Fungal Lipid Biochemistry: Distribution and Metabolism [1 ed.] 978-1-4684-2831-5, 978-1-4684-2829-2

Citation preview

FUNGAL LIPID BIOCHEMISTRY

MONOGRAPHS IN LIPID RESEARCH David Kritchevsky, Series Editor Wistar Institute Philadelphia, Pennsylvania

Volume 1 • FUNGAL LIPID BIOCHEMISTRY 8y John D. Weete • 1974

A Continuation Order Plan is available for this series. A eontinuation order will bring delivery of eaeh new volume immediately upon publieation. Volumes are billed only upon aetual shipment. For further information please eontaet the publisher.

FUNGAL LIPID BIOCHEMISTRY Distribution and Metabolism

John D. Weete Department of Botany and Microbiology Auburn University Auburn, Alabama

With Contributions by

Darreil J. Weber and William M. Hess Department of Botany and Range Science Brigham Young University Provo, Utah

PLENUM PRESS • NEW YORK AND LONDON

Library of Congress Cataloging in Publication Data Weete, John D

1942-

Fungal lipid biochemistry. {Monographs in lipid research, v. 1) Includes bibliographical references. 1. Fungi—Physiology. 2. Lipids. I. Title. [DNLM: 1. Fungi. 2. Lipids. W1 M0567V v. 1 1974 /QK603 W398f 1974] QK601.W44 589'.2'0419247 74-8457 ISBN 978-1-4684-2831-5 ISBN 978-1-4684-2829-2 (eBook) DOI 10.1007/978-1-4684-2829-2

© 1974 Plenum Press, New York Softcover reprint of the hardcover 1st edition 1974 A Division of Plenum Publishing Corporation 227 West 17th Street, New York, N.Y. 10011 United Kingdom edition published by Plenum Press, London A Division of Plenum Publishing Company, Ltd. 4a Lower John Street, London, W1R 3PD, England All rights reserved No part of this book may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise, without written permission from the Publisher

Preface With the development of highly sophisticated analytical techniques and instrumentation during the past 15-20 years, progress in the field of lipid biochemistry has been greatly accelerated. Within this period, there has been an increasing volume of information concerning the distribution and metabolism of lipids in animals and, more recently, in plants. The fungi have played an important role in studies concerning the biochemistry of lipids and, in this text, they are treated separately from the photosynthetic plants. This book is concerned with distribution and biochemistry of lipids in fungi. The text is divided into three sections, beginning with an introduction to fungallipids which includes total lipid abundances in fungal cells and cell fractions and cultural conditions influencing lipid production. In the second section, each chapter deals with the distribution andjor metabolism of a single lipid class as it occurs in fungi. Comparisons with plants and animals are also included. Six major lipid classes are covered which include the aliphatic hydrocarbons, fatty acids, sterols, triacylglycerols, glycerophosphatides, and sphingolipids. The third section contains two chapters concerned with the physiology and ultrastructure of fungal spore formation and germination with particular emphasis on lipids. Although this book is not intended to be a comprehensive review of the literature, the information presented is compiled from over 1000 articles, most of which were published during the past 10-12 years. Although some relevant articles may have been overlooked in the preparation of this text, I believe that each chapter represents an up-to-date and detailed coverage of the subject matter. This work was initiated and partially completed while the author was a Staff Scientist at the Lunar Science Institute, Houston, Texas, which is under the joint support of the Universities Space Research Association and the National Aeronautics and Space Administration, Johnson Spacecraft Center under contract No. NSR 09-051-00l. I would like to thank v

vi

Preface

the Director Dr. l. W. Chamberlain, the librarian Mrs. Fran Waranius, Eloise Williams, and the entire staff of the Lunar Science Institute. Special thanks goes to Mrs. Lila Mager for her contributions to the preparation of the entire manuscript. I would also like to acknowledge Dr. Olivia Thompson, Denise Flournoy, Elaine Turner, Elizabeth Wiggins, and Teresa Glasscoch (artist) of Auburn University for their assistance in the final preparations of this manuscript. Finally, I would like to express my gratitude to Drs. DarreIl l. Weber and William (Bill) M. Hess ofthe Department of Botany and Range Science, Brigham Young University, Provo, Utah, for their contributions of the two final chapters in this text and their reviews of the other chapters. Auburn, Alabama

lOHN

D.

WEETE

Contents

Section I Fungal Lipids

Chapter 1 Introduction to Fungal Lipids

1.1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2. Total Lipids. . . . .. . . . . . . . . . . .. . . . . . . . .. .. . . . .. . . . . . . . . . . 1.2.1. Lipids of Vegetative Hyphae and Yeast Cells. . . . . . . . . . 1.2.2. Spore and Sclerotial Lipids. . . . . . . . . . . . . . . . . . . . . . . . . 1.3. Lipids and Fungal Growth. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4. Cultural Conditions for Lipid Production. . . . . . . . . . . . . . . . . . . 1.4.1. Temperature. ..................................... 1.4.2. Carbon Source. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.3. Inorganic Nutrients. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.4. pH. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.5. Aeration. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.6. Vitamins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5. Cellular Distribution of Lipids. . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5.1. Cell Wall. . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . 1.5.2. Cytoplasm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.6. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vii

3 4 4 12 14 16 16 22 24 25 26 26 27 27 30 32

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Section 11 lipid Classes Chapter 2 Aliphatic Hydrocarbons

2.1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Distribution in the Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Mechanisms of Hydrocarbon Synthesis in Higher Plants. . . . . . 2.3.1. Head-to-Head Condensation Mechanism. . . . . . . . . . . . . 2.3.2. Eiongation-Decarboxylation Mechanisms . . . . . . . . . . . . 2.3.3. Head-to-Head Condensation between Acceptor and Donor Acids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Mechanism of Hydrocarbon Synthesis in Bacteria. . . . . . . . . . . 2.5. BioIogicai Oxidation of Aliphatic Hydrocarbons. . . . . . . . . . . . . 2.5.1. Corynebacterium 7EIC. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.2. Pseudomonas oleovorans. . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.3. Higher Animais. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.4. Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.5. Higher Plants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. References.............................................

39 43 54 54 55 57 57 60 60 62 62 63 64 64

Chapter 3 Fatty Acids

3.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Nomenclature and Structure. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Distribution in the Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1. Myxomycetes and Acrasiales. . . . . . . . . . . . . . . . . . . . . . . . 3.3.2. Phycomycetes..................................... 3.3.3. Ascomycetes and Deuteromycetes (Imperfect Fungi). . . . 3.3.4. Basidiomycetes.................................... 3.4. Cellular and Extracellular Distributions. . . . . . . . . . . . . . . . . . . . 3.4.1. MyceIiai and Yeast Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.1.1. Cell wall . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4.1.2. Protoplast and Membrane. . . . . . . . . . . . . . . . . . . . . . . . .

3.4.2. Extracellular Fatty Acids. . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Fatty Acids of Fungal Spores and Sclerotia. . . . . . . . . . . . . . . . . 3.5.1. Spores. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5.1.1. Phycomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5.1.2. Ascomycetes and Deuteromycetes. . . . . . . . . . . . . . . . . . . . 3.5.1.3. Basidiomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

67 67 70 74 74 79 81 81 81 81 88 89 92 92 92 92 102

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3.5.2. Sclerotia. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6. References.............................................

104 105

Chapter 4

Fatty Acid Metabolism 4.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. Biosynthesis of Saturated Fatty Acids. . . . . . . . . . . . . . . . . . . . . . 4.2.1. Formation of the Initial Reactants in Fatty Acid Biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1.1. Substrate Activation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1.2. Carboxylation of Acetyl CoA. . . . . . . . . . . . . . . . . . . . . . .

