit of detection in GC-MS analysis. Therefore, it is possible that CPA-FA, even if consumed and/or produced by intestinal bacteria at a very low level, can accumulate in adipose tissue. Third, there is possibility that cyclopropaneoctanoic acid 2-hexyl acid could be synthesized by some organ/tissue in human and rat body. However, at present it is difficult to establish the source of CPA-FA in human adipose tissue. The fatty acid composition of adipose tissue is a reliable biomarker for long-term dietary intake of fatty acids

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(Hodson et al., 2008) and one can assume that the main source of CPA-FA in adipose tissue is the consumed fat. Even if ingested at low levels, mainly in the form of plants, dairy products, and ruminant (beef) meat, the CPA-FA could accumulate in adipose tissue and be released into circulation. However, more thorough research is needed to establish unequivocally the main source of CPA-FA in human adipose tissue. The presence of cyclopropaneoctanoic acid 2-hexyl both in the blood and adipose tissue suggests that this CPA-FA can be taken up and released by adipose tissue. The published results indicate that CPA-FA is present not only in bacteria, plants, protozoa, and Myriapoda but also in mammalian tissues including humans, and need further research to gain more information about the source and a possible pathophysiological significance of CPA-FA accumulation in human adipose tissue.

5.7.1 Cyclopropaneoctanoic Acid 2-Hexyl in Patients With Hypertriglyceridemia As described earlier, Sledzinski et al. (2013) reported four CPA-FAs, cyclopropaneoctanoic acid 2-hexyl, cyclopropaneoctanoic acid 2-octyl, cyclopropanenonanoic acid, and 2-[[2-[(2-ethylcyclopropyl)methyl)methyl] cyclopropyl]methyl] acid, in the adipose tissue of obese women. The cyclopropaneoctanoic acid 2-hexyl (also known as cis-9,10-methylenehexadecanoic acid) was the most abundant CPA-FA and the only CPA-FA detectable in their serum. The CPA-FA presents in bacteria and plants is synthesized from unsaturated fatty acid by cyclopropane synthase, an enzyme catalyzing the addition of the methylene group from S-adenosylmethionine to double bond of unsaturated fatty acid (Bao et al., 2002). However, this enzyme has not been identified in animals. The same research group studied whether the presence of CPA-FA was obesity-specific (Mika et al., 2016). To prove this, they determined serum levels of cyclopropaneoctanoic acid 2-hexyl in: (1) nonobese controls, (2) obese patients, (3) obese persons on low-calorie diet for 3 months, and (4) individuals with CKD, that is, individuals with a disease related to dyslipidemia. Obese patients and those with CKD presented with higher serum levels of cyclopropaneoctanoic acid 2-hexyl than controls. Switching obese individuals to a low-calorie (low-lipid) diet resulted in a reduction of cyclopropaneoctanoic acid 2-hexyl concentration to the level observed in controls. Patients with hypertriglyceridemia-related conditions presented with elevated serum levels of cyclopropaneoctanoic acid 2-hexyl. The result showed that hypertriglyceridemia observed during the course of diseases such as CKD and obesity is associated with an increase in serum concentration of cyclopropaneoctanoic acid 2-hexyl. This also suggest that high serum level is related to high serum triacylglycerol concentrations rather than body mass or body mass index.

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5.8 LEISHMANIA CYCLOPROPANE FATTY ACID SYNTHETASE Leishmania are obligate intracellular protozoan parasites that infect humans and other mammalian species causing broad spectrum of diseases called the leishmaniosis. Parasites are transmitted as extracellular flagellated forms by female sandflies during blood feeding (Sacks, 2001). Maintenance of parasites at dermal sites or subsequent dispersal to internal tissues contributes to disease progression causing cutaneous leishmaniasis (CL), mucocutaneous leishmaniasis (MCL), diffuse cutaneous leishmaniasis (DCL), and visceral leishmaniasis (VL) (Murray et al., 2005; Kaye and Scott, 2011). These diseases are associated with particular parasite species, such as Leishmania infantum and Leishmania major usually causing VL and CL, respectively, and Leishmania braziliensis is a major causative agent of MCL. The single gene encoding cyclopropane fatty acid synthetase (CFAS) is present in L. infantum, Leishmania mexicana, and L. braziliensis but absent from L. major, a causative agent of cutaneous leishmaniasis. In L. infantum, the causative agent of visceral leishmaniasis, the CFAS gene is transcribed in both insect (extracellular) and host (intracellular) stages of the parasite life cycle. Lipid analysis of L. infantum wild type, CFAS null, and complemented parasites detect a low abundance CFAS-dependent C19 CPA-FA, in wild type and add-back cell. Subcellular fractionation studies locate the C19 CPA-FA in plasma membraneenriched fractions. This fatty acid was not detectable in wild type L. major, although expression of the L. infantum CFAS gene in L. major generates CPA-FAs, indicating that the substrate for this modification is present in L. major, despite the absence of the modifying enzyme. Following infection in vivo, the L. infantum CFAS nulls exhibit lower parasite burdens in both the liver and spleen of susceptible hosts but it has not been possible to complement this phenotype, suggesting that loss of C19 CPA-FA may lead to irreversible changes in cell physiology that cannot be rescued by reexpression. A CFAS enzyme catalyzes the cyclopropanation of unsaturated fatty acids, and involves the transfer of a methylene group from an S-adenosyl-Lmethionine to a carboncarbon double bond within fatty acyl chain (Yuan and Barry, 1996). Although the position of the cis double bond on the acyl chain is variable in E. coli, Mycobacterium tuberculosis produces several site-specific cyclopropane synthetases that modify mycolic acids (Huang et al., 2002). CFAS-catalyzed membrane modifications have been associated with stress responses to change in pH, temperature, or salinity of the local environment in E. coli (Grogan and Cronan, 1997). Studies in E. coli have shown that increases in CFAS activity, leading to increased CPA-FA content, are associated with changes in environmental conditions such as exposure to high temperature, low pH, high salt concentration, depressed oxygen tension, and support the modification functions as a cellular survival mechanism

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(Knivett and Cullen, 1965; Shabala and Ross, 2008). For example, Helicobactor pylori, which colonizes the mammalian gut and is associated with reduced gastric activity, secretes large amounts of CPA-FA (cis-9,10methyleneoctadecanoic acid) in comparison to other bacterial species that also colonize the intestinal tract (Haque et al., 1996). The CPA-FAs produced by the gastric colonizers help in inhibiting the gastric H 1 /K (1)-ATPase proton pump, and reduced acidity in the infected regions (Beil et al., 1994). Oyola et al. (2012) reported the first functional characterization of CFAS in pathogenic Leishmania species, and focused on L. infantum, causative agent of VL. It is suggested that C18:1 fatty acid substrate may be ubiquitous in Leishmania species, while its modification by cyclopropanation is a species-specific property. The accumulation of cyclopropanated product in an unidentified subcellular compartment in L. infantum may help in the identification of the CFAS substrate in promastigotes. Although the physiological role of cyclopropane modification has not been fully defined in any species, the expression of CFAS in many bacteria and the sporadic distribution of the gene in a few phylogenetically unrelated eukaryotes suggests that different organisms use this modification to facilitate adaptation to environmental conditions and undergo developmental processes requiring changes in membrane structure and function. Cyclopropanation of fatty acids has been associated with drought tolerance in plants (Kuiper and Stuiver, 1972) and egg development in millipedes (Oudejans et al., 1976), while in bacteria the modification has consistently been linked to acid tolerance (Grogan and Cronan, 1997). For intracellular pathogens, cyclopropanation may play a role in survival in physiologically hostile and nutrient-poor compartments within the host cell. This would be of particular relevance to Leishmania species, which have the capacity to survive and replicate in the acidic and nutrient-poor compartments. From the functional analysis, it is clear that only L. major of the Leishmania species currently analyzed has lost the CFAS gene and does not produce this enzyme remains unknown, and suggests that biological aspects of the intracellular survival of this species are uniquely specialized (Oyola et al., 2012).

5.8.1 Leishmania: A Fungal Infection Among the fungal infections, CL is the most prominent type occurring in human where fungi colonize on dead tissue of the stratum corneum and is called dermatophyte. These types of fungi do not produce deep cutaneous or systemic infections. The presence of CPA-FAs in Leishmania species and its direct effect in causing diseases is unknown. This area needs to be explored for future research. Several treatment options are available today for cutaneous fungal infection (Kyle and Dahl, 2004). A number of drugs are available for the treatment of fungal and yeast skin infections. Many antifungal agents are compounded in different type of excipients (vehicles)

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and have been found to be effective. Most commonly, topical drugs are applied to the surface of the skin in the form of cream, lotion, or spray that can be easily penetrated into the skin and prevent them from spreading of infection to the tissues. Amphotericin B has long been a gold standard for the treatment of patience with invasive fungal infections (Sheikh et al., 2010). Amphotericin B has higher affinity for ergosterol than for cholesterol, which results in its binding to fungal, leishmanial, or negleria cells as they contain ergosterol or structurally similar compounds. The resulting ergosterolamphotericin B complex increases membrane permeability of fungal/pathogen cells leading to cell lysis (Neumann et al., 2009). However, amphotericin B maximal utilization in clinical practice is restricted due to its severe toxicity to the kidney and red blood cells that was found to be overcome by using lipid-based delivery system for parental use (Ahmad et al., 1990, 1991). There are certain drawbacks associated with conventional topical delivery of drugs and to circumvent the drawback, the lipid-based carriers of therapeutic products were used as delivery system. The small size of lipid nanoparticles ensures a close contact to the stratum corneum and can increase the amount of drug penetrated into the skin (Dubey et al., 2012). Lipid nanoparticles are able to enhance the chemical stability of compounds sensitive to light, oxidation, and hydrolysis. Usages of lipid as carrier systems for skin administration are related to their physiological nature, which reduces the risk of toxicological problems and local irritancy (Muller et al., 2002). Considering the benefits of using lipids as carrier for topical drug delivery, a novel topical formulation of lipidbased gel formulation of amphotericin B (0.1% amphotericin B gel) was developed in author’s laboratory (Sheikh et al., 2014). In this gel formulation, natural lipids were used to reduce adverse reactions and make it compatible to the skin, and minimize allergic reactions. The newly developed lipid-based amphotericin B gel was highly effective to treat CL and MCL fungal infections, and was found safe, tolerable, and efficacious.

5.9 CONCLUSION Fatty acids containing three-membered carbocyclic rings, namely CPE-FAs and CPA-FAs, are distributed across several plant orders, most notably the Malvales, which includes cottonseed and sterculia. Dihydrosterculic acid is present up to 60% of total fatty acids in L. chinensis seed oil. Long-chain CPA-FAs have also present in various polar lipid classes in root, leaf, stem, and callus tissue in plants of the Malvaceae where they may function in resistance to fungal attack. CPA-FAs have similar melting points and fluidity compared to oleic acid and the absence of the double bond greatly increases their oxidative stability. These features suggest that CPA-FAs may be used as biodegradable lubricants and in other applications including biodiesel. To achieve as a petroleum replacement, these oils must be produced in large quantities at low cost.

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In recent years, cyclopropane derivatives have been attracted more interest because of their biological and pharmaceutical applications. Cyclopropane analogs have known to exhibit diverse pharmacological applications that created enormous interest in bioorganic, medicinal, and pharmaceutical chemistry. The unique reactivity of cyclopropanes due to the high level of strain offers considerable utility in organic synthesis. Designing small molecules that bind to therapeutically important biological targets with high affinity and selectivity is a major goal in contemporary bioorganic and medicinal chemistry. The reactivity of cyclopropanes allows them as versatile intermediates in the synthesis of complex molecules and is frequently employed as versatile building blocks in organic synthesis. Food quality is also becoming an important issue in the food industries because consumers are increasingly interested in the quality and origin of foods. The higher prices of products from PDO, organic foods, or PGI encourage more frequent counterfeiting. CPA-FAs such as lactobacillic acid and dihydrosterculic acid are components of bacterial membranes and have been found in milk cows fed maize silage. The analysis of CPA-FAs may be a useful tool for the quality control for PDO cheeses, whose specifications of production do not allow the use of silages, as well as marker to differentiate high-quality dairy products.

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McCloskey, J.A., Law, J.H., 1967. Ring location in cyclopropane fatty acid esters by a mass spectrometric method. Lipids 2, 225230. McLafferty, F.W., 1959. Mass spectrometric analysis. Molecular rearrangements. Anal. Chem. 31, 8287. Mika, A., Stepnowski, P., Chmielewski, M., Malgorzewicz, S., Kaska, L., Proczko, M., et al., 2016. Increased serum level of cyclopropaneoctanoic acid 2-hexyl in patients with Hypertriglyceridemia-related disorders. Lipids 51, 867873. Minnikin, D.E., 1966. Nuclear magnetic resonance spectra of long chain 1, 2-disubstituted cyclopropane esters. Chem. Ind.2167. Miwa, T.K., 1963. Identification of peaks in gas-liquid chromatography. J. Am. Oil Chem. Soc. 40, 309313. Morris, L.J., Hall, S.W., 1967. Chem. Ind. (Lond.) 32. Muller, R.H., Radtke, M., Wissing, S.A., 2002. Solid lipid nanoparticles (SLN) and nanostructured lipid carriers (NLC) in cosmetic and dermatological preparations. Adv. Drug Deliv. Rev. 54 (Suppl. 1), S131S155. Murray, H.W., Berman, J.D., Davies, C.R., Saravia, N.G., 2005. Advances in leishmaniasis. Lancet 366, 15611577. Mustafa, J., Gupta, A., Ahmad Jr., M.S., Ahmad, F., Osman, S.M., 1986. Cyclopropenoid fatty acids in Gnetum scandens and Sterculia pallens seed oils. J. Am. Oil Chem. Soc. 63, 11911192. Neumann, A., Czub, J., Baginski, M., 2009. On the possibility of the amphotericin B-sterol complex formation in cholesterol- and ergosterol-containing lipid bilayers: a molecular dynamics study. J. Phys. Chem. B. 113, 1587515885. Nolen, G.A., Alexander, J.C., Artman, N.R., 1967. Long-term rat feeding study with used frying fats. J. Nutr. 93, 337349. Nordbay, H.E., Heywang, B.W., Kircher, H.W., Kemmerer, A.R., 1962. Sterculic derivatives and pink egg formation. J. Am. Oil Chem. Soc. 39, 183185. Nunn, J.R., 1952. The structure of Sterculic acid. J. Chem. Soc. 313. Oudejans, R.C., van der Horst, D.J., Opmeer, F.A., Tieleman, W.J., 1976. On the function of cyclopropane fatty acids in millipedes (Diplopode). Comp. Biochem. Physiol. B 54, 227230. Oyola, S.O., Evans, K.J., Smith, T.K., Smith, B.A., Hilley, J.D., Mottram, J.C., et al., 2012. Functional analysis of Leishmania cyclopropane fatty acid synthase. PLoS One. 7 (12), e51300. Pasha, M.K., Ahmad, F., 1992. Analysis of triacylglycerols containing cyclopropane fatty acids in Sterculia fostida (Linn.) seeds lipids. J. Agric. Food. Chem. 40, 626629. Pawlowski, N.E., Eisele, T.A., Ahmad, M.U., Nixon, J.E., Bailey, G.S., 1983. Metabolism of sterculic acid, a cyclopropenoid fatty acid. In: 185th National Meeting, American Chemical Society, AGFD 18. Phelps, R.A., Shenstone, F.S., Kemmerer, A.R., Evans, R.J., 1965. A review of cyclopropenoid compounds; biological effects of some derivatives. Poult. Sci. 44, 358394. Polacheck, J.W., Tropp, B.E., Law, J.H., McCloskey, J.A., 1966. Biosynthesis of cyclopropane compounds. J. Biol. Chem. 241, 33623364. Potteau, B., Lhuissier, M., leelere, J., Custot, F., Mezonnet, R., Cluzan, R., 1970. Composition and physiological effects of heated soybean oil and of its various fractions. 1. Chemical study. Rev. Fr. Corps Gras. 17, 143153. Raju, P.K., Raiser, R., 1966. Gas-liquid chromatographic analysis of cyclopropane fatty acids. Lipids 1, 1015.

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Raju, P.K., Raiser, R., 1967. Inhibition of fatty acyl desaturase by cyclopropene fatty acids. J. Biol. Chem. 242, 379384. Recourt, J.H., Jurriens, G., Schmitz, M., 1967. Analysis of cyclopropenoid acids by gas-liquid. J. Chromatogr. A 30, 3542. Rinehart Jr., K.L., Nilsson, W.A., Whaley, H.A., 1958. Sterculic acid: nuclear magnetic resonance spectrum and structure. J. Am. Chem. Soc. 80, 503504. Rinehart Jr., K.L., Tarimu, C.L., Culbertson, T.P., 1959. Structure of sterculic acid polymer. A cyclopropane rearrangement. J. Am. Chem. Soc. 81, 50075008. Rinehart Jr., K.L., Goldberg, S.I., Tarimu, C.L., Culbertson, T.P., 1961. Cyclopropene rearrangement in the polymerization of sterculic acid. J. Am. Chem. Soc. 83, 225231. Roehm, J.N., Lee, D.J., Wales, S.D., Polityka, S.D., Sinnhuber, R.O., 1970. The effect of dietary sterculic acid on the hepatic lipids of rainbow trout. Lipids 5, 8084. Roomi, M.W., Hopkins, C.Y., 1970. Some reactions of sterculic and malvalic acids. A new source of malvalic acid. Can. J. Biochem. 48, 759762. Sacks, D.L., 2001. Leishmania-sand fly interactions controlling species-specific vector competence. Cell Microbiol. 3, 189196. Sakurada, K., Iwase, H., Takatori, T., Nagao, M., Nakajima, M., Niijima, H., et al., 1999. Identification of cis-9,10-methythylenehexadecanoic acid in submitochondrial particles of bovine heart. Biochim. Biophyss. Acta 1437, 214222. Schmid, K.M., Patterson, G.W., 1988a. Distribution of cyclopropenoid fatty acids in malvaceous plant parts. Phytochemistry 27, 28312834. Schmid, K.M., Patterson, G.W., 1988b. Effects of cyclopropenoid fatty acids on fungal growth and lipid composition. Lipids 23, 248252. Schneider, E.L., Sook, L.P., Hopkins, D.T., 1968. Gas-liquid chromatographic analysis of cyclopropenoid fatty acids. J. Am. Oil Chem. Soc. 45, 585590. Shabala, L., Ross, T., 2008. Cyclopropane fatty acids improve Escherichia coli survival in acidified minimal media by reducing membrane permeability to H 1 and enhanced ability to extrude H 1 . Res. Microbiol. 159, 458461. Shah, S., White, J.M., Williams, S.J., 2014. Total synthesis of cis-cyclopropane fatty acids: dihydromalvalic acid, dihydrosterculic acid, lactobacillic acid, and 9,10-methylenehexadecanoic acid. Org. Biomol. Chem. 12, 94279438. Sheikh, S., Ali, S.M., Ahmad, M.U., Ahmad, A., Mushtaq, M., Ahmad, I., 2010. Nanosomal amphotericin B is an efficacious alternative to Ambisome for fungal therapy. Intl. J. Pharm. 397, 103108. Sheikh, S., Ahmad, A., Ali, S.M., Paithankar, M., Barkate, H., Raval, R.C., et al., 2014. Topical delivery of lipid based amphotericin B gel in the treatment of fungal infection: a clinical efficacy, safety, and tolerability study in patients. J. Clin. Exp. Dermatol. Res. 5, 15 (1000248). Shenstone, F.S., Vickery, J.R., 1956. A biologically active fatty acid in Malvaceae. Nat. Lond. 177, 94. Shenstone, F.S., Vickery, J.R., 1959. Substances in plants of the order Maivales causing pink whites in stored eggs. Poult. Sci. 38, 10551070. Shenstone, F.S., Vickery, J.R., 1961. Occurrence if cyclo-propene acids in some plants of the order Malvales. Nature 190, 168. Simmons, H.E., Smith, R.D., 1959. A new synthesis of cyclopropanes. J. Am. Chem. Soc. 81, 42564264. Sinnhuber, R.O., Wales, J.H., Lee, J.L., Ayres, T.W., Hunter, J., 1968a. Dietery factors and hepatoma in rainbow trout (Salmo gairdneri) II. Cocarcinogenesis by cyclopropenoid fatty acids and the effect of gossypol and altered lipids in Aflatoxin-induced liver cancer. J. Natl Cancer Inst. 41, 12931299.

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Sinnhuber, R.O., Lee, D.J., Wales, J.H., Ayres, J.L., 1968b. Dietary factors and hepatoma in rainbow trout (Salmo gairdnerii). Cocarcinogenesis by cyclopropenoid fatty acids and effect of gossypol and altered lipids on aflatoxin-induced liver cancer. J. Natl. Cancer Inst. 41, 12931299. Sinnhuber, R.O., Lee, D.J., Wales, J.H., Landens, M.K., Keyl, A.C., 1974. Hepatic carcinogenesis of aflatoxin M1 in rainbow trout (Salmo gairdnerii) and its enhancement by cyclopropane fatty acids. Cancer Res. 53, 12581291. Sledzinski, T., Mika, A., Stepnowski, P., Proczko-Markuszewska, M., Kaska, L., Stefaniak, T., et al., 2013. Identification of cyclopropaneoctanoic acid 2-hexyl in human adipose tissue and serum. Lipids. 48, 839848. Smith Jr., C.R., 1970. Occurrence of unusual fatty acids in plants. Progress Chem. Fats Other Lipids 11, 139177. Smith Jr., C.R., Wilson, T.L., Mikolajczak, K.L., 1961. Occurrence of malvalic, sterculic, and dihydrosterculic acids together in seed oils. Chem. Ind. (Lond.) 256257. Smith, G.N., Bu’Lock, J.D., 1964. Biogenesis of cyclopropene acids. Biochem. Biophys. Res. Commun. 17, 433436. Stuart, L.J., Buist, P.H., 2004. The absolute configuration of methyl dihydrosterculate: an unusual phytofatty acid isolated from the seed oil of Litchi clinensis. Tetrahedr. Asymm. 15, 401403. Tashiro, T., Akasaka, K., Ohrui, H., Fattorusso, E., Mori, K., 2002. Determination of the absolute configuration at the two cyclopropane moieties of plakoside A, an immunosuppressive marine galactosphingolipid. Eur. J. Org. Chem. (21), 36593665. Tumbelaka, L.I., Slayden, O.V., Stormshak, F., 1994. Action of cyclopropenoid fatty acid on the Corpus luteum of pregnant and nonpregnant ewes. Biol. Reprod. 50, 253257. Van de Loo, F.J., Fox, B.G., Somerville, C., 1993. Unusual fatty acids. In: Moore Jr., T.S. (Ed.), Lipid Metabolism in Plants. CRC Press, Boca Raton, FL, pp. 91126. Varma, J.P., Nath, B., Aggarwal, J.S., 1955a. Structure of sterculic acid. Nature 175, 84. Varma, J.P., Nath, B., Aggarwal, J.S., 1955b. Structure of sterculic acid. Nature 176, 1082. Varma, J.P., Nath, B., Aggarwal, J.S., 1957. J. Sci. Ind. Res. (India) 16B, 162. Wilson, T.L., Smith Jr., C.R., Mikolajczak, K.L., 1961. Characterization of cyclopropenoid acids in selected seed oils. J. Am. Oil Chem. Soc. 38, 696699. Wood, R., Reiser, R., 1965. Cyclopropane fatty acid metabolism: physical and chemical identification of propane ring metabolic products in the adipose tissue. J. Am. Oil Chem. Soc. 42, 315320. Yano, I., Morris, L.J., Nichols, B.W., James, A.T., 1972b. The biosynthesis of cyclopropane and cyclopropene fatty acids in higher plants (Malvaceae). Lipids 7, 3545. Yano, I., Nichols, B.W., Morris, L.J., James, A.T., 1972a. The distribution of cyclopropane and cyclopropane acids in higher plants (Malvaceae). Lipids 7, 3034. Yuan, Y., Barry III, C.E., 1996. A common mechanism for the biosynthesis of methoxy and cyclopropyl mycolic acids in Mycobacterium tuberculosis. Proc. Natl Acad. Sci. USA 93, 1282812833. Zalkin, H., Law, J.H., Goldfin, H., 1963. Enzymatic synthesis of cyclopropane fatty acids catalyzed by bacterial extracts. J. Biol. Chem. 238, 12421248.

FURTHER READING Villorbine, G., Roura, L., Camps, F., Joglar, J., Fabrias, G., 2003. Enzymatic desaturation of fatty acids: Δ11desaturase activity on cyclopropane acid probes. J. Org. Chem. 68, 28202829.

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Chapter 6

Modification of Oil Crops to Produce Fatty Acids for Industrial Applications John L. Harwood1, Helen K. Woodfield1, Guanqun Chen2 and Randall J. Weselake2 1

Cardiff University, Cardiff, United Kingdom, 2University of Alberta, Edmonton, AB, Canada

Chapter Outline 6.1 Introduction 188 6.2 Key Aspects of Plant Oil Biosynthesis 189 6.3 Major Oil Crops 194 6.3.1 Oil Palm (Elaeis guineensis) 194 6.3.2 Soybean (Glycine max) 197 6.3.3 Brassica Oilseed Species (Brassica napus, Brassica rapa, Brassica oleracea, Brassica carinata) 201 6.3.4 Sunflower (Helianthus annuus) 206 6.4 Minor Oil Crops 208 6.4.1 Alfalfa (Medicago sativa, Medicago falcata) 209 6.4.2 Almond (Prunus dulcis, Prunus amygdalus, Amygdalus communis) 209 6.4.3 Avocado (Persea americana, Persea gratissima) 209 6.4.4 Blackcurrant (Ribes niger) 209 6.4.5 Borage (Borago officinalis) 209 6.4.6 Borneo Tallow (Shorea stenoptera) 209 6.4.7 Camelina (Camelina sativa) 211

6.4.8 6.4.9 6.4.10 6.4.11 6.4.12

6.4.13

6.4.14 6.4.15 6.4.16 6.4.17 6.4.18 6.4.19 6.4.20 6.4.21 6.4.22 6.4.23

Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00005-2 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

Castor (Ricinus communis) 211 Cocoa (Theobroma cacao) 211 Coconut (Cocos nucifera) 212 Coriander (Coriandrum sativum) 212 Cottonseed (Gossypium hirsutum, Gossypium barbadense) 212 Crambe (Crambe abyssinica, Crambe hispanica) (Section 6.5 Also) 212 Cuphea spp. 212 Dimorphotheca (Dimorphotheca pluvialis) 213 Echium (Echium plantagineum) 213 Flax (Linum usitatissimum) 213 Hazelnut (Corylus avellana) 213 Jatropha curcas 213 Jojoba (Simmondsia chinensis) 214 Lesquerella (Lesquerella fendleri) 214 Maize (Corn; Zea mays) 215 Meadowfoam (Limnanthes alba) 215

187

188

Fatty Acids 6.4.24 Mustard (Brassica alba, Brassica carinata, Brassica hirta, Brassica juncea, Brassica nigra) 6.4.25 Oats (Avena sativa) 6.4.26 Olive (Olea europaea) 6.4.27 Peanut (Ground Nut, Arachis hypogaea) 6.4.28 Pine Nuts (Pinus spp.) 6.4.29 Poppy (Papaver somniferum) 6.4.30 Rice (Oryza sativa) Bran Oil

215 215 215 216 216 216 216

6.4.31 Safflower (Carthamus tinctorius) 217 6.4.32 Shea (Butyrospermum parkii, Shea Butter, Karate Butter) 217 6.4.33 Tall 217 6.4.34 Tung (Aleurites fordii) 217 6.4.35 Vernonia Oils 218 6.5 Emerging Industrial Oil Crops 218 6.6 Prospects for Production of Industrial Oils in Vegetative Tissue 222 Acknowledgments 223 References 223

6.1 INTRODUCTION There are over a thousand naturally occurring fatty acids, many of which are present in plants. Broadly speaking, fewer than a dozen are quantitatively important but there is considerable potential to increase the usage of less common acids, especially as specialized chemicals or renewable feedstocks. In this chapter, we focus on the way in which fatty acids, and later, vegetable oils are biosynthesized. We then discuss industrially useful fatty acids and the four main plant sources of oils—oil palm, soybean, rapeseed, and sunflower—followed by notes on other minor crops and their uses. Finally, the future is considered including that of oil crops under development. Much of our discussion relates to the food industry because that is where the vast majority of plant lipids are used. However, in crops such as soybean or oilseed rape, where genetic transformation is relatively easy, their increasing use to produce industrial chemicals is highlighted. Given that consumption of biological oils has been increasing by 5% per year for the last 50 years, it is predictable that demand for oil crops will continue to increase. As a result, boosting oil crop yield and manipulating the quality of oil produced by these crops will be an important research focus in coming years. By understanding how metabolism is controlled and exploiting existing crops better, scientists should be able to make important contributions to the area. This will have environmental as well as economic benefits.

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6.2 KEY ASPECTS OF PLANT OIL BIOSYNTHESIS The biosynthesis of vegetable oil is quite complex. The overall formation can be divided conveniently into fatty acid synthesis and lipid assembly. These biochemical pathways are broadly located in different compartments of the plant cell. De novo fatty acid biosynthesis is localized to the plastids (chloroplasts in green tissue) while lipid assembly through the Kennedy pathway and associated reactions is in the endoplasmic reticulum (ER). Although de novo fatty acid synthesis occurs in the plastids, key fatty acid modification reactions may also be located on the ER. De novo fatty acid formation involves principally the operation of two multienzyme systems—acetyl-CoA carboxylase (ACCase) and fatty acid synthase (FAS) (Fig. 6.1). ACCase catalyzes the conversion of acetyl-CoA to malonyl-CoA, which is then used as the 2C building blocks for fatty acid synthesis. ACCases are found in two distinct forms in plants and, moreover, have two isoforms. One isoform is located in the plastid and is a key for de novo synthesis of fatty acids. The second isoform is presumed to be cytosolic—although that has not been proven conclusively. There is a need for malonyl-CoA in the elongation of fatty acids, which is localized on the ER and this is, clearly, its main function. Once the mechanism of action of graminicides was proven to be through selective inhibition of ACCase in the plastids of monocotyledons

FIGURE 6.1 Simplified depiction of fatty acid biosynthesis of plants. From Gurr et al., 2016. Lipids: Biochemistry, Biotechnology and Health, sixth ed. Wiley-Blackwell, Oxford with permission of the publishers Wiley.

190

Fatty Acids

(Walker et al., 1988), further studies of the molecular structure of plant ACCases took place (Alban et al., 1994). This research revealed that, while all plants contained a multifunctional form of ACCase (molecular mass 200240 KDa), the plastid form could differ. In dicotyledons, the plastid ACCase was a multienzyme complex of four proteins—biotin carboxylase, biotin carboxyl carrier protein, and a heterodimer representing the carboxyltransferase. This multienzyme complex was insensitive to graminicides. In contrast, the graminaceae contained a multifunctional protein, which was sensitive to the selective herbicides (graminicides). The cytosolic ACCase (a multifunctional protein) is poorly sensitive in all plants (Alban et al., 1994; Walker et al., 1988). In plants, the FAS is a Type II enzyme complex (see Fig. 6.2). Both acetyl-CoA and malonyl-CoA:acyl carrier protein (ACP) acyltransferases have been measured but the in vivo function of the former is in doubt since the first condensation reaction utilizes acetyl-CoA as a primer (Jaworski et al., 1989; Walsh et al., 1990), similar to Escherichia coli. There are three isoforms of the condensing enzyme (β-ketoacyl-ACP synthase, KAS). KAS I

FIGURE 6.2 The reactions of FAS.  The first condensation reaction is catalyzed by KAS III, which uses acetyl-CoA and malonylACP substrates. The next six condensations are catalyzed by KAS I and the final reaction between palmitoyl-ACP and malonyl-ACP by KAS II. From Murphy, 2005. Plant Lipids: Biology, Utilisation and Manipulation. Blackwell Publishing Ltd., Oxford with permission of the publishers Blackwell Publishing Ltd.

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catalyzes the bulk of the 2C addition reactions, whereas KAS II has high activity with palmitoyl (16:0)-ACP and, therefore, is key to controlling the ratio of 16C to 18C products from de novo biosynthesis. KAS III catalyzes the first condensation reaction and has low activity thereafter (Harwood, 1996). The sequence of condensation, reduction, dehydration, and a second reduction (Fig. 6.2) proceeds in most plants to give palmitoyl-ACP and stearoyl (18:0)-ACP as the main products of the FAS complex. In certain plants, however, termination can occur at the medium-chain fatty acid level. In the California Bay, this was first demonstrated to be due to the presence of a thioesterase active with 1012C acyl-ACPs (Voelker and Kinney, 2001) and it is presumed that other plants (such as Cuphea spp.) accumulating mediumchain fatty acids have a similar thioesterase present (Hildebrand et al., 2005). Once the basic 16C and 18C fatty acid chains have been synthesized, they can undergo further modifications. The only soluble fatty acid desaturase (FAD) is a stearoyl-ACP Δ9-desaturase, localized to the plastid stroma. The soluble nature of this desaturase allowed Shanklin’s group, in particular, to study its properties, including production of several modified enzymes with markedly different substrate selectivities and reaction characteristics (Cahoon et al., 1997; Shanklin and Cahoon, 1998). In most plants, the activity of stearoyl-ACP desaturase is high enough to prevent stearate accumulating to any degree. Cocoa (Theobroma cacao) is an exception where stearate represents about 35% of the total fatty acids (Griffiths et al., 1993). The fatty acyl-ACPs produced by de novo biosynthesis are hydrolyzed by two thioesterases termed FATA and FATB (with different substrate selectivities). The unesterified fatty acid products are then reesterified by acyl-CoA synthases. The original activity was detected on the chloroplast envelope of spinach (Joyard and Stumpf, 1981) but other enzymes have now been detected in the cytosol, so the location of bulk acyl-CoA formation is unclear at present. Furthermore, the detection of acyl-CoAbinding proteins (ACBPs) in Arabidopsis thaliana (Arabidopsis) (six isoforms) (Lung et al., 2016) and other plants revealed an important intermediate protein involved in oil accumulation. It seems likely that, in Arabidopsis, two of the ACBP isoforms may play an important role during oil accumulation, not only in binding acyl-CoAs but also in delivering fatty acids to phosphoglycerides such as phosphatidylcholine (PtdCho). In general, the role of ACBPs in oil synthesis is ill defined. The acyl-CoA pool is used directly by the three acyltransferases of the Kennedy pathway—glycerol 3-phosphate acyltransferase (GPAT), lysophosphatidate acyltransferase (LPAAT), and diacylglycerol acyltransferase (DGAT) (Fig. 6.3). Phosphatidate phosphohydrolase (PAP) is the fourth enzyme needed. GPAT and LPAAT have quite selective requirements for their acyl substrates. The ER isoforms produced PtdOH with the sn-1 position containing 16C acids and a higher degree of saturation compared to the sn-2 position. The old concept that the enzyme was relatively promiscuous

192

Fatty Acids

FIGURE 6.3 General overview of fatty acid and TAG biosynthesis in maturing seeds of oleaginous crops. Enzymes involved are indicated in boxes. The relative contributions of diacylglycerol acyltransferase (DGAT), diacylglycerol transacylase (DGTA), and phospholipid:diacylglycerol acyltransferase (PDAT) appear to vary between crop species. Additional abbreviations: ACCase, acetyl-CoA carboxylase; CPT, CDP-choline:DAG cholinephosphotransferase; CoA, coenzyme A; DAG, sn-1,2-diacylglycerol; DHAP, dihydroxyacetone phosphate; ER, endoplasmic reticulum; FA, fatty acid; FA-ACP; fatty acyl-acyl carrier protein; FA-CoA, fatty acyl-coenzyme A; Glu6PDH, sn-glucose-6-phosphate dehydrogenase; G3P, sn-glycerol-3-phosphate; GPAT, snglycerol-3-phosphate acyltransferase; LPA, lysophosphatidic acid; LPAAT, lysophosphatidic acid acyltransferase; LPC, lysophosphatidylcholine: LPCAT, lysophosphatidylcholine acyltransferase; MAG, monoacylglycerol; PA, phosphatidic acid; PAP, phosphatidic acid phosphatase; PC/PtdCho, phosphatidycholine; PDCT, phospholipid:DAG acyltransferase; PLA2, phospholipase A2; PUFA, polyunsaturated fatty acid; 16:0, palmitic acid; 18.1, oleic acid. The image was slightly modified to show PDCT action (Chen et al., 2015). From Weselake et al., 2009, Increasing the flow of carbon into seed oil. Biotechnol. Adv. 27, 866878 with permission of Elsevier, Inc.

seems not to be true and, depending on the plant, it can be rather selective. For seeds where unusual fatty acids accumulate, all the acyltransferases seem to have special properties that could be harnessed for plant oil engineering efforts. Previous attempts at genetic engineering to allow common crop species to produce unusual fatty acids have suffered from the inability of the endogenous acyltransferases to utilize such specialized substrates (Bates et al., 2014). While the Kennedy pathway is responsible for the major flow of carbon into triacylglycerol (TAG), there are other important ancillary reactions involved (Fig. 6.3). Since PtdCho is a major substrate for oleate (18:1Δ9cis;

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hereafter 18:1) and linoleate (18:2Δ9cis,12cis; hereafter 18:2) desaturation in the ER, then entry and exit of fatty acids onto this phosphoglyceride are very significant. Entry of fatty acids from the acyl-CoA pool, their desaturation, and further equilibration with (or addition to) sn-1,2-diacylglycerol (DAG) is all part of what has been recently termed acyl editing (Bates et al., 2007). CDPcholine:DAG cholinephosphotransferase (CPT) provides the backbone for PtdCho formation (Gurr et al., 2016) while fatty acids are removed and then replaced by a Lands-type mechanism via lysoPtdCho. PtdCho (either modified or “edited”) is in equilibrium with DAG through the activity of PtdCho:DAG cholinephosphotransferase (PDCT). Alternatively, PtdCho can donate a fatty acid to DAG by the activity of PDAT (PtdCho:DAG acyltransferase) to create TAG in an acyl-CoAindependent pathway. It is well established that the above enzymes are important and there may well be others still undiscovered, especially in different plant species. But the relative contribution of, for example, DGAT verses PDAT for TAG formation is not well established and may well differ significantly in different crops (Zhang et al., 2009). Elongation of preformed fatty acids to produce very long-chain fatty acids (20C or more) takes place on the ER. A cycle of condensation, reduction, dehydration, and a second reduction takes place to add 2C at a time to the hydrocarbon chain, much as with FAS. However, elongation differs from de novo synthesis in several ways. Malonyl-CoA is the source of 2C units (plastid-based FAS uses malonyl-ACP) and the “primer” fatty acid is present as an acyl-CoA. The reactions are also membrane bound in the ER (Leonard et al., 2004). Although desaturation and elongation are the main modification reactions for fatty acids (Fig. 6.1), there are other derivations that can take place. These further enzyme reactions may give rise to “unusual” fatty acids such as those containing hydroxyl, cyclic, or epoxy groups as well as conjugated, acetylenic, or allenic acids. Such products are often major fatty acids in specialized crops (see Section 6.4). As mentioned previously, fatty acids are produced de novo in the plastid. Although reactions such as Δ12- or Δ15-desaturation occur on the ER via the catalytic action of FAD2 and FAD3 and, hence, are pivotal for seed oil formation, there are also desaturases in the plastid. FADs 47 (originally named FADA-D) are plastid-localized (Browse and Somerville, 1991; Wallis and Browse, 2002). FAD4 is responsible for the conversion of palmitate to trans-Δ3-hexadecenoate, which probably takes place on a phosphatidylglycerol molecule (Harwood, 1996; Wallis and Browse, 2002). Successive desaturations of palmitate at the sn-2 position (note the difference from the fatty acid distribution produced on the ER) by FAD5 followed by FAD6/7 yield hexadecenoate, hexadecadienoate, and hexadecatrienoate using (mainly) monogalactosyldiacylglycerol (MGDG) as a substrate (Wallis and Browse, 2002). Oleate at the sn-1 position of MGDG can also be successively desaturated to α-linolenate (18:3Δ9cis,12cis,15cis; hereafter 18:3) via linoleate and the

194

Fatty Acids

catalytic action of FAD5 and FAD6/7. These desaturases are also important for further desaturation of incoming DAG molecules derived from PtdCho in the ER by the so-called “eukaryotic pathway” (Gurr et al., 2016). For general sources of information about plant fatty acid and glycerolipid synthesis, see Bates et al. (2013), Chapman and Ohlrogge (2012), Chen et al. (2015), Li-Beisson et al., (2013), Murphy (2005), and Weselake et al. (2009).

6.3 MAJOR OIL CROPS The four major world oil crops in order of quantitative importance are oil palm, soybean, oilseed rape, and sunflower (Table 6.1). Together these crops account for about 88% of the total vegetable oil output.

6.3.1 Oil Palm (Elaeis guineensis) Oil palm, grown mainly in Indonesia and Malaysia, is a uniquely productive crop capable of yielding in excess of 10 t oil ha21. This is about 10 times the yield of soybean and around 78 times that of oilseed rape. Thus, despite persistent criticism of oil palm plantations from nongovernmental organizations, in terms of the environment oil palm is much better than alternative crops, because it is so productive. TABLE 6.1 Major Oil Crops Productiona (% Total)

Typical Fatty Acid Composition (%) 16:0

18:0

18:1

18:2

18:3

Other

33

44

4

39

10

Tr.

3

Kernel oil

4

8

2

16

3

Tr.

71

Soybean

27

10

4

18

55

13

Tr.

HEARc

1

4

1

15

14

9

57

LEAR

15

4

2

62

22

10

Tr.

Sunflower

8

6

5

22

66

Tr.