4.2.2. Reactions and Enzymes of Fatty Acid Biosynthesis. . . . . 4.2.2.1. 4.2.2.2. 4.2.2.3. 4.2.2.4. 4.2.2.5. 4.2.2.6.

Transacylation. . . . Condensation .... Reduction. . . . . . . Dehydration . . . . . Reduction. . . . . . . Terminal Transfer. .

... ... ... ... ... ...

.... .... .... .... .. .. ....

. . . . . .

........ ........ ........ ........ ........ ........

............. ............. ............. ............. ............. .............

4.2.3. Fatty Acid Synthetase of Yeast. . . . . . . . . . . . . . . . . . . . . . 4.2.4. Mechanisms of Fatty Acid Synthesis by the Yeast Fatty Synthetase Complex. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.5. Control of Fatty Acid Synthesis. . . . . . . . . . . . . . . . . . . . . 4.2.6. Fatty Acid Elongation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Biosynthesis of Unsaturated Fatty Acids. . . . . . . . . . . . . . . . . . . 4.3.1. Monounsaturated Fatty Acids. . . . . . . . . . . . . . . . . . . . . . 4.3.1.1. Aerobic Pathways. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . 4.3.1.2. Anaerobic Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

4.3.2. Polyunsaturated Fatty Acids. . . . . . . . . . . . . . . . . . . . . . . . 4.4. Biosynthesis of Unusual Fatty Acids. . . . . . . . . . . . . . . . . . . . . . . 4.4.1. Nonmethylene Interrupted Ethylenic Acids. . . . . . . . . . . . 4.4.2. Acetylenic Acids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3. Substituted Acids. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3.1. Hydroxy Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3.2. Epoxy Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3.3. Methyl Branched-Chain Acids. . . . . . . . . . . . . . . . . . . . . .

4.5. Fatty Acid Degradation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.1. IX-Oxidation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2. ß-Oxidation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3. w-Oxidation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.4. Lipoxidase.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. References.............................................

109 110 110 110

112 112 114 114 114 115 115 115 116 119 124 126 127 127 127 130 131 134 134 134 136 136 138 139 140 140 141 145 146 146

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Chapter 5 Sterols

5.1. Introduction........................................... 5.2. Structure and Nomenclature. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3. Distribution in the Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.1. Phycomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.2. Ascomycetes and Deuteromycetes. . . . . . . . . . . . . . . . . . . 5.3.3. Basidiomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4. Phylogenetic Relationships. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5. Functions of Sterols. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.1. Higher Plants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5.2. Fungi.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

151 152 155 158 161 163 165 167 167 167 171

Chapter 6 Sterol Biosynthesis

6.1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2. Formation of Squalene. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3. Cyclization of Squalene and the Formation of Lanosterol and Cycloartenol. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4. Conversion of Lanosterol to Ergosterol. . . . . . . . . . . . . . . . . . . . 6.4.1. Demethylation Reactions. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4.2. C-24 Alkylation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4.3. Formation of ｾＸ｟Ｔ＠ Desmethyl Sterols. . . . . .. . . . . . . . . 6.4.4. Formation of Ergosterol from ｾＸ｟Ｔ＠ Desmethyl Sterols 6.5. Biosynthesis of C 27 and C Z9 Sterols by Fungi. . . . . . . . . . . . . . . 6.6. Sterols, Taxonomy, and Fungal Phylogeny. . . . . . . . . . . . . . . . . 6.7. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

175 175 181 184 184 188 190 193 196 197 203

Chapter 7 Acylglycerides, Glycosylglycerides, and Simple Esters

7.1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. Acylglycerides. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.1. Nomenclature and Structure. . . . . . . . . . . . . . . . . . . . . . . . 7.2.2. Distribution. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.3. Biosynthesis. . . . . . . . . .. . . . . . . . . . . . . . . . . . . .. . . . . . ..

209 210 210 211 215

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7.2.4. Lipases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.4.1. Pancreatic Lipase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.4.2. Fungal Lipases .... . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.4.3. Stereospecific Analyses of Acylglycerides. . . . . . . . . . . . . . .

7.3. Glycosylglycerides. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. Simple Esters. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4.1. Methyl and Ethyl Esters of Long-Chain Fatty Acids. . . . 7.4.2. Sterol Esters. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

220 220 221 224 228 229 229 233 235

Chapter 8 Phosphoglycerides

8.1. 8.2. 8.3. 8.4. 8.5.

Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . NomencJature, Structure, and General Distribution. . . . . . . . . . Distribution in the Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phosphoglyceride Biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . Phospholipases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.5.1. Phospholipases A, B, C, D, and Lysophospholipase. . . . 8.5.2. Occurrence of Phospholipases in Fungi. . . . . . . . . . . . . . . 8.6. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

239 239 245 250 259 259 260 262

Chapter 9 Sphingolipids

9.1. Introduction.... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2. Structure and NomencJature. . . . . . . . . . . . . . . . . . .. . . . . . . . . . . 9.3. Distribution............................................ 9.3.1. Plants and Animals. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.2. Fungi.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.2.1. Long-Chain Bases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.2.2. Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3.2.3. Carbohydrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

9.4. Sphingolipid Metabolism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.1. Biosynthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.1.1. Long-Chain Bases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.1.2. Cerebrosides ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4.1.3. Ceramide Oligoglycosides and Other Complex Sphingolipids. .

9.4.2. Degradation...................................... 9.5. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

267 267 271 271 272 272 275 275 278 278 278 281 283 283 285

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Section 111 Physiology and Ultrastructure of Sporogenesis and Spore Germination in Fungi

Chapter 10 Lipid Metabolism and Ultrastructure During Spore Germination

10.1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2. Review of Spore Germination. . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3. Lipid Metabolism and Ultrastructure of Spore Germination in the Various Fungal Classes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.1. Myxomycetes................................... 10.3.2. Zygomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.3. Chytridiomycetes................................ 10.3.4. Oomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.5. Ascomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.5.1. The Aetivation Process . . . . . . . . . . . . . . . . . . . . . . . . 10.3.5.2. Metabolie Changes . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.5.3. Ultrastruetural Changes in Lipid Bodies. . . . . . . . . . . . .

10.3.6. Basidiomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3.6.1. 10.3.6.2. 10.3.6.3. 10.3.6.4.

Stimulators and Inhibitors. . . . . . . . . . . . . . . . . . . . . . Respiratory Aetivities . . . . . . . . . . . . . . . . . . . . . . . . . Metabolie Changes. . . . . . . . . . . . . . . . . . . . . . . . . . . Ultrastruetural Changes in Lipid Bodies. . . . . . . . . . . . .

10.3.7. Oeuteromycetes................................. 10.4. References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

289 290 297 297 298 301 302 303 303 305 312 314 314 315 316 320 321 323

Chapter 11 Lipid Metabolism and Ultrastructure Changes During Sporulation in Fungi

11.1. 11.2. 11.3. 11.4.

Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Review of Sporulation in Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . Electron Microscopy of Fungal Sporulation . . . . . . . . . . . . . . . Specific Changes in Lipids and Ultrastructure Ouring Sporulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.1. Myxomycetes and Cellular Slime Molds. . . . . . . . . . . . 11.4.2. Chytridiomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.3. Oomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.4. Zygomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.5. Trichomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

331 331 334 341 341 343 345 349 352

Contents

xiii

11.4.6. Ascomycetes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.4.7. Basidiomycetes................................. 11.4.8. Deuteromycetes................................. 11.5. References............................................

352 356 358 359

Index to Fungal Species. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

365

Subject Index. . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . .

373

Section I Fungal Lipids

CHAPTER 1

Introduction to Fungal Lipids

1.1 INTRODUCTION To adequately define the term lipid is a difficult task. It has been used to include a large group of compounds that differ widely in their chemical and physical properties. Lipids have been defined on the basis of their relative solubility properties, that is, they include substances which may be partitioned into water-immiscible solvents from water. However, the solubility properties of lipids vary too much for this to be an ade qua te definition and nonlipid substances may possess similar properties. The definition adopted in this text follows closely that of Davenport and Johnson.(1) Lipids are substances that have as part oftheir structure a substituted or unsubstituted aliphatic hydrocarbon chain which confers hydrophobic properties to at least part of the molecule. This definition is extended to include the lipophilic cyclic terpenoid compounds which do not fall into the above category, such as the carotenes, sterols, and steroid hormones. Lipids may be subdivided into smaller groups called lipid classes, each of which includes compounds that have certain aspects of their molecular structure in common and possess similar chemical and physical properties. The classification system used in this text for the lipid classes is shown in Table 1.1. Lipids were among the first natural products to be studied in detail,(2) but significant advances in this field were slow. Within the past 20 years, sophisticated analytical techniques and instrumentation have been developed so that rapid progress is now being made in the area of lipid chemistry. The remainder of this chapter deals with the total abundances of lipids in fungal cells and cell fractions and cultural conditions inftuencing lipid production. 3

4

Introduction to Fungal Lipids

TABLE 1.1. Classification of Lipidsa . b I.

NEUTRAL LIPIDS

A. Hydroearbons-Iong chain, branched and normal, saturated and unsaturated aliphatic hydrocarbons. B. Glyeerides---compounds containing ester, vinyl ether, or saturated ether linkages with the hydroxyl functions of glycerol. C. Fatty Acids-Iong-chain monocarboxylic acids. D. Waxes-esters of long-chain fatty acids and alcohols. E. Estolides-intermolecular lactones of hydroxy fatty acids. F. I soprenoids I. Carotenoids-polyisoprenoid hydrocarbons, alcohols, epoxides, and carboxylic acids containing 40 carbon atoms. 2. Terpenoids-polyisoprenoid compounds varying in carbon number, including vitamin A, squalene, ete. 3. Steroids-alicyclic compounds having the cyclopentanoperhydrophenanthrene carbon skeleton (includes the sterols). G. Other esters (excluding those containing phosphate or sphingosine bases}--these include naturally occurring esters of long-chain fatty acids and short-chain alcohols, such as methyl palmitate, and sterol esters, such as cholesteryl palmitate.

11.

AMPHIPHILIC LIPIDS

A. Glyeerolipids I. Phosphoglycerides--{}erivatives of sn-glycero-3-phosphoric acid that contains at least one O-acyl, O-alkyl, or O-alk-l-enyl group. 2. Glyeosylglyeerides-glycosides of diacylglycerol. B. Sphingolipids I. Phosphosphingolipids-phosphate esters of N-acyl (ceramides) sphinganines. 2. Glyeosphingolipids-glycosides of ceramides. a. Ceramide monoglycosides (I) Cerebrosides-glucosides or galactosides. (2) Sulfatides---contain sulfate ester of galactose. b. Ceramide of oligoglyeosides---contain polysaccharide residues. 3. Glyeophosphosphingolipids---ceramides containing both sugars (polysaccharides) and phosphate esters, phytoglyeolipids in plants and mycoglycolipids in fungi. a b

This classification of lipids is a modification of that outlined by Davenport and Johnson.(I) Severallipid classes not discussed in this text are also omitted from this classification. Reference 164 should be consulted for these lipids (cytosides, globosides, gangliosides, lipoamino acids, lipopeptides, and redox lipids).

1 .2. TOTAL LI PI OS 1.2.1. Lipids of Vegetative Hyphae and Yeast Cells There are numerous reports concerning the production of lipids by various fungal species. A large number of these have been concerned primarily with the cultural conditions favoring fat production and the potential of fungi as a commercial source of fat. The total lipid abundances of many

5

1.2. Total Lipids

of these fungi are given in Table 1.2. Since most of the fungi tested were cuItured under a variety of conditions and the lipids were extracted by several different methods, the values for total lipids are, in most cases, not directIy comparable. However, one can get a general idea of the potential for lipid production by various fungal species. TABLE 1.2. Total Lipid Content of Fungi (Mycelia) Species

PHYCOMYCETES Absidia blakesleeanab A. glauca Absidia ("whorled") A. dauci A. tenuis (PRL 369) Blakeslea trisspora Choanophora curcubitarum C. curcubitarum ( + ) Circinella umbellata C. spinosa Conidiobolus brefeldianus CBS 180/62 C. chlamydosporus CBS 167/55 C. gonimodes CBS 178/61 C. megalotocus CBS C. polytocus CBS 168/55 C. humicola CBS 181/62 C. globuliferous CBS 218/64 C.lomprauges CBS 183/56 C. narodes CBS 183/62 C. paulus CBS 140/57 C. undulatus CBS 142/57 C. heterosporus ATCC 12941 C. heterosporus CBS 543/63

Total Lipid (0;' dry wt.)

Reference

7.2-30.4 18.2-19.5 17.3-21.8 2.3 12.6 15.1-22.1 6.2-16.9 16.1-24.6 6.7-7.4 12.8-21.9 23.7

124 124 124 57 127 124 124 68 124 124 134

15.7

134

17.0

134

12.3

134

17.7

134

18.5

134

21.7

134

14.5

134

18.3

134

18.6

134

8.0

134

15.5

134

13.0

134

6

Introduction to Fungal Lipids

TAßlE 1.2-continued Species

Cunninghamella bertholletiae" Mucor albo-ater M. albo-ater M. plllmhells M. plumbeus M. circinelloides (v. Tieghem) M. circinelloides M. spil10sliS M. spirlOsus M. mucedo M. mucedo M. mucedo M. mucedo M. dispersus M. griseo-cyanus Mucor (N) Mucor(4) Mucor (5) Mucor (7) M. racemosus M. racemosus M. hiemalis M. hiemalis (-) M. hiemalis ( + ) M. mi ehe i M. miehei M. pusillusd M. pusillus M. ramannianus M. ramannianus M. ramannianus Mucor sp. (I) Mucor sp. (11) M. strictus CBS 100.66 M. strictus CBS 576.66 M. strictus CBS 575.66 M. oblongisporus CBS 173.27 M.oblongisporus CBS 220.29

Total Lipid (01" dry wt.)

Reference

12.7-22.0 6.5-41.8 21.8 5.3-14.2 17.9

124 49 126 49 126

15.7-45.4 12.9 28.4-46.2 5.6-12.0 12.0-15.2 19.6-33.3 12.0 2.0-2.9 8.1-10.6 6.0-6.5 2.1-2.9 3.6-5.6 19.4-22.2 3.5-8.5 9.8 8.2-19.0 18.1 15.1-19.3 18.0-18.8 11.9-24.3 7.8-25.1 18.3 23.1-26.2 16.7-55.5 15.2-19.8 15.2 13.2-35.9 8.4-36.4 13.2-20.4

49 126 49 126 33 49 5 124 124 124 124 124 124 124 5 33 5 33 33 5 33 19 5 49 33 5 33 33 33

7.8-24.4

33

11.1-21.4

33

2.5-20.0

33

4.5-17.2

33

7

1.2. Total Lipids

TABLE 1.2-continued Species

M. globosus Parasitella simplex Phycomyces blakesleeanus Pythium ultimum P. irregulare P. vexans Rhizopus arrhizus R. arrhizus R. arrhizus R. nigricans R. nigricans R. nigricans R. oryzea R. oryzea Rhizopus sp. Rhizopus sp. (I) Rhizopus sp. (11) Rhizopus sp. (111) Syncephalastrum sp. Zygorhynchus moelleri Z. moelleri

Total Lipid (% dry wt.)