1

Oil palm Palm oil b

Oilseed rape

a

% of total vegetable oil production 2009/2010. Tr., trace (,0.5). Contains 48% laurate. c Contains 10% eicosenoate and 45% docosenoate (erucate). From Taylor et al., 2011. Metabolic engineering of higher plants to produce bio-industrial oils. In: Moo-Young, M. (Ed.), Comprehensive Biotechnology, second ed., vol. 4. Elsevier, Amsterdam, pp. 6785 and Gunstone et al., 2007. The Lipid Handbook, third ed. CRC, Boca Raton, FL. b

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Oil palm yields two sorts of oils, palm oil from the fruit mesocarp and seed (kernel) oil, each with distinct uses. Palm oil, derived from the fruit mesocarp, is the most globally abundant edible oil, accounting for over 30% of the total vegetable oils used in 2016. The most common cultivar is Tenera, which is a cross between cultivars Tura and Pisifera. Typical oils contain high amounts of palmitate (B45%) and oleate (B40%) with linoleate (B12%) as a significant polyunsaturated fatty acid (PUFA) component. Palm oil usage is greatly extended by fractionation to give oleins, stearins, and mid-fractions. Further details of these fractions (see Table 6.2) and their uses are given in Gunstone et al. (2007). Sambanthamurthi et al. (2000) have reviewed the biochemistry and chemistry of palm oil. Given the supreme efficiency of oil palm for vegetable oil productivity and the inevitable increase in world demand of edible oils, it is obvious that demand for palm oil will continue to rise. This increase in demand combined with the strict laws enacted in Indonesia and, especially, Malaysia to drastically reduce primary forest clearance and planting on peat is galvanizing efforts to increase oil palm yields. Although the average yield of oil in plantations is 34 t ha21, maximums of around 20 t ha21 have been demonstrated, so a doubling of yields should be easily achievable. Current efforts have focused on crop management, replanting with higher-yielding lines, and optimizing harvesting and processing methods. Using traditional breeding and marker-assisted selection, key traits to be focused on for future improvement are dwarf and/or compact varieties, lines yielding high-oleate contents and plants with disease-tolerance, especially against Ganoderma boninense. These subjects are discussed in more detail by Murphy (2014). In terms of transgenic modification, oil palm is still in its infancy. Because E. guineensis is a monocotyledon, use of Agrobacterium for delivery is inefficient and no better than biolistics (Parveez and Bahariah, 2012). Identification of appropriate (palm-sourced) promoters and the long time to

TABLE 6.2 Major Fatty Acid Components of Palm Oil and Its Fractions Palm Oil

Palm Olein

Top Olein

Soft Stearin

Mid-Fraction

14:0

1.1

1.1

1.0

1.1

16:0

44.1

40.9

28.8

49.3

18:0

4.4

4.2

2.5

4.9

18:1 n-9

39.0

41.5

52.0

34.8

32.041.2

18:2 n-6

10.6

11.6

14.6

9.0

2.611.2

18:3 n-3

0.3

0.4

0.4

0.2

Tr.0.2

From Gunstone et al., 2007. The Lipid Handbook, third ed. CRC, Boca Raton, FL.

0.81.4 41.455.5 4.76.7

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first fruiting (B5 years) are also major constraints. Moreover, the ongoing public resistance to genetically engineered (GE) crops in some parts of the world, especially Europe, has reduced funding toward research into genetic manipulation of oil palm. While progress with genetic manipulation of oil palm remains slow, selection of palms from existing stocks seems to be the quickest way forward (Singh et al., 2009). Fortunately, there is a multitude of oil palm varieties from different parts of Africa and South America, some of which have distinct characteristics (Murphy, 2014). The recent publication of genomic sequences (Singh et al., 2013b), identification of the shell gene (Singh et al., 2013a) (Dura is thick-shelled and pisifera is thin shelled), and the reason for mantling in somatic cells (Ong-Abdullah et al., 2015) will enable breeders to use molecular markers for selection (see Murphy, 2014). For recent general reviews on palm oil production, modifications, and oil palm cultivations, see Murphy (2014, 2015). As mentioned earlier, the oil palm yields a second useful fat product, palm kernel oil (Gunstone et al., 2007). The seeds (kernels) of palm fruits contain about 45% oil. Typically, fruits will produce about 8 times as much mesocarp oil as palm kernel oil, nevertheless, the latter is more valuable on a g/g basis. Palm kernel oil is laurate (12:0)-rich (Tables 6.1 and 6.3), has many industrial uses, especially soap production, and is gradually replacing coconut oil as the main laurate-enriched oil. In fact, the production of palm kernel oil in parallel with the emergence of oil palm as the major oil crop meant that the successful transgenic manipulation of canola-type Brassica napus to produce laurate (Voelker et al., 1992) failed commercially (McKeon et al., 2016). Although a significant proportion (16%20%) of

TABLE 6.3 Comparison of the Fatty Acid Compositions (%) of Coconut and Palm Kernel Oils Coconut

Palm Kernel

8:0

510

26

10:0

58

35

12:0

4553

4455

14:0

1721

1418

16:0

810

710

18:0

24

13

18:1

510

1219

18:2

13

14

From Gunstone et al., 2007. The Lipid Handbook, third ed. CRC, Boca Raton, FL.

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palm kernel oil is used for nonfood purposes (Gunstone et al., 2007), it is also widely used in the food industry. Usages include the production of surface-active compounds, spreads, shallow frying, cocoa butter substitutes, filling creams, ice-creams, nondairy whipping creams, and filled milks. It is prone to hydrolytic rancidity and the released laurate results in a soapy flavor. Laurate can be further oxidized and decarboxylated—a process called ketonic rancidity. More details of palm kernel products can be found in Yusolf Basiron (2005). To date, there have been few attempts to genetically modify palm kernel oil or to identify lines with different oil characteristics. Naturally, the difficulties of transgenic modification of kernel oil will be the same as noted earlier for the mesocarp. Nevertheless, knowledge of, for example, the shell gene (Singh et al., 2013a) may be useful in terms of palm kernel oil yields (see Murphy, 2014, 2016a,b).

6.3.2 Soybean (Glycine max) Soybean is the second largest global source of vegetable oil (Vegetable oil production, 2013; Table 6.1). In 2013 the top 10 soybean-growing countries produce about 267 million metric tons of the seed with the United States and Brazil leading production at about 34% and 31%, respectively (Statista, 2013). Commodity soybean seed contains about 40% protein and 20% oil. The oil is enriched in linoleate with lesser amounts of oleate, α-linolenate, palmitate, and stearate (Taylor et al., 2011). For example, in G. max L. cv Thorne, the proportions (wt%) of these fatty acids are 49.4%, 17.9%, 14.4%, 13.1%, and 5.2%, respectively (Buhr et al., 2002). Although soybean has been improved through genetic engineering, it has proven challenging to transform using conventional methods. The two main methods for transforming soybean are Agrobacterium-mediated transformation of cotyledonary node explants from imbibed seeds or young seedlings and particle-bombardment (via gene gun) of somatic embryos (Donaldson and Simmonds, 2000; Maheshwari and Kovalchuk, 2016). Currently, over 90% of globally traded soybean is herbicide-tolerant as a result of genetic engineering (Maheshwari and Kovalchuk, 2016). Soybean somatic embryos have proven to be a useful trait-testing system in research programs aimed at lipid modification (Rao and Hildebrand, 2009). Desired outcomes of metabolic engineering observed at the level of the somatic embryo justify further investment in germination and plant propagation to develop advanced generations of transgenic soybean. Advances in soybean improvement have also benefited from genomic resources (Schmutz et al., 2010). Soybean oil is the main source of biodiesel feedstock in the United States (Karmakar et al., 2010). One advantage of using soybean oil for biodiesel applications is that the crop can be produced with little or no nitrogen (Karmakar et al., 2010). The manufacture of nitrogen fertilizer represents a

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costly input in crop production, which also generates nitrous oxide, a powerful greenhouse gas. Soybean oil is also used to produce industrial materials such as polyurethanes for applications in sealants and molded foams (McVetty et al., 2016). Hydroxylation of vegetable oil produces polyols, which are in turn used in generating polyurethane. The relatively high content of PUFA (. 60%) in soybean oil renders it more oxidatively unstable relative to seed oil enriched in monounsaturated fatty acids. Seed oils enriched in oleate are less easily oxidized while still exhibiting desirable flow properties under cooler environmental conditions (Durrett et al., 2008). High-oleate oil also provides increased uniformity for other industrial applications, which include lubricant formulations, dielectric fluids, plastics, and oleate for oleochemicals (Plenishs High Oleic Soybeans). High-oleate lines are known as those with .70% oleate content in the seed oil (Taylor et al., 2011). Soybean oil enhanced in oleate content was initially produced by downregulation of the gene encoding FAD2 so as to reduce the conversion of 18:1 to 18:2 at the sn-2 position of PtdCho (Kinney et al., 2002; Damude and Kinney, 2008). This approach has also been combined with other strategies aimed at reducing saturated fatty acid content. For example, Buhr et al. (2002) downregulated FAD2-1 along with the FatB gene encoding palmitoyl-thioesterase so as to generate soybean oil with .85% oleate and ,6% saturated fatty acid content. Plenishs High Oleic Soybeans is an example of commercially grown soybean-containing seed oil with .75% oleate and 20% less saturated fatty acid content than commodity soybean oil (Plenishs High Oleic Soybeans). This oil also contains ,3% α-linolenate, which contributes to the increased oxidative stability of the product. Proof-of-concept metabolic engineering studies have been conducted to demonstrate the potential for producing industrially useful conjugated fatty acids (CFAs) or epoxy fatty acids (EFAs) in soybean. Oils enriched in CFAs are even more susceptible to oxidation than oils enriched in PUFA with methylene-interrupted double bonds (Sonntag, 1979). Thus, oils such as tung tree (Vernicia fordii) oil, which is enriched in CFAs, are useful as drying agents in paints, varnishes, and inks. Vernolate (cis-12-epoxyoctadeca-cis-9-enoate), which is an industrially useful EFA, can be used as plasticizer of polyvinyl chloride. In addition, the ability to cross link epoxy groups renders vernolate-enriched oils useful in adhesives and coating materials (Perdue et al., 1986). In maturing seeds that produce oils containing CFAs, fatty acid conjugase (FADX) catalyzes the conversion of linoleic acid into CFA at the sn-2 position of PtdCho (Mietkiewska et al., 2014). Similarly, epoxygenase catalyzes the conversion of sn-2-18:2-PtdCho into sn-2-vernoloyl-PtdCho (Bafor et al., 1993; Yu et al., 2008). Thus, unusual fatty acids, such as CFAs and EFAs, are produced on PtdCho in a similar fashion to PUFA containing methyleneinterrupted double bonds (e.g., α-18:3). FADX enzymes are recognized

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as divergent forms of FAD2. α-Eleostearate (18:3Δ9cis,11trans,13trans) and calendate (18:3Δ8trans,10trans,12cis) are enriched in tung oil and marigold (Calendula officinalis) seed oil, respectively (Mietkiewska et al., 2014) (see Section 6.4). Cahoon et al. (1999, 2001) have engineered soybean somatic embryos to produce α-eleostearate and calendate to levels of 17% and 19%, respectively. Somewhat later, soybean seed oil, with about 20% calendate, was developed through metabolic engineering (Cahoon et al., 2006). Unlike natural species that produce CFAs, metabolically engineered soybean retained a relatively high proportion of CFA at the sn-2 position of PtdCho suggesting that the natural species producing CFAs have effective mechanisms in place for removing CFAs from their site of synthesis in PtdCho (Cahoon et al., 2006). A CYP726A1 gene encoding a cytochrome P450 enzyme from Euphorbia lagascae was used to engineer somatic soybean embryos, which accumulated about 8% of their total fatty acids as Δ12-EFAs (Cahoon et al., 2002). Stokesia laevis and Vernonia galamensis are also high accumulators of vernolate (Section 6.5, Yu et al., 2006). Li et al. (2010a) coexpressed a cDNA (SIEPX)-encoding epoxygenase from S. laevis in combination with cDNAs-encoding DGAT1 or DGAT2 from V. galamensis in soybean. SIEPX expression alone resulted in about 8% vernolate in the seed oil whereas coexpression with VgDGAT1 or VgDGAT2 resulted in about 15% and 26% vernolate, respectively, in the seed oil. DGAT activity in microsomes of developing seeds of V. galamensis or S. laevis was previously demonstrated to exhibit enhanced selectivity for substrates containing vernolate (Yu et al., 2006). Somewhat later, soybean expressing S. laevis SIEPX was shown to exhibit reduced seed oil content and protein content (Li et al., 2012). These transgenic seeds were also shriveled and wrinkled, and exhibited a 11%16% decrease in germination rate compared with the control. Coexpression of SIEPX with VgDGAT1 or VgDGAT2, however, restored normal seed morphology and germination rate, and resulted in normal levels of seed oil and protein content. SIEPX-transformed soybean seeds contained 3%7% vernolate acid in TAG and 11.7%13.5% vernolate in PtdCho. In contrast, seeds coexpressing SIEPX and VgDGAT1 or VgDGAT2 had 17.8% and 27.9% vernolate in TAG, but only about 6% vernolate remained in PtdCho. The data suggested that the effective utilization of vernolatecontaining substrates by VgDGAT1 or VgDGAT2 led to more efficient removal of vernolic acid from PtdCho, which resulted in more stable cellular membrane metabolism. Considerably greater enrichment of soybean oil with CFAs or EFAs will be required to provide suitable oil for industrial applications. Results to date suggest that the expression of multiple genes, encoding lipid biosynthetic enzymes from other sources, in soybean may be required to produce very high levels of unusual fatty acids in the seed oil (Mietkiewska et al., 2014). In addition, it may also be necessary to implement downregulation strategies

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Fatty Acids

to reduce competition between heterologously produced enzymes and endogenous enzymes competing for the same substrates (Van Erp et al., 2015). Given the increasing global demand for more plant oil for food, feed, and nonfood applications (Weselake et al., 2009), it will also be important to boost oil content in soybean seeds. Given the importance of soybean protein, this feat must be achieved without lowering protein content (Roesler et al., 2016). The seed oil content of about 20% for commodity soybean has not changed for several decades and neither conventional breeding nor mappingbased approaches involving quantitative trait loci (QTL) have been effective in increasing soybean seed oil content (Roesler et al., 2016). Seed oil content in soybean is a polygenic trait that is controlled by several QTL (Brim, 1973), the majority of which exhibit genotype-environment interactions (Panthee et al., 2005; Weselake et al., 2009). Within the last decade, however, a number of single gene manipulations have shown promise in bringing about oil content increases by a few percentage points. Rao and Hildebrand (2009) used a yeast SLC1 gene encoding an enzyme with LPAAT activity (Zou et al., 1997) to generate soybean seeds (T3 transgenic lines) exhibiting an absolute increase in seed oil content of about 3.2%. This increase in oil content, however, was accompanied by about a 0.8 percentage point decrease in protein content. Lardizabal et al. (2008) heterologously expressed a codon-optimized DGAT2A from Umbelopsis ramanniana during seed development in soybean, which resulted in a 1.5% absolute increase in seed oil content, which was consistently maintained through consecutive field trials. In a more recent investigation, heterologous overexpression of Sesamum indicum L. cv Wanzhil DGAT1 in soybean, using a CaMV35S promoter, resulted in a mean increase in T3 seed oil content of about 1.4 percentage points (Wang et al., 2014). Seed size was also observed to increase. An investigation of transcript production during seed development in various oil-producing species suggested that DGAT1 is the major enzyme involved in seed oil synthesis in soybean (Li et al., 2010b). Recently, Roesler et al. (2016) used a directed evolution approach to increase the activity of DGAT1 from the American hazelnut shrub (Corylus americana), which exhibited a combination of high oil content (60%) and high-oleate content (79%). Various amino acid substitutions resulted in CaDGAT1 variants with increased activity and improved affinity for oleoylCoA. Both native CaDGAT1 and the CaDGAT1 variants exhibited sigmoidal kinetics. Guided by the relatively high level of amino acid sequence identity between CaDGAT1 and soybean DGAT1, the equivalent of 14 amino acid substitutions promoting higher oil-forming activity in a particular CaDGAT1 variant was installed into soybean DGAT1b. Overexpression of the “super” soybean DGAT1b variant in soybean resulted in a 3 percentage point increase in seed oil content, which was accompanied by about a 2 percentage point decrease in soluble sugars when seed was analyzed from highly replicated, single location, field trials. The greatest absolute changes in fatty acid

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composition showed an increase in oleate and decrease in linoleate. In some cases, increased protein content was also observed.

6.3.3 Brassica Oilseed Species (Brassica napus, Brassica rapa, Brassica oleracea, Brassica carinata) The Brassica genus is a member of the Brassicaceae family, which features plants that produce very long-chain fatty acids in their seed oils and glucosinolates, which occur throughout the plants (McVetty et al., 2016). Glucosinolates appear to be important in conferring tolerance to abiotic stress and can be induced by herbivore (Textor and Gershenzon, 2009) or fungal attack (Abdel-Farid et al., 2010). Brassica oilseed species are generally adapted to cooler temperature regions. Rapeseed, which is mostly B. napus, is the third largest global source of vegetable oil (Vegetable oil production, 2013: Table 6.1). Amphidiploid B. napus originated from interspecific hybridization and spontaneous chromosome doubling of the ancestors of B. rapa (AA, 2n 5 20) and B. oleraceae (CC, 2n 5 18) (U, 1935; Scarth and Tang, 2006). In 2013 nine countries produced over 60 million metric tons of rapeseed with Canada and China leading global production at 29% and 23%, respectively (Statista, 2013). Today’s rapeseed is mainly low in erucic acid (22:1Δ13cis; hereafter 22:1; ,2% of the oil) and glucosinolates (,2 μmol per g of meal at 8.5% water content) and is also known as canola (McVetty et al., 2016). Low erucic acid rapeseed (LEAR) was developed in Canada (Stefansson et al., 1961). The low erucate phenotype is attributable to loss-of-function mutations in FATTY ACID ELONGASE1 genes in the A and C genomes (Fourmann et al., 1998; Katavic et al., 2002; Rahman et al., 2008). The regions surrounding these genes have been explored to provide new tools for high-throughput marker-assisted selection rapeseed breeding programs (Rahman et al., 2008). Globally, high erucic acid rapeseed (HEAR) is about one-tenth of the production of canola (McVetty et al., 2016). The seed oil content of rapeseed averages about 45% (Rahman et al., 2013a). Examples of fatty acid compositions for B. napus enriched in erucate and canola-type B. napus are depicted in Table 6.1. Canola oil is enriched in oleate with lesser amounts of linoleate, α-linolenate, palmitate and stearate, respectively. Most of the B. napus cultivars planted in Canada are herbicide-tolerant though genetic engineering (Weselake, 2011; McVetty et al., 2016). The introduction of foreign DNA into B. napus routinely involves standard protocols in Agrobacterium-mediated transformation (Moloney et al., 1989; Weselake, 2011). The breeding process for Brassica oilseed species has been accelerated by double haploid technology, which involves generation of microspore-derived embryos. These embryos have also proven to be a useful and convenient model system for studying lipid biosynthesis in Brassica oilseed species (Weselake and Taylor, 1999). More recently, advances in

202

Fatty Acids

B. napus improvement have also greatly benefited from genomic resources (Chalhoub et al., 2014). Brassica napus seed oil is the main source of biodiesel feedstock in both Canada and the European Union (Karmakar et al., 2010). Spring habit B. napus is grown in the Canadian prairies and Northern Europe, whereas winter habit B. napus is grown in other parts of Europe (McVetty et al., 2016). In 2013, 433,000 metric tons of oils and fats were used as feedstock for biodiesel production in Canada (Evans, 2013). About 35% of this feedstock was from the oil of canola-type B. napus. In Europe, about 70% of biodiesel feedstock is attributable to B. napus seed oil (McVetty et al., 2016). In contrast to the use of soybean oil for biodiesel, there is concern regarding the use of B. napus seed oil for this purpose because of the high requirement for nitrogen fertilizer (Karmakar et al., 2010). Within the last decade, the increasing use of vegetable oils for biodiesel has also led to higher vegetable oil prices (Durrett et al., 2008). The oils of rapeseed, soybean, and other major oil crops may not be sufficient to satisfy the demand for biodiesel in the long run given the growing global population and the competing demand for food oil (Carlsson, 2009; McVetty et al., 2016). This concern has increased research on developing nonfood oil crops that can grow marginal land and microalgae as potential sources of biodiesel feedstock. Carlsson (2009) has shown that it is in fact more feasible to use vegetable oil as source of feedstock for the production of high-value industrial chemicals and polymers, as a replacement for petrochemical-derived products, rather than to use plant oils for biofuel. Similar to soybean oil, canola oil is also useful as a feedstock to produce various high-value polymers for industrial applications (Hayes and Dumont, 2016; McVetty et al., 2016). High-oleate cultivars of B. napus have been produced through mutagenesis breeding. As examples, the cultivars “Splendor” and “Nexera” contain .75% oleic acid content and reduced α-linolenic acid content in their seed oils (Friedt and Snowdon, 2009). Reduced α-linolenate content can result from mutagenesis of FAD3 genes (Rahman et al., 2013b). Transgenic downregulation of FAD2 gene expression has also been used to generate high-oleate varieties of B. napus (Stoutjeskijk et al., 2000; Kinney et al., 2002; McVetty et al., 2016). High-oleate B. napus seed oil has similar industrial applications to high-oleate soybean oil. High stability and high-oleate B. napus seed oil is particularly useful in hydraulic fluids (McVetty et al., 2016). Brassica seed oils enriched in erucate, however, have largely given rapeseed its reputation as an industrial oil. Currently, HEAR oils contain about 50% erucate, whereas super high erucic acid rapeseed (SHEAR) oils contain .66% erucate (Nath et al., 2009; McVetty et al., 2016). The main interest, however, lies in high erucate cultivars with low glucosinolate content in order to produce nontoxic meal as a by-product of seed oil extraction. Industrial uses of HEAR oils include direct application as green lubricants,

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synthesis of brassylate (C13), and pelargonate (C9) for lubricant and ingredient applications, conversion to erucamide for use as a slip-promoting agent, and conversion to substituted amines for chemical synthesis of surfactants (Sonntag, 1991; Van Dyne and Blase´, 1991; To¨pfer et al., 1995; McVetty et al., 2016). In addition, nylon 13-3 and nylon 9-9 are prepared using brassylate and pelargonate, respectively. SHEAR oils are being developed for specific chemical feedstock applications, where even more increased uniformity is required (McVetty et al., 2016). Development of SHEAR has involved both breeding and genetic engineering-based approaches. Breeding approaches have involved resynthesis of B. napus from parental species of B. oleraceae and B. rapa followed by mutagenesis of microspore-derived embryos to produce double haploid lines exhibiting possible increases in 22:1 content (McVetty et al., 2016). The incorporation of 22:1 into TAG is largely limited by the inability of the resident LPAAT to catalyze the acylation of the sn-2 position of lysophosphatidate (Weselake, 2005). Thus, the theoretical limit for 22:1 content in HEAR oil is about 66% assuming that there are no obstacles to acylating the sn-1 and sn-3 positions with 22:1. Other limitations to 22:1 enrichment include limited activity of the ER fatty acid elongase (FAE) complex and competition of the 18:1-elongation process with FAD2 activity leading to 18:2 (McVetty et al., 2016). The LPAAT limitation was overcome through the introduction of a LPAAT from another plant source (e.g., Limnanthes alba or Tropaeolum majus), which was capable of utilizing 22:1-CoA (Lassner et al., 1995; Xu et al., 2008). In ground-breaking research, Nath et al. (2009) developed a transgenic SHEAR line, which accumulated 72% erucate in the seed oil. These transgenic lines were developed through overexpression of the rapeseed FAE gene combined with expression of LPAAT cDNA from Limnanthes douglasii followed by combination of the transgenic material with mutant alleles for low PUFA (18:2 1 18:3) content. Brassica carinata is being developed as a platform crop for the production of bio-industrial oil feedstocks, because it is well adapted to grow on marginal land and is drought tolerant (Carlsson, 2009; Marillia et al., 2014; McVetty et al., 2016). Both biofuel and industrial chemical applications are being explored. Agrisoma BioSciences (Saskatoon, Canada) has developed B. carinata seed oil as a feedstock for production of 100% drop-in bio-jet fuel (Marillia et al., 2014; McVetty et al., 2016). Jadhav et al. (2005a) further increased the erucate content of high erucate B. carinata by downregulating FAD2 expression so as to provide more 18:1 for elongation to 22:1. B. carinata has also been engineered to produce substantial levels of nervonate (24:1Δ15cis; hereafter 24:1), which is the elongation product of 22:1 (Marillia et al., 2014). Erucate and nervonate-enriched oils have uses in the synthesis of enhanced oil recovery surfactants, which are injected into the earth to promote better recovery of petroleum oil (Marillia et al., 2014). B. carinata seed oils, enriched in nervonate and erucate, have also been fed to

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Fatty Acids

Pseudomonas aeruginosa under conditions of nitrogen-deprivation to generate polyhydroxyalkanoates (bioplastic) for use in biodegradable lacquers (Impallomeni et al., 2011). Guo et al. (2009) introduced cDNA encoding 3ketoacyl-CoA synthase (KCS), from the “money plant” (Lunaria annua L.) into B. carinata to increase elongation of 22:1 to 24:1. The best performing transgenic lines had 30% of 24:1 in the seed oil compared to 28% in untransformed B. carinata. In the same year, Taylor et al. (2009a) reported on B. carinata transgenic lines with up to 44% nervonate and 6% residual erucate in the seed oil, which were generated using a KCS cDNA from bittercress (Cardamine graeca L.). Limnanthes species also produce about 22% docasadienoate (22:2Δ5cis,13cis; hereafter 22:2) in their seed oils. Docasadienoate can be used as a feedstock for producing various lubricants (Marillia et al., 2014). 22:2-CoA is produced by desaturation of 22:1-CoA catalyzed by a Δ15 desaturase (Marillia et al., 2002). Introduction of Des5 cDNA, encoding this desaturase, into B. carinata resulted in 22:2 accounting for up to 15% of the total very long-chain fatty acids in the seed oil (Jadhav et al., 2005b). Various metabolic engineering interventions resulted in B. carinata seed oils containing up to about 65% erucate, 45% nervonate, or just over 10% docasadienoate (Marillia et al., 2014). The earliest successes in the metabolic engineering of B. napus to produce industrially useful fatty acids, not normally synthesized in the developing seed, involved research aimed at enriching accumulation of mediumchain fatty acids (Stoll et al., 2005). Cuphea species produce oils highly enriched in caprylate (8:0), caprate (10:0), and laurate, which are useful in the production of detergents, lubricants, and plasticizers (McKeon, 2016b). Caprate-enriched seed oils are also useful as feedstock for biodiesel. In this regard, considerable breeding efforts have gone into improving the agronomic characteristics of the crop. There has also been interest in using Cuphea acyl-ACP thioesterase to produce caprylate and caprate in canolatype B. napus. Transformation of B. napus with cDNA (CpFatB2) encoding a Cuphea hookeriana thioesterase resulted in seed oil with up to 11 mol% caprylate and 27 mol% caprate (Dehesh et al., 1996). The highest levels of medium-chain fatty acids produced in B. napus, however, were with laurate. Laurate has a long history of industrial use, which includes production of various nitrogen derivatives (Reck, 1985). Canola-type B. napus seed oil with .50% laurate was generated through the heterologous expression of a cDNA encoding the California bay laurel (Umbellularia californica) 12:0-ACP thioesterase during seed development (Voelker et al., 1996). Incorporation of 12:0 into TAG, however, appeared to be limited by induction of β-oxidation and glyoxylate cycle activity (Eccleston and Ohlrogge, 1998) and the inability of B. napus LPAAT activity to utilize 12:0-CoA (Knutzon et al., 1999). Thus, similar to the situation with 22:1 incorporation at the sn-2 position of B. napus TAG, the incorporation of 12:0 at the sn-2 position of TAG required an enzyme from a different plant source. Laurate

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levels in seed oil from transgenic B. napus were further increased via the introduction of 12:0-CoA-preferring LPAAT from coconut (Cocos nucifera) (Knutzon et al., 1999). This seed oil exhibited enhanced production of TAG with 12:0 at the sn-2 position and overall laurate content typically reached levels of about 60%. Unfortunately, as indicated in Section 6.3.1, high laurate B. napus could not compete with other commercial laurate sources such as palm kernel oil. Much like soybean, seed oil content in B. napus is a polygenic trait controlled by numerous gene loci with many genotype X environment interactions (Weselake et al., 2009; Rahman et al., 2013a). Over the last decade or so, there has been incremental progress in raising the seed oil content of B. napus in both Canada and Europe (Rahman et al., 2013a). Although the seed oil content of B. napus germplasm can vary from about 35% to 52% (Rahman et al., 2013a), researchers from China have recently reported development of a B. napus line with about 65% seed oil content (Hu et al., 2013). The line was developed through pyramiding of high oil alleles. Based on oil body analysis, the investigators have proposed that it may be theoretically possible to generate a B. napus line with 75% seed oil content! Despite the known multigene contributions in determining seed oil content, there are several examples of single gene interventions resulting in increased seed oil content in Arabidopsis and/or B. napus. Arabidopsis is a model oilseed species, which also belongs to the Brassicaceae. Metabolic engineering strategies have involved increasing the supply of plastidially derived fatty acids, producing more G3P for the Kennedy pathway, increasing the activity of TAG assembly enzymes, altering carbon partitioning and enhancing production of transcription factors, which upregulate the expression of several genes encoding enzymes involved in glycolysis and fatty acid synthesis (Weselake et al., 2009). More recently, downregulation of SUGAR-DEPENDENT1 TAG LIPASE during seed maturation was shown to increase seed oil content by about 8% on relative basis (Kelly et al., 2013). In wild-type B. napus, seed oil content can decrease by about 10% during the later stages of maturation, which is attributable to induction of lipase activity (Kelly et al., 2013). The DGAT-catalyzed reaction leading to TAG in the Kennedy pathway has been of particular interest as a step for manipulation. Overexpression of either Arabidopsis DGAT1 or B. napus DGAT1 was shown to significantly increased seed oil content in canola-type B. napus under both greenhouse and field conditions (Weselake et al., 2008; Taylor et al., 2009b). Control analysis of lipid biosynthesis in developing seeds of control versus transgenic B. napus L. cv Westar lines indicated that control of TAG assembly decreased from about 70% in the wild type to about 50% in transgenic lines (Weselake et al., 2008). In other words, the TAG assembly process had more influence in regulating seed oil content than reactions involved in de novo fatty acid biosynthesis (Weselake et al., 2008; Harwood et al., 2013; Ramli et al., 2014). These findings suggested that metabolic control analysis may

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Fatty Acids

be helpful in guiding the engineering of oil crops to boost seed oil content. Interestingly, the overexpression of Arabidopsis DGAT1 in B. napus also reduced the penalty on seed oil content caused by drought (Weselake et al., 2008). Therefore, B. napus cultivars (and possibly other oil crops) overexpressing DGAT1 could have a form of built-in insurance so as to minimize reductions in seed oil content under drought conditions (Singer et al., 2016b). This strategy may be particularly useful in the industrial oil crop space, where platform crops, such as B. carinata, are being explored because of their increased tolerance to abiotic stress. In addition, since a major goal in using metabolic engineering to produce industrial oil crops is to increase uniformity with respect to the target fatty acid (e.g., 22:1), it would make sense to utilize DGAT enzymes with enhanced preference for both acyl-CoA and DAG containing the target fatty acyl moieties. Numerous variants of high activity B. napus DGAT1 have also been generated using directed evolution (Siloto et al., 2009), which may prove useful in further increasing seed oil content and possibly altering fatty acid composition associated with changes in DGAT1 substrate selectivity. The recent discovery that the GPAT9 isoenzyme catalyzes the first step in the Kennedy pathway of TAG assembly in Arabidopsis (Shockey et al., 2016; Singer et al., 2016a) could eventually lead to the evaluation of GPAT9 overexpression as means of increasing seed oil content in B. napus, possibly in association with DGAT1 overexpression. Comparative analysis of gene expression in developing embryos of B. napus lines with similar genetic background, but varying in seed oil content, has also been useful in identifying potential genes involved in governing seed oil content (Li et al., 2006; Weselake et al., 2009). Analysis of the metabolome, however, does not always produce results that reflect the relative enzyme levels suggested by transcriptomic data (Schwender et al., 2014). Recent research has further suggested that factors in developing silique and/or seed coat of B. napus may also regulate oil accumulation in the embryo through maternal effects (Liu et al., 2014b; Tan et al., 2015). These investigations will likely lead to new metabolic engineering strategies, which involve metabolic interventions in the silique or seed coat, or possibly combined manipulation of metabolic targets in the embryo, seed coat, and silique.

6.3.4 Sunflower (Helianthus annuus) Sunflower belongs to the Compositae (Asteraceae) family. It was first domesticated and cultivated in North and Central America and was imported into Europe by Spanish explorers in the 16th century. Since then, cultivation has spread all over the world with the main production being in Argentina, Russia, Ukraine and the United States. Presently, it is the fourth largest global vegetable oil product (Table 6.1).

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TABLE 6.4 Composition of Oils From Common Sunflower and Mutant Lines Fatty Acid (%)

Common

HO

HS

HP

UHO

HSHO

16:0

35

5

3

26

4

5

18:0

23

3

30

2

2

18

18:1

3050

75

14

20

91

71

18:2

4060

15

50

51

2

3

.18C

13

2

3

1

1

3

Others

Tr.

Tr.

Tr.

Tr.

Tr.

Tr.

HO, high oleate; HS, high stearate; HP, high palmitate; UHO, ultra high oleate; HSHO, high stearate high oleate; tr., trace (,0.5). From Salas et al., 2014. Biochemistry of high stearic sunflower, a new source of saturated fats. Prog. Lipid Res. 55, 3042.

Traditionally, sunflower oil is high in oleate and, notably, linoleate (Table 6.4). The 40%60% content of linoleate was originally marketed as a particularly attractive feature of sunflower oil but more recently it has been realized that too much n-6 PUFA (such as linoleate) in the diet can have undesirable consequences, especially for chronic inflammatory diseases (see Haslam et al., 2013; Lands, 2014). This has led to the breeding of other sunflower lines with modified fatty acid compositions (Table 6.4). Traditional sunflower varieties yield 40%50% oil. Although it has up to 75% linoleate, it contains virtually no α-linolenate (Table 6.1). Palmitate and stearate are present in approximately equal amounts in standard sunflower oil with oleate at 30%50%. The major TAG molecular species are LLL (27%), LLO (27%), LLP (10%), and LLS (11%) (Gunstone et al., 2007) (where L, linoleate; O, oleate; S, stearate; P, palmitate). In terms of genetic manipulation, sunflower is a difficult subject (Salas et al., 2014). A particular problem is that plants are difficult to regenerate from tissue cultures. Although there have been some recent advances, standard genetic manipulation methods have lagged behind those in other important crops and there are no GE commercial lines currently. On the other hand, sunflower is rather easy to mutagenize by both physical and chemical methods. Chemical mutagenesis with either ethylmethane sulfonate or sodium azide has produced excellent results. In addition, mutagenesis using ionizing radiation followed by screening with targeting induced local lesions in genomes (TILLING) had given useful lines (Kumar et al., 2013). Two aspects of sunflower make trait development through conventional breeding easier compared with other crops. Firstly, it is a diploid plant, which simplifies the genetics, and secondly, there is high genetic variability within the species, which acts as a source for useful traits

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Fatty Acids

(Liu and Burke, 2006; Seiler, 1992). Indeed, a variety of useful lines of sunflower have been produced using a combination of breeding with mutagenesis. Such lines have altered characteristics of oil quality and quantity (Table 6.4) as well as increased resistance to pests, drought, or salinity (Salas et al., 2014). As mentioned earlier, the high concentration of linoleate (40%60%) was originally considered to be an advantage. However, with the realization that the dietary ratio of n-6/n-3 PUFA was important with a value of four being considered desirable, the high linoleate (and very little α-linolenate) of sunflower oil was not ideal (Lands, 2014; Schmitz and Ecker, 2008). Partly for this reason, and because oleate-enriched lines are useful for renewable chemical feedstocks, high-oleate lines have been produced (Table 6.4). Sunola (Highsun) can contain up to 90% oleate while Nuson has somewhat enriched levels of oleate (60%) and is designed to replace standard varieties of sunflower in the United States. Because such lines have not been produced using GM-technology, markets (such as Europe) should not be hostile. Additional sunflower lines have been produced, which contain enhanced palmitate or stearate (Table 6.4). High palmitate has an advantage that it can be used for spreads and other commercial applications (Ferna´ndezMoya et al., 2005; Martı´nez-Force et al., 1998). Solid fats are needed for margarines, shortening, fillings, and confectionary (Gunstone et al., 2007). With the concerns about trans-fatty acids produced during hardening by hydrogenation, alternative sources are needed. While palm oil is an obvious useful product, crops that can be grown in more temperate regions are needed. Moreover, stearate-enriched sunflower varieties are available (Table 6.4). In addition, the high stearate phenotype has been transferred to high-oleate lines to create high stearate, high-oleate lines (HSHO) (Table 6.4). Oils from such lines can be fractionated to produce a number of useful food formulations (Bootello et al., 2011; Salas et al., 2011). These include coatings, confectionary, fillings, and spreads (Bootello et al., 2012). The HSHO lines have good frying stability and contain significant amounts of useful vitamins (e.g., carotenoids, tocopherols) (Salas et al., 2014). Other potential uses for sunflower oil for nonfood purposes are described in Erhan and Adhvargu (2005).

6.4 MINOR OIL CROPS The following are some of the minor oil crops currently on the market or which have the potential to be developed and used for specific applications. For more details, the reader is referred to Murphy (2005) and, in particular, Gunstone et al. (2007) or McKeon et al. (2016). Note that those with significant future potential are covered also in Section 6.5.

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6.4.1 Alfalfa (Medicago sativa, Medicago falcata) Alfalfa seeds are rather low in oil (7.8%), which is highly unsaturated and is enriched in carotenoids. The oil may lower LDL cholesterol and reduce erythema caused by sunburn (Firestone, 2012).

6.4.2 Almond (Prunus dulcis, Prunus amygdalus, Amygdalus communis) An oleate-enriched oil (Table 6.5) (Watkins, 2005), though its fatty acid composition is variable, is commonly used in skin-care products.

6.4.3 Avocado (Persea americana, Persea gratissima) The lipid in avocado is concentrated in the fruit with little in the seed. It is widely used in cosmetic products, being easily absorbed by the skin. It is also sold as a high-oleate oil (Table 6.5) for food use and is marketed in New Zealand as an alternative to olive oil (Birkbeck, 2002; Eyres et al., 2001).

6.4.4 Blackcurrant (Ribes niger) The seed oil from blackcurrant is of interest because it contains appreciable γ-linolenate (18:3Δ6cis,9cis,12cis) acid and a small amount of stearidonic acid (n-3, 18:4) (see Table 6.6). The oil is extracted from seeds that are by-products of juice production from berries and has uses in cosmetics and dietary supplements (Ucciani, 1995).

6.4.5 Borage (Borago officinalis) Borage is notable as a source of γ-linolenate (Table 6.6), which is reported to be of benefit for treatment of a number of diseases such as arthritis and skin complaints (Horrobin, 1992). Other commercial sources include blackcurrant (Ribes niger) and evening primrose (Oenothera biennis). Reports have been made on ways to isolate γ-linolenate or to enhance its level in borage. For general reviews, see Gunstone et al. (2007).

6.4.6 Borneo Tallow (Shorea stenoptera) Also known as illipe butter, this solid fat contains 43% stearate (Table 6.5) and thus has major TAG species with this acid (34% POSt, 47% StOSt). It is one of the permitted tropical fats that can partly replace cocoa butter in chocolate (Campbell, 2002; Timms, 2003).

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Fatty Acids

TABLE 6.5 Fatty Acid Composition of Minor Oils 16:0

18:0

18:1

18:2

18:3

Almond

49

13

6286

2030

Tr.

Avocado

1920

Tr.1

4558

1113

1

Borneo tallow

18

43

37

Tr.

Camelina

8

3

17

23

Cocoa butter

2425

3337

3337

34

Coconut

810

24

510

Coriander

5

1

Cottonseed

23

Tr.

Cuphea

133

Echium

6

E. lagascae

4

Evening primrose

6

3

2

31

12% 20:1, 3% 22:1

13

Tr.

See Tables 6.3 and 6.7

6

15

1

72% petroselinic acid (Table 6.7)

17

56

Tr.

110

133

Different species have up to 95% 8:0, 10:0, 12:0, or 14:0

14

13

See Table 6.6

19

9

See Table 6.7

9

72

See Table 6.6

Flax

57

26

1440

1429

3560

Hazelnut

24

12

3078

1530

Tr.

J. curcas

1017

510

3664

1845

Maize

1215

13

2632

5461

1

23

9

10

1953

2453

15

Mustard

Others

Oats

1328

Olive

820

24

5383

421

49

Peanut

1014

23

4353

2736

Tr.

Pine nuts

6

Tr.

25

46

Tr.

Poppy

10

11

72

5

Rice bran

1228

24

3550

2942

12

Safflower

7

3

14

75

Shea

48

2358

3368

48

Tall

23

3046

3645

43% erucic acid

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TABLE 6.6 Oils Containing Significant Amounts of γ-Linolenic or Stearidonic Acids Seed

Fatty Acids (%) Total 16:0

18:0

18:1

18:2

γ-18:3

18:4

Other

Blackcurrant

7

2

11

47

17

3

13a

Borage

10

4

16

38

23

Tr.

9b

Echium

6

3

14

13

12

17

35c

Evening primrose

6

2

9

72

10

Tr.

1

Tr., trace (,0.5). a α-Linolenate. b Long-chain monoenes. c Includes 33% α-linolenate. From Gunstone et al., 2007. The Lipid Handbook, third ed. CRC, Boca Raton, FL.

6.4.7 Camelina (Camelina sativa) (Section 6.5 Also) Camelina has attracted increasing interest recently (see also Section 6.5), not least because it grows well on marginal land and requires less fertilizer and pesticides than many traditional crops (Leonard, 1998). The seed yield can be up to 3 t ha21 with an oil content of 36%47%. It has high content of linoleic and α-linolenic acids with significant 20 and 22C monoenes (Steinke et al., 2000a,b). Camelina is easy to genetically manipulate and several laboratories have been exploring its potential as a source of unusual fatty acids including “fish oiltype” very long-chain n-3 acids (Petrie et al., 2012, 2014; Ruiz-Lopez et al., 2014).