Reference

16.7 6.1-7.2 15.7-22.1 3.0-48.0 7.9-17.1 5.9-13.3 2.2-15.0 2.7-19.9 0.66-4.9 5.9-13.4 5.3-7.2 15.5-18.3 4.9-35.8 7.0 11.6--25.7 11.6-45.3 20.8-32.8 8.8-32.5 12.1-12.9 7.7-14.0 18.4

19 124 124 74 75 75 8 9 57 49 124 126 49 126 5 33 33 33 124 49 126

14.4-20.7 14.2-14.9 10.5-37.0 39.7 5.7-35.5 4.0 13.1

49 126 51 49 49 121 126

16.0 16.6 12.8-34.9 13.5 14.8-39.8 12.9-25.8 5.2-23.3 45 0.9 2.2 7.6--20.2 10.1-19.0 9.4

50 49 49 126 10 126 53 132 49 127 49 133 19

ASCOMYCETES

Aspergillus clavabus A. clavabus A. fischeri A. flavipes A. flavus A. flavus A. flavus A. flavus (Thom and Church) A. insuetus A. minutus A. minutus A. nidulans A. nidulans A. nidulans A. nidulans A. niger A. niger (NRRL 337) A. nidulans A. sp. Chaetomium thermophile"

8

Introduction to Fungal Lipids

TABlE 1.2-continued Species

Total Lipid (01" dry wt.)

Reference

C. globosum Cladosporium herbarum Claviceps purpureac CyLindrocarpon radicicola Ergot (Claviceps purpurea) Fusarium bulbigenum (2) F. bulbigenum (2) F. bulbigenum (3) F. graminearum F. graminearum F. Lini (1) F. Lini (2) F. lini Bolley (1) F. lycopersici F. lycopersici F. oxysporum F. oxysporum F. solani f. phasevli GLiocladium roseum (PRL 86) Helicostylum piriforme Humicola brevis H. grisea H. grisea var. thermoidead H. insolensd H. linuginosa d H. nigrescens Malbranchea pulchella M. pulchella var. sulfuread M ortierella sp. M. vinacea N eurospora crassa Paecilomyces variati P. aurantiolbrunneum Penicillium chrysogenum (Q-176) P. chrysogenum P. chrysogenum P. chrysogenum (47-133 SLS) P. jlavo-cinereum P. jlavo-cinereum P. jlavo-cinereum (Biourge) P. purpurogenum P. varia ta P. spinulosum

54.1 0.7 1.9-31.3 7.5 30.0 6.9 20.0 16.8 12.6-31.0 10.3-24.4 5.5-28.4 6.8-32.2 5.9-34.6 7.1-16.1 16.2 25.2-33.9 7.7-13.9 2.0 22.4 7.0-9.1 14.6 10.8 13.0 14.2 17.2 8.0 26.5 24.8 34.9 3.2-51.4 6.4-11.9 6.4 6.1 1.8-2.6

19 49 131 114 123 49 126 126 49 126 126 126 49 126 126 49 126 57 127 124 19 19 19 19 19 19 19 19 19 11 6 126 126 52

9.8 1.4 1.2-1.7

19 127 52

12.5 5.3-15.2 28.5 1.3 6.8 5.0-9.5

126 49 50 49 49 53

7.2. Total Lipids

9

TABLE 1.2-continued Species

P. spinulosum P. spinulosum P.oxalicum (Currie and Thorn) P.oxalicum P. lilacinum (NRRL 898) P. cyaneum (S-l1) P. soppi Zal. P. soppi P.luteum P. javanicum P. javanicum P. javanicum P. javanicum (Van Beijrna) P. piscarum (Westling) P. roquefortii (Thorn) P. hirsutum (Dierckx) P. citrinum (Thorn) P. bialowiezense (Zal.) P. dupontid Penicillium sp. Pithomyces chartarum (1) P. chartarum (2) Sclerotinia sclerotiorum Sc/erotium rolfsii Sporotrichum thermophiled S. exile Stemphylium dendriticum Stilbella thermophila Stilbella sp. Trichoderma viride T. viride T. viride Trichosporan cutaneum Trichothecium roseum BASIDIOMYCETES Clitocybe iIIudens Tilletia controversia

Total Lipid (% dry wt.)

Reference

6.8-17.0 7.2-19.0 24.4

126 49 50

15.7-23.2 6.0-47.3

49 58

9.6 (rnean) 20.2 29.7-34.8 16.0 7.9-19.4 5.4 3.5-15.2 17.5-41.5

120 50 126 126 53 126 49 50

26.0-28.0

50

22.9

50

18.4

50

18.1

50

17.0

50

14.8 14.5-15.8 4.6 4.1 1.1-12.4 2.8 15.5 9.5 2.4 38.1 17.0 4.8-7.6 13.3-24.0 4.4-11.2 45.0-56.0 8.1-17.0

19 133 114 114 135 57 19 19 114 19 19 115 126 133 128 125

9.1 5.8

69 4

Introduction to Fungal Lipids

10

TABLE 1.2-continued Species

T. nudum U stilago zeae U. zeae U. zeae (PRL 119)

Total Lipid (°'0 dry wt.)

Reference

0.2-47.2 30.2-36.6 7.0-27.8 10.0

123 49 126 127

Ten races. Twenty-four races. , Many isolates. " Thermophilie. a b

The lipid content of vegetative hyphae varies between 1 and 50 % of the dry weight depending on the species, developmental stage of growth, and cultural conditions. Although many fungi have a high capacity for lipid production, most mycelial species contain between 6 and 9 % lipid when grown under favorable conditions. Considerable variation is present in the lipid content of different species of the same genera and even strains (or isolates) of the same species when cultured under identical conditions. This is illustrated in Table 1.3 where the total lipid abundances of several Fusarium species (and two strains of F. lini) grown in several media are compared. As much as 100% variation in lipid content can occur by growing the fungus on different media which are considered good for fungal growth. TABLE 1.3. Comparison ofthe Total Lipid Abundances of Fungi Grown on Different Mediaa Media b., Fungal Species U stilago zeae Fusarium /ini" Fusarium oxysporum Fusarium /ini" Fusarium graminearum

A

B

C

D

7.02 5.53 7.74 6.77 10.3

18.33 28.36 13.71

27.83 26.35 13.90

16.66 15.00 24.44 32.22 24.44

24.07

This comparison is from Hunter and Rose.(3) The article by Woodbine et al.(49) should be consulted for the exact composition of each of the above media preparations. Each media preparation varied in carbon source concentration and inorganic composition. , All values are reported as the percent of total lipid per gram dry weight of mycelia. "Two different isolates of F. /ini.

a

b

1.2. Total Lipids

11

Yeast lipids are the subject of a review by Hunter and Rose.(3) The total lipid abundances of a number of yeast species and strains are given in Table 1.4. As in the mycelial fungi, considerable variation in lipid content exists among both the yeast species and strains. The lipid content of most yeast fungi ranges between 7 and 15°1" of the dry cell weight. There are several species known as the '"fat yeasts" which produce lipids representing between 30 and over 60 %of their dry cell weight. Rhodotorula and Lipomyces species fall into this group and seem to have the greatest potential for lipid production among the fungi. TABLE 1.4. Total Lipid Contents of Selected Yeast Fungi (Mycelia) Species

Blastomyces dermatitidis Candida albieans (A TeC 10231) C. alhicans 1 Ha 582 (yeastlike) C. lipolytica C. seol/ii AL 25 C. seol/ii 5AAP2 Candida sp. # 5 C. utilis Histoplasma capsulatum Lipomyces starke)'i Pul/ularia pul/ulans Rhodotorula glutinis R. gracilis R. gracilis Rhodotorula sp. Saccharomyees cerevisiae (ATCC 7754) S. cerevisiae (A Tee 7755) S. cereL"isiae Saceharomyces sp. Baker's yeast "Soil yeast" Torulopsis utilis Torulopsis sp. Yeast strain 72

Total Lipid (° 0 dry wt.)

Reference

5.0 0.3-6.3 13.9 6.6-8.5 8.2 10.7 7.8 0.3-D.5 12-18 7.7-31.4 11.0 12 20.3-63.2 43 49.8 68.5-87.1 7.0-10.2 3.3-10.2 17.0 6.0 5.5-65.3 3.0 6.4 25.0-33.0

99 67 130 17 17 17 17 129 99 61 116 17 117 118 159 117-119 117-119 122 111 161 71 105 160 162

The functions of lipids are not weil understood. Historically, lipids have been considered as reserve material which may be converted to energy and carbon skeletons during growth and reproduction. This is true for those lipids which accumulate in globules (sphaerosomes, liposomes) that

72

Introduction to Fungal Lipids

are composed primarily of triacylglycerides. However, as we learn more about the structure and biosynthesis of less abundant lipids, it is becoming apparent that they have more specific roles in cellular growth and reproduction. For example, lipids are essential components ofthe membrane structure and function (transport), lipids may act as stimulators of growth and reproduction, or they may serve as protective coatings. The functions of each lipid dass will be discussed in more detail in the respective chapters of this book.

1 .2.2. Spore and Sclerotial Lipids Fungal spore total lipid abundances vary considerably, depending on the species, but range between 1 and 35 % of the spore dry weight (Table 1.5). The ability of spores to germinate without an exogenous carbon source appears to be related to the abundance of reserve lipid. TABLE 1.5. Total Lipid Contents of Fungal Spores Species

Total Lipid (%drywt.)

Reference

2.65 10.4-16.1 3.7 7.6 4.1 8.4 11.3-19.4 16.1-19.3 9.8

8 5 5 5 5 5 5 5 90

PHYCOMYCETES

Rhizopus arrhizus Rhizopus sp. Mucor mucedo M. ramannianus M. racemosus M. hiema/is M. miehei M. pusillus M. rouxii ASCOMYCETES

Pithomyces chartarum Trichoderma viride Candida albicans" N eurospora crassa Lindegren Sphaerotheca humili var.fu/iginia b Erysiphe graminis

1.4 9.6 20 19.0 10 12

114 115 102 6 139 139

22.0 35.0 20 4.1 12.5 10.2 9.5

136 137 139 138 139 139 139

BASIDIOMYCETES

Flax rust (Melamspora /ini) Tilletia controversac T·foetens· Carnartium fusiforme Cronartium harknessii Uromyces psoraleaee Phragmidium speceosumc

1.2. Total Lipids

13

TABLE 1.5-continued Species

Puccinia hieracii d P. hieracii' P. hefianthi' P. carthami' P. graminis d var. tritici 56 P. graminisd avenae 7A P. triticina d (mixed race) P. CorOl1ata d P. graminis tritici (mixed races) M etampsora fini d 111. medusae d Ustilago zeaeG U. triticiG U. nigra G U. teds' U. bullata' U. maydise Ravillefia hohsolli' Gymllosporallgium juvenescell:f

Total Lipid

(% dry 8.4 7.3

13.3 9.0

18 16 17 17 19.7 14.6 14.6

Reference

wt.)

139 139 139 139 139 139 139 139 141

139 139

22

139

5

139 139 139

4

14.5 0.6 0.4 18.8 6.3

140

140

139 139

Chlamydospore. hConidia. , Teliospores. d Uredospores. e Aeciospores. f Dried gelatinous horns. G

Spores of the rust and smut fungi, which cause diseases of economic crops, seem to contain the greatest abundances of lipid, generally having over 10 % total lipid. Teliospores of the smut fungus Tilletia controversa contain 35 % lipid.(4) With few exceptions, spores produced by other fungi contain less than 10 % lipid. Variation in the total lipid abundances is also present among species of the same genus. For example, chlamydospores of U stilago species range between 4 and 22 % lipid content and similar variations are found among spores of Puccinia and Mucor species. Sumner and Morgan(5) compared the total lipid abundances of spores produced by mesophilic and thermophilie (and thermotolerant) Phycomycetes. They found that the species preferring higher growth temperatures contain higher total lipid levels (10.4 to 19.4 %) in their spores, while those preferring ambient temperatures contain lower lipid levels (3.7 to 8.4 %).

14

Introduction to Fungal Lipids

The spores were found to contain less lipid than the mycelia from which they were harvested. Not all of the spore lipid is ｲ･ｳｴｩ｣ｾ､＠ to lipid globules suspended in the sporoplasm. As in the mycelia and yeast cells, spore lipids are an integral part of the membrane structure. Als0, nonglobule lipids are not randomly distributed throughout the spore, and sporoplasm and spore wall lipid distributions differ (see Chapter 3). Conidia of N eurospora contain 19 % lipid, 95 % of which is phospholipid and carotenoids.(6) The major change in lipid content is found in ungerminated conidia. Presumably this is due to metabolie changes during the aging process. No decrease in lipid content is found during germination, and the principal energy source appears to be the sugar, trehalose.

1.3. LIPIDS AND FUNGAL GROWTH Two patterns of lipid turnover are found in fungi. In some species, the growth cycle is characterized by a rapid synthesis and accumulation phase during logarithmic growth followed by a utilization phase during reproductive and stationary growth (Fig. l.l.a). This general pattern of lipid turnover has been reported for Phycomyces blakesleeanus,(7) Rhizopus arrhizus,(8,9) Aspergillus nidulans,(IO) and Mortierella vinaceaY I) A similar pattern has also been reported for the yeast Candida utilis.(12) The utilization of fat in the later stages of growth may be due to the depletion of certain nutrients in the medium, as shown for Penicillium species,o 3) or to metabolie alterations which occur during the change from vegetative to reproductive growth. On the other hand, some species te nd to accumulate fat as the mycelia ages. Lipids are found as fat globules (sphaerosomes or liposomes) in aging mycelia or mycelia grown under abnormal conditions. The lipid content of Pythium ultimum during 15 days of growth ranges between 3 % at 5 days to 48 % of the dry mycelia at the end of the growth period (Fig. 1.1.b).(14) This also appears to be true for several yeast fungi. The lipid content of Saccharomyces cerevisiae AICC 7755 decreases during the lag phase and the first half of logarithmic growth and then progressively increases during the second half of exponential and post exponential growth. No correlation between respiration and total lipid content has been made. The change in total lipid content of too few fungal species has been determined to know if they can be grouped according to their tendency to utilize or accumulate lipids during the final stages of growth. It is doubtful, however, that this will turn out to be the case, since the growth cycle is influenced considerably by the media composition and environmental conditions of growth.

15

1.3. Lipids and Fungal Growth

15 14 13 12 11 10 9 a

8 7

6

5 4

3 2 (f)

0 Q.

0

Ｚｊｾ＠

2

3

4

5

6

7

Ｎｊｾ＠

;:!

0

I-

50 ·10 30 b

20 10

0

5

10

15

TIME (DAYSI Fig. 1.1. (a) Production of lipid by Rhizopus arrhizus during 7 days of growth.(S) (b) Production of lipid by Pythium ultimum during 15 days of growth.(14)

76

Introduction to FungBI Lipids

It is apparent that culture age is an important factor in determining the total lipid content offungi. Not only the total abundances oflipid change with age, but significant changes in the relative composition of individual lipid c1asses also occur.