6.4.8 Castor (Ricinus communis) Castor oil contains a very high concentration of ricinoleate and is grown mainly in India, China, and Brazil. It has a number of uses and is a starting point for the production of specialized chemicals. However, the presence of an acute poison, ricin, in the plant (not the oil) has led to a reduction in its growth in many countries (see Caupin, 1997; Gunstone et al., 2007; McKeon, 2016a).

6.4.9 Cocoa (Theobroma cacao) The fat from cocoa beans has been exploited since the Aztecs. Small differences in fat quality are found in different growing regions, with South American crops being more unsaturated. The fat is rich in palmitate (32%39%), oleate (32%39%) and, notably, stearate (30%36%).

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Fatty Acids

Seventeen TAG species are reported, which are crucial in producing desirable melting properties. Because of its importance in chocolate, the properties of cocoa butter have been extensively studied (Beckett, 2000; Padley, 1997; Shukla, 1997, Timms, 2003).

6.4.10 Coconut (Cocos nucifera) Grown mainly in the Philippines and Indonesia, coconut oil is rich in laurate (Table 6.3). It has great utility for small holders (Cassiday, 2016; Nguyen et al., 2015). For general reviews, see Canapi et al. (2005) and Pantzaris and Basiron (2002).

6.4.11 Coriander (Coriandrum sativum) Because of its high content of petroselenate, attempts have been made to develop coriander as an agricultural crop. Alternatively, the desaturase required for the production of petroselenic acid could be transferred to rape or soybean (Cahoon et al., 2006; Firestone, 2012; Gunstone et al., 2007).

6.4.12 Cottonseed (Gossypium hirsutum, Gossypium barbadense) Cottonseed oil is a by-product of cotton manufacture and used to be much more important as a vegetable oil than it is now. China is the main producer and consumer. The oil is high in linoleate (56%). It has been genetically engineered to produce high-oleate oils or those with high saturates so they could be used to form spreads, but these lines are not yet commercially available (Gunstone et al., 2007). Cottonseed oil is generally used for the production of cooking fats and spreads (O’Brien, 2002, 2005).

6.4.13 Crambe (Crambe abyssinica, Crambe hispanica) (Section 6.5 Also) Crambe seeds contain high levels of the industrially useful erucic acid (50%55%). Now that most modern varieties of rapeseed are low in erucate content (e.g., canola), Crambe may be a good alternative source of erucate. Its oil can be used for chemicals and surfactants used in the production of plastic bags, personal care products, and detergents (Erhan and Adhvargu, 2005).

6.4.14 Cuphea spp. Different varieties of Cuphea accumulate large amounts of 814C saturated fatty acids. The initial problems with seed dormancy and shattering that

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prevented their development in agriculture have now been solved so Cuphea has considerable potential as a speciality crop in the future (Firestone, 2012).

6.4.15 Dimorphotheca (Dimorphotheca pluvialis) Seeds of this plant typically have about 20% oil but about 60% of this is dimorphecolate, an unusual hydroxy fatty acid, which is a convenient source of hydroxyl- and 9-oxostearate and of hydroxyl epoxy esters (Firestone, 2012).

6.4.16 Echium (Echium plantagineum) Echium contains significant amounts of stearidonic acid, a potentially valuable nutraceutical (Table 6.6) (Kallio, 2003). Attempts are being made to domesticate it for this reason—the only other useful source being blackcurrant seed oil, where levels of stearidonate (18:46cis,9cis,12cis,15cis) are only about 3% (Gunstone et al., 2007).

6.4.17 Flax (Linum usitatissimum) Flax (or linseed) is a traditional source of α-linolenic acid (35%60%, Table 6.5), although production has declined somewhat in recent years. It is used as a drying oil for paints and floor coverings (linoleum) and to produce epoxydized products (Erhan and Adhvargu, 2005). Recent interest in n-3 PUFAs has led to significant use in nutraceutical products. A form through random chemical mutagenesis with 72% linoleate is traded as “linola” (Green and Dribneuki, 1994).

6.4.18 Hazelnut (Corylus avellana) Hazelnut oil is rich in oleate (Table 6.5) and is sometimes used to adulterate the more expensive olive oil (Aparicio and Harwood, 2013). Methods for detecting such adulterations have been described, many being based on the presence of filbertone in hazelnut (Bewadt and Aparicio, 2003). For general comments, see Crews et al. (2005) and Watkins (2005).

6.4.19 Jatropha curcas (See Section 6.5) Jatropha curcas (Table 6.5) is grown predominantly in India, Indonesia, and Nicaragua. The kernel (called the physic nut) is 50% oil, rich in palmitate (16%), oleate (51%), and linoleate (23%). Recent interest centers on its use for biodiesel, especially in India (see Gunstone et al., 2007).

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Fatty Acids

6.4.20 Jojoba (Simmondsia chinensis) Jojoba is a desert plant very resistant to drought and heat. It is grown in Southwestern United States, Mexico, Latin America, Israel, South Africa, and Australia. It takes at least 10 years to come into harvest but can then be used for 100 years. The oil is almost exclusively wax esters (Table 6.7) of 4044C, composed of monounsaturated acids and alcohols. It is a replacement for sperm whale oil, but due to its high value, it is mainly used in cosmetics. If the supply increases and the price drops, jojoba oil could be a superior lubricant (see Wisniak, 1987; Firestone, 2012; Erhan and Adhvargu, 2005).

6.4.21 Lesquerella (Lesquerella fendleri) (See Section 6.5) Although this is not yet a commercial crop, Lesquerella is being considered as a supplement to castor oil because the oil contains 54% lesquerolate (14-OH, Δ11-20:1, Table 6.7) and 4% auricolate (14-OH, Δ11,17-20:2). Processing of lesqueroate yields useful industrial chemicals and partial acylation of lesquerella oil with cinnamic acid (or 4-methoxycinnamic acid) produces compounds useful as sunscreens (Compton, 2005). Lesquerella plants are salt-tolerant and this property is being utilized and TABLE 6.7 Oils With Unusual Fatty Acids Castor (R. communis)

90% ricinoleic acid

Cocoa butter (T. cacao)

About 35% stearic acid

Coconut (C. nucifera)

72% medium-chain acids (mainly lauric)

Coriander (C. sativum)

72% petroselinic acid

Cuphea

Different species have up to 95% 8:014:0

D. pluvialis

About 60% dimorphecolic acid

E. plantagineum

12% γ-18:3, 17% stearidonic acid

E. lagascae

64% vernolic acid

Jojoba (S. chinensis)

Wax esters of 20 and 22C monounsaturated acids and alcohols

L. fendleri

54% lesquerolic acid

Meadowfoam (L. alba)

90% 20 and 22C monounsaturated (often Δ5) acids

Palm kernel (Elaeis guinensis)

67% medium-chain acids (mainly lauric)

Tung (A. fordii)

70% α-eleostearic acid

Vernonia

52%80% vernolic acid, depending on species

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developed further (see Abbott, 1997; Firestone, 2012; Isbell and Cermak, 2002; Erhan and Adhvargu, 2005).

6.4.22 Maize (Corn; Zea mays) Corn oil is a by-product of the starch industry with more than half of the total amount (2.05 MT) produced in the United States. The oil has palmitate (12%15%), oleate (22%23%), and linoleate (52%62%) as major constituents, with less than 1% α-linolenate. It is marketed as a healthy oil, low in saturates, and high in linoleate although, in the light of recent research about dietary fatty acids, this claim could be debated (Gurr et al., 2016). It does, however, show good oxidative stability (Gunstone et al., 2007).

6.4.23 Meadowfoam (Limnanthes alba) The oil from this plant contains .95% 20 and 22C acids (Table 6.7). The main components are 63%67% Δ5-20:1 and 16%18% Δ13-22:1. It is grown in the United States and winter cultivars suitable for northern Europe are being developed (Firestone, 2012; Isbell, 1997). The oil is used in cosmetics with potential as a lubricant, a source of chemical derivatives from reactions with the Δ5 bond of 20:1 (5-icosenoate) (Isbell, 1997, 1998; Isbell and Cermak, 2002), and can be converted to a solid wax (Erhan and Adhvargu, 2005).

6.4.24 Mustard (Brassica alba, Brassica carinata, Brassica hirta, Brassica juncea, Brassica nigra) All these plants accumulate erucate-enriched oil (25%40%). Brassica juncea (oriental mustard) has been bred to give low erucate and low glucosinolates [similar to canola lines (LEAR) of oilseed rape]. This could potentially expand the canola growing area in Canada (see Firestone, 2012; Gunstone et al., 2007 and Section 6.3 covering Brassica oilseed species).

6.4.25 Oats (Avena sativa) This cereal contains significant amount of oil (4%8% with some lines higher). Its fatty acid composition is shown in Table 6.5. Although TAG is the major oil constituent (51%), partial glycerides (7%), fatty acids (7%), glycolipids (8%), and phospholipid (20%) are also found in the oil. It has several uses in the food industry (Firestone, 2012; Herslof, 2000; Peterson, 2002).

6.4.26 Olive (Olea europaea) Olive oil only accounts for about 1.5% of total vegetable oils but it is highly prized, especially as a key component of the “Mediterranean diet.” The best

216

Fatty Acids

grades (Extra Virgin, Virgin) are obtained by pressing the fruits so that the oil [rich (56%83%) in oleate] contains many antioxidants. In part, these antioxidants contribute to the flavor. Because of its high price, olive oil is sometimes adulterated, occasionally with severe and fatal consequences. General aspects of olive oil are covered by Aparicio and Harwood (2013).

6.4.27 Peanut (Ground Nut, Arachis hypogaea) Peanuts are used as snack foods and in animal feeds. If stored poorly, they are prone to Aspergillus infection, which will produce the carcinogenic aflatoxin. There is also a significant risk of allergic reactions. The oil does not have these problems; it is present at 40%50% in the nuts and is rich in oleate (43%) and linoleate (36%, Table 6.5). These values vary with different varieties, all containing saturated and unsaturated very long-chain fatty acids (7%8%). The molecular species of TAG have been analyzed (Dorschel, 2002) and the phospholipid content defined (Singleton and Strikeleather, 1995). The main producers of ground nut oil are China (45%) and India (25%). Although only about half the peanuts harvested are used for oil extraction, the latter still represents about 3.5% of world vegetable oils (Gunstone et al., 2007, see also Pattee, 2005).

6.4.28 Pine Nuts (Pinus spp.) There are a large number of species of pine (Wolff and Bayard, 1995) with oil contents in the range 13%35% for 18 species examined. Notable amounts of Δ5 unsaturated fatty acids are found with pinoleate and sciadonate being common. Pinoleic acid has uses as an appetite suppressant (Watkins, 2005).

6.4.29 Poppy (Papaver somniferum) Poppy seeds have uses as birdseed and in baking but the oil (40%70%) is used as a semidrying oil by artists and as an edible oil (Table 6.5). Unlike the plant, the oil does not contain opium. It is linoleate-enriched (Gunstone et al., 2007).

6.4.30 Rice (Oryza sativa) Bran Oil Rice feeds half the world’s population and to produce white rice, the bran layer is removed. This bran layer makes up 8%10% of the rice grain so it is a sizable portion of the total crop harvest. Rice bran contains 18%24% oil, consisting of palmitate (12%28%), oleate (35%50%), and linoleate (29%45%) (Table 6.5), with a significant quantity of unesterified fatty acids, especially when the bran is stored (due to endogenous lipases).

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Although there is a potential for up to 8 MT of rice bran oil to be produced per year, present production is less than 1 MT. India (59%), China (14%), and Japan (10%) are the main countries producing rice bran oil. Although TAG is the main class, there are glycerolipids, phospholipids, waxes, and the aforementioned nonesterified fatty acids. Notable amounts of tocols and oryzanols give rice bran oil high oxidative stability so it is good to use as a salad or frying oil, or as a coating oil for biscuits and nuts (see Armughan et al. 2004; Kochar, 2001).

6.4.31 Safflower (Carthamus tinctorius) Produced mainly in the United States, Mexico, and India, safflower seeds have 38%48% oil, which is normally rich in linoleate (75%) (Table 6.5). High-oleate varieties have been produced. The regular oil is used as a starting point for the preparation of conjugated 18:2 acids and as a nonyellowing drying oil. The florets are used as a source of red or yellow coloring for foods and as dyes (see Gunstone et al., 2007).

6.4.32 Shea (Butyrospermum parkii, Shea Butter, Karate Butter) These trees, grown mainly in Western Africa, produce an oil with about 11% unsaponified components, including polyisoprene hydrocarbons. The oil also contains an exceptional level of stearate (23%58%) (Table 6.5) and can be fractionated to give a stearin, of use as a cocoa butter equivalent (Firestone, 2012; Shukla, 1996).

6.4.33 Tall Tall oil fatty acids are a by-product of the wood pulp industry when pine wood chips are digested and then chemically treated (Gunstone et al., 2007). It is produced mainly in North America and Scandinavia with the two oils differing somewhat in composition due to the tree species used. Either oil, however, is used to produce dimer acids, alkyols and coatings, detergents, and lubricants. There are future possibilities for use as solvents, inks and for biodiesel production (see Gunstone et al., 2007; Hase and Pojakkala, 1994).

6.4.34 Tung (Aleurites fordii) Tung oil (also called Chinese wood oil) contains a conjugated triene acid, α-eleostearic acid at 69% (Table 6.7). The oil dries quicker than linseed with less oxygen incorporation. Tung oil is mainly exported from China to other countries in the Far East or the United States (Ucciani, 1995). Nonfood uses include paints, coatings, varnishes, resins, and sealers. Of especial note is its use as coatings for food, drink, or medicine containers and as insulation for

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wires and metallic surfaces in communication hardware (Erhan and Adhvargu, 2005).

6.4.35 Vernonia Oils Various species of Vernonia contain seed oils with large amounts (up to 80%) of EFAs (especially vernolate) (Table 6.7). Attempts are being made to domesticate V. galamensis and E. lagascae [which also has high (52% 62%) vernolate] (Sherringham et al., 2003). EFAs have various industrial uses such as for adhesives, plasticizers, paints, and resins (see Section 6.3.2).

6.5 EMERGING INDUSTRIAL OIL CROPS Hundreds of oil plants have been identified for their seed oils containing fatty acids with unusual chemical properties. Although some of the plants have been domesticated as oil crops and used primarily for industrial application, many others were not well-suited to cultivation and required breeding in order to be cultivated as crops (McKeon et al., 2016). In this section, five representative emerging oil crops, camelina, crambe, pennycress, jatropha, and Physaria (Syn. Lesquerella), are briefly described. Camelina (C. sativa) is a member in the Brassicaceae family adapted to growth in temperate climates such as the northern portions of the United States and southern Canada. Camelina seed oil contains α-linolenate (over 30%), linolate (B23%), and monounsaturated fatty acids (18:1, 20:1, and 22:1; B28%). It is native to Europe and was an important crop historically. In North America, the first intentional planting of Camelina was recorded as early as 1863 (Scoggan, 1957). This oil-bearing plant is widely recognized as an excellent feedstock for second-generation biofuels and industrial feedstocks due to its many attractive features. These include high seed oil content, strong adaptability to many different environmental conditions, low input requirement, drought resistance and high-value meal with sufficient residual lipid content, and a protein profile similar to soy meal (Bansal and Durrett, 2016; Gugel and Falk, 2006; Murphy, 2016a,b). Camelina is amenable to rapid genetic modification, and compared with other emerging crops, there has been rapid progress in Camelina metabolic engineering. Camelina transcriptomic and genomic data are abundant (Bansal and Durrett, 2016; Kagale et al., 2014). Camelina can be efficiently transformed by Agrobacterium with a similar floral dip procedure as is used for Arabidopsis (Lu and Kang, 2008). Various studies have proven that fatty acid composition of camelina seeds can be modified by genetic engineering without the penalty of severe impacts on either the phenotype or germination of the seed. For example, transgenic Camelina has been developed, which can produce wax esters similar to that derived from sperm whale oil, hydroxy fatty

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acids and high levels of docosahexaenoate (22:6Δ4cis,7cis,10cis,13cis,16cis,19cis), and/or eicosapentaenoic acid (20:5Δ5cis,8cis,11cis,14cis,17cis) (Iven et al., 2016; Mansour et al., 2014; Petrie et al., 2014; Ruiz-Lopez et al., 2014; Snapp et al., 2014). Engineering Camelina can also produce a high level of cis-vaccenic acid (18:1Δ11cis), which has high value in pharmaceutical and industrial applications (Nguyen et al., 2013). In another study, when Camelina was transformed with the Euonymus alatus DIACYLGLYCEROL ACETYLTRANSFERASE(EaDAcT) gene with RNAi suppression of endogenous DGAT1, the seeds accumulated acetyl-TAG up to 85 mol% in fieldgrown trial (Liu et al., 2015). The Camelina acetyl-TAG oils have reduced viscosity, freezing point, and caloric content, enabling use of this oil in several industrial applications. Overall, Camelina is an ideal platform for production of specific fatty acids for use in industrial applications. Crambe (C. abyssinica Hochst) is another member of the Brassicaceae family, which originated in the Mediterranean region and parts of Eastern Africa (Weiss, 2000). Although Crambe has been planted as an annual oilseed crop for some time in certain regions, the interest in developing this plant as an industrial crop has increased recently (Zhu, 2016). Crambe seeds contain about 22%26% protein and 26%38% oil. The major fatty acid of Crambe seed oils is erucate (50%65%) (Finiguerra et al., 2001; Lazzeri et al., 1994; Wang et al., 1995). Erucate is an important feedstock in the oleochemical industry and has potential uses for hydraulic fluids, lubricants, additives, and starting material for plastics and nylon (Carlsson et al., 2011; Zhu, 2016). Currently, HEAR is the major commercial source of erucate (Table 6.1). Crambe has some advantages compared with rapeseed: its seed oil naturally contains higher erucate, it does not cross-pollinate with food oilseed crops and it is generally more resistant to diseases and insects (Li et al., 2016; Piazza and Foglia, 2001). Crambe seed cake contains high levels of glucosinolates (3%6%, w/w) and thus cannot be directly used as animal feeds, but the high protein and fiber contents make Crambe seed cake valuable in potential nonfood applications. Crambe has low genetic variability (Warwick and Gugel, 2003), which limits its genetic improvement through conventional breeding methods. Concerted efforts to domesticate Crambe have only begun within the past decade. Some progress has been achieved recently, which demonstrated the feasibility of developing Crambe into a bioplatform for industrial feedstock production. For example, several efficient transformation methods have been established (Chhikara et al., 2012; Gła˛b et al., 2013; Li et al., 2013; Qi et al., 2014). Transgenic Crambe lines with high erucate (73%), high-oleate, or even wax esters were also generated in proof-of-concept metabolic engineering studies (Li et al., 2016; Zhu, 2016). Another interesting plant in the Brassicaceae family is pennycress (Thlaspi spp.). This plant was recently identified as a potential oilseed crop for biofuel production and other industrial applications because it has high

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productivity potential (up to 840 L ha21 oil and 1470 kg ha21 press-cake) and high seed oil content (up to 38%) (Sedbrook et al., 2014). The seed oil contains erucate (27.5%38.4%), linolenic acid (8.4%15.2%), and oleate (7.7%17.1%) (Phippen and Phippen, 2013). In addition, since pennycress has a short life cycle and good cold tolerance, it could serve as a winter oilseed producing cover crop. A breeding program for pennycress was recently initiated in an effort to develop alternative energy sources, and some advances have been made. Recently, the de novo assembly of the comprehensive gene expression profile (transcriptome) in pennycress and a draft pennycress genome sequence were published, which would benefit the direct molecular breeding of pennycress (Dorn et al., 2013, 2015). Moreover, over 100 populations of pennycress have been evaluated for various agronomic traits, which indicated that pennycress is a tremendous potential oil crop (Phippen et al., 2010a,b; Phippen and Phippen, 2013; Sedbrook et al., 2014). In order to breed pennycress into a readily cultivated crop plant, additional breeding efforts are necessary to solve some agronomic problems regarding seed dormancy, oil quality, seed glucosinolates, flowering time and maturation, pod shatter, and seed size. Jatropha (J. curcas), family Euphorbiaceae, is grown widely in tropical and subtropical areas in Latin America, Asia, and Africa and is native to Mexico (Dias et al., 2012). Jatropha seeds contain 27%40% oil, which is rich in palmitic acid (C16:0, 13.4%15.3%), oleate (34.3%45.8%), and linoleate (29.0%44.2%) (Meher et al., 2013) (Table 6.5). This oil-bearing plant has some positive attributes that make it unique among oilseed crops such as drought tolerance, pest resistance, rapid growth, and short development period (Meher et al., 2013). Jatropha was believed to be able to grow and fruit on marginal or nonagricultural areas with relatively low inputs required. However, Jatropha seeds generally contain toxic components including curcin, which is a ribosome inactivating protein and phorbol esters that are irritants and promote tumors (He et al., 2011; Nakao et al., 2015). Due to the toxic nature of the plant and the extracted oil, Jatropha oil is inedible. It is, however, valuable for biodiesel production, especially in countries where the food/fuel debate has plagued the use of food oil crops in industrial applications. Jatropha has generated much excitement: over 1500 articles have been published in the past decade (Edrisi et al., 2015). Due to the recent upsurge in developing biodiesel, large-scale planting of Jatropha has been initiated (over 10 million ha globally) even before comprehensive agronomic improvement and evaluation (Singh et al., 2014). Unfortunately, the growth performance and oil yield of Jatropha on largescale plantations were far lower than expected. For instance, the actual seed yields range from 0.5 to 2 t ha21, which is much lower than being expected (212 t ha21) (Edrisi et al., 2015; Singh et al., 2014). The plant is also not very resistant to diseases, and when planted on marginal lands, Jatropha cannot grow well with high oil yield as commonly believed (Sanderson, 2009;

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Singh et al., 2014). The seed oil yield of Jatropha is much lower than other oil-producing plants such as oil palm (e.g., E. guineensis), rapeseed (B. napus), and coconut (C. nucifera) (Yue et al., 2013). Domestication of Jatropha is ongoing with both conventional and molecular breeding approaches being used to increase the oil yield and solve other agronomic problems such as seed yield, oil content and composition, female to male flower ratio, synchronous flowering and fruiting, oil quality, and branch number, resulting in some improvements (Carels, 2009; Edrisi et al., 2015). The potential for Jatropha to become a significant industrial oil crop is vast but additional efforts in the domestication of Jatropha and improvement in agronomic practices are crucial. Hydroxy fatty acids (see Table 6.7) have hundreds of applications in the production of industrial materials including lubricants, functional fluids, solvents, plastics, inks, paints, and cosmetics (Dyer et al., 2008; Napier, 2007). Currently, the only commercial hydroxy fatty acid is ricinoleic acid (12-OH 18:1Δ9cis; hereafter 12-OH 18:1) from castor (R. communis) oil. However, castor bean is not suited to large-scale agricultural production due to the presence of the highly toxic protein ricin in the seeds (Lee et al., 2015). As a result, the annual production of castor oil is only 645,000 t, which significantly limits the application of hydroxy fatty acids in industrial applications (McKeon, 2016a). Physaria fendleri (also known as L. fendleri), a Brassicaceae family plant native to the Americas, accumulates about 24% of oil in seeds in which over 60% of the fatty acids are lesquerolate (14-OH 20:1Δ11cis; hereafter 14-OH 20:1) (Dierig and Ray, 2009; Dierig et al., 2011). Recent studies indicated that P. fendleri has good agronomical characteristics (Dierig et al., 2011). This plant grows well in areas with 250400 mm of rainfall and thus is ideal for the semiarid regions of North America. P. fendleri can tolerate freezing temperatures and thus can be planted in fall and harvested in June throughout the Southwestern United States. This plant will not compete with current commodity crops but can be placed in rotation with them. Therefore, P. fendleri is considered as an emerging oil crop for the production of hydroxy fatty acids (Dierig and Ray, 2009; Dierig et al., 2011). Since the main use of the Physaria plant is lesquerolate production, traits that enhance the oil content are of great interest. Breeding efforts are also concentrating on agronomic characteristics such as soil temperature, moisture, depth of planting, planting dates, planting methods, and nitrogen fertilizer (Cruz et al., 2012, 2013a,b, 2014; Dierig and Crafts-Brandner, 2011; Dierig et al., 2011, 2012; Liu et al., 2014a; Pastor-Pastor et al., 2015; Windauer et al., 2013). Several significant advances have been recently achieved. For example, seed oil content of P. fendleri has been improved from 24% to over 30% and the current seed yield is approximately 1800 kg ha21. Modern molecular technologies and genetic engineering have contributed to Physaria research. For instance, an effective transformation system was

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established for the stable genetic transformation of P. fendleri (Chen, 2011), and a P. fendleri seed transcriptome has been established for discovering genes encoding enzymes involved in the synthesis of TAGs (Kim and Chen, 2015). Moreover, a recent study indicated that although lesquerolic acid is mostly at the sn-1 and sn-3 positions of TAG in seeds, the expression of a castor LPAAT2 gene could increase 12-OH 18:1 at the sn-2 position of TAG from 2% to 14%17% (Chen et al., 2016). The results indicated that metabolic engineering can further optimize P. fendleri for the production of hydroxy fatty acids. In addition, Physaria genes have been used in engineering other plants to produce hydroxy fatty acids (Broun et al., 1998; Lee et al., 2015). In summary, Camelina, Crambe, pennycress, Jatropha, and Physaria are potential new oil crops for industrial applications and considerable breeding efforts have been made in their domestication. Additional breeding advances, however, will be needed to bring them into large-scale agricultural production. In addition, many plants, such as chia (Salvia hispanica), Cuphea spp., D. pluvialis, E. lagascae, and Lunaria annua, produce valuable fatty acids but have not yet been successfully domesticated into large-scale growing (see Section 6.4). These plants may eventually be cultivated for industrial oil production or merely serve as genetic sources for plant biotechnology (McKeon et al., 2016; Vanhercke et al., 2013).

6.6 PROSPECTS FOR PRODUCTION OF INDUSTRIAL OILS IN VEGETATIVE TISSUE The large-scale production of TAG in vegetative tissue has the potential to provide an enormous global supply of oil for liquid biofuel and other industrial applications without affecting the supply of seed oil for food and feed applications. Unlike developing seeds of oleaginous crops or mesocarp tissue, leaves generally produce low amounts of TAG (up to about 0.5% dry wt) (Lin and Oliver, 2008). In chloroplasts, de novo fatty acid synthesis supplies acyl chains in support of membrane lipid production and active development of leaves (Chapman et al., 2013; Vanhercke et al., 2014a). TAG only appears to act as a transient storage depot for sequestering free fatty acids in leaves during membrane alteration and turnover. In contrast, developing seeds act as a “sink” organ, where TAG accumulates to high levels. In recent years, great advances have been made in reprogramming vegetative tissue to accumulate TAG to levels .15% (dry wt) (reviewed by Chapman et al., 2013; Vanhercke et al., 2014a; Xu and Shanklin, 2016; Weselake, 2016). Much of the inspiration for these initiatives appears to have come from the knowledge and insight gained in exploring different metabolic engineering strategies to boost seed oil content. Strategies for increasing oil content in vegetative tissue have included increasing the supply of building blocks for TAG production, increasing TAG production, and

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reduction of TAG turnover. The most advanced engineering experiments have combined the earlier interventions. Enriching leaf TAG in monounsaturated fatty acids could result in a suitable feedstock for the production of biodiesel. For example, combined expression of cDNAs encoding the WRINKLED1 transcription factor, DGAT1, and oleosin in tobacco (Nicotiana tabacum) resulted in leaf TAG enriched in 18:1 with decreased α-18:3 content (Vanhercke et al., 2014b). Proof-of-concept studies by Reynolds et al. (2015) using a transient leaf expression system in N. benthamiana have shown that combined expression of cDNAs encoding WRINKLED1, DGAT1, medium-chain thioesterase, and coconut LPAAT resulted in significant levels of medium-chain fatty acids in the leaf TAG that was produced. Perennial C4 grasses, such as switchgrass (Panicum virgatum L.) and sugarcane (Saccharum spp.), are of particular interest because of their efficient production of high levels of biomass. Zale et al. (2016) implemented several metabolic interventions to engineer sugarcane, which accumulated about 1% and 2% TAG in leaf and stem tissue, respectively. It was estimated that each percentage of TAG produced in transgenic sugarcane corresponded to the TAG produced in the same land area as B. napus. Investigations are also underway to engineer rapidly growing trees, such as poplar (Populus spp.), to generate TAG in their woody stems (Nookaraju et al., 2014), once again suggesting the potential for yet another high biomass source of plant oil.

ACKNOWLEDGMENTS J.L.H. acknowledges the support of the Biotechnology and Biological Sciences Research Council (United Kingdom), The Malaysian Palm Oil Board, and Arcadia BioSciences. R.J.W. acknowledges the support of Alberta Enterprise and Advanced Education, Alberta Innovates Bio Solutions, AVAC Ltd., the Canada Foundation for Innovation, the Canada Research Chairs Program, and the Natural Sciences and Engineering Research Council of Canada.

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FURTHER READING Ackman, R.G., 1990. Canola fatty acids—an ideal mixture for health, nutrition, and food use. In: Shahidi, F. (Ed.), Canola and Rapeseed: Production, Chemistry, Nutrition, and Processing Technology. Van Nostrand Reinhold, New York, pp. 8198.

Chapter 7

Microbial Production of Fatty Acids Colin Ratledge1 and Casey Lippmeier2 1

University of Hull, Hull, United Kingdom, 2DSM Nutritional Products, Columbia, MD, United States

Chapter Outline 7.1 Introduction 237 7.2 The Process of Lipid Accumulation in Oleaginous Microorganisms 241 7.3 Economic Considerations— Heterotrophic Microorganisms 244 7.4 Economic Considerations— Phototrophic Microorganisms 248 7.5 Production of PUFAs 251 7.5.1 Nutritionally Important Fatty Acids—Background Information 251 7.5.2 Production of Gamma-Linolenic Acid (GLA 18:3 n-6) 255

7.5.3 Production of Arachidonic Acid (ARA 20:4 n-6) 258 7.5.4 Production of Docosahexaenoic Acid (DHA 22:6 n-3) 259 7.5.5 Production of Eicosapentaenoic Acid (EPA 20:5 n-3) 260 7.5.6 Production of EPA/DHA Mixtures as Alternatives to Fish Oils 264 7.6 Safety Aspects 266 7.7 Future Prospects 268 References 270

7.1 INTRODUCTION The majority of the world’s supply of oils and fats are derived from plants and animals. These are, almost invariably, in the form of triacylglycerols, colloquially known as triglycerides. Only a tiny proportion of the total is produced by microorganisms simply because the means of producing them is much more expensive than obtaining them from plants. Animal fats, which are mainly produced as by-products from the meat industry, have historically also been relatively inexpensive. Thus, if we have to rely upon the technology of large-scale fermentations to grow microorganisms in sufficient quantities to provide realistic and useful amounts of their triglyceride oils, there is an overriding need to produce high-value oils and fats to offset the high costs of production. Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00006-4 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

237

238

Fatty Acids

Although small amounts of specialty fats are made in commercial quantities by bacterial fermentation, most microorganisms that will be described in this chapter are representatives of the more complex eukaryotic class of microbes that encompass yeasts, fungi, and microalgae. Among eukaryotes, the occurrence of oils and fats in yeasts and fungi has been known since the end of the 19th century. Early studies on microbial oils in the first decades of the 20th century established equivalence between their fatty acids and those found in plants and animals (Woodbine, 1959). By the time of the Second World War (193945), scientists in Germany, who had pioneered much of the earlier studies, considered that microbial oils could be used for human consumption though, in practice, this did not take place. Instead, the microorganisms that were grown on a modest industrial scale for oil production were fed as the entire biomass without oil extraction to army horses with no ill effects (Bunker, 1945; see also Ratledge, 2005). With the advent of gasliquid chromatography analysis in the 1960s, it quickly became apparent that the fatty acids present in most microorganisms were the same as those found in higher organisms and there was no a priori reason to assume that these would be in any way harmful to the humans or animals that might ingest them. Analysis also established that fatty acids were stored in microorganisms in the same form of triacylglycerols that occurred in plants and animals (Shaw, 1966). Thus, to all intents and purposes, microbial oils were regarded equivalent to those already being produced commercially. Those microorganisms that accumulate oils within their cells have been termed “oleaginous”—simply meaning “oil-bearing” (Thorpe and Ratledge, 1972). Originally (Ratledge, 1982), it was suggested that the term should only be applied to any microorganism accumulating more than 20% of its biomass as storage lipid as amounts slightly less than this could simply be due to peculiarities of the growth of the organism in question. The minimum limit of 20% lipid has, in retrospect, been found to be a useful empirical standard to define oleaginicity. Examples of various oleaginous microorganisms are given in Table 7.1 together with their lipid contents and fatty acids. It should, though, be appreciated that there can be considerable variation in the lipid levels in an oleaginous species of microorganism that will depend on the particular strain being used as well as how it is being cultivated (for example, see Sitepui et al., 2013). A very useful overview of the oil contents of yeasts and other fungi as well as thraustochytrids and other chromalveolates as the principal oleaginous microorganisms has been complied by Ochsenreither et al. (2016) and an equally useful summary of the lipids of microalgae has been presented by Bellou et al. (2014). The term “oleaginicity” can be applied both to heterotrophically growing microorganisms, that is those that need a fixed carbon source (usually glucose or sucrose) for growth, and also to photosynthetically growing organisms, such as the microalgae that can use sunlight as their energy source and

TABLE 7.1 Oil Contents and Lipid Profiles of Selected Oleaginous Yeasts, Fungi, and Microalgaea Maximum Lipid Content (% w/w)

14:0

16:0

Cryptococcus curvatus

58

trace

32

Lipomyces starkeyi

63

trace

34

Rhodotorula glutinis

72

trace

Rhodosporidium toruloides

66

Yarrowia lipolytica

16:1

18:0

18:1

18:2

18:3 (n-3)

18:3 (n-6)

15

44

8

6

5

51

3

37

1

3

47

8

trace

18

3

3

66

36

trace

11

6

1

28

51

1

Aspergillus terreus

57

2

23

trace

14

40

21

Cunninghamella echinulata

24

trace

13

2

46

16

19.5

Mortierella alpina

50

11

14

7

14

Mucor circinelloides

25

18

Pythium irregulare

.25

20:4 (n-6)

20:5 (n-3)

22:6 (n-3)

Yeasts

Fungi

1

8 1

22

1

6

40

11

17

7

2

14

18

49

11

14 (Continued )

TABLE 7.1 (Continued) Maximum Lipid Content (% w/w)

14:0

16:0

16:1

18:0

18:1

Crypthecodinium cohnii b

B50

20

18

2

,0.5

15

Isochrysis galbana

B2230

12

1

11

3

2

Nannochloropsis oculata

3035

4

15

22

3

Porphyridium cruentum

B10

30

5

,1

Schizochytrium sp.b

40

22

,0.5

18:2

18:3 (n-3)

18:3 (n-6)

20:4 (n-6)

20:5 (n-3)

22:6 (n-3)

Microalgaeb

a

8

0.5

Data from Ratledge (2013). Grown phototrophically except for the nonphotosynthesizing C. cohnii and Schizochytrium sp. Also contains B17% 22:5 (n-6).

b c

1

40 25 1

4

11

38

16 41c

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CO2 as their carbon source. But, phototrophic algae need to be considered separately from heterotrophic microorganisms as there are considerable difficulties in ensuring a plentiful supply of CO2 for them. The oils obtained from microbes have been termed as “single-cell oils” (SCOs) in a deliberate attempt to mimic the term “single-cell proteins” that was used in the 1960s and 1970s to indicate the proteins produced by microorganisms that were destined for animal consumption (Ratledge, 1976). This term is now in general use and refers to oils that are intended for both human and animal consumption as well as those being considered for use in the biodiesel industry for the production of methyl fatty acid esters. As no such commercial process has yet been developed specifically for producing SCOs for the biofuel market, this chapter will not cover these aspects.

7.2 THE PROCESS OF LIPID ACCUMULATION IN OLEAGINOUS MICROORGANISMS To engender lipid accumulation in a microorganism, it is essential to manipulate its metabolic pathways so that cells do not continue to multiply beyond a certain limit. This is usually accomplished by growing the organism in a culture medium with a limiting amount of available nitrogen in it. Other nutrients besides nitrogen can be used but nitrogen limitation is the usual choice. The culture medium, however, also needs to have a plentiful supply of carbon, usually in the form of glucose although, again, other carbohydrate feedstocks can be used if these are relatively cheap. The course of lipid accumulation in an oleaginous microorganism is shown in Fig. 7.1. There are essentially two distinct phases of the culture of an oleaginous microorganism. In the first phase, the balanced phase of growth, the cells have all nutrients available to them and therefore grow as rapidly as possible. This phase ends when the nutrient chosen to be growth limiting is exhausted. When this happens, as for example with N limitation, cells are unable to synthesize further amounts of proteins and nucleic acids as both these essential components of the cell require N for their synthesis. Thus, cells are no longer able to multiply but they continue to be metabolically active. They continue to take up the carbon source still remaining in the medium and enter the second phase of the process—lipid accumulation. For an oleaginous microorganism, the carbon source is now preferentially channeled into lipid biosynthesis. For a nonoleaginous organism placed in the same culture medium with a high ratio of C:N, they may convert the remaining glucose into some form of storage polysaccharide material or may even use it to produce increased amounts of various metabolites that could then spill out of the cells into the culture medium. A simple example of the latter would be citric acid from the fungus Aspergillus niger, which is used for the commercial production of this organic acid.

242

Fatty Acids

Balanced growth

100

Lipid accumulation 60

75 40

Nitrogen

Lipid

50

20 25 Glucose

0

0

25

50 Time (arbitrary scale)

75

Biomass (dry wt) Lipid (% dry wt)

Glucose/Nitrogen (arbitrary values)

Biomass

0 100

FIGURE 7.1 Course of lipid accumulation of an oleaginous microorganism during growth.

How the oleaginous microorganisms are able to affect the conversion of the feedstock substrate, e.g., glucose, into triacylglycerols has been studied over the past three to four decades with there now being a reasonable view of how this is accomplished at the biochemical level (see, for example, Botham and Ratledge, 1979; Ratledge and Wynn, 2002; Ratledge, 2014). An outline of this aspect of metabolism is given in Fig. 7.2. The key to the process is that when the cells switch from the balanced phase of growth into the lipid accumulation phase, there is no longer a need for the cells to produce a high amount of metabolically available energy, which is in the form of adenosine triphosphate (ATP) to keep synthesizing new cells and cellular components. The production of ATP occurs in the mitochondrion of the cells, and this involves both the action of the tricarboxylic acid (TCA) cycle (or the Krebs cycle) and the process of oxidative phosphorylation whereby the reduced cofactors involved in the enzyme reactions are converted to the oxidized counterparts and simultaneously form ATP from adenosine diphosphate (ADP). One of the key reactions is that catalyzed by isocitrate dehydrogenase (ICDH) (see Fig. 7.2). In the oleaginous microorganism, this enzyme becomes deactivated as it has a specific requirement for a high concentration of adenosine monophosphate (AMP) to be available (Botham and Ratledge, 1979). The concentration of AMP in the mitochondrion drops very rapidly when the cells exhaust their N source and is caused by a sudden increase in activity of a deaminating enzyme, AMP deaminase. This appears to be an attempt by the cells to obtain additional N from their own internal resources. AMP deaminase effectively deaminates

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FIGURE 7.2 Outline of the main sequence of events leading to lipid accumulation in oleaginous microorganisms. Lipid accumulation is triggered by a sequence of events described in the text. The concentration of AMP is controlled by AD which converts it into IMP. ICDH, isocitrate dehydrogenase (AMP dependent); TCA cycle, tricarboxylic acid cycle; ACL, ATP:citrate lyase; FAS, fatty acid synthase; MDH, malate dehydrogenase; ME, malic enzyme; AD, AMP deaminase; IMP, inosine monophosphate (see also Figure 7.3).

AMP into inosine monophosphate (IMP) with the consequence that ICDH is no longer active. This, in turn, stops the citric acid cycle from operating and causes the substrate of the enzyme, isocitrate, to build up in the mitochondrion. Isocitrate equilibrates with citrate and citrate then exits from the mitochondrion into the cytosol of the cell. Some citrate also escapes from certain cell types into the culture medium. Citrate now becomes the substrate of what is the second key enzyme involved in lipid accumulation: ATP:citrate lyase (ACL). This enzyme is uniquely only found in oleaginous cells and is not present in the nonoleaginous strains; it can therefore be regarded as a marker enzyme for oleaginicity. It converts citrate into acetyl-coenzyme A (acetyl-CoA) and oxaloacetate. The acetyl-CoA then becomes the substrate for lipid biosynthesis using the ubiquitous fatty acid synthase (FAS) system. At the same time (see Fig. 7.2), the other product from ACL, oxaloacetate, is reduced to malate using malate dehydrogenase that involves the participation of NADH (reduced nicotinamide adenine dinucleotide). The malate is further converted into pyruvate by another key enzyme of the oleaginous cell, malic enzyme (ME).

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This latter enzyme decarboxylates malate into pyruvate with the simultaneous conversion of NADP1 into NADPH. The pyruvate can then be reused to form further amounts of citrate and the NADPH then becomes the requisite reductant to drive fatty acid biosynthesis. The sequence of reactions, as shown in outline in Fig. 7.2, then provides the basis for understanding the process of oleaginicity (Ratledge and Wynn, 2002; Ratledge, 2014). Although some variations on this may occur, as not every oleaginous microorganism contains an active ME, and there may be alternative routes to deactivating ICDH and controlling the activity of the TCA cycle, the route provides a basis for calculating the maximum lipid yield that can be obtained by using glucose as feedstock in a fermentation process. The stoichiometry of the conversion of glucose to triacylglycerol is given in Fig. 7.3. This indicates that the theoretical maximum yield is 31.6 g triacylglycerol from 100 g glucose (see Ratledge, 2014). For those microorganisms lacking ME, an alternative means of recycling NADP1 back to NADPH has to be used and this is considered to be by the reactions involved in the pentose phosphate cycle used for the metabolism of glucose into pentoses and tetroses. This, however, is not as efficient as the ME system and the theoretical yield now drops to 27.6 g triacylglycerol from 100 g glucose. Other means of generating NADPH, although, cannot be ruled out (see Dulermo et al., 2015). The values given above are, however, theoretical yields based on an idealized fat profile and do not take into account that some glucose must be used to produce all the other components within a cell. The maximum practical yields that have been attained have mainly used continuous culture systems that maximize the efficiency of the growth process. Yields of 2022 g TAG/100 g glucose have been reported, which might be as high as can be achieved using growing cells (Gill et al, 1977; Hassan et al., 1993; Ykema et al., 1988). A higher yield of 27 g oil/100 g glucose has, although, been reported when this is calculated for the cells after the onset of nitrogen exhaustion from the medium—in other words, in these conditions, the glucose is only being used for lipid biosynthesis and not for the production of new cells (Tai and Stephanopoulos, 2013). This cannot be attained in practice as some glucose must be used to produce nonlipid cell components. Thus, in general terms, some 5 tons of glucose are needed to produce 1 ton of lipid (i.e., a 20% conversion yield). This then places a severe economic restraint as to which microbial oils may be produced economically.

7.3 ECONOMIC CONSIDERATIONS—HETEROTROPHIC MICROORGANISMS Large-scale production of microorganisms was developed by some of the major petroleum (gasoline) companies during the 1960s and 1970s for the conversion of n-alkanes into SCP to be used as animal feed. BP Co. Ltd in

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FIGURE 7.3 Stoichiometry for the conversion of glucose to lipid involving the participation of malic enzyme. Numbers in parentheses indicate the required stoichiometry. Overall conversion: 4.5 glucose 1 CoA 1 9 NAD117 NADPH117 ATP 5 C18-fatty acyl-CoA 1 9 CO2 1 9 NADH17 NADP1117 ADP117 Pi. ACC, acetyl-CoA carboxylase; ACL, ATP:citrate lyase; CS, citrate synthase; EMP, reactions of the EmbdenMeyerhofParnas pathway; FAS, fatty acid synthase; MDH, malate dehydrogenase; ME, malic enzyme (NADP-dependent); PC, pyruvate carboxylase; PDH, pyruvate dehydrogenase; G3P, glycerol 3-phosphate; ?, unspecified reactions needed to provide the additional NADPH. From Ratledge, C., 2014. The role of malic enzyme as the provider of NADPH in oleaginous microorganisms: a reappraisal and unsolved problems. Biotechnol. Lett. 36, 15571568; with a minor correction.

the UK pioneered the production of SCP using a yeast, Candida (now Yarrowia) lipolytica, and went so far as constructing biofermenters up to 250 m3 for this purpose. Four such fermenters as part of a dedicated site were built in Sardinia, Italy, and were aimed at producing upwards of 10,000 tons of biomass rich in protein per year. But challenges to the long-term

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safety of the biomass being fed to cattle led to the process being halted by the Italian government. This, coupled with the rising cost of petroleum during the oil crisis of the 1970s and the falling price of soybean protein, as the main rival to SCP, then persuaded BP Co. Ltd to abandon this approach. ICI Co. Ltd in the United Kingdom pioneered the conversion of methanol (derived from methane being extracted as natural gas) into SCP using a bacterium. This involved one single reactor of 1500 m3 that represented, at the time, the largest single bioreactor in the world. Again, this process did not realize its potential and was also abandoned after a short period of production. These advancements in large-scale fermentation technology, however, had considerable repercussions throughout the biotechnology industries. They had a massive effect on the thinking of biotechnologists as to what might now be possible if such technology was transferable for the manufacture of other products. It obviously brought about considerable interest in how the technology might be applied to the production of microbial oils on a large scale. But a simple examination of the likely economics quickly led to the conclusion that it was unlikely to be useful for producing microbial oils of similar composition to the major commodity plant oils: soybean oil, palm oil, sunflower oil, etc. The major problem with the production of cheap microbial oils is not only the high cost of fermentation but also the cost of the feedstock material that has to be used. As discussed in Section 2, the very best conversions of glucose to lipid that can be achieved are unlikely to exceed a sugar to oil conversion rate of 22%. If we assume therefore that about 5 tons of sugar are needed to produce 1 ton of oil, we can quickly see the impossibility of economically producing a microbial that would rival the selling price of a plant commodity oil. Assuming we have an oleaginous microorganism that could use sucrose, which is derived from sugarcane and is the cheapest available sugar (but not every oleaginous microorganism can use sucrose which being a disaccharide requires either invertase or acidification to hydrolyze it into its component hexose sugars, glucose and fructose), then with the current (2016) cost of sucrose being about $400/ton, the cost of feedstock alone for the fermentation will be $2000 to produce 1 ton of oil. Adding to this the cost of operating the fermentation process and also the cost of oil extraction, the final price of the oil could be easily doubled: $4000/ton. As plant oils currently sell for $700800/ton depending on their source—soybean, sunflower, palm—it is evident that only the most expensive of microbial oils could be considered as economically viable. Thus, only the highest value microbial oils are realistic targets and these oils are currently the highly unsaturated oils used principally for nutritional purposes (see Section 5.1). These are discussed in the sections below. There have been numerous papers written on the topic of producing microbial lipids from cheap or even waste materials. If the objective of these

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researches is to produce nutritionally important fatty acids, then it is axiomatic that the feedstock being used must also be of food-grade quality. You cannot use impure materials or ones of uncertain provenance to produce any biotechnological product that is intended for human consumption. This immediately rules out the possible use of raw glycerol derived from the biodiesel industry. This substrate has been the favorite feedstock for numerous papers focusing on microbial lipid production. Unfortunately, raw glycerol contains residual and toxic methanol (used in the transesterification of the original oil into methyl fatty acid esters), has a high salt content as well as free fatty acids, and whatever other materials of a deleterious nature that might have been in the original oil or in the sulfuric acid being used in the process (Ciriminna et al., 2014). The final raw glycerol is usually very dark brown. Crude glycerol, even at 80% purity, cannot be used by traditional oleochemical refiners because it is clearly corrosive and would damage storage equipment and pipework (Ciriminna et al., 2014). Refinement of the glycerol is therefore essential for it to be used in fermentations but this then places a premium price on it. The current price of purified glycerol is about $1400/ton. This is about 33.5 times the price of sucrose and therefore immediately precludes its use as a fermentation feedstock. Other substrates that have been suggested have included a number of agricultural waste materials. Almost all of these are unsuitable for largescale process. For a waste material to be of use, it has to be available throughout the year as it is uneconomic to use a fermenter otherwise. It must also be storable as not all the material that becomes available can be used immediately. Finally, the effluent fermentation broth arising from the use of waste material will have a very high biological oxygen demand (BOD) by virtue of all the unused materials still remaining after the fermentation. Disposal of such broths with a high BOD is not cost-free and, at best, will involve using a purpose-built anaerobic digester. Proponents of the use of raw glycerol, however, point out that this material might be tolerated by microorganisms without having to undergo any refinement. (This is possibly an unrealistic hope as very few studies have been done to establish the usability of the authentic material as a feedstock.) As such, this material could then be used for producing microbial lipids not destined for human or animal consumption. Such a use would be the production of biodiesel. But, biodiesel is currently produced from cheap plant oils such as palm oil that currently sells for $700/ton. This then places a minimum price for the microbial oil to be produced and, in our opinion, it will not be economical to produce an oil for such a low price. It has been proposed that some decrease in process costs may be realized by the manufacture of sidestream products in a biorefinery model. Unlike petroleum refineries, however, fermentation-based biorefineries have far more flexibility to focus their processes on the highest value products in response to market demands. One has to also remember that biodiesel itself competes with

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petroleum (gasoline) and all the indications are that the price of crude petroleum on the world market will remain low at about $60$80 per barrel (about $420$600/ton) for the foreseeable future. The margins for profit in producing microbial lipids for anything other than use as high-value nutritional oils are therefore too low to be of any current commercial interest. The costs of producing a given SCO vary considerably depending on the natural productivity of the strain used, the design of the process, the facilities used to grow the cultures, the desired extraction, purity, refinement, and stability of the oil, and the regulatory requirements of the finished product. The first SCO produced, an oil rich in gamma-linolenic acid (GLA) derived from the fungus Mucor circinelloides (see Section 5.2), was aimed at being a direct competitor to evening primrose oil. At the time of production in the mid-1980s, this oil was selling for approximately $50/kg. Thus, we can speculate that the total costs of its production by fermentation technology would be somewhat less than this: e.g., $25/kg or $25,000/ton. More recent information about the price of nutritional oils from heterotrophic microbes has been reviewed by Borowitzka (2013). The reported price of these oils [containing polyunsaturated fatty acids (PUFA), sterols, xanthophylls, or carotenoids] is generally an order higher than the oil derived from M. circinelloides. The least expensive, low-grade microbial oils may be those proposed for use as feedstocks for biodiesel with profiles similar to those of plant oils currently used for biodiesel. However, it would be difficult to lower the production cost of any microbial oil to less than $10/kg even for the most efficient organism and process. In 2008, it was suggested that the minimum possible cost of producing a microbial oil would be well over $3/kg (Ratledge and Cohen, 2008); today’s value would therefore be double or treble that value.

7.4 ECONOMIC CONSIDERATIONS—PHOTOTROPHIC MICROORGANISMS The attractiveness of using microalgae for the production of oils, or indeed, any high-value product, lies in the simplicity of their growth requirements. They use sunlight as their energy source and CO2 for their carbon source. Both are free at the point of use. On paper, they should be the cheapest way of producing oils. But there are major problems. Many people have examined lipid production by algae growing in photobioreactors. These systems use either glass or clear plastic vessels or tubing as the basic fermentor and then are illuminated by fluorescent light. The systems are fine for small-scale investigations at the laboratory level but are completely impractical for scale-up to accommodate, e.g., 100 m3 of culture. Large arrays of clear plastic tubing have been tried out of doors but a basic problem with algal cultivation is the so-called phenomenon of “self-shading” whereby the increasing density of cells in the culture decreases the

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penetration of light, thus limiting the energy available for cell growth and carbon assimilation to achieve the biophysically defined maximum (Zaslavskaia et al., 2001). The biomass density that is attained with strictly photosynthetic cultures does not normally exceed about 5 g/L (Brennan and Owende, 2010). If this is compared with heterotrophically grown microorganisms that can reach up to 200 g/L in less than 3 days cultivation (see, for example, Barclay et al., 2010), then the first disadvantage of using photosynthetic algae becomes clear. The option for the cheapest growth of algae must then be those that minimize capital and process costs: in practice this means open ponds, man-made lagoons, or even sheltered coves next to the ocean. Use of the latter would, though, mean that marine algae would have to be used rather than those that require freshwater and cannot tolerate growth in seawater. If freshwater algae are used, then the supply of water becomes crucial: when algae grow outdoors in ponds or lagoons, they need warm conditions (B30 C) and as much sunshine as possible. This means that the locations have to be in dry areas in near-desert conditions located in the warmer areas of the world. However, locations cannot include much of the tropics as the advent of the usual storms in these areas would cause major disruption of the growth systems. But dry locations are, by definition, continuously short of water; water loss by evaporation then becomes a key factor as the supply of water in these areas is not a simple matter. Fig. 7.4 provides a view of a relatively small-scale open pond array being used for the cultivation of oleaginous microalgae. The next difficulty comes in providing a culture medium with a high ratio of C:N. Oleaginous algae are just like any other oleaginous microorganism and, for lipid accumulation to occur, they need a high content of C in their growth medium. This, for a photosynthetic organism, means CO2. But the content of CO2 in the atmosphere is far too low to promote high levels of lipid accumulation. Hence an additional supply of the concentrated gas is needed and this is expensive. There is also the problem as to how to maintain a high concentration of CO2 in the growth medium in an outdoor location. Where CO2 has been used to promote lipid accumulation, this has been in laboratory bioreactors or in tubular arrays placed outside in a warm and sunny location. It is usually inefficient to add CO2 in any meaningful manner to an outdoor system for algal cultivation. The gas is bubbled into the system but the majority (.95%) then is often simply lost into the atmosphere. Thus, it is almost impossible to engender high lipid contents in algae being grown outdoors no matter what type of growth system is being used. If algae are grown without providing CO2, then the total amount of lipid that is produced is no more than 10% of the biomass. In other words, 90% of what is produced is not lipid. After the lipid has been extracted from the cells, very little remains in the residual biomass that might be of primary commercial value and which could remain stable through the extraction procedures. It is therefore untenable to try growing photosynthetic algae for the production of

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FIGURE 7.4 Cellana open raceway facility for the R & D production of lipids, biomass and other products using phototrophically grown microalgae. The site is located on six acres of volcanic plain at the westernmost edge of Kona, Hawaii.

anything less than the very highest valued lipid products. This would include carotenoids, such as astaxanthin and beta-carotene, as well as the higher valued PUFAs. For example, Milledge (2011) cited an estimate of $920/kg for eicosapentaenoic acid (EPA)containing oil from the photosynthetic Phaeodactylum tricornutum as being economically viable. Finally, there is the problem with the types of lipid that algae produce. For the heterotrophic oleaginous microorganism, the oil that is produced is usually in the form of triacylglycerols and is therefore immediately acceptable for human consumption. With photosynthetically grown algae, they produce a range of lipid types of which most are associated with the photosynthetic apparatus. Thus, only a very small amount of triacylglycerol is produced; the majority (.90%) of the total extractable lipids are polar lipids: phospholipids and glycolipids. However, as is indicated in Section 5.5, polar lipids may, in certain circumstances, be an acceptable form of lipid to be used for nutritional purposes. Some companies using open pond systems for growing phototrophs have developed procedures to circumvent some of the aforementioned issues. For example, Cellana, whose Kona facility is pictured in Fig. 7.4, takes advantage of the biphasic nature of oleaginous cultures by growing seed cultures in closed bioreactors before inoculating to ponds for the oil accumulation

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phase. This system reduces issues with contamination and also enables greater delivery of concentrated CO2 during the initial growth phase. Recently, Cellana reported a demonstration conducted with the diatom Staurosira and the green alga Desmodesmus in which both species produced 75 metric tons of biomass per hectare per year and 30 metric tons of lipid per hectare per year (Huntley et al., 2015). However, only a relatively small number of companies are commercially producing lipids with phototrophs and, aside from the aforementioned study, very little information of the manufacturing costs has been released. In conclusion, the use of photosynthetic algae as producers of economic quantities of high-value lipids is limited. There seems little prospect that algae could be used for any purpose other than the production of high-value neutraceuticals and it is, in our opinion, quite unrealistic for them to be considered as competitive sources of biofuels or other products of equally low value in current market conditions.

7.5 PRODUCTION OF PUFAs 7.5.1 Nutritionally Important Fatty Acids—Background Information It has been known for over 90 years that some fatty acids are essential for the well-being of humans and other animals (Osborne and Mendel, 1920; Evans and Burr, 1927, 1928). These are the PUFAs. Some of these fatty acids, moreover, have to be provided in the diet as they cannot be synthesized de novo in the human body. This is shown diagrammatically in Fig. 7.5A: fatty acids of the n-6 and n-3 series cannot be synthesized in humans from oleic acid (18:1) and therefore both linoleic acid (LIN, 18:2 n-6) and alpha-linolenic acid (ALA, 18:3 n-3) must be obtained from the diet. Normally, there is no difficulty in this as both these fatty acids occur in most plant and animal products that make up our diet. Severe deficiencies of these fatty acids are rare but have been observed in experimental animals receiving very restricted diets (see Evans and Burr, 1927, for example) and among congenital sufferers of certain peroxisomal biogenesis disorders (reviewed by Klouwer et al., 2015). The conversion of ALA into the longer chain PUFAs may, however, proceed more slowly than it should for the required amounts of both EPA (20:5 n-3) and docosahexaenoic acid (DHA, 22:6 n-3) to satisfy our physiological requirements for them (Sinclair et al., 2002). Both EPA and DHA carry out important physiological roles in the body (Calder, 2006, 2009, 2013). They are both converted into a series of prostaglandins and related materials, including resolvins, thromoboxanes, leukotrienes, and protectins, which are anti-inflammatory, prevent platelet aggregation, and promote vasodilation (Serhan, 2005; Cottin et al., 2011;

(A) Fatty acid synthase

16:0 elongase Δ9 DS 18:0

Malonyl-CoA

Δ12 DS* 18:1(Δ9) Δ6 DS 18:2(Δ6,9) elongase 20:2(Δ8,11) Δ5 DS 20:3(Δ5,8,11)

Acetyl-CoA

Glucose

Acetyl-CoA Δ15 DS* 18:2(Δ9,12) (LIN)

18:3(Δ9,12,15)(ALA)

Δ6 DS 18:3(Δ6,9,12) (GLA)

Δ6 DS 18:4(Δ6,9,12,15)(STA)

elongase

elongase

20:3(Δ8,11,14)

20:4(Δ8,11,14,17)

Δ5 DS 20:4(Δ5,8,11,14) (ARA)

Δ5 DS 20:5(Δ5,8,11,14,17)(EPA) elongase 22:5(Δ7,10,13,16,19) Δ4 DS 22:6(Δ4,7,10,13,16,19) (DHA)

n-9 series

n-6 series

n-3 series

DS = desaturase; the position where the double bond is introduced is indicated by Δx, where x is the C atom numbered from the carboxylic acid group of the fatty acid. * indicates desaturases that are not present in humans and other animals.

(B) ARA (20:4 n-6)

Prostaglandins: PGD2, PGE2, PGF2, PGI2 Thromboxanes: TXA2, TXB2 Leukotrienes: LTA4, LTB4, LTC4, LTD4, LTE4 Lipoxin: LXA4

ARA (20:4 n-6)

Prostaglandins: PGD2, PGE2, PGF2, PGI2 Thromboxanes: TXA2, TXB2 Leukotrienes: LTA4, LTB4, LTC4, LTD4, LTE4 Lipoxin: LXA4

EPA (20:5 n-3)

Prostaglandins:PGD3, PGE3, PGI3, PGF3α Thromboxanes: TXA3 Leukotrienes: LTA5, LTB5, LTC5, LTD5, LTE5 Resolvin: RE1

DHA (22:6 n-3)

Resolvin: D5, Protectin: D1

FIGURE 7.5 (A) Pathways of biosynthesis of fatty acids showing formation of the principle unsaturated and polyunsaturated fatty acids of the n-3, n-6, and n-9 series. The conversions of oleic acid (18:1 n-9) into linoleic acid (18:2 n-6) and then into alpha-linolenic acid (18:3 n-3) do not take place in animals. Consequently both linoleic and linolenic acids must be provided in the diet. (B) Eicosanoid lipids derived from the principal n-3 and n-6 polyunsaturated fatty acids. The lipids produced from arachidonic acid (20:4 n-6) lead mainly to inflammatory or proinflammatory responses in animals. They are also proarrhythmic, activate platelet aggregation, and lead to vasoconstriction. The lipids from eicosapentaenoic acid (22:6 n-3) and docosahexaenoic acid (22:6 n-3) are anti-inflammatory; they are also antiarrhythmic, inhibit platelet aggregation, and lead to vasodilation. (For further details see Schmitz and Ecker, 2008, and Adkins and Kelley, 2010.) Acetyl-CoA, acetyl-coenzyme A; LIN, linoleic acid; GLA, gamma-linolenic acid; ALA, alpha-linolenic acid; EPA, eicosapentaenoic acid; DHA, docosahexaenoic acid; ARA, arachidonic acid.

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Lands, 2014; see Fig. 7.5B). Their role in preventing heart disease has also received much attention over the past two decades (Mozaffarian and Wu, 2011, 2012; Mozaffarian et al., 2013; Harris et al., 2013; Sperling and Nelson, 2016; Jump et al., 2012). The mechanisms whereby these two fatty acids exert their effects have been ably reviewed by Adkins and Kelley (2010) and Lands (2014) highlighting the key roles of the eicosanoid and proresolvin mediators in the various protective processes. Besides their protective effects, there are also beneficial effects for treating patients who have already suffered an acute myocardial infarction with high doses of n-3 fatty acids (Heydari et al., 2016). There is also some indication that intake of ALA may be beneficial for the prevention of cardiovascular disease (Fleming and Kris-Etherton, 2014) although, as the authors indicate, there is a lack of sufficient evidence from well-controlled clinical trials to make any recommendation about the amounts of ALA that would be needed. Oils containing both EPA and DHA have also been recommended for the treatment of age-related macular degeneration and other eye diseases (Sangiovanni et al., 2008, 2009). This is perhaps not surprising in view of the well-established occurrence of these fatty acids in the retinal membranes of eyes. Strong indications have also been given to suggest a positive role for them in the prevention of the onset of Alzheimer’s disease. Excellent reviews on this topic have been published by Hooijmans and Kiliaan (2008) and Huang (2010) with additional information in the papers of Ma et al. (2007) and Daiello et al. (2015). There have also been numerous indications of a preventive role of both n-3 and n-6 PUFAs in various cancers (Currie et al., 2013; Zheng et al., 2014; Xu and Qian, 2014). Strong though some of these claims might be (Zheng et al., 2014), there is no indication that PUFAs should be replacement therapies for existing chemotherapy of cancers. At best, DHA, and possibly EPA, should be regarded as possibly beneficial adjuncts to existing treatments. As the principal fatty acids involved in cardio-protective effects, EPA and DHA, occur in fish oils, which are their principal source, it is now recommended by many governmental agencies that our diet should include eating oily fish (salmon, trout, herring, mackerel, etc.) to ensure a sufficient supply. Some cautionary note must be added here as fish oils may contain undesirable amounts of environmental pollutants ingested by the fish. Contents of heavy metals, including mercury, and dioxins have been reported in fish oils. As a result, the American Heart Association had recommended that children and pregnant women consume the most popular oily fish no more than twice per week (Lichtenstein et al., 2006) but with the caveat that for middle-aged or older men and postmenopausal women the benefits of eating Food and Drug Administration (FDA)suggested amounts of oily fish outweigh the risks of associated contaminants (Krauss and Pruitt, 2014). A daily intake of DHA 1 EPA between 250 and 500 mg/day has been recommended (Kris-Etherton and Hill, 2008; Harris et al., 2009; Holub, 2009), but

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whether these PUFAs are derived from fish or from microorganisms is immaterial though as clearly microbial oils will not contain any of the deleterious materials found in some fish oils, particularly poorly refined ones. There is also very strong evidence for the role of DHA in the development of memory and eyesight especially in newly born children (Birch et al., 1992; Saldanha et al., 2009; O’Connor et al., 2001; Vanderhoof et al., 1999; Sinclair et al., 2005; Sinclair and Jayasooriya, 2010). DHA itself is a major component of brain lipids and is also found in the membranes of the eye. This then provides the scientific basis of including DHA in the diet of premature and newly born babies and also young children. Such are the clear benefits of including DHA in infant formulas that it is now incorporated into these preparations in over 70 countries of the world (Kyle, 2010). As EPA is not involved in these processes and may even be counterindicated (Sinclair and Jayasooriya, 2010), it is not desirable to use fish oils for infant formulas. It is also not possible to exclusively produce DHA cost-effectively from fish oils as this requires the use of very large-scale high-performance liquid chromatography (HPLC) or molecular distillation, both of which are prohibitively expensive for this purpose. Thus, alternative sources of DHA need to be identified and, as indicated below (Section 5.4), several microorganisms have been identified that are able to produce it in sufficient yields to be used on a commercial scale. It was discovered that when added to infant formulas, some of the DHA can be retro-converted into EPA using essentially the reverse of the reactions described in Fig. 7.5A. As indicated above, EPA is an undesirable PUFA to give to infants; thus, the retro-conversion had to be prevented. The easiest way of accomplishing this was to add arachidonic acid (ARA, 20:4 n-6) into the formula as this effectively prevents the conversion of DHA into EPA. The supply of ARA, fortunately, was not problematic as this was known to be a microbially produced PUFA coming from the fungus Mortierella alpina. The development of the fungus as a source of ARA was pioneered by Japanese scientists in the 1980s (Totani et al., 1987). Thus, simultaneously with the development of microbial processes to produce DHA-rich oils came the development of a large-scale process to produce ARA-rich oils. This process is described in Section 5.3. In addition to its ability to prevent the undesirable conversion of DHA into EPA when included in infant formula, ARA is an immediate precursor of the more biologically active series 2 prostaglandins, and series 4 leukotrines (see Fig. 7.5B) that are signaling molecules mediating the development of the immune system in infants. ARA is more abundant in human breast milk than DHA, a fact that correlates well with observations that the demand for ARA is not sufficiently met by biosynthesis alone in developing infants (reviewed by Hadley et al., 2016). Although EPA is not recommended for infants, there is some interest in it being used along with DHA for the treatment in prevention of cardiac problems. EPA is also a substrate for producing various prostaglandins, thromboxanes, leukotrienes, and lipoxins

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(see Fig. 7.5B), which are complementary to those of DHA. It is also known to have positive effects on blood pressure, platelet aggregation, and inflammation. Thus, as indicated above, fish oils, that contain both EPA and DHA in approximately equal amounts, are strongly recommended as nutritional supplements for adults to diminish the incidence of heart problems. Where there may be some objections from vegetarians and vegans, and possibly some religious groups, to the intake of fish products into their diets, this has then opened up opportunities for algal oils to be produced as fish oil substitutes. EPA, often given as its ethyl ester and independently of its role in preventing heart disease, has been suggested for the treatment of several neurological disorders including schizophrenia, bipolar disorder, depression, and attention deficit/hyperactivity disorder in children (Puri and Richardson, 1998; Peet and Stokes, 2005; Mazza et al., 2007; Ross et al., 2007; Konigs and Kiliaan, 2016). Suggestions have also been made for it having a positive role in treating obesity, metabolic syndrome, non-alcoholic steatohepatitis, and type 2 diabetes (see Zhu et al., 2010). At the time of writing, no largescale trials of these effects of EPA appear to have been carried out at the clinical level. However, it is being used for the treatment of hypertriglyceridemia which is characterized by a high level of triacylglycerols circulating in the blood (Bays et al., 2011). Treatment with ethyl EPA is now prescribed and commercial preparations of Lovaza (produced by GlaxoSmithKline) and Vascepa—previously known as AMR101 (produced by Amarin Corp.)—are available. Both are derived from fish oils with the fatty acids being esterified and then separated by large-scale HPLC. Because this is very expensive, a number of companies are now actively exploring the possibilities of producing oils rich in EPA using microbial routes. If these ventures are successful, then microbial EPA should be cheaper than that obtained via the fish oil route.

7.5.2 Production of Gamma-Linolenic Acid (GLA 18:3 n-6) The first microbial oil, Single Cell Oil (SCO), that was produced commercially was an oil rich in GLA and was produced using the fungus, Mucor circinelloides. GLA is regarded as an essential fatty acid in that it cannot be synthesized de novo from dietary oleic acid (see Fig. 7.5A). But it can be synthesized from linoleic acid (LIN 18:2 n-6) even though LIN must be obtained in the diet as it too cannot be synthesized from oleic acid. In effect both LIN and GLA should be regarded as essential fatty acids and, indeed, these have formerly been known as vitamins F and FF, respectively (Ratledge, 2016). In addition, alpha-linolenic acid (ALA 18:3 n-3), which also must be provided in the diet, can also be regarded as another essential fatty acid. Confusingly, it too was given the name of vitamin F leading to the terms vitamin F and FF being abandoned entirely. The dietary role of LIN appears to be solely as the precursor of the n-6 series of fatty acids but GLA itself has been credited with numerous

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Fatty Acids

nutritional properties with claims for its benefits including prevention of hardening of the arteries, heart disease, cirrhosis, rheumatoid arthritis, and high blood pressure (Kapoor and Nair, 2005; Wanasundara and Wanasundara, 2006). There has also been an indication of the usefulness of GLA in helping to treat breast cancer alongside antimitotic drugs (Menendez et al., 2004). The focus of commercial microbial oil production was an alternative and cheaper oil to that from the seeds of evening primrose (Oenothera biennis) which was, at that time, the sole source of GLA. Claims for the benefits of evening primrose oil, besides those already given above, included treatment of multiple sclerosis, a claim that has since been discounted. Evening primrose oil was, and still is, sold as an over-the-counter supplement for the relief of premenstrual tension, a claim being based on its content of GLA. It is also considered of benefit for the treatment of childhood eczema. As the microbial oil contained twice the level of GLA as evening primrose oil, its entry into the marketplace seemed assured. But, as was to be realized only too late due to poor marketing, people who bought evening primrose oil did not understand the link to its content of GLA. Therefore they continued to buy the plant oil even though the SCO was cheaper and had a higher content of the active ingredient—GLA. Nevertheless, the production of the GLA-SCO continued for 6 years (198590) and was undertaken by J & E Sturge Ltd at Selby, North Yorkshire, England, using their expertise in large-scale fermentation technology developed for the production of citric acid using the filamentous fungus, Aspergillus niger. At the time, GLA was produced in 220 m3 fermenters. The process took 7296 hours with about 60 kg cell dry weight/m3 being accumulated with the cells having an oil content of 25%. Over the years of production about 50 tons of oil were generated. The oil was given the trade name “Oil of Javanicus”—taken from the original name of the mold Mucor javanicus—a name that clearly indicated its geographical origins. A detailed description of the process and of the research work that was carried out to produce the SCOGLA has been given by one of the authors of this chapter (Ratledge, 2006) and should be referred to for further details and information. As this was the very first SCO offered for sale and human consumption, extensive trials to establish its safety had to be carried out (see Section 6). These trials indicated that the oil was safe and was given tacit approval by the regulatory UK authorities for its sale to the general public. This decision was, in part, supported by M. circinelloides having been used in the traditional oriental food of Tempe for centuries, if not millennia, thereby establishing that it was a microorganism of established food use without any toxicity of it ever having been reported. The demise of the GLA-SCO process was due to its poor take up by the market and, concomitantly, its low profitability. There was also a new rival plant oil introduced into the market that had a slightly higher content of

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GLA than the microbial oil. This was the oil from the seeds of borage (Borago officinalis)—a wayside weed that was developed solely for its production of GLA. A comparison of the oils from M. circinelloides and the seeds of the two plants is given in Table 7.2. Although the process was short-lived, it did establish some very useful guidelines. It was entirely possible to grow oleaginous microorganisms on a large scale to produce an edible oil. The process of extracting the oil from the fungus was not difficult and followed the same principles and, indeed, used the same equipment that was used for the extraction of small quantities of plant materials for the production of specialty oils. Some refinement and deodorization of the oil was necessary to produce the final bright, golden oil. But again the techniques were the same as already used by the plant oil industry. Finally, the toxicity trials that the oil had to undergo to establish its safety were not excessively difficult. It has to be said, however, that with M. circinelloides having associations with traditional Asian fermented foods, the acceptance of its oil was considerably enhanced. Today, there is only a limited demand for GLA-rich oils. Evening primrose oil remains the main source with about 10% GLA content, and this is sold principally in the United Kingdom and some other European countries mainly for the relief of premenstrual tension. Some small use remains for the treatment of childhood eczema. Borage oil, with a GLA content of 22%, also finds a small niche market for the same applications. However, should GLA ever be required in large quantities then it could be produced very easily using genetically modified safflower (Carthamus tinctorius) which produces TABLE 7.2 Fatty Acid Profiles of Oils Rich in Gamma-Linolenic Acid (18:3 n-6) from Mucor circinelloides and Two Plant Oils Produced Commercially Major Fatty Acids Organism

Oil Content

16:0

18:0

(% w/w)

18:1

18:2

GLA

18:3 (n-3)

(Rel. % w/w)

Mucor circinelloides (Oil of Javanicus)

25

22

6

40

11

18

Oenothera biennis (evening primrose)

16

6

2

8

75

810

0.2

Borago officinalis (borage)

30

10

4

16

40

22

0.5

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Fatty Acids

an oil with GLA at 70% of the total fatty acids (Nykiforuk et al., 2012; Knauf et al., 2011). Thus, the prospects of reviving the Mucor-derived oil would seem to be extremely remote. Nevertheless, the experience gained in developing this SCO proved to be invaluable for the next generation of microbial oils.

7.5.3 Production of Arachidonic Acid (ARA 20:4 n-6) ARA is a polyunsaturated fat containing 20 carbons and four methyleneinterrupted cis unsaturations. It is arguably the most nutritionally important omega-6 fatty acid, being the primary precursor of the proinflammatory prostaglandins and leukotrienes (see Fig. 7.5B). Although not generally recommended for supplementation in adults (who ingest omega-6 fats in excess), ARA is considered essential for the development of immune functions in newborn babies. This is the primary reason for its inclusion along with DHA in most infant formulas (mentioned in Section 5.1). As previously discussed, inclusion of ARA in infant formulas prevents retro-conversion of DHA to EPA. Another reason for the pairing of ARA and DHA in formula is rooted in the observation that any individual 20- or 22-carbon PUFA (ARA or DHA) in the diet of model animals causes repression of a key desaturase involved in the conversion of 18-carbon PUFAs to their 20- or 22-carbon products (reviewed by Brenna, 2016, and Hadley et al., 2016). Thus, the inclusion of DHA or ARA alone may inhibit synthesis of the other from its precursor 18-carbon omega-6 or omega-3 fatty acid. Since both fatty acids have been known for their benefits for infant development and both are naturally found in mothers’ breast milk (Jensen and Lammi-Keefe, 1998), it follows that these fatty acids should similarly be included as a pair in formulas. Industrially, ARA is produced by submerged fermentation of the filamentous, zygomycete fungus Mortierella alpina. DSM and Nissui (in partnership with Suntory) are the largest producers of an ARA-rich oil using M. alpina. Other suppliers of this oil include Cargill Alking Bioengineering, producing an oil known as CABIO, and Hubei Fuxing Biotechnology. The basic fermentation process is similar for all manufacturers. An agitated, fed-batch fermentation can produce more than 50 g dry cell mass per liter in 57 days. Some manufacturers have claimed oil levels in the cell mass of higher than 50%, and of this, as much as 65% may be ARA (Singh and Ward, 1997). As with all fermentation processes for oleaginous organisms, that with M. alpina occurs in two stages. The initial phase is targeted towards accumulating cell mass by constant feeding of nitrogen, air, and a carbon source (usually glucose). In the second stage, the nitrogen feed (yeast extract or an ammonium salt) is halted and the cells are induced to convert the supplied glucose to fat. Among SCO processes, one distinguishing feature of M. alpina fermentation is the need to control fungal morphology. Because Mortierella is filamentous, submerged fermentation causes it to grow in “fuzzy” looking

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TABLE 7.3 Fatty Acid Profiles of Commercial Oils Rich in Arachidonic Acid Using Mortierella alpina Fermentations Major Fatty Acids 16:0

18:0

18:1

18:2

18:3 (n-6)

20:3 (n-6)

20:4 (n-6)

22:0

24:0

(Relative % w/w) ARASCOa

8

11

14

7

4

4

49

CABIO oilb

7.5

6

9

6

25

4

43

1 3

9.5

a

Oil produced by DSM (the Netherlands). Oil produced by Cargill Alking Co. Ltd (from Casterton et al., 2009) and Kusumoto et al. (2007).

b

pellets, or as a pulp, in which nutrient uptake may be impeded. Generally, smaller, looser pellets are more desirable and enable faster cell division and mass transfer of nutrients (Totani et al., 2002). When the fermentation is complete, cells are harvested, dried, and the oil is extracted from the biomass and refined, again using techniques similar to those used for extraction of oils from oilseed crops (Bresson et al., 2008). Profiles of the fatty acids from the various strains of M. alpina that are in production are given in Table 7.3.

7.5.4 Production of Docosahexaenoic Acid (DHA 22:6 n-3) DHA is the largest and most complex of the nutritionally important PUFAs with 22 carbons and 6 methylene-interrupted cis-double bonds. The first-tomarket fermentable source of DHA was the heterotrophic microalgae Crypthecodinium cohnii. The C. cohnii process was developed by Martek Biosciences in the 1990s to furnish DHA for inclusion in infant formula. The organism was chosen primarily because of its very simple fatty acid profile, in which DHA is often the sole PUFA comprising 40% or more of the total fat (Behrens and Kyle, 1996). C. cohnii is a dinoflagellate and, as the name of its phylum implies, vegetative cells swim by the action of two flagella, one radial flagella for steering and another posterior flagella for propulsion (Bean and Himes, 1982). Unfortunately, this means that C. cohnii has to consume more glucose to produce the extra energy needed for swimming and thus fermentations evolve far more CO2 and yields of fat from glucose are much lower than comparable processes based on other species that do not swim during vegetative growth. C. cohnii is still used for production of DHA for infant formula. However, a more efficient DHA-producing organism has been

260

Fatty Acids

identified and adapted for industrial fermentation to produce a DHA-rich oil that may be suitable for inclusion in infant formulas (Mehta et al., 2016). Several members of the class Labyrinthulomycetes and in particular the families Thraustochytriaceae and Labyrinthulaceae, colloquially known as the “Thraustochytrids” and “Labyrinthulids” or simply algal “chytrids,” have been used for industrial production of DHA and other PUFA by fermentation. Chytrids are perhaps most interesting for their ability to make PUFA using not only the classic standard pathway of desaturases and elongases, but also a heterotrimeric synthase related to polyketide and fatty acid synthases (Matsuda et al., 2012; Metz et al., 2001). In at least one species, this PUFA synthase is the exclusive means of de novo DHA production (Lippmeier et al., 2009). Because of this, chytrids may be grown with low levels of dissolved oxygen during the lipid accumulation phase of fermentation, as the oxygen requirements of the desaturases are greatly diminished and thereby result in a saving on fermentation energy costs (Qu et al., 2011). Because chytrids only accumulate fat in vegetative cells which do not swim, they are typically capable of achieving much higher yields of fat from glucose than C. cohnii. Lastly, chytrids have cell walls that are more amenable to rupture via hydrolysis, enabling manufacturers to avoid the use of more expensive organic solvents for oil extraction (Ruecker et al., 2004). Today, the largest manufacturer of DHA oils from both C. cohnii and certain species of chytrids is DSM Nutritional Products. Other companies producing DHA from chytrids and other microalgae include Runke Bioengineering, Daesang, Cargill, Cellana, Synthetic Genomics, and TerraVia (formerly known as Solazyme). The latter company, in a joint venture with Bunge Ltd, have now launched their DHA product derived from a Schizochytrium sp. under the name of AlgaPrime—see Table 7.4. This Schizochytrium sp. is grown in Brazil mainly on sucrose together with sugarcane wastes (see http://algaprime.com/Algaprime_Product_Sheet.pdf). The product is being aimed at the fish feed market. The fatty acid profiles of the DHA-rich oils that are in commercial production are given in Table 7.4.

7.5.5 Production of Eicosapentaenoic Acid (EPA 20:5 n-3) EPA, like DHA, is a highly active PUFA. It gives rise to a number of metabolic derivatives that have key physiological roles in human and animal metabolism. Claims for its benefits have included treatment of various neuropsychiatric disorders including bipolar disorder, depression, and schizophrenia (Peet and Stokes, 2005; Riediger et al., 2009; Lin et al., 2010; Sublette et al., 2011). Clinical trials of EPA have usually involved oral administration of its ethyl ester. EPA ethyl esters are prepared from fish oils as no commercially viable source of it, as the sole PUFA in the profile, is known to occur naturally. Of the three currently produced EPA-containing treatments for hypertriglyceridemia, Lovaza (from GSK), Vascepa (from Amarin Corp. plc),

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TABLE 7.4 Profiles of the Principal Fatty Acids in Commercially Produced Microbial Oils Rich in Docosahexaenoic Acid (DHA)

DHA-SCOa b

Schizo-SCO

c

14:0

16:0

16:1

18:0

18:1

22:5 (n-6)

22:6 (n-3)

26

18

2

,0.5

15



40

7

16

,0.5

1

16

16

39

2

13

27

Schizo-TKd

6

18

AlgaPrimee

?

34

Schizo-ONC

?

1

,1

8

40

0.5

0.5

19

49

1.5

?

12.6

48

a

From Crypthecodinium cohnii produced by DSM and sold as life’s DHA. From Schizochytrium sp. (produced by DSM) with trade names of DHASCO-S and DHA-Gold. From Schizochytrium sp. as produced by Ocean Nutrition Canada Ltd (see Burja et al., 2006) and now owned by DSM. d From Schizochytrium-TK as produced by Jiangsu TianKai Biotechnology Co. Ltd (Nanjing, China)—see Ren et al. (2010). e From Schizochytrium sp. Produced by TerraVia Inc.—Bunge Ltd; incomplete fatty acid analysis available. Source: Ratledge, C., 2016. Microbial production of vitamin F and other polyunsaturated fatty acids. In: Vandamme, E.J., Revuelta, J.L. (Eds.), Industrial Biotechnology of Vitamins, Biopigments and Antioxidants. Wiley-VCH. pp. 287320. b c

and Epanova (from AstraZeneca), only Vascepa appears to be single EPA, the other two preparations also contain DHA along with the EPA. EPA-rich oils are not found in microorganisms in the same way as ARA and DHA have been found as the sole PUFAs in several microbial species. EPA invariably occurs along with either ARA or DHA or sometimes both. Where it does occur in some relative abundance is in microalgae. Table 7.5 gives the EPA contents of a number of algae that have been grown photosynthetically (see Bellou et al., 2014). Of these Nannochloropsis oculata and N. salina appear to offer the best possibilities for commercialization. Only a few companies are currently involved in developing algal cultivation systems for EPA production, including Qponics, Qualitas Health, and DSM. Of these, only the latter two companies appear to have attained any commercial product (detailed further in Section 5.6). Several other companies have tried to develop EPA products using photosynthetically grown microalgae but have withdrawn from this field due to commercial difficulties. Heterotrophic cultivation of some of the microalgae that produce EPA may be an alternative to phototrophic cultivation as, when all factors are taken into account, this may be the cheapest method of production. While there is no plant or microorganism that produces EPA as a single, dominant PUFA, DuPont at Wilmington, DE, decided that the market prospects for a SCO-EPA were promising enough for the company to embark on an ambitious program to transform an oleaginous microorganism to

262

Fatty Acids

TABLE 7.5 Contents of EPA in the Fatty Acids of Various Microalgae Grown Phototrophically Organism

% EPA in Total Fatty Acids

Amphidinium sp.

17a

Chlamydomonas sp.

19

Chroomonas salina

13

Pavlova lutheri

18

Pavlova salina

19

Pavlova sp. H

29

Pavlova sp. L

23

Asterionella sp.

26

Chaetoceros constrictus

19

Nannochlopsis oceanica

23

Nannochlopsis oculata

3036

Nannochlopsis salina

26

Nannochlopsis spp.