1.4. CULTURAL CONDITIONS FOR LIPID PRODUCTION The metabolie activities of all Iiving organisms are influenced by the environment in which they grow, and these activities are reflected in the overall growth characteristics of the organism. Such cultural parameters as temperature, carbon source, inorganic nutrients, pH, aeration, and vitamins are discussed below as they re la te to the growth and total lipid content of fungi.

1.4.1. Temperature Fungi differ in their optimum growth temperatures, and, since total lipid abundances are influenced by culture temperature, this growth characteristic should be determined for a species or isolate before meaningful studies are carried out. Comparisons ofthe lipid content offungi as a function of culture temperature must be based on the lipid levels produced at the optimum growth temperatures. Fungi are generally c1assified into three groups according to their optimum growth temperature requirements. Mesophilic fungi have growth temperature optima between 25 and 33°C and psychrophilic fungi range between 10 and 20°C in growth temperature optima. Thermophilie fungi are defined on the basis of maximum and minimum growth temperaures, i.e., maximum temperature for growth at or above 50°C and minimum at or below 20°c.(15) Thermotolerant is a term used to designate fungi that can grow at maxima near 50°C but minima below 20°C. There are several reports on the influence of growth temperature on the lipid content of fungi, and, as shown in Table 1.6, species differ in their response to variations in growth temperatures. Lowering the growth temperature of Rhodotorula gracilis from 28°C to 22°C results in a reduction by greater than half in total lipid produced. ( 6 ) Other studies involving yeasts showed that decreasing growth temperatures results in increased lipid content,ll 7,18) which may be attributed to the reduced growth rate associated with the lower temperaturesY8) It is difficult to generalize, based on the reported studies, but it appears that increasing growth temperatures are accompanied by increasing lipid levels. This is probably true only within certain temperature ranges, which may differ depending on the species.

7.4. Cultural Conditions for Lipid Production

77

The total lipid eontents of closely related mesophilie and thermophilie fungi are eompared and range between 8 and 54 % of the fungus dry weight (Table 1.2).(19) Although there is a wide range of lipid abundanees in these fungi, most speeies range between 8 and 18 %, and no signifieant differenees are noted among these fungi having different growth temperature optima. Sumner and Morgan,(5) on the other hand, found that the lipid eontent of spores and myeelia of mesophilie Mucorales fungi is lower than that of closely related thermotolerant and thermophilie species. Growth temperatures have a more striking and consistent influence on the degree of lipid unsaturation. A relatively high degree of unsaturation is found in lipids from organisms grown under low-temperature conditions. This has been observed in higher plants and animals,(20-23) insects,(24) and certain microorganisms.(2S-29) Since increasing unsaturation alters the physical properties oflipids, particularly melting point, it has been postulated that this phenomenon represents a factor important to adaptation to cold environments. It follows that the order of increasing unsaturation in lipids from fungi which have different temperature optima should be psychrophile > mesophile > thermophile. It should also follow that if these organisms are grown at temperatures at either extreme of their optima, the degree of unsaturation adjusts accordingly. Studies with Asperigillus niger and Rhizopus nigricans, (28) Rhodotorula gracilis,(30) and Aspergillus nidulans(31) ha ve shown that increases in the degree of lipid unsaturation do oceur at reduced temperatures, while the lipids of Saccharomyces cerevisiae(32) beeome more saturated when grown at elevated temperatures. Similar inereases in the degree ofunsaturation were not obtained in studies with Pyhtium ultimum,04) Cunninghamella blakesleeana and Rhizopus arrhizus.(9) The influenee of growth temperature on the degree of unsaturation in several fungal speeies is shown in Table 1.7. Mumma et al.(19) compared the degree ofunsaturation in lipids extraeted from mesophilic and thermophilie fungi grown at their temperature optima. They found that the lipids of mesophilie fungi eontain the highest degree of unsaturation at 0.96 to 1.60 double bonds per mole as eompared to that of thermophiles at 0.65 to 1.01 double bonds per mole (see Table 1.7 for method of ealculating Mmole). Sumner and Morgan(S) reported a similar degree of unsaturation, which appears to be typical of most fungi, in the lipids of several mesophilic Mucor species whieh grow best at moderate temperatures (Table 1.7). They also reported that the degrees of unsaturation in lipids of mesophilic and psychrophilie fungi are similar, while that of thermotolerant fungi grown at 28°C is not significantly different.(33) Factors other than culture temperature seem to influence the degree oflipid unsaturation during growth, since lipids of younger cultures are more unsaturated at higher tempera tures.

Cunninghamella blakesleeanus Rhizopus arrhizus R. species b R. species BI d Pythium irregulareO P. vexans· P. ultimum M ueor miehei b M. pusil/us M. strie/us' (CBS 100.66)

PHYCOMYCETES

Fungus

16.9

10

15

21.5

16

20

12.6 8.6

22

14.2

24

24.3 26.2

25.7

3.1

25

13.9 5.9

27

28

48

4.1 19.5

30

Growth Temperature (0C)

14.9 11.3

32

8.8

36

5.0

37

3.4

38

I\.9 23.1

I\.6 13.1

48

25-30 25-30 20

37 30

(0C)

33

9 9 5 33 75 75 14 5 5

Optimum Reference Growth Temp.

TABLE 1.6. Effect of Culture Temperature on the Growth and Lipid Production in Fungi a

ｾ＠

-6'

,..

-

ｾ＠ =:! 'g

Cl

(')

:;-

ä ｾ＠ ...0' =:! ...

Co

-

16.0 10.3

20.0

10.8

14.4

2 \.4

16.2

8.5

24.4

19.8

6.6 12.5

22.6 19.4 7.8 35.9 8.4 I\.9

25

3

17

33 33 33 33

33

33

33

a

Values are expressed as a pereentage of the fungal dry weight. h pH was also a parameter in this study, but had little effeet on myeelial weight produeed. Cultures grown at pH 6.8 were seleeted for this table. Effeet of pH on lipids will be diseussed below. 'Thermotolerant of thermophilie fungi. d Psyehrophilie fungi. ,. Thermophilie fungi.

Candida lipolytica Saccharomyces cerevisiae

ASCOMYCETES

M. strictus' (CBS 576.66) M. strictus' (CBS 575.66) M. oblongisporus' (CBS 173.27) M. oblongisporus' (CBS 220.29) Mucor sp. (I)d M ucor sp. (lI)d Mucor sp. (lII)d

c:: c:: ii1 .....

...o·

10

...

::J

...o·

ｾ＠ I')

()

ｾ＠

Q:

.... ,... 'tj.

::J

.c;-

ｾ＠

::J

()

C')

;;-

C')

ｾ＠

...

Introduction to Fungal Lipids

20

TAßlE 1.7. The Effect of Growth Temperature on the Degree of Unsaturation in Fungal lipids Fungi

U nsaturation L'l./mole"

Temp.