3033

Phaeodactylum tricornutum

1430

Porphyridium cruentum

2037.5

a Also contains 26% DHA. Source: From Bellou, S., Baeshen, M.N., Elazzazy, A.M., Aggeli, D., Sayegh, F., Aggelis, G., 2014. Microalgal lipids, biochemistry and biotechnological perspectives. Biotechnol. Adv. 32, 14761493.

produce EPA via genetic engineering. The choice of organism was Yarrowia lipolytica which, at the time of commencement of the project (B2000), was the only oleaginous organism whose genome had been sequenced. The yeast in its natural form only produces LIN (18:2) as the longest chain length and most unsaturated fatty acid (see Table 7.6). A number of genes therefore had to be added to achieve conversion of 18:2 into 20:5. In addition, other genes were found to be necessary to control the expression of those genes actively coding for the enzymes involved in fatty acid desaturation and elongation (see Fig. 7.5A). In all, some 30 copies of nine different genes had to be incorporated into the genome of the yeast (Xue et al., 2013; Xie et al., 2015; Zhu and Jackson, 2015). The final recombinant strain (known as Y4305) was able to produce a lipid content in the cells of about 30% (w/w) with EPA accounting for 56% of the total fatty acids (see Table 7.6). Some 90 patents (see, for example, Hong et al., 2014) were filed on the process and the yeast went into commercial production in 2013 with the initial product being sold as an over-the-counter nutraceutical through a wholly

TABLE 7.6 Profiles of Major Fatty Acids in the Various Microbial Oils Being Produced with a High Content of Eicosapentaenoic Acid (20:5 n-3) and Compared to Krill Oil Major Fatty Acids 16:0

16:1

18:0

18:1

18:2

18

16

6

45

15

Y. lipolytica y4305

3

0.7

1

4

17

Omega-3c

21.5

1

2.5

Y. lipolytica WTa b

d

Almega PL e

Krill oil a

20:2

3.5

20:4 (n-6)

20:4 (n-3)

0.6

2

EPA

22:5 (n-6)

22:5 (n-3)

22:6 (n-3)

1.6

3.5

40

0.3

7

56.6

1.6

21.7

9.6

12

0.1

1

1

3

25

12

3

0.6

6

1

0.4

13.6

Yarrowia lipolytica wild type (from Xue et al., 2013). Genetically engineered stain derived from wild type (from Xue et al., 2013). c Produced by Amerifit Inc., Cromwell, CT (a wholly owned subsidiary company of DSM) using a Schizochytrium sp. (see Gilles et al., 2011). d Polar lipids from Nannochloropsis oculata fatty acids given as percentage of oil (from Kagan et al., 2013). e Given as percentage of oil (from Kagan et al., 2015). b

264

Fatty Acids

owned subsidiary company, New Harvest. It was sold simply as Omega-3 vegetarian EPA in 600 mg capsules intended for one-a-day consumption. However, sales were disappointing and the product has now been discontinued. The subsequent application has been to use the entire yeast biomass, without extraction of the oil, as a fish feed material. This clearly decreased the cost of downstream processing by eliminating most of the downstream steps. All that was now needed was to harvest the yeast from the fermenters and then dry it into a stable powder. The fish chosen for this application was salmon. In 2013, DuPont established a joint venture company, Verlasso, in cooperation with the Chilean salmon producer, AquaChile, who agreed to use the yeast in their farmed salmon process (http://www.verlasso.com/farming/ fish-in-fish-out/about-our-yeast/). The addition of the EPA-rich yeast to the salmon feed meant that it could significantly decrease the amount of fish oil and fish meal that needed to be fed to the salmon, reducing the amount of wild fish needed for feedstock by 75%, from 4 kg per kg salmon to just 1 kg. At the same time, the added yeast led to a substantial improvement in the flavor of the salmon so much so that the ocean-farmed salmon could command a premium price of about $4$10 per kg when sold in the United States. It is not known, however, for how much longer the yeast will be produced as, at the beginning of 2016, DuPont merged with Corning-Dow, and this has meant considerable rethinking of the new company’s products as well as their research and development plans.

7.5.6 Production of EPA/DHA Mixtures as Alternatives to Fish Oils As previously discussed in Section 5.4, strains of Schizochytrium have been used for commercial production of DHA since the 1990s. Certain species of Schizochytrium and related Thraustochytrids, Aurantiochytrids, Oblongiochytrids, and Aplanochytrids may produce DHA alone but most make at least two or more PUFAs besides DHA including docosapentaenoic acid (usually n-6 but also n-3), EPA, and ARA (Yokoyama et al., 2007). Of interest in the development of EPA-containing oils has been the recent introduction of a new oil being produced with another Schizochytrium strain. This oil was launched by DSM and has been given the trade name of Ovega-3. The fatty acid profile of the oil from this microorganism is given in Table 7.6. Ovega-3 contains both DHA and EPA in a triacylglycerol form (Gillies et al., 2011). The oil of these capsules is extracted from the alga grown under heterotrophic conditions in South Carolina (Fig. 7.6). The extraction process is based on cellular hydrolysis and does not use organic solvents (Ruecker et al., 2004). Recently, Skretting, which is one of the three leading companies producing aquaculture feeds, announced an interest in using algal oil containing EPA and DHA supplied by DSM and the German company, Evonik, for

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265

FIGURE 7.6 Photograph of DSM’s fermentation facility at Kingstree, SC. This is used for the cultivation of oleaginous algae and fungi. The microbial oils containing DHA, EPA and ARA are extracted and refined on site. The largest vessels may hold more than 250,000 liters of broth.

inclusion in their products. This in anticipation of a near-future gap rising between the demand for fish and the current flat supply of fish oils and fish meal for feed (http://www.feednavigator.com/R-D/Skretting-gets-behindalgal-oil-breakthrough-from-Evonik-and-DSM). Other partnerships entering this arena are TerraVia (Solazyme) along with Bunge joining with BioMar, and EWOS now becoming part of the Cargill group. Thus, we are now seeing massive interest in developing either microbial biomass or the extracted PUFA-rich oil being used for fish feeding. The market is likely to be an expanding one and also potentially very lucrative. Another commercial oil containing EPA as the principal PUFA is sold by Qualitas Health under the trade name of Almega PL, where PL stands for polar lipids. The production organism, N. oculata, is grown in large open raceways in West Texas. The lipid is extracted and then fractionated to give a mixture of polar lipids with EPA at about 25% of the total fatty acids (Table 7.6). The product has been extensively examined from a safety viewpoint (Kagan et al., 2014) and has received FDA approval for its sale to the public. These algal lipids have been compared favorably with krill oil with respect to their fatty acid profiles and, especially, to the relative contents of

266

Fatty Acids

EPA (Kagan et al., 2013). Somewhat controversially, Qualitas also claims that the digestibility of its product and thus the uptake of the PUFAs, in particular that of EPA, is superior to that found for the uptake of EPA from fish oils that are often in the form of fatty acid ethyl esters (FAEE). Studies have shown equivalent bioavailabity of PUFAs from PL, FAEE, and TAGs (Kagan et al., 2015).

7.6 SAFETY ASPECTS With the launch of the first SCO in 1985—see Section 5.2—came the appreciation that, with this being a novel product, it would have to undergo stringent trials before it could be sold to the general public. The first batch of oil that was produced using M. circinelloides proved to be toxic when fed to brine shrimps. This did not augur well but it was quickly appreciated that the toxicity of the Oil of Javanicus product was due to the presence of free fatty acids in the oil. It is well known that these entities are cytotoxic to living cells and, once they were removed from the oil, the next preparation gave no signs of toxicity to the shrimps. The fatty acids had arisen in the oil during the downstream processes: harvesting the cells from the fermenter, their drying, and subsequent solvent extraction. It was found that the lipases were activated as soon as cells were separated from their culture medium and were thereby deprived of an extracellular carbon source (glucose). The cells, then sensing carbon deprivation, began to consume their stored intracellular lipids, hydrolyzing the triacylglycerols into the component fatty acids by the action of these activated lipases. The solution to the problem was to prevent the activation of the lipases. This was done by heating the cells to about 60 C65 C before the harvesting process took place. This then led to an oil being produced in which the free fatty acids could scarcely be detected. Further feeding trials of the GLA-SCO for 90 days to experimental animals (mice, rats, and rabbits) indicated that the microbial oil was of no toxicological concern and showed no adverse effects when given at many times the recommended nutritional dosage to the animals. Indeed, it could be claimed that the oil was safer than corresponding commercial plant oils in that it had extremely low levels of herbicide and pesticide residues, much lower than are found in the plant oils where these agents are routinely sprayed on the crops. The minute traces of these residues that were found in the microbial oil could be traced back to their occurrence in the glucose syrups being used as feedstock. The glucose was being produced by the hydrolysis of the starch derived from corn, which is clearly sprayed with several agricultural agents during its growth. It should though be stated that the amounts of spray residues found in all commercial plant oils are well below the regulatory limits. (Further information is given in the specific review related to GLA production by Ratledge, 2006.) The studies proved valuable to support the approval of the first wave of SCOs that became commercially available.

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The safety of the oil from C. cohnii, used to fortify formula for term and preterm infant formulas, clearly required considerable assurances that it was suitable for long-term consumption in sensitive populations. Initial trials (see Kyle and Arterburn, 1998) indicated that the oil was well tolerated and was not harmful. The safety of this oil was considerably helped by the organism having no known toxicity to humans nor did it produce any toxin or reveal a taxonomical identity with any toxin-producing microalgae. Further stringent testing (see Zeller, 2005; Ryan et al., 2010; Sinclair and Jayasooriya, 2010) has confirmed the safety of the oil. Similar conclusions have been given for the ARA-rich oil that is included in infant formulas along with the DHA oil from C. cohnii. The specific tests have included in vitro mutagenicity and genotoxicological trials plus many studies with rodents and other animals. In the United States, two paths currently exist for regulating new substances for use in foods. The first path established in 1958 is the food additive petition (FAP) process in which the US FDA evaluates nonpublic data submitted by the applicant. No oils derived from any microalgae have been submitted for approval through the FAP process. Instead, all new edible algal oils in the market have thus far been regulated by the second path, the “Generally Recognized as Safe” (GRAS) self-affirmation process in which a company or individual assesses the safety of the food substance. The assessment is subsequently evaluated by a panel of qualified experts who review the publicly available literature and determine whether the food substance is safe for the intended use based on scientific procedures. The company that concluded the self-affirmation has the option to notify the FDA. The Agency will then evaluate the notice and determine if it provides sufficient basis for a GRAS determination. The agency may ask further questions or issue a letter of no objection. If safety was not established, the agency will ask the notifying organization to withdraw or “cease to evaluate further.” This GRAS self-affirmation process was established in 1997 for human foods, with final rules formalized in August and effective in October 2016. (https://www.federalregister.gov/documents/2016/08/17/2016-19164/substances-generally-recognized-as-safe). The first algal oil for which a basis for GRAS status was determined was a blended oil containing both DHA from C. cohnii and ARA from M. alpina (http:// www.fda.gov/Food/IngredientsPackagingLabeling/GRAS/NoticeInventory/ ucm154126.htm). This determination was issued in May 2001, paving the way for the inclusion of oils in infant formulas globally. Other oils from the heterotrophic alga Schizochytrium sp., were established as GRAS in February 2004 (http://www.fda.gov/Food/IngredientsPackagingLabeling/GRAS/ NoticeInventory/ucm153961.htm) and June 2015 (http://http://www.fda.gov/ Food/IngredientsPackagingLabeling/GRAS/NoticeInventory/ucm462744.htm). The former of these two oils was commercialized primarily to supply DHA alone for human dietary supplements and the latter to offer DHA and EPA in a ratio analogous to a “vegetarian fish oil”. It can therefore be concluded that the safety of the microbial oils is exactly the same, or even better, than any other plant or animal oil that is

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routinely consumed by humans, including newly born babies. Even at very high doses, where up to 7 g DHASCO oil (about 25 times the recommended daily dose) has been given to human volunteers, no adverse effects were observed except for slight “fishy burps” (Wynn and Ratledge, 2006). Lewis et al. (2016). After a detailed toxicological examination of the ARA-rich oil from M. alpina and the DHA-rich oil from Schizochytrium sp. in rats for up to 90 days, Lewis et al. (2016) concluded that neither oil produced any significant changes in physical, physiological, biochemical, hematological, or histopathological parameters. The safety of microbial oils is now accepted by all regulatory authorities around the world. Each new oil that comes on to the market and requires evaluation can therefore use the existing claims of safety to their advantage. Microbial oils have been one of the most tested of all oils. They are as safe as any other commodity oil or fat.

7.7 FUTURE PROSPECTS With the arrival of microbial oils into the marketplace in 1985, they have gradually become of increasing importance and value in the niche market of high-value nutraceuticals. The market for PUFAs continues to grow. The demand is principally for DHA and ARA to be incorporated into infant formulas and for DHA 1 EPA oils as over-the-counter nutraceuticals. The latter combination is usually derived from fish oils with rigorous quality control precautions now being taken to monitor the oils very carefully for the possible presence of undesirable dioxins and heavy metals. There are, however, a small minority of people who dislike taking fish oil capsules principally because of the “fishy burps” that arise. Other groups who do not want to take these oils include vegetarians, vegans, and some religious groups. Together, these represent a small but significant number of people who are looking for non-fish-derived sources of DHA/EPA. Only microbial oils can therefore fulfill these requirements. The current principal sources are then based on using C. cohnii and various species of Schizochytrium or Thraustochytrium. The recent arrival into the marketplace of Ovega oil with a content of 22% EPA and 40% DHA (see Table 7.5) is the first of what will doubtless prove to be several such oils. Other sources that are likely to appear within the next few years will be oils derived from photosynthetically grown algae. Algal growers have, at last, begun to appreciate that there is very little prospect of being able to produce lipids as biofuels in an economic manner. The current and likely future prices of crude petroleum oil will continue to depress the market demand for alternative sources of liquid biofuels. Many global markets for biodiesel exist, but these are supplied by plant oils as they are currently cheaper to produce than any microbial oil no matter how the microorganism is grown. Algal oils simply cost too much to produce for this purpose, but the tremendous interest in using these microorganisms over the past 10 years or more has been so intense that the technologies developed for them cannot be abandoned without incurring huge financial losses. Some

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rescue products are clearly needed. These can only be very high-value products and include carotenoids and xanthophylls such as astaxanthin, as an antioxidant and as a colorant, as well as the PUFAs. Thus, we can expect to see several oils derived from photosynthetically grown algae coming on to the market that will provide mixtures of EPA and DHA. The test for these oils will be to see how their prices compare with similar oils coming from heterotrophically grown Schizochytrium spp. and related organisms. This will then reveal once and for all whether phototrophic cultivation is substantially cheaper than heterotrophic cultivation. But the future may not belong to microbial oils indefinitely. Plant geneticists have, for the past two decades, reported on the development of genetically modified plants that will be able to produce PUFAs longer than C18. Apart from mosses (Kaewsuwan et al., 2006), there are no plants that are able to produce PUFAs longer than stearidonic acid (18:4 n-3). The possibilities of using oils containing relatively high levels of this acid, that can be obtained from Echium plantagineum, have been explored as a means of indirectly boosting the levels of EPA and DHA in humans (see Fig. 7.5A). However, the nutritional benefits derived from an intake of Echium oil appear to be marginal. The use of Echium oil for fish feeding has also been considered and would, if successful, be a possible cheaper alternative to the microbial oils that are currently being used or considered for such use. For the production of plant oils with useful amounts of ARA, EPA, or DHA, or combinations of these fatty acids, it is therefore necessary to use genetic engineering techniques. Two principal groups are involved: CSIRO in Australia and Rothamsted Research Laboratory in the United Kingdom. Both laboratories are government funded and both groups have achieved limited success. Each is using the established oilseed crop plant of Camelina sativa; the former group has achieved an oil with a content of DHA at 10% of the total fatty acids (Petri et al., 2012, 2014), whereas the latter group has achieved a content of EPA/DHA at 14.5% in a field-grown trial of the genetically modified (GM) plant (Ruiz-Lopez et al., 2014, 2015; Usher et al., 2015). While the oil content of the plant seeds is about 30% (w/w), no indication has been given by either group of the amount of oil produced per unit area of land which is clearly a key economic factor. Although there was considerable interest in the possible developments of the GM plant in 2014 and 2015 (Napier et al., 2015), very little new information has been provided over the past 2 years about larger scale cultivations of the GM crop. It is also possibly significant that neither research group is looking toward providing oils for the nutraceutical market. This, perhaps, is not surprising in view of the relatively low levels of the key PUFAs in the oils. The stated aim of the work is now to provide a plant oil that can be used for fish feeding (Napier et al., 2015; Betancor et al., 2015a, 2015b, 2016a, 2016b), which is therefore a low value application. Initial trials of the GM oils with feeding to sea bream (Betancor et al., 2016b), have shown that the GM oil with DHA (6.9% of the total fatty acids) was as good as fish oil (with 9.7% DHA) as a feed

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supplement but the GM oil with EPA (13.5%) was slightly less effective. Both oils were well tolerated by the fish. Similar results were obtained when the GM oils were fed to Atlantic salmon (Betancor et al., 2016b; and reviewed by Sprague et al., 2016) thereby leading Betancor et al. (2016a) to conclude “. . . that genetically modified oilseed crops are a potential solution to fill the gap between demand and supply of EPA and DHA and, specially, are a viable alternative to fish oil for the supply of n-3 LC-PUFA in aquaculture.” Similar work to produce transgenic Canola plants that contain DHA in the oil has been performed in a collaboration between DSM and Dow (Walsh et al., 2016). Although Canola is a commodity oilseed crop and its oil would be well accepted and tolerated, the amount of DHA was relatively small at only 3.7% of the total fatty acids. EPA was also present at 0.7%. The authors indicated that some 14 g of oil would have to be ingested to achieve an intake of 600 mg DHA, which would be slightly above the daily dietary recommendation. A possible (but unexpressed) application of the oil or the entire seed meal would be, as with the work with C. sativa, into fish feeding. Also declaring an interest in this expanding market has been the announcement in 2011 by Cargill that they have formed a partnership with BASF to also use Canola to develop a GM crop containing sufficient amounts of DHA to enter the fish feed market. The economics of producing these GM plants for fish feeding always has to be compared with the prevailing price of fish meal, which is the standard feed material and currently is about $1600/ton. As non-GM Canola oil sells for about $800$900/ton, this would indicate that there is probably sufficient profit margin in the GM oil for it to outcompete fish meal. The key, as always, will be whether the rather low content of DHA/EPA in these GM oils will be sufficient to satisfy the fish farmers when they could be purchasing oil from Schizochyrium with over 40% DHA. The future for fish feeding is clearly interesting and will be watched with interest as to how an economic breakthrough might be made. And, of course, the yields today may be low, but tomorrow may be a different story. Serious attempts to produce plant oils that could rival the current microbial oils for their high contents of the very long chain PUFAs, however, seem as distant today as they did 25 years ago when the first thoughts of being able to genetically manipulate plants into producing DHA and EPA were proposed. Microbial oils seem likely to dominate the marketplace for the supply of high-value single PUFAs, whether this is ARA, EPA, or DHA, into the neutraceutical and infant formula markets for many decades to come.

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Chapter 8

Chemical Derivatization of Castor Oil and Their Industrial Utilization Rachapudi B.N. Prasad and Bhamidipati V.S.K. Rao CSIR-Indian Institute of Chemical Technology, Hyderabad, Telangana, India

Chapter Outline 8.1 Introduction 280 8.2 Derivatives of Castor Oil Based on Unsaturation of Ricinoleic Acid 282 8.2.1 Hydrogenated Castor Oil 282 8.2.2 Epoxy Castor Oil 282 8.2.3 Ozonolysis of Castor Oil 284 8.2.4 Preparation of 9,10,12Trihydroxy Octadecanoic Acid 285 8.2.5 Halogenated Derivatives of Castor Oil 285 8.2.6 Novel Derivatives of Ricinoleic Acid Employing Metathesis Reaction 285 8.3 Derivatives of Castor Oil Based on Hydroxy Functionality of Ricinoleic Acid 286 8.3.1 Dehydrated Castor Oil and Dehydrated Castor Oil Fatty Acids 286 8.3.2 Sulfated Castor Oil (Turkey Red Oil) 288 8.3.3 Acetylated Castor Oil 288 8.3.4 Castor OilBased Estolides 289

8.3.5 Castor OilBased Polymer Products 8.3.6 Potent Hydroxy Derivatives of Ricinoleic Acid 8.4 Derivatives Based on Ester Functionality of Castor Oil 8.4.1 Hydroxy Fatty Acid Esters 8.4.2 Castor OilBased Biodiesel 8.4.3 Preparation of Ricinoleyl Alcohol 8.4.4 Ricinoleic AcidBased Amides 8.4.5 Ethanolamides of Castor Oil Fatty Acids 8.5 Unique Derivatives of Castor Oil 8.5.1 Castor OilBased Dimer Acids 8.5.2 10-Undecenoic Acid and Heptaldehyde 8.5.3 Sebacic Acid and 2Octanol References

Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00008-8 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

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8.1 INTRODUCTION Castor plant (Ricinus communis) belongs to the family Euphorbiaceae and grows wild in varied climatic conditions. This plant was originated in India as well as in Africa. The size, appearance, and its parts vary depending on the variety, environment, and agronomical practices of the plant. Castor oil is initially domesticated in Eastern Africa and later introduced to China from India about 1400 years ago (Patel et al., 2016). China and Brazil are the major growing countries of castor cultivation up to 90% of the global production, even though it is being grown in about 30 countries. However, India produces 85% of global production of castor oil and dominates in international trade (Ogunniyi, 2006). India is a leading exporter of castor oil over 90% valuing up to US$ 1 billion per annum and the United States, European Union, Japan, Brazil, and China are the major importers, accounting up to 84% of imported castor oil (Patel et al., 2016). Castor crop cultivation involves various challenges and climate adaptability restricts the castor plantation in the United States in addition to the presence of toxic protein, namely ricin in the plant. The crop also involves labor-intensive harvesting process, which warrants the United States and other developed countries to pursue castor plantation (Patel et al., 2016). Castor leaves provide the necessary nutrients required for the growth of silkworm as a host plant. The silk produced from the castor plantbased silkworm is known as eri silk. The by-product of this industry is eri pupae, which is a good source for protein and nutrient oil. The eri silkworm pupae contain about 18%20% (dry basis) oil and found to contain alpha linolenic acid (ALA) up to 43%. The regiospecific analysis of the oil showed a higher level of ALA (47.3%) at the sn-2 position (Shiv Shankar et al., 2006). The oil found to contain about 2.5% of phospholipids and phosphatidylethanolamine is the major phospholipid (64%) followed by phosphatidylcholine (19.2%). Cardiolipin and phosphatidylinositol also contain in minor quantities (Ravinder et al., 2016). The same group has reported the refining process for eri pupal oil (Ravinder et al., 2015). Oil extraction is usually carried out by mechanical expression or solvent extraction, or both and average oil content is about 45%55% by weight depending on the castor varieties and geographical location (Ogunniyi, 2006). Castor seeds report to contain three toxic constituents, namely ricin (glycoprotein), ricinine (alkaloid), and allergen (proteincarbohydrate complex), and these three components retain in the deoiled cake during extraction and oil is free of these components. Due to this reason, the castor deoiled cake cannot be utilized for edible applications, even though it contains significant amounts of protein and hence restricted to low-value applications like biofertilizer. However, the protein isolate was extracted from the castor deoiled cake and two different products, namely N-acyl amino acids (Prasad et al., 1988) and diethanolamides (Lakshminarayana et al., 1992),

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were reported with good surfactant properties for possible use in industrial applications. Since ages, castor oil has been used in variety of medicinal applications including as a purgative laxative stimulant and it is classified by the U.S. Food and Drug Administration (FDA) as generally recognized as safe and effective (GRASE). Ricinoleic acid (RA) has been shown to be effective in preventing the growth of numerous species of viruses, bacteria, yeasts, and molds. Castor oil is an ancient and popular nonedible oil with significant industrial and medicinal value (Anjani, 2012). The oil possesses most unusual physical and chemical properties compared to other traditional vegetable oils, due to the presence of hydroxy unsaturated fatty acid called RA [(12R,9Z)-hydroxyoctadecenoic acid] ranges from 87% to 92% (BorchJensen et al., 1997; Binder et al., 1962). The other fatty acids, namely palmitic (0.81.1), stearic (0.71.0), oleic (2.23.3), linoleic (4.14.7), and linolenic (0.50.7), are present in minor quantities in the oil. RA is an 18-carbon straight chain acid with a cis-double bond between 9th and 10th carbon and a hydroxy group at 12th carbon. Due to the presence of hydroxy functionality, castor oil exhibits unique combination of physical properties such as high viscosity [889.3 centistokes (cSt)], density (0.959g/ml at 25 C), thermal conductivity (4.727 W m C21), pour point (2.7 C), melting point (22 to 25 C), boiling point (313 C), excellent solubility in alcohols, and ability to plasticize a wide variety of natural and synthetic resins, waxes, polymers, and elastomers (Kazeem et al., 2014). Castor oil maintains its fluidity at both extremely high and low temperatures and due to this nature, it is considered as an attractive lubricant and in addition it is also an excellent as feedstock for the preparation of variety of biolubricant base stocks. Due to the presence of hydroxy fatty acid (HFA), castor oil is a well-known industrial multifunctional molecule with a variety of applications such as specialty soaps, adhesives, surfactants, cosmetics and personal care products, wax substitutes, inks, perfumes, plasticizers, paints and coatings, variety of lubricants, and greases, as well as in the food, fine chemicals, and pharmaceuticals industries (Achaya, 1971, Borg et al., 2009). Since castor oil is a polar dielectric with a relatively high dielectric constant, the dried castor oil is used as a dielectric fluid within high-performance highvoltage capacitors. RA and 12-hydroxy stearic acid (12-HSA) are derived from castor oil and hydrogenated castor oil (HCO), respectively. The three functionalities present in RA made this molecule very unique in the chemical world. Ester functionality of castor oil can involve in the hydrolysis, esterification, alcoholysis, saponification, hydrogenolysis, amidation, and halogenation, and generate final products like fatty acids, glycerol esters, partial esters, soluble/ insoluble soaps, alcohols, amine salts, amides, acid chlorides, etc. The unsaturation of castor oil particularly that of RA can involve in the reactions like

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oxidation, hydrogenation, epoxidation, halogenation, sulfonation, addition reactions resulting in polymerized oils, hydroxy stearates, epoxidized oil, halogenated oils, sulfonated oils, etc. In similar way the hydroxy functionality can participate in reactions like dehydration, caustic fusion, halogenation, alkoxylation, esterification, sulfation, and urethane resulting in dehydrated castor oil (DCO) and its fatty acids, sebacic acid, 2-octanol, 10-undecenoic acid (UDA), heptaldehyde, halogenated oils, alkoxylated oils, phosphate esters, turkey red oil, urethane polymers, etc. Due to this uniqueness, castor oil has become a potential alternative to petroleum-based products and also projected as a best candidate to exploit in biorefinery mode as thousands of derivatives can be prepared from it. In addition, castor oil is completely biodegradable and renewable feedstock. Several interesting reviews have been published in the literature related to castor oil production, chemistry, and value-added products (Achaya, 1971; Borg et al., 2009; Gayki et al., 2015; Mubofu, 2016; Mutlu and Meir, 2010; Pabi´s and Kula, 2016; Patel et al., 2016).

8.2 DERIVATIVES OF CASTOR OIL BASED ON UNSATURATION OF RICINOLEIC ACID 8.2.1 Hydrogenated Castor Oil A variety of products like HCO, 12-HSA, and methyl-12-HSA are produced by the hydrogenation of castor oil followed by hydrolysis and esterification (Fig. 8.1). Raney nickel is the most commonly used catalyst for the hydrogenation at a temperature of 150 C under hydrogen pressure of around 150 psi for complete hydrogenation (Naughton, 1974). Hydrogenation of castor oil improves the melting point, stability, and thermal characteristics of the oil. HCO is widely used in grease formulations, leather polishes, paint additives, wax, rubber and plastic manufacture, fruit coating, carbon paper, candles and crayons, and cosmetic and pharmaceutical preparations. 12-HSA is produced after saponification and acid hydrolysis of HCO and it is converted into methyl-12-HSA. All these products are useful for the preparation of metallic soaps such as lithium, calcium, etc. for use in multipurpose greases and lubricants (Ishchuk et al., 1974, 1986).

8.2.2 Epoxy Castor Oil The double bonds present in fatty acid chains of castor oil or its alkyl esters undergo reaction with peracids in presence of a catalyst to form epoxides. Epoxidized castor oil was prepared (Fig. 8.2) by reacting castor oil with 30% hydrogen peroxide using Amberlite and glacial acetic acid in toluene with 84% yield (Park et al., 2004). Epoxidized castor oil was also prepared using

Chemical Derivatization of Castor Oil Chapter | 8

FIGURE 8.1 Preparation of HCO and its products.

FIGURE 8.2 Preparation of epoxidized castor oil.

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peracetic acid generated in situ from acetic acid and 30% hydrogen peroxide in the presence of an ion exchange resin as catalyst (Amberlite IR-120) with an epoxy yield of 78% (Fiser et al., 2012). Most common catalysts employed for epoxidation reaction are mineral acid like sulfuric acid, acidic cation exchange resin such as the sulfonated polystyrene-type Amberlite, tungstenbased catalyst, Ti(IV)-grafted silica catalyst (Abdullah and Salimon, 2010), and methyl-tri-n-octylammonium diperoxotungstophosphate (Chakrapani and Crivello, 1998). Epoxidized castor oil used directly as plasticizers and polymer stabilizers, paint and coating components, and lubricants. The epoxy ring is reactive and is useful as an potential intermediate for the preparation of alcohols, glycols, alkanolamines, polyols, and several polymers. Coatings were prepared by reacting tetraethoxysilane with castor oil or epoxidized castor oil (de Luca et al., 2006). Regioisomers of azido diol were prepared from ricinoleic epoxide and sodium azide and investigated for its biological activity (Furmeier and Metzger, 2003). Butyl 10,12-dihydroxy-9-behenoxystearate and butyl 10,12-dihydroxy-9-octyloxystearate were prepared from epoxy RA with low-temperature property and higher oxidation stability, respectively, compared with common synthetic esters. This observation reveals that the increasing chain length of the mid-chain ester has a positive influence on the low-temperature properties and better oxidation stability is achieved when the chain length of the mid-chain ester decreases (Salimon et al., 2011). Polymeric nanocomposites were synthesized from epoxidized castor oilbased polymer and montmorillonite clay employing an in situ polymerization route with efficient mechanical properties compared with traditional polymers (Thamaraichelvi et al., 2016). Fatty acidbased cyclic carbonates were reported by intramolecular rearrangement of RAbased epoxy carbonate ester with Lewis acids without the use of carbon dioxide. Initially, methyl-8-(3-(2-(methoxy carbonyloxy)octyl)oxiran-2-yl)octanoate and methyl-8-(3-(2-(ethoxycarbonyloxy)octyl)oxiran-2-yl) octanoate were prepared by the carbonate interchange reaction between methyl ricinoleate and dialkyl carbonates followed by epoxidation. These products are converted into five- and six-membered cyclic carbonates through a spiroorthocarbonate intermediate by intramolecular rearrangement in presence of Lewis acids (Naganna et al., 2016).

8.2.3 Ozonolysis of Castor Oil Triglycerides having 9-carbon fatty acids with terminal hydroxyl groups were prepared by ozonolyis of castor oil followed by sodium borohydride or lithium aluminum hydridebased reduction (Fig. 8.3). Condensation polymers like polyurethane (PU), polyethers, and polyesters can be prepared by employing these fatty acids as monomers (Mubofu, 2016).

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FIGURE 8.3 Ozonolyis and reduction products of castor oil.

8.2.4 Preparation of 9,10,12-Trihydroxy Octadecanoic Acid 9,10,12-Trihydroxy octadecanoic acid-rich fatty acid mixture was prepared from castor oil and converted it into their alkyl esters followed by acylation of hydroxyl groups with fatty acid anhydrides (C2C5). These products exhibited wide viscosity range and excellent low pour point and can be used as potential lubricant base stocks (Rao et al., 2013a) for variety of applications.

8.2.5 Halogenated Derivatives of Castor Oil Halogenated derivatives were reported by reacting chlorine or bromine or NBS, etc., with the double bond of RA (Yousef et al., 2001) for use in nitrile rubber formulations.

8.2.6 Novel Derivatives of Ricinoleic Acid Employing Metathesis Reaction Olefin metathesis of methyl ricinoleate was exploited for the development of potent intermediates for variety of applications (Fig. 8.4) as described here. Self-metathesis of methyl ricinoleate results in diester and diol, which are useful for the development of lubricants, surfactants, polymers, etc. (Kroha, 2004). Cross metathesis of methyl ricinoleate with methyl acrylate generates linear bifunctional derivatives. Methyl ricinoleate reacts with different aliphatic acids to produce branched acids, which on further ring closing metathesis results in lactones along with mono- and diesters. These intermediates find several synthetic application possibilities for renewable polyesters and polyanhydrides (Rybak and Meier, 2007; Ngo et al., 2006).

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FIGURE 8.4 Products generated from methyl ricinoleate via metathesis route.

8.3 DERIVATIVES OF CASTOR OIL BASED ON HYDROXY FUNCTIONALITY OF RICINOLEIC ACID 8.3.1 Dehydrated Castor Oil and Dehydrated Castor Oil Fatty Acids Castor oil is classified as a nondrying oil, and after dehydration process, this can be converted into semidrying or drying oil known as DCO. The catalytic dehydration processes are generally carried out at a temperature of 250260 C in the presence of catalysts under an inert atmosphere or vacuum, and most widely used catalysts reported are sulfuric acid, sodium bisulfate, phosphoric acid, phthalic anhydride, and acid-activated clays (Priest and von Mikusch, 1940; Terrill, 1940; Don, 1959; Bhowmick and Sarma, 1977; Forbes and Neville, 1940; Grummitt and Marsh, 1953). The dehydration process involves the removal of hydroxyl group along with one hydrogen present on the either of the carbons attached to the carbon bearing the hydroxyl group forming conjugated and nonconjugated fatty acids. In the similar way, RA, a hydrolyzed product of castor oil, can also be dehydrated into conjugated and nonconjugated linoleic acids (Fig. 8.5). Polymerization of conjugated RA and estolide formation are most common side reactions during the process. Polymerization can effectively be controlled by the addition of small quantities of antipolymerization reagents depending on the nature of catalyst employed (Achaya, 1971). The DCO and DCO fatty acids are extensively used in the preparation of coatings, vanishes, lubricants, paints, inks, alkyd resins, and primers (Ogunniyi, 2006; Mutlu

FIGURE 8.5 Preparation of DCO and DCO fatty acids.

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and Meir, 2010; Ramamurthi et al., 1998; Shende et al., 2002; Onukwlo and Igbokwe, 2008). DCO was reacted with glycerol and phthalic anhydride using 0.3% (wt%) NaOH for preparing alkyd resins with acid value of 6.6. The physicochemical properties and high chemical resistance of alkyd resin film revealed that they have promising applications in formulating paints (Hlaing and Mya, 2008).

8.3.2 Sulfated Castor Oil (Turkey Red Oil) Sulfated castor oil, popularly known as turkey red oil, is produced by reacting castor oil with sulfuric acid (Rangarajan and Palaniappan, 1958). Castor oil can be sulfonated directly with sulfur trioxide in continuous or batch equipment (Zamiri et al., 2011). Sulfation takes place on the hydroxy group of RA to form OSO3H group (Fig. 8.6). In a typical batch process, the reaction is carried out at # 35 C by reacting castor oil with concentrated H2SO4 for 34 hours. Turkey red oil is known for its wetting, emulsifying, and dispersing properties and used in the preparation of synthetic detergents, shampoos, lubricants, softeners, dyes, etc. (Gherca et al., 2012; Bishai and Hakim, 1955).

8.3.3 Acetylated Castor Oil Acetylated castor oil is an important derivative of castor oil and it is prepared by reacting the hydroxy group of RA with acetylating agent like acetic anhydride in quantitative yields (Mukherjea et al., 1978). Acetylated castor oil is well known as a secondary plasticizer for polyvinyl chloride (PVC) with low volatility, good dielectric properties, and lubricity with extensive

FIGURE 8.6 Preparation of sulfated castor oil (turkey red oil).

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use in PVC-based electrical cable industries. Acetylated castor oil is also used as plasticizer for other polymers like nitrocellulose, polyurethane, and ethylene-vinyl acetate copolymer. In another study, the acylated derivatives of castor oil and castor oil fatty acid methyl and 2-ethylhexyl esters were prepared employing a series of anhydrides (C1C6) with excellent lowtemperature properties and attractive flash points, air release value, NOACK volatilities, load-carrying capacity, emulsion stability, etc. These lubricant base stocks will have lot of potential for use in hydraulic- and metal-working fluids and other industrial fluids with their wide range of properties (Geethanjali et al., 2016) after appropriate formulations.

8.3.4 Castor OilBased Estolides HFA-based estolides are generally formed by esterification between the carboxyl group of a fatty acid molecule with the hydroxy group of another fatty acid molecule, and the resulting compounds exhibit excellent lowtemperature properties (Isbell et al., 2006; Naughton, 1974). RAbased estolides were prepared by controlled homopolymerization of two molecules of castor oil fatty acids with about 95 acid value (Bhaskar et al., 2014). The free carboxylic group of the estolide was further esterified with liner or branched chain alcohols along with acetylation of free hydroxyl group to obtain RAbased estolide esters and their acetates. These novel classes of biodegradable compounds exhibit superior low-temperature and viscometric properties with very good oxidative stability. The secondary linkages of the estolides are more resistant to hydrolysis compared with triglycerides, and the unique structure of the estolide results in materials that have physical properties far superior to those of vegetable and mineral oils for certain applications like lubricants, greases, plastics, inks, cosmetics, and surfactants. A new class of estolides were prepared by esterifying methyl ricinoleate with dicarboxylic acids (C6, C8, and C10) under solvent- and catalyst-free conditions to obtain dimer estolide and monoacid estolide with one unreacted carboxyl group (Sammaiah et al., 2016). The estolide esters exhibited wide viscosity range varying from ISO VG 15 to 46 with outstanding lowtemperature property (248 to , 2 60 C), and these products will have potential as lubricant base stocks for environmentally friendly industrial oils and automotive applications including low-temperature applications.

8.3.5 Castor OilBased Polymer Products A variety of polymers like polyamides, polyethers, polyesters, poly(2-hydroxyethylmethacrylate), and polyurethanes are synthesized by making use of the hydroxyl group of RA (Mubofu, 2016). As the RA contains secondary hydroxyl group, castor oilbased PUs exhibit semiflexible or semirigid properties (Mutlu and Meir, 2010). The foam derived from castor oil becomes

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exhibited outstanding biodegradable characteristics compared with petroleum PU foam (Cangemi et al., 2008). These polymers have been exploited for several biomedical applications like drug delivery systems (Kunduru et al., 2015; Shikanov et al., 2004), tissue engineering (Baheiraei et al., 2016), bioadhesives (Ferreira et al., 2007), and wound dressing (Yari et al., 2014). PUs are a class of compounds with a urethane functionality (NHCOO) in the polymer molecule and prepared from organic isocyanates and hydroxy group containing molecules such as castor oil. Castor oilbased PUs have been employed for variety of applications like thermoplastic elastomers, rigid, semirigid and flexible foams, sealants, elastomers, biomedical implants, adhesives, and coatings (Mubofu, 2016). Epoxyterminated PU prepolymers were synthesized by reacting castor oil by using a curing agent like 1,6-hexamethylene diamine (Yeganeh and Hojati-Talemi, 2007). Castor oil, in combination with recycled polyethylene terephthalate, adipic acid, and polyethylene glycol (PEG), has been used to prepare PU coatings for insulation applications (Yeganeh and Moeini, 2007). PUsorganoclay nanocomposites were prepared with a combination of polypropylene glycol polyol and DCO (15%), enforced with C30B nanofillers for use in coatings, adhesives, and automotive applications (Alaa et al., 2015). Polymer electrolyte films were prepared by adding LiI and NaI to castor oilbased PU for applying as a host in polymer electrolyte for electrochemical devices, and this study revealed that the castor oil PU can be used as an alternative bio-based polymer membrane in polymer electrolytes (Ibrahim et al., 2015). Hyperbranched polyester-/bitumen-based nanocomposites were prepared employing monoglyceride of castor oilbased carboxyl terminated prepolymer and 2,2-bis (hydroxymethyl) propionic acid for low-cost high-performing surface-coating binder materials applications (Bhagawati et al., 2016). A series of polyesters were synthesized from RA and 10-UDA with antibacterial activity (Totaro et al., 2014). Hydrorphobicity imparts to the polyesters of RA due to its dangling chains and this property influences the mechanical and physical properties of the polymers. These polymers produce different types of products ranging from solid implants to in situ injectable hydrophobic gels (Kunduru et al., 2015; Petrovi´c et al., 2008). Castor oil was used for coating applications by preparing β-ketoesters by reacting hydroxy functionality of castor oil with t-butyl acetoacetate in high yields and the products exhibited good glosses to the films (Trevino and Trumbo, 2002). Ethoxylates of castor oil and HCO or their respective free fatty acids and alkyl esters are well-known surfactants. Ethoxylation is a chemical process in which ethylene oxide is reacted in desired molar ratios with an alcohol or acid or triglyceride oils to produce surfactants. Ethoxylated products were prepared by directly reacting ethylene oxide with the ester group and hydroxyl group of castor oil fatty acid methyl esters in the presence of an

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alkaline catalyst, and these products are useful as nonionic surfactants with good properties (Zhang et al., 2015). Rhizomucor miehei lipase-catalyzed esterification was employed for the preparation of PEG esters of castor oil fatty acids to overcome problems associated with chemical processes (Ghosh and Bhattacharyya, 1998).

8.3.6 Potent Hydroxy Derivatives of Ricinoleic Acid (Z)-Ethyl 12-nitrooxy-octadec-9-enoate (NCOE) was prepared from RA, and the studies revealed that NCOE induces short-lasting hypotension and bradycardia, and promotes vasorelaxation due to NO release through the compound metabolism (Machado et al., 2014). A series of RAbased glycosides were prepared from methyl ricinoleate under KoenigKnorr conditions with good antibacterial activity (Kuppala et al., 2016). RAbased Schiff bases with good antimicrobial and antibiofilm activities were prepared by reacting methyl-12-aminooctadec-9-enoate with different substituted aromatic aldehydes (Mohini et al., 2014). A series of RAbased lipoamino acid derivatives were synthesized from methyl-12-aminooctadec-9-enoate and L-amino acids, namely glycine, alanine, phenyl alanine, valine, leucine, isoleucine, proline, and tryptophan, and some of the products exhibited promising antibacterial and antifungal activities (Mohini et al., 2016).

8.4 DERIVATIVES BASED ON ESTER FUNCTIONALITY OF CASTOR OIL 8.4.1 Hydroxy Fatty Acid Esters HFA esters find variety of uses as emollients, emulsifiers, plasticizers, mold-release agents, foaming agents for cosmetics, effective wetting agents, pigment dispersants and stabilizers, pearlizing and opacifying agents, solubilizes for hydrophobic components in cosmetics, moisturizers for winterized and sun-dried skin, waxes, and chemical intermediates (Hayes, 1996). Simple fatty acid esters are generally prepared either by esterification of fatty acids or by transesterification of fatty acid methyl esters with monohydric alcohols. However, in case of HFA, there is a chance of homopolymerization resulting in estolide (Hayes and Kleiman, 1996). In this context, 1,3-specific lipases like Mucor miehei lipase (Lipozyme TM) have proved to be valuable catalysts for esterification of RA or 12-HSA with monohydric alcohols like decyl, dodecyl, tetradecyl, hexadecyl, octadecyl, etc., without the formation of estolides and lactones (Hayes, 1996; Vorderwulbecke et al., 1992; Ghosh and Bhattacharya, 1992). An efficient and simple lipase-mediated synthesis of alkyl ricinoleates and 12-hydroxy stearates was carried out by transesterification of methyl ricinoleate/12hydroxy stearate and various alcohols (octyl, decyl, undecenyl, dodecyl,

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hexadecyl, octadecyl, and octadecenyl alcohols) in a solvent-free system without estolide formation (Karuna et al., 2005). These esters were converted to sulfated sodium salts and evaluated for surfactant properties, and among the compounds, dodecyl ricinoleate and dodecyl 12-hydroxy stearates were found to be the best in their respective series and were also found comparable to sodium dodecyl sulfate (SDS). In another study, esterification of acylated castor fatty acids with branched monoalcohol, 2-ethylhexanol, and polyols, namely neopentyl glycol (NPG), trimethylolpropane (TMP), and pentaerythritol (PE), was carried out, and these compounds exhibited very low pour points (230 to 245 C), broad viscosity ranges 20.27370.73 cSt, higher viscosity indices (144171), good thermal and oxidative stabilities, and high weld load capacities suitable for multirange industrial applications such as hydraulic fluids, metal-working fluids, gear oil, forging, and aviation applications (Kamalakar et al., 2015). Two bioactive esters were prepared from RA by reacting with 2,4- or 2,6-diisopropylphenol (Khan et al., 2012). Castor oil fatty acids are being used for the preparation of almost all the derivatives without separating the non-HFAs from RA. In this context, the process reported for the enrichment of methyl ricinoleate up to 98% purity from castor oil fatty acid methyl esters using liquidliquid extraction is a very attractive method for avoiding non-HFAs during the production of RAbased derivatives (Rao et al., 2013b). Enzymatic hydrolysis is an attractive route for the production of free RA from castor oil or methyl ricinoleate without formation of estolide. A first order reversible kinetic model was proposed for the enzymatic hydrolysis of methyl ricinoleate employing Candida antarctica lipase (Neeharika et al., 2015). In a similar way, enzymatic hydrolysis of enriched castor oil fatty acid methyl esters was optimized using response surface methodology and the predicted results matched with the experimental values (Neeharika et al., 2014).

8.4.2 Castor OilBased Biodiesel Even though biodiesel can also be prepared from castor oil in quantitative yields by reacting with methanol (Jeong and Park, 2009; Franc¸a et al., 2009), it may not be commercially viable due to the high cost of castor oil compared to other vegetable oils. Due to its high density, viscosity, and hygroscopicity, castor oilbased biodiesel may not be directly used in place of diesel in internal combustion engines (Scholz and Silva, 2008). However, it may be useful if it is blended with petrodiesel or biodiesel made from other common vegetable oils like cotton seed or soybean oils in certain proportions due to its specific lubricity property in concentrations up to 0.2% and its

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high energy value and positive properties and good oxidative stability (Canoira et al., 2010; Berman et al., 2011; Albuquerque et al., 2009).

8.4.3 Preparation of Ricinoleyl Alcohol Castor oil and its methyl esters are reduced to hydroxy fatty alcohols employing catalysts like copper/cadmium (3:1 wt/wt) (Pantulu and Achaya, 1964) and sodium in presence of secondary alcohols (Hansley, 1947), respectively. The saturated alcohol, namely 12-hydroxy stearyl alcohol, is produced from methyl-12-hydroxy stearate using nonselective catalysts. Ricinoleyl alcohol is a base material for the preparation of surface active compounds. The disulfates of 12-hydroxy ricinoleyl alcohol found to exhibit a good wetting, detersive, and emulsifying properties (Subrahmanyam and Achaya, 1961; Pantulu and Achaya, 1971).

8.4.4 Ricinoleic AcidBased Amides Several RAbased amide derivatives were reported in the literature. Phenylacetylrinvanil is one such derivative prepared from rinvanil, which in turn prepared from enzymatic amidation of methyl ricinoleate with vanillamine hydrochloride (Castillo et al., 2008; Appendino et al., 2002). This potent molecule was exploited for possible applications like analgesic, antiinflammatory, and anticancer efficacies. Number of biologically active amides were prepared by reacting methyl ricinoleate with benzyloamines (Dos Santos et al., 2015) or RA chloride with different amines (Narasimhan et al., 2007). Hydrazide derivatives of isoniazid were also reported with antimicrobial activity by making use of RA (Rodrigues et al., 2013).

8.4.5 Ethanolamides of Castor Oil Fatty Acids Fatty mono- and diethanolamides were prepared from castor oil fatty acids by reacting with monoethanolamine and diethanolamine. The ethanolamides were further sulfated using chlorosulfonic acid and the sulfated sodium salts with good surfactant properties, namely surface tension, critical micelle concentration, emulsifying property, wetting, foaming power, and calcium tolerance, which are almost comparable to sodium dodecyl sulfate (SDS), a popular anionic surfactant (Kamalakar et al., 2014).

8.5 UNIQUE DERIVATIVES OF CASTOR OIL 8.5.1 Castor OilBased Dimer Acids Dimer acids are a variety of substituted cyclohexenedicarboxylic acids formed by a DielsAlder type reaction from self condensation of unsaturated

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straight chain fatty acids, and these are the highest molecular weight dibasic acids commercially available (Cowan, 1962). In case of castor oil, dimer acids are industrially prepared by thermal polymerization of DCO fatty acids (Breakey and Rowe, 1960). Catalysts such as acid clays, iodine, and sulfur, and sodium or potassium bisulfates were used to improve the yields of dimer acids (Den Otter, 1970; Silverstone, 1967). Dimer acids are predominantly isomers of 36-carbon dibasic acids with smaller amounts of 54-carbon tribasic acids or higher molecular weight polycarboxylic acids, and pure dimer acids are prepared by fractionation of the commercial dimer acids. Dimer acids are capable of undergoing three types of chemical reactions, which involve the carboxyl group, double bonds, and the carbon atoms adjacent to the carboxyl groups (alpha carbon) and from the practical point of view dimer acids possessing a high degree of flexibility by virtue of the longchain part in the molecule. The dimer acids have several broad applications in the manufacture of polymer technology and contribute to the thermal and hydrolytic stability, water repellency, and pigment wetting properties of the polymers like polyamides, polyesters, and epoxy resins. Dimer acids are also used in several other applications like corrosion inhibition activity in petroleum-processing equipment, enhancing lubricating property of hydraulic fluids, reduce grain less, and increase the stability in greases (Kale et al., 1994; Bajpai and Khare, 2004; Islam et al., 2014).

8.5.2 10-Undecenoic Acid and Heptaldehyde Pyrolysis of castor oil or its fatty acids or its fatty acid esters under appropriate experimental conditions produce UDA or its ester and heptaldehyde as major products (Fig. 8.7). During the pyrolysis, the RA, which is present in the castor oil fatty acids, undergoes McLafferty-type rearrangement to produce UDA and heptaldehyde. The pyrolysis of methyl ricinoleate is the preferred method since castor oil has a higher viscosity, thus resulting in polymerized materials and poisonous gases during its direct pyrolysis (Naughton, 1974; Flinn, 1978). A number of studies have been performed considering different pyrolysis temperatures (Vernon and Ross, 1936; Gupta and Aggarwal, 1954) and temperature of 400500 C is essential for effective conversion. Pyrolysis of castor oil requires higher temperatures than its corresponding fatty acid methyl esters. The pyrolysis reaction is generally carried out in the absence of air to minimize the oxidative degradation. The reaction has been done with and without a catalyst, and steam is used as diluent to improve the product quality while minimizing the charring of the products (Vishwanadham et al., 1995; Naughton, 1974; Domsch, 1994; Hunting, 1981). Heptaldehyde is used as a solvent for rubber, resins, and plastics, and UDA can directly be used for applications as bactericides and fungicides (Naughton, 1974). 10-UDA was shown to be a valuable precursor for the

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FIGURE 8.7 Preparation of 10-UDA and heptaldehyde from castor oil.

synthesis of antitumor compounds, antibiotics, and nylon 11 (Rahman et al., 2005; Mustafa et al., 2005; Van der Steen et al., 2008; Green and Wittcoff, 2003; Van der Steen and Stevens, 2009).

8.5.3 Sebacic Acid and 2-Octanol Sebacic acid, a C-1,10 dicarboxylic acid, is an industrially important oleochemical produced from castor oil by alkali fission reaction. Castor oil or its split upon treatment with excess alkali at elevated temperatures in presence of a suitable catalyst produces sebacic acid and 2-octanol as major products (Fig. 8.8). Different batch and continuous processes were reported in the literature (Hargreaves and Owen, 1947; Naughton, 1974; Dytham and Weedon, 1960), and the reaction is temperature and alkali dependent. When the reaction performed at 180200 C with equimolar ratio of castor oil:split:methyl ester and alkali (NaOH/KOH), 10-hydroxydecanoic acid and 2-octanone found to be the major products, whereas sebacic acid and 2-octanol are the major products when the temperatures are in the range of 250270 C using 2 molar excess of alkali (Naughton, 1974; Flinn, 1978; Diamond et al., 1965). Chemically inert heat transfer fluids with high boiling points ( . 300 C) such as mineral oil, therminol, m-cresol, Paratherm NF, glycol

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FIGURE 8.8 Preparation of sebacic acid and 2-octanol from castor oil.

oil, or petroleum oil were also employed in the reaction mixture to reduce foaming and solidification of the reaction mixture while improving the product yield and purity (Vasishtha, et al., 1990; Ries and Totah, 1999; Logan and Udeshi, 2002). Sebacic acid and its derivatives have a variety of industrial uses in plasticizers, lubricants, hydraulic and break fluids, coatings, perfumes, cosmetics, nylon 610, candles, and specific medical applications (Tang et al., 2006; Kim et al., 2009). 2-Octanol is used as solvent and finds applications as plasticizer, dehydrator, antibubbling agent, and floating agent in coal industry (Datta et al., 2011).

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Chapter 9

Chemical Modification of High Free Fatty Acid Oils for Biodiesel Production Godlisten G. Kombe The University of Dodoma, Dodoma, Tanzania

Chapter Outline 9.1 Introduction 9.2 Production of Biodiesel 9.2.1 Types of Feedstocks 9.2.2 The Potential of High FFA Feedstocks in Biodiesel Production 9.2.3 Challenges of Processing High FFA Feedstocks

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9.3 Chemical Modification of High FFA Feedstocks for Biodiesel 9.3.1 Potential Processes for Modification of High FFA Feedstocks 9.4 Conclusion and Recommendations References Further Reading

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9.1 INTRODUCTION Petroleum as a fossil fuel is a nonrenewable resource that will persevere in driving up energy costs as oil global production declines. The world energy needs are expected to rise significantly in the future due to growth in population and economic development (IEA, 2007). One of the energy sectors that deserve exceptional attention in regard to consumption of fossil fuel is the transport sector. The transport sector is growing rapidly and therefore an improvement in efficiencies and diversification in this area will be essential. According to Goldemberg and Johansson (2004), fossil fuel accounts for 97% of transportation energy in the industrialized countries, with natural gas (2%) and electricity (1%) accounting for the rest. Alternative energy sources are under research to supplement diesel fossil fuel consumption. Within the alternative energy segment, biodiesel has captured an increasing interest due to its technical and environmental credentials such as reduced global Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00009-X Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

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warming escalation, renewability, biodegradability, clean combustion behavior, good lubricity, and higher cetane number, which gives better ignition quality (Adaileh and AlQdah, 2012). Furthermore, biodiesel development can create new job opportunities and reduce the total dependency on fossil fuels and save expenditure on diesel for countries that depend greatly on the importation of petroleum diesel for energy. Biodiesel is a fatty acid alkyl ester formed from the reaction of triacylglycerol (edible or nonedible oil feedstocks) with a monohydric alcohol. The feedstock from edible oil tends to compete with food supply and result into high biodiesel prices. This is due to the facts that the cost of the feedstocks accounts for 70%90% of the total cost of biodiesel production (Khandelwal and Chauhan, 2012). Nonedible oil feedstocks are cheap since they may not compete with either food supply or land for food cultivation (Sawangkeaw and Ngamprasertsith, 2013). Unfortunately, most nonedible oil feedstocks have free fatty acid (FFA) contents of above 3%, which hinder the use of traditional biodiesel production processes, base catalyzed transesterification, which is kinetically much faster and recognized to be viable economically (Dorado et al., 2002; Helwani et al., 2009). The high FFA tends to form concurrent soap with the catalyst, which reduces the reaction yield and significantly hinders the washing process by developing emulsions, thus leading to extensive yield losses. The challenges posed by high FFA calls for their modifications before alkali transesterification. Three potential FFA technologies for high feedstock modifications, namely, acid esterification, chemical reesterification/glycerolysis, and neutralization prior to alkali transesterification process are discussed in this chapter.

9.2 PRODUCTION OF BIODIESEL 9.2.1 Types of Feedstocks Fats and oils as potential feedstock for biodiesel production are made up of triacylglycerol (triglyceride) molecules. Each triglyceride has three longchain fatty acids of 822 carbons attached to a glycerol backbone. Once the fatty acid chains are detached from the triacylglycerol, they tend to be free from their mother bond and become “FFAs.” The FFAs are the ones that are responsible for the acidity in the oil. As stated earlier, these FFAs are responsible for low biodiesel yield in the alkali-catalyzed transesterification process. Biodiesel feedstock can be categorized easily based on the amount of FFA produced. According to Kinast (2003), biodiesel feedstock can be refined oils with less than 1.5% FFA, low FFA oil with ,4% FFA, and high FFA oil with $ 20% FFA. Depending on the amount of FFA, the oil source can either be from edible or nonedible oils. Several fats and oil sources from plant and animal species have been recognized as the possible edible grade feedstocks oil for biodiesel production

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in different countries. Palm oil, soybean oil, rapeseed oil, sunflower, and beef tallow are the most common feedstocks in this category. Most of these edible grade oils are said to have low FFA, which makes them a good candidate for biodiesel production by alkali transesterification. Unfortunately, their use competes directly with food sources and may cause food insecurity. Yet again, their competition with the edible oil market makes the cost of biodiesel unattractive. Nonedible oil feedstocks may be obtained from plant oil, waste oil, and other sources such as algae, microalgae, fungi, and terpenes. The potential plant oil source includes castor oil, jatropha oil, Pongamia, Karanja oil, neem oil, and Mahua oil. Initially, most of these plants species were regarded as wild species. Waste oil sources include oil from used vegetable oils, yellow grease, brown grease, and soap stock (which is a by-product of vegetable oil refining process).

9.2.2 The Potential of High FFA Feedstocks in Biodiesel Production As stated earlier, the high cost of biodiesel is mainly contributed by the cost of feedstock, which usually accounts for 70%90% of the total cost of biodiesel. This can be attributed to the use of edible grade oil, which has high market demand. The cost of edible grade oil is likely to rise as the world population keeps growing with an increase in their market demand. At the same time, production of biodiesel from edible grade oil may compete directly with food sources and cause food insecurity (Chhetri et al., 2008). Therefore, it is challenging to justify the use of these oils for fuel for the sustainable development of biodiesel. To overcome these challenges, several researchers have worked on nonedible grade oil as a good alternative (Chhetri et al., 2008; Gui et al., 2008; Khandelwal and Chauhan, 2012; Mathiyazhagan et al., 2011; Murugesan et al., 2009; Patil and Deng, 2009). Nonedible grade oils are not suitable for food and therefore they do not compete with edible oils. Their sources may be plant, animal, algae, fungi, and other waste oil sources. Nonedible grade oil from plant species can be grown and survive in marginal land and wastelands, which are not suitable for food crops. Subramanian et al. (2005) identified more than 300 tree species as a good source of nonedible grade oil, this means that nonedible grade oils from these plants species can be a significant potential feedstock for biodiesel production. Nonedible plants can be cultivated under different environment and climatic conditions, including on waste, sandy, and saline soils. They can still produce high crop yield with minimum inputs and lower the cultivation cost (Gui et al., 2008; Kumar et al., 2007). Some nonedible plant species like Karanja can be cultivated to produce seed oils and improve the soil quality. Karanja is a nitrogen-fixing tree, which can

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improve the exhausted soil and make it reusable for the agricultural purpose (Gui et al., 2008). Waste oil is other potential feedstocks for biodiesel production; they are cheaper than other types of oil. Several researches have worked on the production of biodiesel from waste oil (Kulkarni and Dalai, 2006; Mustafa, 2007; Predojevic, 2008; Van Kasteren and Nisworo, 2007; Wang et al., 2007; Yuan et al., 2008; Zhang et al., 2003). The price of virgin vegetable oils is normally around 23 times that of waste oil (Kulkarni and Dalai, 2006). Their source differs from country to country depend on the vegetable oil consumptions of a given country (Bankovi´c-Ili´c et al., 2012). Substantial quantities of used cooking oil are available globally. Math et al. (2010) estimated the total amount of waste oil produced per year from the United States (US), EU countries, Canada, and India to about 1.675 million tons. If only 80% can be recovered for biodiesel production, then about 1.34 million tons of biodiesel can be produced from this source. The available quantities of oil from plant, animal, and waste oil cannot meet the worldwide biodiesel production demand. This gives room to the search for other feedstocks sources such as those of algae. Algae are another good source of nonedible oil for biodiesel production. Algae are easy to cultivate, can grow in different places as long as there are enough sunshine and some nutrients under little or even no care with water that is unsuitable for human consumption. Their growth rates can be improved by the addition of specific nutrients and sufficient aeration. Since algae species can be prepared to grow in different environmental conditions, it is, therefore, possible to find species appropriate to local environments or specific growth characteristics. Algae can contribute to a reduction in land requirements for farming since they are recognized for higher energy yield per hectare. Sheehan et al. (1998) assessed the yield (per acre) of oil from microalgae to be above 200 times the yield from the best-cultivated plant oil. The microalgae oil has much higher biomass yields than land plants, some species can accumulate up to 20%50% dry weight of biomass (Bankovi´c-Ili´c et al., 2012).

9.2.3 Challenges of Processing High FFA Feedstocks A good number of studies have been conducted on various transesterification routes for biodiesel production. The studied transesterification routes include noncatalyzed, alkali-catalyzed, acid-catalyzed, or enzyme-catalyzed (Atabani et al., 2012; Ayhan, 2003; Helwani et al., 2009; Leung et al., 2010; Meher et al., 2006). The alkali-catalyzed transesterification has been widely accepted and used as traditional biodiesel production method (de Lima da Silva et al., 2009; de Oliveira et al., 2005; Dorado et al., 2002, 2004a,b; Nakpong and Wootthikanokkhan, 2010b; Rashid and Anwar, 2008). In this transesterification reaction, methanol/ethanol as monohydric alcohol is allowed to react with triglycerides in the presence of alkali catalyst to

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produce biodiesel and glycerol as a by-product. The catalyst used may be heterogeneous or homogeneous base catalyst. The homogeneously catalyzed reaction is kinetically much faster, requires low temperature (60 C) with capabilities of achieving high conversion (98%) when compared with other transesterifications routes (Atadashi et al., 2011; Ejikeme et al., 2010; Helwani et al., 2009). It has been recognized to be economically feasible (Helwani et al., 2009). Bacovsky et al. (2007) conducted a study on the status of biodiesel production technology and concluded that the alkali transesterification is the most of the commercialized biodiesel production technology. Unfortunately, the main challenge facing this technology is its sensitivity to the purity of reactants. It is very sensitive to both water and FFAs content (Encinar et al., 2005; Van Kasteren and Nisworo, 2007; Zhang et al., 2003). The reaction works for feedstock with FFA content up to 3% without affecting the process negatively (Knothe et al., 2005). Therefore, in order to achieve high biodiesel yield, the FFA in the feedstock should be less than 3% (Dorado et al., 2002). Most of the nonedible oil feedstocks have high FFA of about 3%, which hinder their direct application in the alkali transesterification. The high FFA tends to form concurrent soap catalyst and lower the biodiesel yield. Furthermore, the soap formed affects significantly the washing process by forming emulsions, thus leading to substantial yield losses (Canakci and Van Gerpen, 1999; Freedman et al., 1984; Hanna and Ma, 1999; Marchetti et al., 2007; Tongurai et al., 2001). These challenges call for necessary modification of FFA in these feedstocks prior to alkali transesterification process. The FFA in the feedstock can be modified by neutralization to soap stock and separated from oil or modified into an ester or reesterified back into the oil by neutralization, esterification, and reesterification/glycerolysis process, respectively. These three potential FFA modification processes are the subject of discussion in this chapter.

9.3 CHEMICAL MODIFICATION OF HIGH FFA FEEDSTOCKS FOR BIODIESEL 9.3.1 Potential Processes for Modification of High FFA Feedstocks As seen earlier, most of the less expensive biodiesel feedstocks available in the market contain a high amount of FFA, that is, above 3%, which is required for alkali-catalyzed transesterification. The high FFA in nonedible oil can be modified and converted easily by using the alkali-catalyzed transesterification. The modification of FFA can be done by neutralizing with any strong base, esterifying with a monohydric alcohol in the presence of an acidic catalyst, or reesterifying with glycerol with or without a catalyst.

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9.3.1.1 Neutralization One of the oldest methods for modifying FFA in oil is the neutralization process. In this process, an alkali (normally NaOH) is added to the acidic oil and thereby precipitating the FFA as soap stock; the latter is then removed by gravity or mechanical separation from the neutral oil. Neutralization has been one of the most commonly used methods for lowering the FFA in oil during the chemical refining of vegetable oil. It lowers the FFA, along with substantial quantities of mucilaginous substances, phospholipids, and color pigments (Bhosle and Subramanian, 2005). Neutralization is the process of removing the natural acidity of the oil emanating from the presence of FFAs. According to Gunstone et al. (1994), vegetable oils are neutralized to remove the FFAs, some of the dirty and denatured phosphatides, saponifiable impurities and pigments. The neutralization process depends on the initial amount of FFA in oil, the type of alkali used, strength of the alkali solution, the amount of excess alkali used over the stoichiometrically required, the temperature during addition of alkali, and the degree of agitation. The degummed or crude oil is mixed with a proportioned amount of dilute caustic soda solution. The mixture is then adequately blended at room temperature for 515 minutes to ensure sufficient contact of NaOH (caustic soda) or any alkali with the FFA, phosphatides, and color pigments. The gums are hydrolyzed by the water in the caustic solution and become oil insoluble. The mixture is then heated to 74 C to provide the thermal shock necessary to break the emulsion and then centrifuged to remove the soap stock. The separation is accomplished by allowing a small amount of the soap stock phase to pass along with the refined oil for removal by the water wash centrifuge (Sullivan, 1968). After separation of the soap stock from the oil, the oil still contains some soap, which needs to be washed (Gunstone et al., 1994). This is done by adding hot water at 90 C; the quantity of water used is normally 8% of the weight of the oil. Neutralization process has the capability of reducing the high FFA in oil to the value less than 0.03% depending on the characteristics of oil (Hodgson, 1996). However, the main challenge facing the vegetable oil neutralization process is the loss of the neutral oils due to (1) formation of emulsions of oil in water, these emulsions may be very stable so that on separation of the soap solution, oil may be entrained; (2) entrainment of oil droplets with the soap solution, the high viscosity of the soap solution hinders the settling of oil droplets; and (3) saponification of the neutral oil under the influence of the alkali. The waste water from the washing process requires treatment to meet the statutory standards; this is normally expensive and increases the cost of the neutralized oil. The process has also been faced with the challenge of getting the right amount of NaOH to be used and process conditions to minimize loss of neutral oil (Gunstone et al., 2007).

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TABLE 9.1 Optimized Conditions for Jatropha and Castor Oil Neutralization Unit Initial FFA

Jatropha Oil

Castor Oil

4.54

6.5

63

64

Temperature



C

Strength of NaOH



Be (Baume)

16

15

Excess NaOH amount

%

0.30

33

Oil loss

%

10.05

18.64

Final FFA

%

0.209

0.38

The loss of neutral oil due to the formation of soap limits the use of neutralization process for oils with more than 5% FFA. For oils with more than 5% of FFA, neutralization results into higher losses of neutral oil, this can be attributed to saponification and emulsification (Bhosle and Subramanian, 2005). Aryusuk et al. (2008) studied the effects of crude rice bran oil components on the neutralization losses and reported the average refining loss in the range of 13.2%13.4% and 16.9%17.9% for the oil with 6.8% and 10% FFA, respectively. In the effort of trying to reduce neutral oil losses, Kombe (2013) tried to optimize the neutralization of 6.5% FFA castor oil and 4.54% FFA jatropha oil by using the surface response methodology. Temperature, strength of NaOH, and excess NaOH amount to be added for neutralization were the analyzed factors for neutralization process. Table 9.1 summarizes the obtained results. Although, it was not easy to get a comparable oil loss for jatropha and castor oil, but refining losses of 46 times the FFA content have been reported for oils with FFA of 2%6.3% (Mishra et al., 1988). The use of surface response methodology has led to a low loss in neutral oil and allows the model to be used for jatropha and castor oils neutralization, which gives less than 1% FFA for efficient production of biodiesel. In spite of mentioning challenges posed by the chemical modification of FFA, the process is used commercially by many edible oil grade refining industries for effective reduction of FFA to the desired level regardless of the FFA content in the crude oil (Bhosle and Subramanian, 2005). This process is still useful industrially, and calls for further research in optimization; uses of various nonfood chemicals that could lower the loss in neutral oil; and makes the process more attractive for low-cost high-FFA feedstock regardless of the sensory and other edible grade quality parameters that are normally imposed and limit the current neutralization technology.

312

Fatty Acids

9.3.1.2 Esterification The earlier mentioned FFA modification process modifies the FFA by forming soap with alkali materials. This can result in biodiesel yield loss, more alkaline consumption, and a potentially difficult phase separation by the excess soap formation from the process. To overcome the challenges posed by neutralization process, the acid-catalyzed transesterification can be applied to directly convert both FFA and oil into biodiesel. However, it has not been adopted by the biodiesel producers due to the long reaction time, high amount of alcohol requirements, and lower yield (Canacki and Gerpen, 1999). Alternatively, a two-step transesterification process has been revealed to work well in the production of biodiesel from feedstocks with high FFA content (Canakci and Van Gerpen, 1999; Corro et al., 2011; Wang et al., 2007; Zullaikah et al., 2005). The basic idea behind the two-step process is that the high FFAs in the oil are first modified into esters and reduce the FFA to less than 1% by acid catalysis esterification, thereafter; the alkali transesterification could be used to convert the triglycerides, which are the neutral oil into biodiesel. Many scholars have found two-step acid-catalyzed esterification very effective for transesterification of feedstocks possessing high FFA content, with the yield of biodiesel in the overall process exceeding 90% (Berchmans and Hirata, 2008; Canakci and Van Gerpen, 1999; Meher et al., 2006). The modification of FFA into ester using acid-catalyzed esterification involves the reaction of FFAs and alcohol in the presence of an acidic catalyst to give fatty acid alkyl ester (biodiesel) and water. An acidic catalyst can withstand high FFAs and moisture content in the feedstocks (Atadashi et al., 2012; Cardoso et al., 2008; Chongkhong et al., 2009). The esterification reaction is presented in Fig. 9.1. In this equation, R1 represents a linear chain of 1117 carbon atoms with a variable number of unsaturated hydrocarbon depending on the origin of the feedstock, and R2 is a methyl radical (Arora et al., 2016). Several researchers have worked on the modification of high FFA (5%40%) from different feedstock prior alkali-catalyzed transesterification. The reported feedstocks include crude jatropha oil, crude Mahua oil, waste cooking oil, crude rubber seed oil, fryer grease, crude tobacco oil, crude coconut oil, crude rice bran oil, chicken fat, and mixed crude palm oil whereby the initial FFA in these feedstocks was reduced to less than 12 wt % (Alptekin et al., 2011; Arora et al., 2016; Canakci and Van Gerpen, 2001a; Crabbe et al., 2001; Ghadge and Raheman, 2005; Jansri and O

H+ O

OH + H3C

O

OH

R1 FIGURE 9.1 Esterification reaction.

C R2

O

+ CH3

H2O

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Prateepchaikul, 2011; Kumar Tiwari et al., 2007; Marchetti et al., 2007; Nakpong and Wootthikanokkhan, 2010a; Ramadhas et al., 2005; Veljkovi´c et al., 2006). After acid esterification method, the transesterification of top oil phase with less than 2 wt% of FFA gave over 90 wt% of methyl ester in 12 hours using a methanol-to-oil molar ratio of 39:1, with alkaline catalyst (sodium hydroxide or potassium hydroxide) around 12 wt% at 60 C (Jansri and Prateepchaikul, 2011). It has also been reported that, in order to get a complete modification of FFA in the feedstock to esters with less than 1% FFA, then the reaction will take about 60 minutes at 50 C with a concentration H2SO4 to oil ratio 1% w/w (Ghadge and Raheman, 2005; Veljkovi´c et al., 2006). Chai et al. (2014) summarized optimum conditions for the acid esterification of various biodiesel feedstock and pointed out that the industrial and laboratory values of alcohol and catalyst vary with the initial amount of FFA. The variation was significant for the feedstock with less than 15%. Studies have shown that the process of modifying FFA for transesterification using acid esterification reaction is affected by factors such as the reaction time, the amount and type of alcohol, the amount and type of acid catalyst, the moisture content of the feedstock, and reaction temperature. 9.3.1.2.1 The Effect of Amount and Type of Catalyst Many researchers have studied the effect of the amount and type of acid in the acid esterification process for FFA modification. Alptekin et al. (2011) compared three different homogeneous acid catalyst namely sulfuric, hydrochloric, and sulfamic acids in modifying the 15% FFA in chicken fat prior to transesterification. In their study, the type and amount of acid were found to have effect in modifying the FFA in the chicken fat. The amount of acids (3%, 6%, 15%, 20%, and 35%) based on the FFA of the chicken fat was observed for 1 hour at the temperature of 60 C. The 6% concentration of acidic catalyst was not shown to be active in lowering the level of FFA in the feedstock for all of the three catalysts. The effect of sulfuric and hydrochloric acids was alike in the esterification. Sulfamic acid did not show any significant effect on the reduction of the FFA in the feedstock. The 15% FFA in the chicken fat was modified to below 1% when using 20% sulfuric acid and methanol in a molar ratio of 40:1 for 60, 70, and 80 minutes at 60 C. Canakci and Van Gerpen (2001b) observed that an increase in the amount of catalyst to be effective in lowering the acid value of the oil. Using a 10:1 molar ratio and 30 minutes of reaction time, the acid value of the feedstock was dropped from 41.33 to 1.37 mg KOH g21 with 15% H2SO4 catalyst. Nakpong and Wootthikanokkhan (2010b) studied the effect of catalyst concentration and reaction time in modifying the 12.8% FFA in coconut oil using acid esterification. The reaction was performed with different catalyst

314

Fatty Acids

concentrations (0.5%, 0.6%, 0.7%, and 0.8%, v/v, of oil) and reaction times (30, 60, 90, and 120 minutes) under constant methanol-to-oil ratio of 0.35 v/v and reaction temperature of 60 C, respectively. Their results indicate that ester formation rate increased with increasing catalyst concentration. They also noted that at a lower catalyst concentration of 0.5% (v/v) of oil, the FFA was not reduced to below 2% even after 120 minutes. The acid concentration of 0.7% (v/v) of oil was found to be optimum in FFA modification within a short time. The amount of acid catalyst used has also been found to have effects on the conversion of 12% FFA in rice bran oil. Using H2SO4 as a catalyst in the range of 0.151.0 wt%, the FFA modification increases with increased catalyst quantity up to 0.5% of the catalyst. Below 0.5% concentration, the final acid value of oil remains above 2% FFA. The optimum amount of H2SO4 catalyst was 0.5% (v/v) of oil (Arora et al., 2016). Similar results have been observed in varying the amount of H2SO4 in the range of 0.25%2% when modifying 17% FFA in crude rubber oil, the catalyst reaches maximum conversion efficiency at 0.5% (Ramadhas et al., 2005). Zhang et al. (2010) used solid ferric sulfate (catalyst) and noted that when methanol-to-FFA molar ratio was 40.91:1 and catalyst amount was 7.31%, the FFA of Zanthoxylum bungeanum seed oil dropped to 0. 73% from 8.05% after 3 hours of reaction. On varying catalyst in the range of 2.44%12.19%, the FFA of Z. bungeanum seed oil could not go below 1% FFA during 1.5 hours of reaction. Little effect was observed when the catalyst amount exceeded 7.31%. Similar results have been reported by Wang et al. (2007) when using ferric sulfate catalyst in modifying 38.15% FFA in waste cooking oil. The modification of FFA was slow without a catalyst, but when 1 wt% of ferric sulfate was added, 94.4% of FFA was converted into FAME in 3 hours. However, an addition of catalyst above 2 wt% shows little effect on the rate of reaction.

9.3.1.2.2

The Effect of Reaction Time

The effect of reaction time on FFA modification prior to transesterification was studied by Wang et al. (2007); ferric sulfate was used to catalyze the esterification in the presence of methanol. The modification of FFA was shown to be divided into three phases: in the first phase, over 85% of FFA was converted into fatty acid methyl ester within 30 minutes; in the second phase, which was between 30 and 120 minutes of reaction time, the reaction rate was slow with the conversion of FFA of over 95%. The reaction reached to equilibrium after 120 minutes, which is the third phase. Further extending reaction time did not increase the conversion of FFA significantly beyond this phase. Alptekin et al. (2011) investigated the effect of reaction time on the modification of 15% FFA in the chicken fat. The selected reaction time was 60, 70, and 80 minutes at 60 C with 20% sulfuric acid and

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methanol-to-oil molar ratio of 40:1. The reduction of FFA from 15% to 0.93%, 0.80%, and 0.67% was reported in 60, 70, and 80 minutes, respectively. Jain et al. (2011) studied the modification of 21.84% FFA in waste cooking oil by using 1 wt% of H2SO4 at 65 C with methanol-to-oil ratio of 3:7 (v/v). The modification of FFA to fatty acid methyl ester was faster within the first 100 minutes of reaction and thereafter the rate becomes constant. Chai et al. (2014) show that the reaction time of 120 minutes was enough to lower the 15%25% FFA in oil to less than 0.5% and that prolonging reaction time was not necessary as it will just increase the reaction cost. 9.3.1.2.3 The Effect of Temperature FFA modification reaction has also been shown to be sensitive to reaction temperature. Poor FFA modifications have been reported when running the process at low temperature (Ramadhas et al., 2005). When running the reaction at room temperature, the modification of FFA to fatty acid methyl ester was low, even after 2 hours of reaction with continuous stirring. Only 10% of the 17% FFA in rubber seed oil could be modified to fatty acid methyl ester. Elevated temperature above room temperature has been shown to increase the reaction rates but higher reaction temperatures above the boiling point of methanol increase the chance of loss of methanol and increase in dark color of the product. Ramadhas et al. (2005) proposed an optimum temperature range of 4558 C for high conversion. The reaction temperature has also been shown to influence the reaction by Arora et al. (2016). The maximum FFA conversion was reported at 60 C. The use of higher temperature above 60 C increased the reaction rate further, but the elevated rate cannot be compensated by the rate of loss in methanol. In general, numerous authors proposed the temperature range of 5560 C as the best for modification of FFA (Ghadge and Raheman, 2005; Kumar et al., 2007; Lin et al., 2009; Thiruvengadaravi et al., 2009; Veljkovi´c et al., 2006). 9.3.1.2.4

The Effect of Oil-to-Methanol Molar Ratio

The esterification reaction for modification of FFA into esters is reversible, therefore the amount of alcohol required affects the esterification efficiency as well as the cost of biodiesel. In order to obtain a good equilibrium conversion, the backwards reaction can be reduced by the use of excess methanol. Using H2SO4 catalyst in modifying the 34.6% FFA in waste cooking oil, the modification efficiency has shown to increase from 80.43% to 94.54% when the methanol to waste cooking oil ratio changes from 10% to 20% (v/v). Further increase in methanol-to-oil ratio from 20% to 30% (v/v) shows only a slightly change 1.11% in efficiency (Ding et al., 2012). Similarly, Arora et al. (2016) observed an increase in conversion when oil-to-methanol ratio raised from 1:5 to 1:30. There was no significant increase in conversion was

316

Fatty Acids

observed when the molar ratio increased beyond 1:20. Kombe (2013) reported that high modification efficiency can be obtained within a short reaction time at higher methanol-to-oil ratio. Furthermore, the use of a low amount of methanol (less than 10 wt%) did not produce good modification efficiency, even after long reaction time. The reason being a dissolution effect of the methanol over the reaction mixture that takes place with a stronger effect than that provided by the kinetics, and this gives a smaller reaction rate (Marchetti and Errazu, 2008). Even though an excess methanol speeds up the reaction, it also has a consequence on then the operating cost and the size of the reactor (Khan et al., 2010).

9.3.1.3 Reesterification/Glycerolysis The chemical reesterification/glycerolysis is another FFA modification process, which has been in existence for more than centuries for the production of monoglycerides (MG) and diglycerides (DG) (Bhosle and Subramanian, 2005). The process can be dated back to 1924 when Gun proposed production of monoglyceride by using fat glycerolysis as the first step in the formation of a “synthetic butter” (Sonntag, 1982). It produces MG and DG, which have variety uses from surfactants to emulsifiers in foods, paints, cosmetics, and pharmaceutical products. In glycerolysis, oil or fat that is a source of fatty acid is allowed to react with glycerol at high temperatures of about 180 C with or without catalyst (Kumoro, 2012). Unlike esterification, glycerolysis modifies the fatty acid into neutral acyl glycerol by reesterifying them with free hydroxyl groups in the parent oil (or with added hydroxyl groups from glycerol) (Anderson, 1962). The glycerolysis reaction starts with the formation of MG, and the MG are further esterified to DG and finally to a triglyceride (Blanco et al., 2004). The reaction is capable of utilizing fatty acid and FFA in the oil and reesterifying them to MG, DG, and triglycerides, which are neutral oil. In biodiesel production, the same reaction can be used to lower the FFA by reesterifying them with glycerol and produced a neutralized oil with less than 1% FFA, which qualifies it for alkali transesterification. Unlike acid esterification process, glycerolysis requires no alcohol, and the water formed in the reaction can be instantly vaporized and vented from the reaction mixture with the aid of vacuum and nitrogen purging (Noureddini et al., 2004). The water produced in the reaction should be removed to avoid the establishment of equilibrium between the reactants under the experimental conditions. The use of an inert gas or air and maintain vacuum have been recommended methods to eliminate water from the reaction mixture (Bhosle and Subramanian, 2005). The glycerolysis has the potential to use the purified glycerol from the transesterification and thereby lower the cost of biodiesel (Kombe, 2015).

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The glycerolysis of FFA results into three primary reactions as shown in Fig. 9.2. The reaction of FFA and glycerol to form MG and water is initially the primary reaction during glycerolysis and is responsible for the majority of FFA reduction. The glycerol then reacts with triglyceride in the second reaction, whereas 1 mol of triglyceride reacts with 1 mol of free glycerol to form 1 mol each of monoglyceride and diglyceride. The final reaction is that of FFA with MG to form DG and water; this reaction may become dominant as monoglyceride concentrations increase and free glycerol concentration R C OH

R

O

H 2C

O C

H 2O (1)

OH HC

OH

H 2C

OH

Catalyst

H 2C OH Glycerol

FFA

+

CH2

CH

+

OH

O

Monoglyceride

Water

R C O

O R

H2C +

CH

C

C O

O

CH

H2C

+

CH2

OH

CH R

H2C

HC

OH

O C

Glycerol

OH

Diglyceride

R C

O

O

C

O OH +

OH

Monoglyceride

R

C

OH

H2C

O

O CH2

+

H2C Catalyst

HC

OH

H 2C

OH

H 2O (3)

CH R H2C

FFA

(2)

H2C

O

R Triglyceride

R

O

O

Catalyst

CH2 O

C

OH CH2

R

R

O

Monoglyceride

FIGURE 9.2 Glycerolysis reaction.