1.16-1.47 1.33-1.50 1.23-1.28 0.96-1.29 0.95 0.76 1.02 0.93 0.83 0.96 1.13-1.26 0.98-1.24 1.03-1.18 0.94-1.04 0.99-1.19 0.89-1.06 0.95-1.12 0.84-1.07 1.00-1.29 1.17-1.37 0.96-1.04 0.91-1.04 0.96-1.06 0.79-1.00 0.93-D.98 0.76 0.76-D.84 0.6O-D.86 1.02-1.22 0.8O-D.95 0.99-1.24 0.94-1.29 1.22 0.94 0.85 0.93 0.69 1.29 0.83 0.82

25 25 25 25 25 48 25 48 45 25 10 20 10 20 10 20 10 25 10 25 28 48 28 48 28 48 36 48 28 48 28 48 25 48 15 20 25 30 20 30

1.00 1.38

25 10

Reference

(0C)

PHYCOMYCETES

Mucor mucedo b M. ramannianus b M. racemosus b M. hiemalis b M. miehei M. pusillus M. pusillusd M. globosus b 1'.1. strictus' (eBS 100.66) M. slriclus' (eBS 576.66) M. strictus' (eBS 575.66) M. oblongisporus' (eBS 173.27) M.oblongisporus' (eBS 220.29) Mucor sp. Jd

Mucor sp. II d M. sp. m d (Mucor miehei) Rhizopus sp. IIJ d

Rhizopus sp.

Je

Rhizopus sp. IIe Rhizopus sp. R. arrhizus

PYlhium ultimum

5 5 5 5 5

19 19 33 33 33 33 33 33 33 33 33 33 33 5 57

32

ASCOMYCETES

Candida lipolyticab Candida sp. (# 5)'

17 17

1.4. Cultural Conditions for Lipid Production

21

TABLE 1.7-continued Fungi

C. scoltii (AL25) C. scoltii (5AAP 2 ) C. lipolytica b C. lipolYlica C. ulilisb,} NCYC 321 C. utilis b.• NCYC 321 C. utilis b.! NCYC 321 C. urilis NCYC 321 Chaelomium thermophiled C. globosum b Rhodotorula glubinis b Humicola grisea d var. thermoidea H. insolensd H. lanuginosa d H. grisea b H. nigrescensb H. brevis b M. pulchellab Penicillil1m dllPOl1ti d P. chrysogenum b Sporotrichum thermophile S. exile b Stilbella thermophilad Stilbella Sp.b Saccharomyces cerel'isiae S. ceret'isiae Sclerotinia sc/erot;orum (14 day old) S. sclerotiorum (sclerotia) S. sclerotiorum (sclerotial exudate)

t.jmole = 1.0 x (% monoene)!100 Mesophile. , Psychrophile. d Thermophile. e Thermotolerant. f 75 mm Hg0 2 tension. • 1 mm Hg0 2 tension. a

+

2.0 x

Unsaturation t.jmole"

Temp.

1.72 1.72 0.86 1.12 1.40 1.27 1.51 1.08 0.65 0.96 0.63 0.97 1.01 0.82 1.54 1.34 1.17 1.27 0.86 1.60 1.00 1.27 0.56 1.47 calc. calc. 2.09 1.66 0.44

10 10 25 10 30 30 15 15 45 25 25 45 45 45 25 25 25 25 45 25 45 25 45 25 26 40 5 20 20

CC)

Reference

17 17 17 17 142 142 142 142 19 19 17 19 19 19 19 19 19 19 19 19 19 19 19 19 32 32 135 163 163

(% dienes)/I00 + 3.0 x (% trienes)/100.

b

When representative mesophilie, psyehrophilie, thermo tolerant, and thermophilie speeies are grown at some temperature other than their optimum for growth, the expeeted ehanges in the degree oflipid unsaturation oeeur, i.e., a deerease in temperature is aeeompanied by an inerease in the degree of lipid unsaturation. For example, lowering the growth temperature from 48 to 25°C of the thermotolerant Rhizopus sp. and the thermophilie

22

Introduction to Fungal Lipids

Mueor miehei and M. pusil/us results in an increase ofboth spore and mycelial lipid unsaturation.(5) Similar results are obtained when the lipids of the mesophilic Candida lipolytiea and psychrophilic C. seottii grown at 25° and 10°C are compared.(17) It is also noted that the psychrophilic yeasts are characterized by a high degree oflipid unsaturation which remains relatively constant during the growth cycle. The effects of temperature on the degree of lipid unsaturation in an organism may be exerted in its influence on the oxygen tension of the media. (Effects of aeration on lipid production will be discussed in more detail below.) It is weil known that oxygen is required during the dehydrogenation of fatty acids (see Chapter 4), and factors influencing its a vailability certainly affect the rate of desaturation. Brown and Rose(34) pointed out that the effects of temperature on the fatty-acid composition of cells may be due to changes in the metabolie balance rather than a specific effect. Meyer and Bloch(35) suggested that the effect of temperature is primarily on synthesis of the desaturase enzyme. As an explanation for a higher degree of unsaturation at lower temperatures, Kates and Baxter( 1 7) proposed that the rates of synthesis and degradation of unsaturated fatty acids are both temperature-dependent and that these rates have different temperature coefficients. They assumed that at lower temperatures the synthesis of unsaturated acids is retarded less than degradation, so that under these conditions linoleic acid accumulates with a consequent increase in the degree of unsaturation.

1 .4.2. Carbon Source Carbohydrates are the best carbon substrates for fungal growth, and, in 1878, Naegeli and Loew(36) were the first to show that these compounds can be converted to lipid by yeast. Smedley-MacLean(37) has also shown that reserve carbohydrates of yeast are converted to lipid in aerated cultures. Numerous studies have been carried out to survey microorganisms as primary sources of lipid to supplement the rapidly depleting world food supply, and, in this connection, severallaboratories have conducted searches for fat-producing fungi.(38-42) The potential of an organism to be an economic producer of fat is related to its ability to efficiently utilize the available carbon substrate, particularly industrial waste products. According to Rippel(43) and others,(44-48) the maximum carbohydrate utilization efficiency by fungi appears to be between 15 and 18 % of the available sugar in the media. A number offungal species, particularly ofthe genera Penieillium, Aspergillus, and Fusarium, have been examined for their ability to utilize various monoand disaccharides., Glucose is the most commonly used carbon substrate

7.4. Cultural Conditions tor Lipid Production

23

for fungi grown for experimental purposes, and it is efficiently converted to lipid by a number of species. Woodbine ef al.(49) found that, of the 40 strains examined, A. jlavipes and F. lini produce the greatest amount of fat from glucose at 39.7 and 34.6 % of the dry mycelial weight, respectively. Fungal species also appear to vary considerably in the optimum glucose concentration required for the greatest yield and efficiency ofutilization. Ward ef al.(SO) surveyed the fat accumulation of 60 fungi grown on glucose and found that Penicillium javanicum van Beijma yields the highest lipid content at 41.5 %. Forty percent glucose in the media gave the highest yields offat in the fungi studied. Prill et al.(S!) studied the effect of glucose concentrations of 1 to on the fat content of A. fischeri and found that lipid abundances 70 increase almost linearly from 10.4 to 36.0 % with increasing glucose concentrations. Gaby et al.(S2) found that glucose is more readily oxidized by P. chrysogenum than other substrates, and Gregory and Woodbine(53) found that reducing the glucose in the media by half lowers the lipid yield by A. nidulans and P. spincilosum but not P. javanicum. Several comparative studies have been conducted to determine the best carbohydrate carbon source for fat production by fungi. Chesters and Peberdy(!l) found that fungi grown with glucose or maltose produce the highest fat yields. The same is true for P. lilacinum(54) and A. nidulans.(55) Both sucrose and fructose are excellent carbon sources for fat production by P. chrysogenum (sucrose :> fructose> glucose)(S6) and Rhizopus arrhizus (fructose> sucrose).(57) In a study to evaluate the efficiency of sucrose as a carbon source for fungal growth and fat production, maximal fat content was found at concentrations of 22.5 %. In this study, P. soppi and F. lini(!) had the highest fat content at 34.8 and 28.4 %, respectively. Osman et al.(58) also found that sucrose is the best carbon substrate for fat production by P. lilacinum at concentrations of 17 %. Gad and Hassen(59) found sucrose to be the best carbon substrate for fat production by A. fisheri, and Singh and Singh(60) found the same to be true for P. aurantio-brunneum at a 40 % sucrose concentration. Although lactose is used as the carbon substrate for the superior fat-producing strains of Lipomyces starkeyi and L. lipojer, sucrose > fructose. Other compounds, particularly hydrocarbons, have been evaluated for their value as sole carbon substrates for fungal growth, but little concern has been given to their value as efficient substrates for fat production. In one such study, when Candida lipolytica was grown on hydrocarbons as the sole carbon source, the lipid content was increased.(62.63) ljhe oxidation of hydrocarbons by fungi is discussed in Chapter 2. %