OH Diglyceride

Water

318

Fatty Acids

diminishes with time (Anderson, 2014). The glycerolysis reaction has been reported to be affected by the reaction temperature, amount, type of catalyst, and the amount of glycerol. 9.3.1.3.1 The Effect of Temperature The glycerolysis process can occur at different temperatures, depending on the type of oil used. Felizardo et al. (2011) used temperatures of 180, 220, and 230 C in glycerolysis of acidulated soap stock with 50% FFA. The temperature increase was found to favor the reaction kinetics at 230 C. Conversely, high FFA conversion into glyceride with the significant difference in FFA drop occurred when the temperature increases from 180 to 220 C. The FFA content of the acidulated soap stocks was reduced from 50% to 5% after 3 hours of reaction at 200 C. Similarly, the effect of temperature has also been reported by various researchers (Bhattacharyya and Bhattacharyya, 1987; Bhosle and Subramanian, 2005; Singh and Singh, 2009) in glycerolysis of high FFA rice bran oil. The maximum FFA conversion was observed between 180 and 200 C. The reaction temperature of 210 C was found to be more effective than below 200 C in glycerolysis of rice bran oil FFA (9.5%35.0%) using MG. Ebewele et al. (2010) reported that the glycerolysis of rubber seed oil containing 37.69% FFA, at a low temperature of 150 C, and the FFA were reduced to 7.03% in 6 hours of reaction time. When the temperature is raised to 200 C, the FFAs were reduced to 1.5% in the same 6 hours of reaction time. Upon further increasing the temperature to 250 C, the rate of modification of FFA increases within the first 2 hours. The application of 250 C temperature was not shown to be optimum. The final FFA was only 3.88% after 6 hours of reaction. The optimum temperature for maximum FFA modification was found to be in between 200 and 250 C. Anderson (2014) compared glycerolysis at various operational temperatures using batch-wise glycerolysis reactions on brown grease with 50% FFA at 177 and 238 C. At 238 C, the FFA concentration dropped rapidly to less than 1% within 1 hour. The temperature of 177 C lowers the FFA to 2% after 9 hours of reaction. It was further observed that the glycerolysis at 238 C reaches an equilibrium state within 12 hours, while the reaction at 177 C continues well beyond 5 hours. Kombe et al. (2013) and Kombe (2015) reported the novel lowtemperature glycerolysis for 6.50% FFA in crude castor oil and 4.54% FFA in crude jatropha oil. In the glycerolysis process, they used NaOH as a homogeneous base catalyst, which improves the miscibility between glycerol and oil and allows the reactions to be carried at temperature below 100 C. In both studies, the temperature range was 3090 C. In the glycerolysis of jatropha oil, it was found that, at 90 C, the high glycerolysis efficiency can be obtained within short reaction time. This was due to the variation of

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temperature with mass transfer in the phase containing triglycerides to the glycerin, this increases the solubility of both phases (Felizardo et al., 2011). 9.3.1.3.2

The Effect of Amount and Type of Catalyst

The glycerolysis of FFA is affected by the type and amount of catalyst used, even though the reaction can progress without catalyst (Bhosle and Subramanian, 2005). Feuge et al. (1945) tried various types of catalyst in glycerolysis of mixed fatty acids with 90.3% FFA in peanut oil obtained by saponification under reduced pressure (20 mmHg) and at 200 C. They tried catalyst like AICI2  6H2O, Al2O3, SnO2, SbCl3, HgCl2, FeO, NiCl2  6H2O, NaOH, MgCl2  6H2O, MgO, MnCl2  4H2O, PbCl2, ZnO, FeCl3  6H2O, CdCl2  2.5H2O, PbO, MnO2, ZnCl2, SnCl2  2H2O, SnCl4  5H2O, and HCl. SnCl2  2H2O, SnCl4  5H2O, and ZnCl2 and all were excellent in catalytic activity. The FFA in the oil reduced from 90.3% to 2.8%, 2.4%, and 3.5%, respectively, in 6 hours. The FFA drops to 5.34% after 8 hours and at an elevated temperature of 241 C without using a catalyst. Similarly, Jansri (2015) tried Zn, ZnCl2, ZnO and ZnSO4  7H2O, SnCl4  5H2O, and SnCl2  2H2O as catalysts in the glycerolysis of vegetable oil with 20 wt% FFA at 150 C under atmospheric conditions. In this work, ZnO was the most suitable catalyst for effective reduction of FFA to 1.416% within 3 hours of reaction time. Ebewele et al. (2010) noted slow reaction kinetics in the absence of a catalyst. The 37.69% FFA in rubber seed oil was only reduced to 15.38% in 6 hours. Yet, on using 0.25% (w/w) zinc dust and 0.15% (w/w) of ZnCl2, the significant reduction in FFA was achieved. Zinc dust dropped the FFA in the rubber seed oil from 37.69% to 1.50% while ZnCl2 to about 1.27% within 6 hours of reaction time. The trial on the combination of the two catalysts did not show any significant reduction in FFA. Felizardo et al. (2011) use metallic zinc and dehydrated zinc acetate as glycerolysis catalysts for 50% FFA of acidulated soap stock. The amount of catalysts used was 0.1%, 0.2%, and 0.3% (w/w). There was no significant effect on the reaction kinetics for both catalysts. By increasing the amount of catalyst lead to increase in the reaction kinetic up to 1 hour. After 1 hour, the final acidity was not seemed to be affected. The same drop of FFA in the oil was observed after 2 hours of reaction time without using a catalyst. Singh and Singh (2009) studied the effect of the amount of catalyst on glycerolysis of rice bran oil with 50% and 70% excess glycerol and for 7 hours of reaction time. SnCl2 catalyst concentrations of 0.1%, 0.15%, 0.2%, 0.25%, and 0.3% (w/w) were used. The 0.2% (w/w) catalyst was established as optimum in reducing the FFA in the rice bran oil for 50% and 70% excess of glycerol. Bhattacharyya and Bhattacharyya (1987) tried SnCl2 and an aromatic sulfonic acid (p-toluene sulfonic acid) as a catalyst to examine the extent of glycerolysis of FFA in rice bran oil. Both catalysts were shown to influence the glycerolysis rate during the first 2 hours of reaction.

320

Fatty Acids

The p-toluene sulfonic acid was more effective in reducing the rice bran oil with 15%30% FFA to low levels of 1.6%4.0%. Wang et al. (2012) tried the super acid solid catalyst SO422/ZrO2Al2O3 in the glycerolysis before homogeneous base transesterification. The 46.5% FFA in the waste cooking oil was reduced to 0.7% FFA. The glycerolysis efficiency was found to be 98.4%. The catalyst showed good activity in the glycerolysis of waste cooking oil by glycerol. Their work also shows the advantages of easy separation of excess glycerol and less catalyst loading (0.3%, w/w). 9.3.1.3.3

The Effect of the Amount of Glycerol

The effects of amount glycerol on the glycerolysis reaction of oil with 37.69% FFA were studied by Felizardo et al. (2011). The experiments were performed at 220 C with a glycerol excess of 4%, 11%, and 52%. The use of more than 10% (molar ratio glycerol/FFA 5 1.10) excess glycerol did not show significant improvements in reaction kinetics at a temperature of 220 C. The stoichiometric amount of glycerol (4.3%, w/w, of oil) resulted in significant FFA reduction as compared with when no glycerol was used in the reaction. Similarly, Ebewele et al. (2010) used 30% excess of glycerol under similar reaction conditions, observed insignificant progress in FFA reduction. The 30% excess of glycerol affects the FFA reduction rate during the first 2 hours of reaction and thereafter the rate decreases significantly. Bhosle and Subramanian (2005) observed the reduction in FFA from 37.69% to 1.5% after 6 hours of reaction with 4.3% (w/w) glycerol as the stoichiometric amounts at 200 C. However, when no glycerol was used, the FFA dropped from 37.69% to about 15% under the same reaction conditions. Thus, the reduction in FFA was linked to the reaction between FFA and the free hydroxyl groups remaining in the oil. Bhattacharyya and Bhattacharyya (1987) reported the effect of the amount glycerol on the extent of glycerolysis of crude rice bran oil with 15.3% FFA. Excess required amount of glycerol 10%, 30%, and 50% was tried. After 6 hours of reaction, the FFA was lowered to 6%, 5.6%, and 4.8% by using 10%, 20%, and 50% excess glycerol, respectively. The high amount of glycerol was shown to increase the rate of reaction. Singh and Singh (2009) used 50%, 70%, and 100% as the excess amount of theoretical glycerol required in glycerolysis of rice bran oil with an acid value of 24.3 mg KOH g21. When using 50% excess glycerol, the drop in acid value was about 19.3% at 200 C for 6 hours. On increasing the excess glycerol up to 70%, the glycerolysis rate was faster with a drop in the acid value of 20.2% at 200 C within 4 hours. When using 100% excess glycerol, the effect was almost similar to that of 70% excess glycerol. Yet, the impact of using a high amount of glycerol was not promising since the maximum drop in acid value was only 20% after 5 hours.

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9.3.1.3.4

321

Glycerolysis for Biodiesel Production

The glycerolysis process has proven to be capable of modifying the high FFA feedstock to less than 1% FFA, which can be transesterified easily as summarized in Table 9.2. Sousa et al. (2010) tried to use the glycerol (from transesterification) for the glycerolysis of 2.36% FFA in castor oil without catalyst for 2 hours at 120 C. Their results proved that glycerol from transesterification can be used to lower the FFA to 0.22%. This is one of the promising results, which offers the market for glycerol and hence lowers the cost of producing biodiesel especially from high FFA feedstocks that are normally available at a cheap price. Unlike other FFA modification process like acid esterification, glycerolysis requires no acid or methanol and the water formed can be easily vacuumed out. The acid esterification produces water and wets the acidic methanol needs to be purified and recovered. Furthermore, using acid esterification for feedstock with more than 40% FFA may require multiple steps to lower the FFA to accepted level; this will end up generating even more acidic, wet methanol. The neutralization of the produced acidic methanol will also need drying using multistage distillation with significant reflux rates and cause high energy use. Kombe (2015) and Kombe et al. (2013) tried the homogeneous base glycerolysis which can be done at a low temperature of less than 90 C. Most of the existing literature as summarized in Table 9.1 has been on the utilization of glycerolysis for producing edible grade products (MG and DG) whereby sensory properties and color are of importance. Such applications of glycerolysis hinder further investigation on different catalysts, which are not good for edible grade product but can work well for biodiesel production. Unless further research is conducted in understanding the kinetics, application of different catalysts, and optimizing process, the glycerolysis process is still regarded as an expensive process for modification of FFA due to high heat involved and requires a high-pressure boiler and the application of vacuum while heating to eliminate water produced in the reaction.

9.4 CONCLUSION AND RECOMMENDATIONS Despite challenges, biodiesel feedstocks from oil and fats with high FFA have shown to be a potential feedstock for low-cost biodiesel production. The effect of high free fatty acid in the base transesterification process can be eliminated by modifying them into soap stock by neutralization process, converting them into esters by acid esterification process, or reesterifying them into glycerides by glycerolysis process. The three presented chemical modification technologies have shown to lowers the free fatty acids less than 3% and eliminate the challenges posed by high free fatty acid in the transesterification process. Further research for the use of various nonfood-based

TABLE 9.2 Summary of the Effect of Glycerolysis on the Final Amount of FFA Oil Type

Time (Hours)

Temperature ( C)

Catalyst

Amount of Excess Glycerol

Initial FFA (%)

Final FFA (%)

Sources

Mixture of oil with palmitic acid

3

150

ZnO

50%

20%

1.9

Jansri (2015)

Jatropha oil

1.2

65

NaOH

2.24 g/g glycerol to oil

4.54

0.07

Kombe et al. (2013)

Castro oil

1.4

56

NaOH

2.34 g/g glycerol/ oil

6.50

0.06

Kombe (2015)

Rice bran oil

6

200

p-Toluene sulfonic acid

50%

15.3

1.6

Bhattacharyya and Bhattacharyya (1987)

Waste cooking oil

4

200

SO22 4 /ZrO2Al2O3

Mole ratio of glycerol to FFA (1.4:1)

43.4

0.66

Wang et al. (2012)

Rice bran oil

6

200

p-Toluene sulfonic acid

50%

20.5

3.1

Singh and Singh (2009)

Rice bran oil

4

200

SnCl2

70%

24.3

3.0

Rice bran oil

6

200

SnCl2

0%

64.7

0.9

Bhosle and Subramanian (2005)

Rubber seed oil

6

200

ZnCl2

4.3%

37.69

1.5

Ebewele et al. (2010)

Mixed fatty acids

4

200

SnC14. 5H2O

Stoichiometric amount

90.3

1.8

Feuge et al. (1945)

Waste cooking oil

4

200

SO22 4 /ZrO2Al2O3

70%

44.42

0.707

Wang et al. (2012)

Castor oil

2

120

No catalyst

100%

2.36%

0.22%

Sousa et al. (2010)

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chemicals and catalysts in chemical modification of high free fatty acid feedstock, economic evaluation of each process, chemical kinetics, and process optimization have been recommended.

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FURTHER READING Thiam, M., unkown. Characterisation of a biodiesel from an alkali transesterification of jatropha curcas oil. Polytechnic Institute, Department of Chemical Engineering.

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Chapter 10

Synthesis of Sugar Fatty Acid Esters and Their Industrial Utilizations Bianca Pe´rez, Sampson Anankanbil and Zheng Guo Aarhus University, Aarhus, Denmark

Chapter Outline 10.1 Introduction 329 10.2 Synthesis of Sugar Fatty Acid Esters 331 10.2.1 Chemical Synthesis of Sugar Fatty Acid Esters 331 10.2.2 Enzymatic Synthesis of Sugar Fatty Acid Esters 333 10.3 Physicochemical Properties of Sugar Fatty Acid Esters 343 10.3.1 Emulsifying Stability and Foaming Ability 344

10.3.2 Toxicity and Biodegradability 10.4 Industrial Applications of Sugar Fatty Acid Esters 10.5 Conclusion Acknowledgment Abbreviations References Further Reading

345 346 347 348 348 348 354

10.1 INTRODUCTION Sugar fatty acid esters (SFAEs) are nonionic surfactants, which contain one or more saccharide rings, for example, sucrose, linked to one or multiple hydrophobic fatty acid chains (Scheme 10.1). The most common fatty acids observed in sugar esters are lauric (C12:0), myristic (C14:0), palmitic (C16:0), stearic (18:0), oleic (C18:1), behenic (C22:0), and erucic acid (C22:1). SFAE can be synthetically tailored for a specific application, and presents a variety of hydrophilic-lipophilic balance (HLB) values ranging from 1 to 16. The HLB values determine their physicochemical properties for specific uses. Sugar esters with low HLB values (HLB 5 36) are good water-in-oil emulsifier, with medium HLB values (79) are good wetting agent, and with high HLB values (1016) are appropriate emulsifier for oil-in-water emulsion. For instance, sugar esters with high HLB values yield low viscosity emulsions suitable for Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00010-6 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

329

330

Fatty Acids

thin skin lotions. Moreover, their tasteless, odorless, nontoxic, and biodegradable features make them excellent biocompatible food emulsifiers (Ducret et al., 1995). In addition, since they are not irritating to the skin or eyes, SFAEs are extensively used in skin-care products to generate deodorant and eyelash, among other cosmetics. For example, sucrose esters of coconut oil-derived fatty acids are widely used as emulsifiers in skin moisturizers. Furthermore, the antimicrobial and antitumor properties of SFAEs have demonstrated their relevance for the pharmaceutical industry (Ferrer et al., 2005a,b).

SCHEME 10.1 The structure of sucrose monolaurate. The hydrophilic sucrose moiety is printed in red (gray in print versions) and the hydrophobic lauric acyl moiety is in blue (black in print versions).

The synthesis of SFAE can be carried out by chemical or enzymatic methods. The latter involves one reaction step and is environmentally friendly. On the other hand, the chemical synthesis generally involves severe reaction conditions, which lead to high energy consumption, use of hazardous chemicals, and generation of undesirable products. Chemical synthesis of sugar esters is mainly carried out at high temperature in the presence of alkaline catalysts. Van Der Plank and Rozendaal (1991) patented a chemical process to obtain sucrose polyesters, which involved mixing the polyol with an alkaline catalyst such as KOH, NaOH, or their carbonates at temperatures above 100 C in aqueous solution, ketones, or C1-5 alcohols. However, chemical methods are generally not selective and are not preferred in the food industry due to possible traces of organic solvents. Enzymatic methods are generally more selective and have a preference to react first with the less sterically impeded fatty acyl chain. Lipases are commonly used enzymes in the synthesis of SFAEs due to their ability to catalyze esterification and transesterification reactions among others (Davis and Boyer, 2001). A disadvantage of enzymatic techniques is that enzymes mainly work at moderate temperature, which could be a problem to create a reaction system that solubilizes both hydrophilic sugar and hydrophobic fatty acid. Nonetheless, some new processes have proved to be effective to target this issue; for instance, the use of ionic liquids (ILs) in combination with enzyme technology or using supercritical CO2 to enable enzymatic processes that do not take place at mild conditions (Habulin et al., 2008). This chapter reviews the classic approach and latest progress in chemical and enzymatic synthesis of sugar esters, their physicochemical properties,

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and their relevance in cosmetic, pharmaceutical, and food industries. Most recent reports in the area and novel technique available to ensure the solubility of both sugar and fatty acids and enable selective enzymatic reaction are also highlighted. Furthermore, the toxicity and biodegradability of SFAE and structural activity relationships relevance for the design of future generation of sugar esters are discussed.

10.2 SYNTHESIS OF SUGAR FATTY ACID ESTERS SFAEs can be synthesized by chemical or enzymatic methods. Most recent studies are focusing on developing enzyme technology for the synthesis of SFAEs, which is a more preferable approach for the products used in food, cosmetic, and pharmaceutical industries. On the other hand, chemical synthesis of SFAEs requires higher energy consumption and may lead to the generation of side-products, which might be toxic, allergenic, or even carcinogenic (Yan, 2001; Puterka et al., 2003). The main technical concern about chemical synthesis is its low selectivity, which leads to complex mixtures of mono-, di-, and triesters of sugars. The use of protection and deprotection techniques is quite attractive in chemical synthesis of SFAEs but these are not viable on industrial scale. Nevertheless, chemical synthesis has been the source of most SFAE products in the market, at least until recently (Yan, 2001).

10.2.1 Chemical Synthesis of Sugar Fatty Acid Esters Sucrose is one of the most used sugars in SFAE. Sucrose is a disaccharide that has nine chiral carbon centers including three primary hydroxyl groups at positions 6, 10 , and 60 and five secondary hydroxyl groups at carbons 2, 3, 4, 30 , and 4 (Scheme 10.2) (Plat and Linhardt, 2001). Therefore, regioselective acylation of sucrose is very important to yield highly pure single product. However, this is difficult due to similar reactivity of the hydroxyl groups in the molecule and the intramolecular acyl migration in unprotected derivatives. Furthermore, the high temperatures and alkaline catalysts required for esterification of sugars generally cause discoloration, polymerization, cyclization, dehydration of products, and thereby lowering yield. There are a number of examples in the literature that describe chemical synthesis of SFAEs (Chauvin and Plusquellec, 1991; Baczko et al., 1995; Chauvin et al., 1993; Vlahov et al., 1997).

SCHEME 10.2 Molecular structure of sucrose.

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Chauvin and Plusquellec (1991) reported a method for the regioselective modifications of sucrose using 3-acyl-5-methyl-1,3,4-thiadiazole-2(3H)thiones (Scheme 10.3) in N,N-dimethylformamide (DMF) and 1,4-diazobicyclo-[2.2.2] octane at low temperatures, which yield predominantly 60 -acyl sucrose (Scheme 10.3) with an 35%43% isolation yield. They also reported the modification of sucrose to obtain 2-O-acylsucroses 3a and 2-0-(N-alkyl carbamoyl)sucrose 3b (Scheme 10.4) in high yields using 3-acyl-thiazoledine-2-thione 4 in the presence of sodium hydride (Chauvin et al., 1993). They determined that the ratio of sodium hydride (NaH) to substrate has to be kept as low as possible to avoid the reaction of multiple hydroxyl groups and that the acylating reagent should not form electrophilic acylium intermediate as this also influences the regioselectivity of the reaction. In terms of selectivity, 3-acylthiazolidine-2-thiones (Scheme 10.4) seems to be better acylating reagent than 3-acyl-bmethyl-1,3,4-thiadiazole-2(3H)-thiones (Scheme 10.4), which can generate electrophilic ion-pairs in the presence of an organic base.

SCHEME 10.3 Chemical structure of 3-acyl-5-methyl-1,3,4-thiadiazole-2(3H)-thiones and the 60 -acyl sucrose.

Other examples, as reported by Baczko et al. (1995) for the synthesis of 6O-acylsucroses, involve the initial generation of 2-O-acylsucroses using sodium hydride and the appropriate 3-acylthiazolidine-2-thiones or 3-acyl-5-methyl1,3,4-thiadiazole-2(3H)-thiones, followed by the intramolecular isomerization in the presence of 1,8-diazabicyclo [5.4.0] undec-7-ene (DBU) or an aqueous solution of trimethylamine (Scheme 10.5). Their results show that 2-O-acyl migrates to yield 3-O-acylsucrose and then the latter isomerized to generate 6O-acylsucrose. Otherwise, 6-O-acylsucroses were directly obtained when the unprotected sucrose was acylated in the presence of DBU (Scheme 10.5).

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SCHEME 10.4 Molecular structure of 2-O-acyl sucrose 3-acylthiazolidine-2-thiones 4,3-acylbmethyl-1,3,4-thiadiazole-2(3H)-thiones 5.

A different approach for the regioselective synthesis of sucrose monoesters consists of first converting sucrose into a dibutylstannylene acetal using di-n-butyltin oxide in methanol (Vlahov et al., 1997) (Scheme 10.6). Posteriorly, the respective acetal is made to react with the anhydride of the fatty acid in DMF at room temperature, which yields a single product after 48 hours. It could be inferred that the sucrose forms a six-member stannylene acetal and it later reacts with the anhydrous species to yield the desired 6-Oacylsucroses.

10.2.2 Enzymatic Synthesis of Sugar Fatty Acid Esters Enzymatic synthesis of SFAEs is a more environmentally friendly and food compatible approach. Hydrolases (lipases and proteases) are the most common biocatalysts for the synthesis of SFAEs in organic solvent systems (Theil, 1995; Gotor, 1999). Lipases are enzymes that can catalyze the

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SCHEME 10.5 One step synthetic pathway to obtain 6-O-acylsucrose.

esterification of sugar (Scheme 10.7) (Wei et al., 2015). They can also catalyze backwards reaction, such as hydrolysis of sugar esters. Lipases possess unique properties, for instance, they are regiospecific and stereoselective (HKittikun et al., 2012). Moreover, lipases can catalyze heterogeneous reactions at the interface of water soluble and water insoluble systems, in organic solvents, and even at high temperatures. The most used lipases are extracellular lipases generated by microorganisms, for example, Rhizopus delemar (Ac¸ıkel et al., 2010), Aspergillus terreus (Gulati et al., 1999), Streptomyces cinnamomeus (Sommer et al., 1997), Bacillus stearothermophilus MC7 (Kambourova et al., 2003), Acinetobacter sp. RAG-1 (Snellman et al., 2002), Microbacterium sp. 7-1W (Honda et al., 2002), Bacillus sp. H-257 (Imamura and Kitaura, 2000), Geobacillus sp. TW1 (Li and Zhang, 2005), and Streptomyces rimosus R6-554 W (Abrami´c et al., 1999).

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SCHEME 10.6 Synthesis of 6-O-acylsucroses using n-butyl oxide.

SCHEME 10.7 Lipase-catalyzed acylation of sorbitol using behenic acid.

10.2.2.1 Synthesis of Sugar Fatty Acid Esters in Conventional Solvents Although the regioselective modifications of sugars represents one of the main challenges for the synthesis of sugar esters because of their polyhydroxy nature, a number of commercially available enzymes have been used for the regioselective acylation of sugars (Ferrer et al., 2005b; Khaled et al., 1991). For example, CAL B is reported to be able to mediate the acylation of fructose with oleic acid in the presence of 2-methyl-2-butanol (2M2B) as solvent (Khaled et al., 1991); and a high yield of 80% could be obtained by effluent recycling and drying. Ferrer et al. (2005b) used lipase from

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Thermomyces lanuginosus to catalyze the regioselective acylation of secondary hydroxyl groups in a mixture of tert-amyl alcohol: dimethylsulfoxide (DMSO; 4:1, v/v). In comparison with lipase from Candida antarctica, which was very useful for the preparation of 6,6-di-acylsucrose, T. lanuginosus preferentially catalyzed the synthesis of 6-O-acylsucrose. Other example of lipases catalyzed esterification constituted the work performed by Chaiyaso et al. (2006). They reported yields of glucose palmitate at around 74% using acetone as solvent and C. antarctica lipase B as biocatalyst. In addition, the enzymatic modification of disaccharide is also possible. Riva et al. (1988) used Bacillus subtilis protease to catalyze the acylation of maltose, cellobiose, sucrose, and lactose with trichloroethyl butanoate in anhydrous DMF. The yields of sugar monoesters obtained were approximately 50%. Carrea et al. (1989) using subtilisin (Protease N from from B. subtilis) achieved the transesterification of the 1-O-hexyl derivative of sucrose with activated lauric acid in acetone. In another study, crude subtilisin (Protease N) and Bacillus protease (Bioenzyme-240) were found to be the most effective enzyme used for the synthesis of butyric acid esters of sucrose in anhydrous pyridine (Patil et al., 1991). Similarly, lipase from Byssochlamys fulva NTG9 was used to synthesize sugar esters from sucrose and oleic acid in tert-butyl alcohol, a less toxic alternative to pyridine and DMF (Akoh, 1994). Tert-butyl alcohol partially solubilizes disaccharides and has been applied for the synthesis of a number of sugar esters at high conversion rates. The conversion rates of disaccharides to corresponding sugar esters have been found to increase with increasing temperature. For instance, conversion rate of sucrose is only 1% at 40 C but is 18% at 80 C. Likewise, the conversion rates of maltose were increased by a factor of more than 5.6 under refluxing tert-butyl alcohol compared with at 40 C. Also, fatty acid chain length affects both initial rates and regioselectivity in the esterification of disaccharides. Short-chain acyl donors have higher initial rates (Pedersen et al., 2002b). Although most studies on sugar ester synthesis focus on total ester yields or regioselective esterification of sugar, it is also pertinent to examine the degree of esterification (DE) of sugar. This would enable the determination of exact HLB values, which could ensure a proper application. In the determination of DE of synthetic fructose laurate catalyzed by CAL B, a significant difference was observed between esterification in 2M2B and methylethyl ketone (MEK) (Li et al., 2015a, b). A preferential formation of diesters in MEK was observed compared to in 2M2B. Changes in conformation of CAL B binding, as noted by Fourier transform infrared spectroscopy, were suggested as the reason for the differences in DE observed for synthesis of sugar esters in the different organic solvents. Furthermore, lipases can mediate transesterification as well. Sin et al. (1998) reported the synthesis of fructose esters from fructose and vinyl esters. Accordingly, the yield of the reaction increases with the increase of the chain length of fatty acid in the vinyl esters. Similarly, Zhang et al. (2015) recently reported that the Lipozyme TL IM catalyzed synthesis of various sugar monoesters in a mixture of 2-methyl-butanol and DMSO (8:2, v/v) using vinyl fatty acid esters with chain lengths from C8 to C12 as acyl donors. Though

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good yields were obtained for the different acyl donors, the lipase seemed to prefer long-chain fatty acids. In contrast, previous reports (Carrea et al., 1989; Chahid et al., 1992) demonstrated that the efficacy of synthesis of fructose esters diminishes when the chain length of fatty acids in vinyl esters increases. Apart from the chain length of the acyl donor, the structure of sugars also influences the esterification yields. Ku and Hang (1995) studied the esterification of various sugars with fatty acids in tert-butyl alcohol catalyzed by lipase from B. fulva. Linoleic acid was the acyl donor giving the highest yield (65.5%) among the fatty acids tested. The highest percentage of esterification was achieved with fructose (71.3%) followed by maltose (67%) and glucose (47.8%). No esterification reaction took place with lactose as acyl acceptor (0%); however, sucrose is observed 36.6% of esterification. Sharma and Chattopadhyay (1993) studied the acetylation of fructose, glucose, and arabinose monosaccharides by using lipase from porcine pancreas (PPL). The sugars were preadsorbed on silica gel and treated with vinyl acetate in the presence of PPL and molecular sieves in diisopropyl ether to yield the monoacetylated derivatives. The products obtained for glucose, fructose, and arabinose were 6-O-acetylglucopyranoside, 1-O-acetylfructoside (mixture of β-pyranose and α-furanose), and 5-O-acetylarabinofuranoside and in 62%, 70%, and 68% yields, respectively (Scheme 10.8).

SCHEME 10.8 Porcine pancreatic lipase-catalyzed transesterification of glucose, fructose, and arabinose with vinyl acetate in diisopropyl ether.

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The use of biocatalyst also plays an important role in determining reaction yields and products. Akoh and Mutua (1994) observed that the lipase from C. antarctica (SP-382) was most effective in catalyzing transesterification of methyl α-D-glucopyranoside, methyl β-D-galactopyranoside, and octyl β-D-glucopyranoside with methyl oleate (Akoh and Mutua 1994). Compared with the immobilized Mucor miehei (mol% of fatty acid incorporation 5 18) and the other nonimmobilized lipases (mol% of fatty acid incorporation 5 037), the immobilized Candida lipases SP-382 and 2001 yieled higher incorporation of fatty acids chains (4477 mol%). Moreover, lipase from C. antarctica was able to catalyze the acylation of the rare sugar, allose (a C-3 epimer of glucose) with vinyl esters in acetonitrile to give allose-6-alkanoates (Afach et al., 2005). Other enzymes have also been reported to be capable of catalyzing synthesis of sugar esters via transesterification. For example, Pedersen et al. (2002a) achieved the transesterification of sucrose with vinyl laurate in DMSO using the metalloprotease, thermolysin (Scheme 10.9). This novel activity of thermolysin widens the window of opportunity for the synthesis of sugar esters and related compounds.

SCHEME 10.9 Synthesis of lauroyl-sucrose using thermolysin and DMSO.

Organic solvents may affect enzyme-catalyzed synthesis of SFAE through their effects on the hydration status of enzymes and consequently influence reaction parameters such as reaction rate, catalyst turnover (Kcat), maximum reaction velocity (Vmax), the substrate’s affinity for the enzyme (Km), and the specificity constant (Kcat/Km) (Kumar, 2016). Condensation reactions such as esterification are ideally carried out in a solvent with low water activity but capable of solubilizing fatty acids and sugars. For instance, by changing solvent from hexane to tertiary alcohols, there was a preferential synthesis of monoesters of oleoyl xylitol compared with di- and triesters (Castillo et al., 2003), demonstrating the effect of solvent polarity on catalytic specificity. In addition, Watanabe et al. (2001) observed differences in the equilibrium constants (Kc) for the synthesis of lauroyl mannose and vinylacetyl glucose in some organic solvents, which are resulted from different activity coefficient of substrates and products. The Kc values for the formation of lauroyl mannose were lowest in 2-methyl-2-propanol and 2M2B, intermediate in acetone but highest in acetonitrile. A useful measure of the hydrophobic/hydrophilic property of an organic solvent to influence enzyme catalysis is the log P value, where P is the

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partition coefficient between 1-octanol and water phases. Solvents with high log P value are hydrophobic while those with low log P values are hydrophilic. It is also reported that log P is well correlated to the apparent catalytic activity and specificity of hydrolases in organic solvents (Lu et al., 2008) but some researchers argue that log P does not necessarily correlate very well with enzyme activity in organic solvents (Ghatorae et al., 1994; Narayan and Klibanov, 1993). Dimroth-Reichardt solvent parameter (ET) can also assess the suitability of a solvent for catalysis. ET is an empirical measure of the polarity of solvents, solvent dielectric constant, electron acceptance index, and the Hildebrand solubility parameter (Valivety et al., 1994; Brink and Tramper, 1985; Affleck et al., 1992a,b). For example, Castillo et al. (2003) found that when synthesizing oleoyl xylitols in mixed solvents, the equilibrium conversion to a diester was found to increase with decreasing ET(30). A good solvent for synthesis of SFAE should be able to dissolve appreciable amounts of both sugar and fatty acids. However, the extremely different properties of sugars and fatty acids complicate the selection of a suitable solvent for catalysis. Moreover, the inertness of the solvent, in terms of its effect on the activity and stability of enzymes, is important too. Studies have shown that the choices of solvent impact both the enantioselectivity and specificity of lipase-catalyzed reactions (Liu et al., 2009; Hudson et al., 2005; Rubio et al., 1991; Paula et al., 2005; Klibanov, 1990; Sakurai et al., 1988; Wescott and Klibanov, 1994). Both hydrophilic and hydrophobic solvents have been used in enzymecatalyzed esterification reactions with quite opposite effects on reaction parameters. Generally, the use of water-miscible organic solvents such as acetone, acetonitrile, and tertiary alcohols have the merit of being able to solubilize sugars without the need for further solubilization reagents (Watanabe et al., 2000; Degn and Zimmerman, 2001; Castillo et al., 2003). A drawback of these solvents, however, is their ability to dehydrate the hydration layer surrounding enzymes bringing about enzyme inactivation. On the other hand, the highest enzyme stability and activity have been recorded in hydrophobic solvents (Ryu and Dordick, 1989). Despite the latter hydrophobic solvents may not always be a good choice for lipase-catalyzed esterification since sugars are hydrophilic and require complete dissolution (Degn and Zimmerman, 2001; Chang and Shaw, 2009). The art of blending two or more solvents could be an alternative to modify the polarity and ionization capacity of water-immiscible organic solvent for lipase-catalyzed esterification reactions (Jia et al., 2010). Solvents that can solubilize sugars are pyridine, dimethylpyrolidone, and DMF, nonetheless, these are very toxic solvents and known to inactivate the enzyme (Ganske and Bornscheuer, 2005a,b). The environmental and safety issues associated with organic solvents have limited their use in modern day enzymatic esterification. However,

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there are multiple examples in the literature of the generation of sugar ester using enzymatic reaction in these solvents. For example, Yan et al. (1999) obtained good yields of glucose caprylate using 2-methyl ketone (66%) and acetone (90%) as solvents. In addition, Degn and Zimmerman (2001) improved the yield of glucose myristate from 222 to 1212 mmol g21 h21 by switching from tert-butanol to a mixture of tert-butanol and pyridine (55:45, v/v). The synthetic activity for cellubiose, sucrose, maltose, or lactose was not reported, probably owing to the low solubility of these sugars in the solvent systems (Degn and Zimmerman, 2001). Recently, conversions of 88%96% of various xylitol monoesters of fatty acids were obtained by Adnani et al. (2011) using hexane as solvent and immobilized Novozym 435 lipase as biocatalyst. Also, Jia et al. (2010) obtained good yields of sugar esters (dilauroyl maltose) in a mixture of hexane and acetone. Several other researchers have investigated for the synthesis of sugar esters in organic solvents with excellent results (Neta et al., 2012; Zaidan et al., 2012; Walsh et al., 2009; Yu et al., 2008; Sakaki et al., 2006; Oosterom et al., 1996; Coulon et al., 1998). One strategy to overcome the solubility problems of sugars constitutes using sugar derivatives instead of native sugars but this often requires several protection and deprotection steps because of the modified properties of the sugar derivatives (Chang and Shaw, 2009). Other approach is the suspension of undissolved sugars, which provides a continuous supply of sugars to the reaction for synthesis of SFAEs (Paradkar and Dordick, 1994; Degn and Zimmerman, 2001). In addition, organoboronic acids have also proved to help solubilizing sugars (Ferrier, 1972; Park et al., 1992). Organoboronic acids have the ability to solubilize sugars by forming carbohydrateboronate complexes through reversible condensation reactions with carbohydrates. These complexes are soluble in nonpolar solvents and are hydrolysable by minimal amounts of water, thereby making esterification of sugars possible in organic solvents. Schlotterbeck et al. (1993) studied the esterification of fructose with stearic acid in hexane at 60 C using phenylboronic acid as solubilizing agent to achieve two isomeric monoacylated esters (Scheme 10.10). These isomers, although inseparable on TLC, were confirmed by proton and carbon-13 NMR. It was also possible to regioselectively monoacylate glucose and galactose using the same method. Other example constituted the work carried out by Oguntimein et al. (1993). They reported the monoacylation of fructose and glucose with stearic acid (C-6 or C-1 esters) in tert-butyl alcohol. Yields of up to 10%24% were obtained with immobilized lipozyme TM 20 (Rhizomucor miehei) or SP-382 (Candida sp.) lipase. Organoboronic acids were used as solubilizing agents for sugars and the syntheses were carried out in different organic solvents including benzene, 1,4dioxane, heptane, hexane, pyridine, tert-butylether, and toluene. An increase in extent of esterification was observed with increasing hydrophobicity of the solvents corroborating both recent and earlier studies.

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SCHEME 10.10 Lipozyme-catalyzed esterification of sucrose with stearic acid in hexane using phenylboronic acid as solubilizing agent.

10.2.2.2 Synthesis of Sugar Fatty Acid Esters in Green Solvents An attractive alternative to conventional organic solvents as medium for enzyme catalysis including synthesis of sugar esters is ILs. ILs are nonvolatile, nonflammable, and thermally and chemically stable organic salts. In contrast to conventional solvents, ILs are composed of molecular ions (see Scheme 10.11 examples of cations and anions of ILs). ILs have low melting points (,100 C) and remain in the liquid state even at temperatures as high as 300 C (Lue et al., 2007). Moreover, ILs have high polarity and can dissolve a wide variety of substrates.

SCHEME 10.11 Examples of cations and anions found in ILs.

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The use of ILs can be considered “green” and environmentally benign owing, primarily, to their negligible vapor pressures. ILs have been used for enzyme catalysis with some excellent results (Lue et al., 2007). It is not, therefore, surprising that ILs have become the substitutes for conventional organic solvents over the last decade or so. The premier use of ILs in lipasecatalyzed synthesis of sugar esters was first reported by Madeira et al. (2000) and confirmed by Park and Kazlauskas (2001) in the acylation of glucose and maltose in ILs (Scheme 10.12). Excellent results were obtained in 1-methoxyethyl-3-methylimidazolium tetrafluoroborate [MOEMIm][BF4]. This IL dissolves glucose more than a 100 times better than acetone. Particularly, ILs based on dicyanamide anions are very effective nonprotic solvents that can dissolve large quantities of sugars from glucose to even cellulose. Glucose and sucrose can, for example, be dissolved to extents of 145 and 195 g L21, respectively in 1-butyl-3-methylimidazolium dicyanamide [BMIm][dca] (Liu et al., 2005).

SCHEME 10.12 CAL Bcatalyzed transesterification of glucose with vinyl acetate in 1-methoxyethyl-3-methylimidazolium tetrafluoroborate [MOEMIm][BF4].

ILs have proven to achieve better regioselectivity in synthesis of sugar esters compared with conventional solvents. For instance, the yield of monoesters from the acylation of glucose using CAL B was 53% in tetrahydrofuran, while it was 93% in [MOEMIm][BF4] (Park and Kazlauskas, 2001). In addition, Kim et al. (2003) observed significantly enhanced reactivity and regioselectivity in the acylation of glycosides in [BMIM]1PF62 ([BMIM]1 1-butyl-3-methylimidazolium) and 1 2 [MOEMIM] PF6 ([MOEMIM]1 1-methoxyethyl-3-methylimidazolium), respectively, in comparison to using chloroform or tetrahydrofuran as solvents. The reactions were catalyzed by lipase from Candida rugosa with vinyl acetate as acyl donor and at room temperature. Furthermore, the mixtures of ILs have been observed to have improved productivity compared to pure ILs. Lee et al. (2008) studied the enzymatic synthesis of 6-O-lauroyl-D-glucose in the mixtures of two ILs. They observed a higher lipase activity in the water-miscible IL, [Bmim][TfO], whereas an improved stability of Novozyme 435 lipase was noted in the hydrophobic IL, [Bmim][Tf2N]. A ratio of 1:1 (v/v) of the two ILs was found as the best for optimal activity and stability of Novozym 435 lipase.

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Moreover, improved conversion rates have been obtained for lipasecatalyzed synthesis of sugar esters in mixtures of ILs and organic solvents, which could function as cosolvents as well as solubilizing agents. Abdulmalek et al. (2012), for instance, obtained conversion rates as high as 87% after only 2 hours of reaction during the synthesis of galactose oleate using Lipozyme RM IM, DMSO, and 1-butyl-3-methylimidazolium tetrafluoroborate ([Bmim] [BF4]). The optimal ratio of the solvents was DMSO: ([Bmim][BF4]) 5 1:20 (v/v). Other researchers also have obtained good results in the synthesis of sugar esters using ILs (Ganske and Bornscheuer, 2005a,b; Li et al., 2015b; Findrik et al., 2016). Thus it becomes obvious that ILs have several advantages over conventional solvents and held much promise for the future. Another group of “green solvents” that are becoming popular is supercritical fluids (SCFs). SCFs are compounds, which exist at temperatures and pressures above their corresponding critical values. They have become attractive for biocatalysis due to their low viscosity and surface tension as well as their high diffusivity, which makes them similar in behavior to gases, thereby promoting the solubility of a wide range of solutes. Another special characteristic of SCFs is that the solubility of solutes could be changed by adjusting temperatures and pressures, particularly close to the critical values (Vermue and Tramper, 1995). SCFs, just as ILs, have gained increased application due to their perceived “green” nature and environmental friendliness. Supercritical carbon dioxide is always the favored SCF system due to its nontoxicity, nonflammability, and the fact that it is inexpensive. Lipases are the common enzymes used as catalysts in SCFs, and more so, when the fluid is carbon dioxide (Mesiano et al., 1999). The first reported use of SCFs in ˇ enzyme catalysis was in 1985 (Hammond et al., 1985). Sabeder et al. (2006) noticed an improved conversion rate of palmitic acid during the synthesis of fructose palmitate in supercritical carbon dioxide catalyzed by lipase from C. antarctica. A conversion rate of 67% was obtained in supercritical carbon dioxide compared to 65% in 2M2B. Habulin et al. (2008) obtained high yields of various sugar esters in supercritical carbon dioxide at 10 MPa catalyzed by immobilized lipase from C. antarctica (CAL B). Sucrose laurate was found to be effective as an antimicrobial agent against Bacillus cereus at a concentration of 9.375 mg mL21. The main advantages of using SCFs in sugar ester synthesis are: (1) easy downstream processing of sugar esters and (2) increased reaction rates due to increased mass transfer rates.

10.3 PHYSICOCHEMICAL PROPERTIES OF SUGAR FATTY ACID ESTERS Sugar esters are highly interesting for industry applications due to their surface active properties and the fact that they are generated from renewable resources (Yanke et al., 2004). Although the melting point of sugar is high,

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depending on the DE, the melting point of sugar esters can vary between 40 and 79 C. The melting point is very important to help predicting the thermal behavior of sugar esters for storage or industrial processes. Moreover, the length of the hydrophobic chain and size of the hydrophilic group on sugar esters provide a wide range of HLB values, high HLB value representing water soluble surfactants or a low HLB resulting oil soluble emulsifiers. The HLB values can range between 0 and 16. For instance, if eight hydroxyl groups in sucrose were to be esterified, the product would be highly hydrophobic and soluble in oil. However, partial esterification will generate an amphiphilic sugar ester, which can be used as emulsifiers in the food, cosmetic, and pharmaceutical industries. The longer the fatty acid chain and the higher the DE and the lower is the HLB value. Moreover, depending on the degree of acylation, they can present different properties such as critical micelle concentration (CMC) and foaming ability. Some typical physicochemical properties, toxicity, and biodegradability of SFAEs are presented in the following section.

10.3.1 Emulsifying Stability and Foaming Ability Amphiphilic sugar esters can form thermodynamically stable molecular aggregates named micelles when in an aqueous solution. The surfactant molecular structure and experimental conditions will determine the CMC value, which is the specific concentration at which the micelles will start forming. For instance, increasing alkyl chain decreases CMC value (Becerra et al., 2008; Ko¨nnecker et al., 2011). Becerra et al. (2008) showed that both fluorescence measurements and surface tension measurements displayed the same tendency of CMC variation to decrease as the number of methylene units increases. The CMC value is of high relevance as it represents the amount of surfactant required to solubilize hydrophobic compounds in water. Adding more surfactant after the CMC is reached yields more micelles and promotes the growth of aggregates. Moreover, temperature has also an influence on the formation of micelles and surface activity as increasing temperature leads to a lower CMC and a larger micelles size (Cristo´bal Carnero and Jose´, 2008). Sugar esters are capable of reducing the surface tension of water, which is highly valuable for industry applications. For instance, coconut milk, as other emulsions, is not stable and prompt to phase separation. Coconut milk emulsions have larger droplet size and lack of good emulsifiers, which lead to unfavorable contacts between water and oil. Thus using surfactants capable of reducing surface tension will improve the emulsion stability. In fact, Akoh and Nwosu (1992) demonstrated that the greater the ability of a surfactant to reduce surface tension, the greater the emulsion stability formed in coconut milk. Other example constituted the one reported by Neta et al. (2012), who synthesized a series of sugar esters including fructose, sucrose,

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and lactose esters; and demonstrated that lactose esters are the best biosurfactant reducing surface tension from 52.0 to 38.0 N m21, displaying an emulsification index of 54.1%. Decreasing the surface tension is also of relevance for generating foods, which consist of foam because the airwater interfacial area can be enlarged by decreasing the surface tension. For instance, Tual et al. (2006) demonstrated that the percent of sugar ester was relevant when preparing dairy foam using oil-in-water emulsions generated by homogenization of anhydrous milk fat (20 wt%) with an aqueous phase of skim milk powder (6.5 wt %), sucrose (15 wt%), hydrocolloids (2 wt%), and sucrose esters at high pressure. They tested different content of sugar esters, which varied from 0 to 0.35 wt% and found that the most stable and firm foam were formed when using c.0.1 wt% of sugar ester. Furthermore, they showed that that emulsion droplets disrupt in the presence of surfactant at the airwater interface, which suggests that some destabilization of droplets by surfactants at the interface can lead to firmness and stability, and consequently, good foam.

10.3.2 Toxicity and Biodegradability U.S. Food and Drug Administration y172.859 established that sucrose esters are permitted for Good Manufacturing Practice as long as they meet certain specifications. For example, sugar esters must have residue of ignition (sulfate ash) below 2%, their heavy metal content must not be higher than 50 parts per million (ppm) and their use must not exceed the amount required to accomplish the intended purpose. Accordingly, addition of sucrose fatty acid esters as emulsifiers, texturizer, and stabilizer is permitted in different foods including dairy and nondairy products, confectionary, and bakery. Recently, Kurti et al. (2012) studied the toxicity and permeability of sucrose esters on a culture model of the nasal barrier because of the interest to use sugar esters as excipients for nasal drug delivery. They determined that 0.1 mg mL21 laurate or myristate sucrose esters could be safely used on cells for 1 hour. Furthermore, they demonstrated the potential of sucrose esters as permeability enhancers for nasal drug delivery. Others works report the nontoxic concentrations of palmitate (P-1695), myristate (M-1695), and laurate (D-1216) on Caco-2 epithelial cells (Kiss et al., 2014). This group of researchers reported the following safe concentrations on Caco-2 epithelial cells 30, 60, and 100 μg mL21 for P-1695 and M-1695, respectively; being D-1216 the less toxic of the sucrose esters tested. Moreover, P-1695 and M1695 showed to reduce metabolic activity of Caco-2 cells at concentrations higher than the nontoxic doses. The toxicity of sucrose esters on the skin has also been investigated. Le´merya et al. (2015) determined the skin toxicity in vitro using lactate dehydrogenase and 3-(4,5-dimethylthiazol-2-yl)2,5-diphenyltetrazolium bromide tetrazolium dye test of cell viability.

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Accordingly, sucrose esters containing a C12 alkyl chain were the most toxic of the nonionic surfactants evaluated. Interestingly, they found that the CMC values did not result a relevant parameter to account for the skin toxicity of the compounds tested. Sugar esters biodegradability is also a relevant factor as it helps determining if the concentration of sugar esters remains below the detrimental levels to the environment (Baker et al., 2000). For instance, house-hold cleaning products, which contain surface active molecules such as sugar esters, are normally disposed through the drain and, therefore, the biodegradability of sugar esters becomes of interest as the detergent surfactants residues are linked to foaming incident in sewage treatment plants. However, the biodegradability of nonionic surfactants is more difficult to predict because of the wide variety of molecular structure and the lack of a common functional groups. Nonetheless, Sturm (1973) developed a method to predict the rate and degree of biodegradation. By measuring CO2 production, they were able to carry out rapid screening of organic materials without the need of specific analytical techniques and measure rate and degree of biodegradation. Regardless, in general, sugar esters are known for their excellent biodegradability that does not generate environmental pollution.

10.4 INDUSTRIAL APPLICATIONS OF SUGAR FATTY ACID ESTERS Sugar esters are commercially available and have found a wide range of use in the food, cosmetic, and pharmaceutical industries (Chang and Shaw, 2009). In fact, the U.S. Food and Drug Administration has allowed the addition of sucrose esters to certain processed foods (21CFR 172.859). Sugar esters have also demonstrated to possess antitumor (Ferrer et al., 2005a), insecticidal (Puterka et al., 2003), antifungal (Zhao, 2014), and antibacterial properties (Zhao et al., 2015). Their antibacterial properties are believed to be a result of their interaction with cell membrane of bacteria causing autolysis. The antimicrobial activity depends on the sugar, the fatty acid chain linked to the sugar, and the DE. Wagh et al. (2012) demonstrated that lactose monolaurate and sucrose monolaurate were effective against Gram-positive bacteria. Furthermore, lactose monolaurate displayed minimal bactericidal concentrations that range from 5 to 9.5 mM for Listeria monocytogenes isolates and from 0.2 to 2 mM for Mycobacterium isolates. Posteriorly, Chen et al. (2014) found that lactose monolaurate can inhibit the growth of a fivestrain cocktail of L. monocytogenes in yogurt, milk cottage, and cheese. The lactose monolaurate results demonstrate the highly attractive properties of sugar esters to prevent food contamination. Sugar esters properties made these agents highly valuable for the pharmaceutical, food, and cosmetic industries. In the pharmaceutical industry, sugar ester can help in the solubilization of poorly water soluble drugs (Sz˝uts and

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Szabo´-Re´ve´sz, 2012). They can improve drug release profile, consequently enhance drug bioavailability, and reduce side effects. For instance, Kiss et al. (2014) demonstrated the potential of laurate ester as absorption enhancer by testing three different sugar esters with HLB values of 16 (laurate, myristate, and palmitate esters) on Caco-2 cell. In the food industry, sugar esters chemically resemble triglycerides and are stable for food processing. For example, olestra is a triglyceride substitute that contains a sucrose ring with six to eight of its hydroxyl groups esterified by long-chain fatty acid (Jandacek, 2012). Procter & Gamble (Ohio, USA) industrialized olestra as a noncaloric substitute for diary fat as olestra is not absorbed from the small intestine to the blood and tissues and has a fat-like taste. In addition in the food industry, sucrose esters can be used as delivery system such as nanoemulsions and microemulsions, which are composed of oil, surfactant, and water. Sucrose laurate constitutes an example for delivering flavor into food and beverages. Since sucrose esters are nontoxic and compatible with the skin, their low or nonirritable properties made them highly attractive for the cosmetic industry. Other example of the use of sugar ester to develop novel technologies relevant for industry application constituted the one reported by Park et al. (2007). They patented a skin external preparation containing stratum corneum intracellular lipids and sorbitan stearate, sucrose cocoate, or a mixture thereof emulsifying agents. The skin external preparation presents a hexagonal gel structure, which is similar to the lamellar structure formed by the lipid matrix of the stratum corneum. Thus, this invention provides a very useful technology to study damage skin barrier function and develop novel moisturizers for various skin diseases.

10.5 CONCLUSION Sugar esters are nonionic surfactant that can be tailored for a specific purpose. Being nontoxic, biodegradable, and nonirritable to the skin or eyes make these products attractive for food, pharmaceutical, and cosmetic industries. Therefore, it does not come into a surprise that the sucrose esters market is projected to be valued USD 74.6 million by 2020 (Rohan, 2016). All combined highlight the relevance of these compounds for the scientific and industrial community. Nonetheless, many questions remain for scientific research and applied studies regarding the nature of these compounds. Their wide range of HLB value leaves room for the design and synthesis of novel sugar esters for food, pharmaceutical, and cosmetic applications. For instance, the design of sugar esters that resemble triglycerides and act as noncalorie fats makes sugar esters of interest for the food industry. In addition, a paradigm shift has emerged, where the interest lies in the use of methods that are more environmentally benign (Chang and Shaw, 2009). The use of natural biocatalysts, otherwise called enzymes, provides an alternative to chemical synthesis. Enzymatic synthesis of SFAE requires

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mild conditions and has high regioselectivity. Moreover, subsequent downstream processing is minimal owing to the absence of side-products (Hill and Rhode, 1999). Consequently, the interest in enzyme-catalyzed synthesis of SFAE has increased and a vast amount of literature exists on the use of enzymes for the synthesis of SFAEs, reporting high conversion rates and high productivities. Therefore, enzymatic synthesis of sugar esters proves to be a valuable technology for the production of individual SFAE and there is still room for further research since there is paucity of information on the functional properties of SFAEs. This chapter describes the chemical and enzymatic pathways designed to obtain sugar esters, the physicochemical properties of these compounds, their toxicity and biodegradability, and the applications in the food, pharmaceutical, and cosmetic industries. Accordingly, it is expected that knowledge/technology presented in this chapter could motivate the design of novel lipidbased surface-active compounds valuable for the food, pharmaceutical, and cosmetic applications.

ACKNOWLEDGMENT B. Pe´rez thanks the Danish Council for Independent Research for postdoctoral grant 505400062B.

ABBREVIATIONS CMC DBU DE HLB ILs 2M2B MEK DMF DMSO PPL SCFs SFAE

critical micelle concentration 1,8-diazabicyclo [5.4.0] undec-7-ene degree of esterification hydrophilic-lipophilic balance Ionic liquids 2-methyl-2-butanol methylethyl ketone N,N-dimethylformamide dimethylsulfoxide porcine pancreas lipase supercritical fluids sugar fatty acid esters

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FURTHER READING Ruiz, C.C., Jose´, Molina-Bolı´var, A.M., 2008. Self-assembly and micellar structures of sugar-based surfactants. In: Ruiz, C.C. (Ed.), Sugar-Based Surfactants. CRC Press, Boca Raton, FL.

Chapter 11

Fatty AcidsBased Surfactants and Their Uses Douglas G. Hayes University of Tennessee, Knoxville, TN, United States

Chapter Outline 11.1 Introduction 11.1.1 Biobased Surfactants: A Growing Market 11.2 Biobased Surfactants Are a Robust Product for an Oleochemical-Based Biorefinery 11.3 Oleochemical Feedstocks for Surfactant Synthesis 11.4 Sustainability of OleochemicalBased Surfactants: Truths and Myths 11.5 Green Manufacturing of Biobased Surfactants 11.6 Ionic Surfactants 11.6.1 Methyl Ester Sulfonates 11.6.2 Esterquats 11.6.3 Amino AcidBased Surfactants 11.6.4 Others

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359 361

367 368 369 369 369 370 371

11.7 Ester-Based Nonionic Surfactants 372 11.7.1 Glyceride Esters 372 11.7.2 Ethoxylates of Fatty Acids and Partial Glycerides 372 11.7.3 Sugar Esters 372 11.7.4 Polyol Esters 373 11.8 Ether and Amide-Based Nonionic Surfactants 373 11.8.1 Alkyl Polyglucosides 373 11.8.2 N-Alkyl N-Methyl Glucamine 374 11.8.3 Others 374 11.9 Zwitterionic (Amphoteric) Surfactants 374 11.9.1 Phospholipids 374 11.9.2 Betaines 375 11.10 Glycolipid Biosurfactants 376 11.11 Conclusion 378 References 379

11.1 INTRODUCTION 11.1.1 Biobased Surfactants: A Growing Market Surfactants and detergents, molecules that adhere to interfaces (e.g., wateroil, liquidgas, and solidliquid or gas) and lower their surface energy, have numerous applications in our everyday lives, including foods, Fatty Acids. DOI: http://dx.doi.org/10.1016/B978-0-12-809521-8.00013-1 Copyright © 2017 AOCS Press. Published by Elsevier Inc. All rights reserved.

355

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Fatty Acids

medicines, toiletries, cleaners, automotive fluids, paints and coatings, etc. Their surface activity is enabled by their molecular structure, consisting of separated hydrophilic and lipophilic domains. As one adds surfactant to a two-phase system (e.g., water and oil), the concentration of surfactant adsorbed at the liquidliquid interface increases and concurrently the surface energy (i.e., interfacial tension) decreases until the interface becomes saturated in surfactant, known as the critical micelle concentration (CMC). As surfactant addition crosses the CMC, the surface tension is not further decreased, and the excess surfactant frequently forms self-assembly systems such as micelles. The interfacial tension for a liquidgas system is commonly referred to as the “surface tension.” Surfactants can be categorized by their chemistry, particularly that of their polar moiety, or “head group”: cationic, anionic, zwitterionic, or nonionic. The relative strength of surfactants’ hydrophilic and lipophilic moieties (known as their hydrophiliclipophilic balance, or HLB) determines the nature of their surface activity, whether they are able to dissolve water into oil (more lipophilic), oil into water (more hydrophilic), or nearly balanced in hydrophilicity and lipophilicity, allowing them to form bicontinuous and lamellar structures. Surfactants’ HLB can be tuned by environmental factors such as temperature (which increases the polarity of ionic surfactants and the lipophilicity of alkyl ethoxylate nonionic surfactants) and salinity (which decreases the hydrophilicity of ionic surfactants via Debye shielding). The ideal surfactant is described as inducing a low surface or interfacial tension and possessing low Krafft-point temperature (where the latter is a critical temperature below which surfactants form crystalline structures and above which surfactants can form micelles and related self-assembly systems), high solubility in water or oil, insensitivity of its surface activity to temperature, salinity, or other environmental factors, fast kinetics for their self-assembly, high biodegradability and biocompatibility, an excellent environmental profile, and a low cost-to-performance ratio (Scheibel, 2007). Biobased surfactants are derived “. . . in whole or significant part of biological products or renewable domestic agricultural materials (including plant, animal, and marine materials) or forestry materials” (US Senate Committee on Agriculture Nutrition and Forestry, 2006), are frequently fatty acidbased, and are valuable products attracting increased interest. In 2012 biobased surfactants comprised 24.2% of the $27.0 billion/15.6 million metric tons (MMT) surfactant and detergent industry (i.e., $6.6 billion/3.8 MMT) (Markets and Markets, 2012). The majority of the biobased surfactants market is in Europe and North America (47.4% and 26.6%, respectively, in 2012) (Markets and Markets, 2012). The predictions for 2017 are $36.5 billion/20.9 MMT for the overall surfactant and detergent market, and $10.1 billion/5.7 MMT for biobased surfactants, accounting for 27.5% of the overall market, with the greatest region of growth being Asia/Pacific (Markets and Markets, 2012). The increase has occurred despite the decrease

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Fatty AcidsBased Surfactants and Their Uses Chapter | 11

in cost for petroleum feedstocks in recent years. The molecular structure of several biobased surfactants described in this chapter is depicted in Figs. 11.111.4. Use of biobased surfactants spans all market sectors associated with surfactants and detergents: laundry detergents, foods, cosmetics, personal care products, pharmaceuticals, paints and coatings, and environmental remediation to name a few. A major driver for the increased demand by consumers and retailers for products that represent increased sustainability described as “. . .nature. . .not [being] subject to increasing concentrations of substances extracted from the earth’s crust, to high concentrations of substances produced by society, or to physical degradation” (McCoy, 2008a). Sustainability therefore focuses upon the entire lifecycle of a material, from cradle-to-grave-to-cradle, and includes energy and water conservation, absence of environmental impact (including production of pollutants) and minimal production of wastes. Unlike fossil fuelbased feedstocks, the use of biobased feedstocks such as fatty acids and their derivatives in the preparation of surfactants leads to no net increase of atmospheric carbon dioxide, a greenhouse gas associated with climate change and global warming by climatology experts (Maslin, 2014; Stott et al., 2016). In this chapter, biobased surfactants involving the fatty acid group (which comprises the majority of biobased surfactants) will be reviewed in terms of the oleochemical biorefinery model and sustainability, including their preparation via green manufacturing principles. This chapter subsequently describes the synthesis, properties, and applications of several ionic surfactants, such as O

O O Palmitic acid methyl ester sulfonate (MES)

S

O– Na+

O O O

O O

OH N+

O S

–O

O

O Didodecyl esterquat O

O O Phosphatidylcholine

O O

O

N+

P

O O



O–

N H

N+

Cocamidopropyl betaine O

FIGURE 11.1 Molecular structure of ionic and zwitterionic (amphoteric) biobased surfactants.

358

Fatty Acids O

OH

OH OH

N

O OH

Decyl N-methyl glucamine

OH

ONa N

O O NH

Sodium lauroyl sarcosinate

ONa

Sodium lauroyl glutamate

O

O

O

O

O

O

+H N 3

O

C11H23

O

ONa

Cl

+ – H3N

(3,3'-Di-Lauroyl-N ε,N ε-bis-2,3dihydroxypropyl)-lysine methyl ester hydrochloride

O O C11H23

+H N 2

O

1,2-Di-O-Lauryl-rac-glycero3-O-L-Arginine hydrochloride

NH

NH2+

O N C11H23

O

C11H23

O OH

Cl-

OH

FIGURE 11.2 Amino acidbased surfactants.

OH H OH H O

β-Dodecyl maltoside (β-C12G2)

O

H

H

OH OH

O

OH

O

O H

O

H

H

6'

1'

OH

O

6

H

HO H

H

3

H

H

HO OH

3

O

H

Sucrose 6-Monooleate H

OH

O OH

H

Lauroyl-N-methylglucamide

H

HO OH

HO

O

H

H

OH

OH OH

N OH

OH

O

O O

3

3 O

j OH

O

O

O

O

n

O

3

3

O

O

k m

O

OH

OH

Acetylated monoolein O

Polysorbate (ethoxylated sorbitanoleic acid ester)

FIGURE 11.3 Molecular structure of nonionic biobased surfactants.

Fatty AcidsBased Surfactants and Their Uses Chapter | 11

359

OH O O

O

O

O OH

O OH

O O O

O OH

O

Mono-rhamnolipid

O H

O

O

H OH

OH

Sophorolipid OH

O HO

OR

OH

H OH

O

O

H

O O

H O

OH

H

OH

O O

O H

OH

OH O

OH

H

O

OH OH

OH

H

H

H

Mannosylerythritol n lipid

R= H, or O

m

H

Trehalose lipid OR

O

OH

m+n =27-31

FIGURE 11.4 Molecular structure of biosurfactants.

methyl ethyl sulfonates (MES) and esterquats, nonionic surfactants [a category of surfactant undergoing an increase in market sector (Patel, 2004)], including sugar and polyol esters and alkyl glycosides, and zwitterionic (amphoteric) surfactants such as phospholipids and betaines. Thereafter, glycolipid biosurfactants is discussed. The term biosurfactant refers to the surfactants produced directly by microorganisms that typically consists of lipid, protein, and/or carbohydrate moieties and are frequently associated with cell walls or membranes (Kitamoto et al., 2009; Pinzon et al., 2009). Biosurfactants are divided into four categories: fatty acidtype (e.g., phospholipids, fatty acid soaps, etc.), glycolipid-type, lipopeptide-type, and polymer-type (Kitamoto et al., 2009). The latter two are not discussed in this review. The oldest known biobased surfactant, soap (saponified fatty acids), and polymeric surfactants are also not discussed herein.

11.2 BIOBASED SURFACTANTS ARE A ROBUST PRODUCT FOR AN OLEOCHEMICAL-BASED BIOREFINERY Fig. 11.5 depicts the possible process streams that could be leveraged for production of biobased surfactants in an oleochemical biorefinery. The biorefinery concept entails the utilization of biobased feedstocks

360

Fatty Acids

Nutraceuticals & food products

Sterols / minor components Proteins Meal Polysaccharides (starch) Oilseed crops

Soapstock Refined oil

Glucose (sugars)

Amino acids Ethanol (alcohols)

Alcoholamines Biofuels

Sorbitol Phospholipids FFA

Biosurfactants Lysolecithin Fatty amines Fatty alcohols

FAME / FAEE

Diethyl carbonate Carbonates

Biodiesel Propanediols

MAG / DAG Polyglycerol Glycerine

Chemicals & materials

Glyceric acid

FIGURE 11.5 Production of biobased surfactants according to an oleochemical biorefinery model.

(e.g., lignocellulosic biomass, oilseed crops, and aquatic organisms) for the production of fuels, chemical intermediates, fine chemicals, and materials resulting from the proper fractionation of the feedstock, analogous to the fractionation of petroleum at a petrochemical refinery (e.g., by distillation) into short- and medium-chain alkanes for fuels, long-chain alkanes for lubrication, and aromatics for preparation of chemicals and polymeric materials (a broad description of a complex process) (Hatti-Kaul et al., 2007; Hill, 2007; Johansson and Svensson, 2001). As described earlier, fatty acyl groups serve as the principal biobased surfactant feedstock. [However, sterols and lignin can also serve as sources for the lipophilic portion of surfactants (Holladay et al., 2007; Johansson and Svensson, 2001).] The fatty acyl groups are typically derived from oilseeds in triacylglycerol (TAG) form, but also can be derived from oleochemical coproducts such as free fatty acid (FFA) or phospholipids (e.g., in soapstock, a coproduct formed during degumming) obtained during the refining process. Fatty acyl groups used as lipophilic building blocks for surfactants are typically in the form of FFA or FA esters, obtained via hydrolysis or alcoholysis of TAG, respectively. Particularly attractive as acyl donors are FA methyl esters (FAME) due to relative abundance in biodiesel preparation. Many fatty acidbased surfactants contain an ester bond to conjugate the hydrophilic and lipophilic

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361

compounds. Ester bonds allow for biodegradability and biocompatibility, which are useful properties for surfactants used in foods, cosmetics, personal care products, and pharmaceuticals. However, ester bonds are quite labile, which prevents their utility for many product sectors, such as laundry detergents. More robust are ether, amide, and carbonate linkages. To enable formation of more stable biobased surfactants, fatty acyl groups can be reduced to fatty alcohols or fatty amines (Egan, 1968; Giraldo et al., 2010). Longchain carbonates are prepared through conjugation of fatty alcohol and diethyl carbonate (Banno et al., 2007), a chemical prepared from catalytic oxidation of ethanol (Rudnick, 2006). Conversion of fatty acids into acid chlorides (Bauer, 1946; Busch et al., 2004) allows for additional reactions to occur. In addition, the hydrophilic moiety of the surfactant can also be derived from oilseed components, such as polysaccharides and proteins. A desirable feedstock for the hydrophile is glycerol, an inexpensive coproduct derived from biodiesel production. Glycerol can be used directly (e.g., producing monoacylglycerols, or MAG, or their acylated or ethoxylated form) and converted into other glycols such as glyceric acid (Habe et al., 2009), 1,2- and 1,3-propanediol, glycerol carbonate, or polymerized into polyglycerol (Barrault et al., 2004), to enhance the diversity of biobased surfactant products that can be prepared. Sugars are a common biorefinery feedstock useful for preparing surfactant hydrophiles, including their derivatives, such as sugar alcohols (e.g., sorbitol and sorbitan, the latter produced from dehydration of the former) (Liu et al., 2010; Tang et al., 2004; Wen et al., 2004), furfuryl (De Jong and Marcotullio, 2010) and levoglucosanyl (Lakshmanan and Hoelscher, 1970) derivatives, and glucaric acid (Anonymous, 2014). Amino acids (Husmann, 2008; Infante et al., 2009) [or ethanolamine and isopropylamine, derived from serine and threonine, respectively (Scott et al., 2007)] and DNA (Leal et al., 2006) are also useful biorefinery streams that can be used as feedstocks for the hydrophile. Alternatively, oleochemical feedstocks can be utilized as carbon-energy sources for microorganisms that produce glycolipid biosurfactants such as sophorolipids (SLs) and rhamnolipids (RLs). Although ethoxylate groups are generally derived from petrochemicals, these important groups contained in nonionic surfactants can potentially be derived from biobased ethylene, produced from bioethanol that derived from sugar cane (Gielen et al., 2008). Several commercially available biobased surfactants are listed in Table 11.1.

11.3 OLEOCHEMICAL FEEDSTOCKS FOR SURFACTANT SYNTHESIS For nonfood applications, the optimal fatty acyl feedstock will contain 1014 carbons and no double bonds, the latter to enhance oxidative stability. The composition of common high-lauric oils employed for biobased

362

Fatty Acids

TABLE 11.1 Selected Commercially Available Biobased Surfactants Manufacturer

Product

Ionic Surfactants BASF (Ludwigshafen, Germany)

Sulfopon Sodium coco sulfates

Clariant (Muttenz, Switzerland)

Hostapon CT (Sodium Methyl Cocoyl Taurate)

Evonik (Essen, Germany)

Adogen, Arosurf, Carspray, Rewoquat esterquats

Huish Detergents, Inc. (Salt Lake City, UT, United States)

MES

Lion Corp. (Tokyo, Japan)

MES

Longkey (Guangzhou, China)

MES

Stepan (Northfield, IL, United States)

Alpha-Step MES

Undesa (Barcelona, Spain)

Khemifluid esterquats

Amino Acid Surfactants Stepan (Northfield, IL, United States)

BergaSoft SCG 22 sodium Nα-cocoyl glutamate

Schill 1 Seilacher (Boeblingen, Germany)

Perlastan Nα-acyl glutamate and sarcosinate

Zschimmer and Schwarz (Lahnstein, Germany)

Protelan AGL sodium Nα-cocoyl glutamate

Zwitterionic (Amphoteric) Surfactants American Lecithin Company (Oxford, CT, United States)

Lecithins

BASF (Ludwigshafen, Germany)

Dehyton cocamidopropyl betaine

Cargill (Minneapolis, MN, United States)

Various soy lecithin grades and products

Fresenius Kabi (Homburg, Germany)

Lecithins, from egg yolk for pharmaceutical applications

Nonionic Surfactants BASF (Ludwigshafen, Germany)

Dehymuls PGPH (polyglycerol polyhydroxystearate), Plantacare, Glucopon, and Plantaren APGs

Brenntag Specialties (Mulheim an der Ruhr, Germany)

Mono- and diglyceride mixtures (and acetic, citric, and lactic acid esters thereof); sucrose esters, polyglycerol esters, polyglycerol polyricinoleate (Continued )

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363

TABLE 11.1 (Continued) Manufacturer

Product

Clariant (Muttenz, Switzerland)

Genapol fatty alcohol ethoxylates, Genagen fatty acid ethoxylates, Genamin amine ethoxylates, Genamin castor oil ethoxylates, GlucoTain alkanoyl-N-methylglucamide

Croda (Snaith, United Kingdom)

Castor oilbased ethoxylates: Etocas and Croduret series; fatty alcohol ethoxylates: Brij series; castor oilbased ethoxylates: Etocas and Croduret series; fatty acid ethoxylates: Myrj series, sorbitan(ol) ester ethoxylates

Danisco (Copenhagen, Denmark)

Grinsted acetam and citrem (acetic acid and lactic acid, respectively) esters of MAG, PGPR (polyglycerol polyricinoleic acid esters), PGMS (propylene glycol monostearate ester), SMS sorbitan monostearate

Environmental Fluids, Inc. (Scottsdale, AZ, United States)

Enviracide fatty acid ethoxylates, Enviramine Coco amine ethoxylates, Envirotan sorbitan esters

Esterchem (Leekbrook, United Kingdom)

Sorbitan esters, MAG, ethylene and propylene glycol esters, fatty acid ethoxylates

Evonik (Essen, Germany)

Isolan GPS polyglycerol esters

Guangxi Gaotong Food Technology Co., Ltd. (Liuzhou, China)

Sucrose esters of hydrogenated palm oil fatty acids

Huntsman (The Woodlands, TX, United States)

Ecoteric fatty acid ethoxylates and sorbitan esters, Alkadet APGs, Empilan ethylene glycol esters

Hychem Corp. (Belmar, NJ, United States)

Monoglycerides, polyglycerol esters, propylene glycol esters

Kerry Ingredients and Flavours (Beloit, WI, United States)

Myverol monoglycerides, Admul polyglycerol esters

Mitsubishi-Kagaku Foods Corp. (Tokyo, Japan)

Ryoto sugar esters

Nikkol (Tokyo, Japan)

Decaglyn polyglycerol polyricinoleate and polyesters

Nippon Fine Chemicals (Tokyo, Japan)

Sucraph AG-8 caprylyl glucoside

Riken Vitamin (Tokyo, Japan)

Rikemal and Poem monoglycerides (and acetylated and citrate esters thereof), di- and polyglycerol esters, polyglycerol polyricinoleate, sorbitan esters, propylene glycol esters (Continued )

364

Fatty Acids

TABLE 11.1 (Continued) Manufacturer

Product

Saibaba Surfactants P Ltd (Gujarat, India)

COCO amine ethoxylates

Zhejiang Deyer Chemicals Co (Zhejiang, China)

Polyglycerol esters, sorbitan monostearate, sucrose esters

Biosurfactants BASF (Ludwigshafen, Germany)

Rhamnolipids, Rewoform SL 446 Sophorolipids

Brenntag Specialties (Mulheim an der Ruhr, Germany)

Sophorolipids (modified)

Clariant (Muttenz, Switzerland)

NatSurFact rhamnolipids, sophorolipids

Ecover (Malle, Belgium)

Eco-Surfactant Sophorolipids

Evonik (Essen, Germany)

Rhamnolipids, sophorolipids

GlycoSurf, LLC (Park City, UT, United States)

Rhamnolipids (synthetically produced)

Groupe Soliance, Division of Givaudan Active Beauty, Pomacle France

Sophogreen Sophorolipids

Lion Corp. (Tokyo, Japan)

Sophorolipids

Nikkol (Tokyo, Japan)

Rhamnolipids and Trehalose Lipids

Schill 1 Seilacher (Boeblingen, Germany)

Sophorolipids

Stepan (Northfield, IL, United States)

ACS-Sophor sophorolipids from mahua oil

Wheatoleo (Reims, France)

Sophorolipids

surfactant synthesis is given in Table 11.2. The most frequently used sources are palm kernel (Chempro Gujarat India, 2016), palm stearin [a palmitic acidrich coproduct produced from palm oil (Sellami et al., 2012)], and coconut (Pham, 2016) oils. Currently there are no currently available high-lauric oils produced in North America or Europe, although cuphea, which contains 77%84% capric acid in its oil (Table 11.2), has been investigated as a potentially valuable new oilseed crop (McKeon, 2016b). However, an emerging source of high-lauric acid content is algal oil. Interest in oil expression by algae increased greatly in the first decade of the 21st century for the production of biodiesel and related biofuels. Due to the decrease of petroleum prices, interest toward this application has waned.

TABLE 11.2 Fatty Acid Composition of Selected Feedstocks Used to Prepare Biobased Surfactants Fatty Acid

Palm Kernela

Coconutb

Cupheac

10:0

37

4.4

7784

12:0

4052

44.5

23

14:0

1418

18.6

24

16:0

79

12.0

3245

18:0

13

4.8

18:1

1119

18:2

0.52

a

Palma

c

Beef Tallowe

Palm Stearinf

0.10 0.52

1.01.5

1.1

13.016.0

24.028.0

55.9

27

6.08.0

20.024.0

3.7

11.0

3852

39.041.3

38.043.5

32.4

2.2

511

37.038.0

2.04.0

6.7

Chempro Gujarat India (2016). Pham (2016). McKeon (2016b) (PSR23, an interspecific hybrid of Cuphea viscosissima and Cuphea lanceolate). d Barros et al. (2015) (Jatropha curcas). e Alm (2017). f Sellami et al. (2012). b

Jatrophad

366

Fatty Acids

Therefore, companies that have focused upon algal oils such as Solazyme (South San Francisco, CA, United States) have leveraged their intellectual property relating to recombinant DNA expression in algae, and bioprocessing of algae, to prepare oils tailored in their fatty acyl composition to target specific applications, such as high-lauriccapric and myristic oils for biobased surfactants and other ingredients used in cosmetics (AlgaPur). Sugar cane, an inexpensive renewable resource, is commonly used as a carbon source for the fermentative AlgaPur product. Microalgae oils are produced with low carbon, water, and land use impact. Another potential route to preparing medium-chain fatty acids or their alkyl esters is olefin metathesis, a genre of reactions involving the cleavage and reformation of carboncarbon double bonds, enabled by the development of homogeneous transition metal carbine catalysts in recent years, particularly Grubbs and Schrock catalysts, which led to Nobel prizes in chemistry for the inventors in 2005 (Montero de Espinosa and Meier, 2012). For example, cross-metathesis of oleic acid and ethene (the latter of which can be biobased, i.e., derived from sugar cane) will produce 9-dodecenoic acid (Fig. 11.6) (Rybak et al., 2008). 10-Undecenoic acid is readily prepared from ricinoleic acid (Van der Steen and Stevens Christian, 2009). Lard and tallow are inexpensive sources of C16- and C18-rich saturates (Table 11.2). Hydroxy acidrich oils, particularly castor and lesquerella oils (which contain ricinoleic [R-18:1-9c, OH-12] acid and its C20 homolog, lesquerolic acid, as their prominent fatty acyl group, respectively), have several specific applications as surfactants (described later) (Chen, 2016; McKeon, 2016a,c). Similarly, epoxy fatty acids may serve similar applications as hydroxyl acids, and can occur naturally (e.g., vernolic acid, 18:1-9, epoxy-12S,13R) from Vernonia galamensis oil (McKeon, 2016c) or via chemical epoxidation of TAG containing unsaturated FA (Tan and Chow, 2010).

O

OH

Ethylene

Oleic acid

Grubbs catalyst O

1-Decene

OH

9-Decenoic acid FIGURE 11.6 Production of medium-chain fatty acids via cross-metathesis between oleic acid and ethylene.

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367

For food-related applications, high-oleic oils such as corn, olive, cottonseed, palm, or soybean oils are commonly used as sources of lipophilic building blocks for biobased surfactants. However, jatropha and soapnut (Sapindus) oils, derived from a plant native to India that can be cultivated inexpensively on marginal agricultural land, may be a viable replacement for the common high-oleic oils described earlier (Table 11.2). Jatropha oil has attracted interest since 2000 as an economically viable feedstock for biodiesel (Barros et al., 2015). If these oils are to be used in nonfood applications, they need to be hydrogenated (e.g., using Ni catalysts) to improve their oxidative stability (Veldsink et al., 1997).

11.4 SUSTAINABILITY OF OLEOCHEMICAL-BASED SURFACTANTS: TRUTHS AND MYTHS The main advantage of biobased surfactants with regard to environmental sustainability is their replacement of surfactants derived from fossil fuelbased feedstocks, to reduce the net production of CO2. It should be noted that many biobased surfactants contain only partial biobased content. The latter is quantified by determining the percentages of surfactant carbon atoms that are derived from renewable resources. For instance, for the fatty alcohol ethoxylate C12E5, its molecular structure consists of 12 carbons derived from seed oils (comprising its fatty alkyl group) and 10 carbons in its ethoxylate group derived from fossil fuels. (However, as noted earlier, the ethoxylate group may be derived in the future from biobased sources.) Therefore, the biobased content of C12E5 is (12/22) 3 100% 5 54.5%. The biobased content is more formally determined via analysis of the stable isotopes of carbon [e.g., via ASTM D6866 (ASTM International, 2012) or the equivalent]. Moreover, the 14C content of renewable resources is significantly high in contrast to near-zero 14C content for fossil fuels. A common myth is that biobased surfactants are more biodegradable than surfactants derived from fossil fuels. This hypothesis is simply not true; moreover, a chemical’s beginning- and end-of-life are uncoupled. A chemical’s biodegradability property is solely a function of its molecular structure, and has no relationship with its origin. For example, alkylphenyl ethoxylates, derived from petroleum, possess excellent environmental profiles (Balson and Felix, 1995). In contrast, biobased feedstocks that undergo chemical modification to a significant extent are poorly biodegradable (McCoy, 2007a). In terms of economic sustainability, the main difference between biobased and fossil fuelderived surfactants is the costs associated with their respective feedstocks. Moreover, there are typically no significant differences in manufacturing costs between the two (US Department of Energy, 1999); or, biobased surfactants may be slightly less expensive (McCoy, 2007b).

368

Fatty Acids

However, their biobased origin serves as an incentive for consumers interested in purchasing more environmentally friendly products, particularly in Europe and North America. Many companies pursue eco-friendly labeling of their biobased surfactant products to further attract consumers. A common label pursued in the United States is “Biopreferred,” a program regulated by the US Department of Agriculture. Canada and the European Union have similar programs (Canadian EcoLogo and Eco-Label, respectively). Many labels are certified by nongovernmental organizations such as Underwriter’s Laboratory (ECOLOGO). Although many of the labels are based in part on international standards (e.g., ISO 14024 and 14001, respectively), differences exist between regulating agencies and organizations and in regulations between years, which creates difficulty for surfactant manufacturers and consumers (McCoy, 2008b; Schalitz, 2007). Palm and palm kernel oils have been a particular target of environmental activists, particularly in Europe, due to concerns of deforestation in countries such as Indonesia and Malaysia to obtain new cultivatable land for palm plantations. To address this issue, the Roundtable for Sustainable Palm Oil (RSPO) council has been established. This organization, through involvement with several stakeholder groups (growers and plantations, users and manufacturers, and nongovernment organizations advocating environmental sustainability), has developed effective regulations for land usage and farming practices on farm oil plantations to minimize environmental impact. Users of palm oil such as Unilever, a founding member of the RSPO in 2004, specify certification by RSPO for its purchase and use in their products. Recently, the Malaysian (and Indonesian) governments have prepared their own certifications, Malaysian (Indonesian) Sustainable Palm Oil Board (MSPO and ISPO, respectively), that unlike the RSPO would be affordable for mid-sized palm plantations, but are less stringent than those of the RSPO.

11.5 GREEN MANUFACTURING OF BIOBASED SURFACTANTS To further enhance sustainability, green manufacturing principles are of interest to manufacturers of biobased surfactants. A potentially valuable green manufacturing approach in the future will be the use of enzymes, to lower energy costs, to increase reaction selectivity (thereby reducing the amount of by-products, which will facilitate downstream purification and reduce that amount of generated waste). Particularly, lipases are the most potentially valuable, due to their relatively low costs, high stability (when operated under proper conditions, noting the availability of many thermostable lipases), and their ability to form ester bonds, which are prominent in many surfactants [reviewed in Hayes (2011)].

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11.6 IONIC SURFACTANTS 11.6.1 Methyl Ester Sulfonates MES (Fig. 11.1) are perhaps the most widely used biobased surfactant, with its major application being in powder and liquid laundry detergents, the largest market sector for surfactants, particularly as a replacement of the fossil fuelderived surfactants, linear alkyl sulfonates. The major producers of MES in the world are Lion Corp. (Tokyo, Japan, 40 kton/yr), Stepan (Northfield, IL, United States, 50 kton/yr), Huish Detergents, Inc. (Salt Lake City, UT, United States, 80 kton/yr), and Lonkey Industrial Co LTD (Guangzhou, China) (Table 11.1). MES is produced in a multistage process (Ahmad et al., 2007; Edser, 2006; Foster, 2006a,b,c; Roberts et al., 2008). First, methyl esters are reacted with SO3 to produce methyl ester sulfonic acid. The latter is sent to an acid digester, then is bleached, and then is neutralized with NaOH to produce a crude MES. The final step is removal of methanol. A common feedstock for MES is palm stearin, particularly a C16-rich fraction thereof, after their conversion to methyl esters. Also employed are palm kernel oil, coconut oil, and tallow. MES are highly biodegradable, can withstand calcium hardness, possess outstanding detergency properties and stability in hot and cold water, do not inactivate enzymes used in laundry products (e.g., amylases, proteases, and lipases), but also are poor at foaming and are susceptible to hydrolysis under high pH conditions (Bognolo, 2008; Foster, 2006b).

11.6.2 Esterquats Esterquats, quaternary ammonium compounds containing cleavable ester bonds to conjugate fatty acyl groups to a polar group containing the quaternary ammonium head group, are the most commonly used biobased cationic surfactant. They are produced by transesterification of FAME (e.g., from animal fats or vegetable oils) with triethanolamine (for the esterquat shown in Fig. 11.1), methyldiethanolamine, or other hydroxyl amines at a controlled stoichiometric ratio (e.g., 2:1 mole ratio to prepare the diesterquat shown in Fig. 11.1, with smaller and larger ratios producing mono- and triesterquats, respectively) at 250