24

Introduction to Fungal Lipids

1.4.3. Inorganic Nutrients Defined media used for the laboratory culture of fungi are composed of a carbon source and a mixture of mineral salts which generally inc1ude as major elements calcium, magnesium, potassium, phosphorous, a nitrogen source, and a few minor elements (iron, manganese, and zinc). The factors influencing lipid production by fungi have been reviewed by Woodbine,(64) and it appears that the importance of the amount and composition of mineral salts in fungal growth media is not well understood. Stein berg and Ordal(16) reported that the addition of Ca, Na, and Fe to the growth media is not necessary for fat production in yeast. Phosphate seems to influence lipid accumulation, since S. cerevisiae growp in a phosphate-deficient medium contains less lipid than cells grown in a phosphate-sufficient medium.(65,66) Increasing NaCl concentrations from 0 to 10% in the medium results in almost linear increases in lipid content from 0.32 to 6.29 %.(67) Similar results are obtained with S. cerevisiae(68) and Endomyces vernalis. Other than the carbon substrate, nitrogen has the most pronounced effect on lipid production by fungi. Fungi vary as to the best nitrogen source for growth, but inorganic nitrogen in its most reduced form, NH;, is more efficiently utilized by a greater number of fungi than other nitrogen forms. Prill et al.(51) compared the effects of several NH 4N0 3 concentrations (0.2 to 10 %) on the growth and lipid content of A. fischeri and found the greatest lipid content and mycelia (per gram of glucose utilized) are produced at the 0.2 % nitrogen concentration. However, the greatest amount of glucose is extracted from the medium at 2 % NH 4N0 3 . In some species, organic nitrogen is best for mycelial growth, but is inferior to the inorganic form for fat production. Some exceptions do occur, however, since A. nidulans contains higher lipid levels when grown with asparagine as the nitrogen source than when grown with nitrate or ammonium salts.(70) Asparagine is second to (NH 4hHP0 4 for lipid production in M ortierella vinacea.( 11) Glycine is a more efficient nitrogen source than NaN0 3 for fat production by P. lilacinum. Other organic and inorganic nitrogen sources have proved less efficient for fat production. The most important nutritional parameter for lipid production by fungi is the carbon: nitrogen (C: N) ratio in the media. Increasing the carbon and nitrogen content of the media while holding the C: N ratio constant results in increased mycelial production, but not fat accumulation. In Aspergillus and Penicillium species, low C: N ratios favor protein synthesis and high C: N ratios favor fat production when NH 4N0 3 is used as the nitrogen source. The optimum by P. lilacinum is 65: 1 at NaN0 3 concentrations of 0.65°1".(58) A carbon : nitrogen ratio of 80: 1 was found to be best for the growth and fat production by M. vinacea grown on (NH4hHP04

1.4. Cultural Conditions for Lipid Production

25

as the nitrogen source. On the other hand, certain unidentified yeasts isolated from the soil by Starkey(71) contained the highest lipid content when grown on nitrogen-deficient media. Fat production can be regulated in fungi by altering the C: N ratio of the medium. Rhodotorula gracilis can be induced to produce lipid concentrations, having a fat coefficient of 15.0 to 21.0°;", up to 60°;" ofthe dry cell weight.(4S.72-74)

1.4.4. pH During fungal growth, the pH of the culture media changes, and the degree of change is dependent on the media composition. Hence, it is difficult to draw conclusions concerning the effect of pH on fungal growth and fat production. Optimum pH values for the growth of most fungi are between 6.0 and 7.0. In unbuffered media, the pH level falls with increasing age of the culture until a value is reached which results in growth retardation. When ammonium salts of mineral acids are used as the nitrogen source, pH lowering is caused by the removal of NR;'" from the media during growth. Prill er al.(Sl) studied the effect of acidity and alkalinity of the culture media on growth and fat accumulation by A. fischer i as a function of the nitrogen source. Controls included buffered media using calcium carbonate and potassium acetate. Generally, lipid accumulation, utilization of glucose, and, in so me ca ses, growth were retarded in unbuffered media. Mycelia grown in buffered media contained a higher content of fat and showed a more efficient utilization of glucose. In mycelia (A. fischeri) grown in media having an initial low pH of 2.0 to 4.6, the fat content remained relatively constant during growth at levels between 17.0 and 19.5 '%; of the dry mycelial weight. At initial pH levels between 6.0 and 8.0, mycelial fat abundances were considerably higher (23.9 to 37.0 %) at the end of the growth period. Torulopsis /ipofera contained the greatest fat abundances when grown in media having a pH of 5.5 to 6.0. The rate oflipid accumulation by Rhodotorula graci/is increased linearly between pH 3.0 and 8.5 and increased from 2.1 to 3.1 gof fat per 100 gof the yeast per hOUr.(16) Cantrell and Dowler!7S) compared the effects of temperature and pH on the growth and lipid composition of Pythium vexans and P. irregulare. They reported that at a given temperature the best pH for lipid production va ries with the species, but at high pH va lues (pH 5.9, 6.8, and 7.5) no variations in lipid content are observed. There are several reports that there is little correlation between lipid production by fungi and pH ofthe growth media. For example, no differences in fat accumulation were noted in P. lilacinum when grown at pH values of 4.0 to 8.0. Maximum mycelial weights for P. lilacinum are produced at pH

26

Introduction to Fungal Lipids

4.0, and the best pH values for fat accumulation are from 4.0 to 6.8. Similar

results have been reported by other investigators.(76.77) Castelli et al.(78) studied the effect of pH and CO 2 concentration on the lipid content of Saccharomyces cerevisiae and reported that, at a constant pH of 5.5, a threefold increase in bicarbonate concentration and pC0 2 causes a 27 % increase in total lipid content. If the pH level is 6.0 and the pC0 2 is held constant by the addition of bicarbonate, no significant increase in the total lipid occurs.

1 .4.5. Aeration The method of culturing is an important factor in the production of lipids by fungi. Several techniques are commonly employed and include still or shake cultures in different types of flasks, semisolid agar medium in petri dishes, batch cultures, and others. The degree of aeration differs for each of these methods. There are differing opinions on the importance of culture aeration for lipid production. Prill et al.(51) reported that aerated cultures of A. fischer i grow more rapidly and utilize most of the available glucose by the end of the growth period. Nonaerated cultures do not grow as fast and do not use as much ofthe available carbon substrate. However, no significant differences in the amount or nature offat was noted in the aerated and nonaerated fungi. Other in vestiga tors ha ve reported similar results. (71, 79) Enebo et al.,(44) however, reported that it is possible to obtain higher fat yields in the yeast Rhodotorula gracilis with culture aeration. In addition to mechanical agitation of the fungal hyphae, culture aeration influences the temperature and oxygen tension of the media. These environmental parameters were discussed above.

1.4.6. Vitamins Most fungi are able to grow and reproduce on a carbohydrate and mineral salts media, while others require such additives as thiamine, biotin, pyridoxine, certain purines, amino acids, and/or inositol. The relationship between lipid production and vitamin deficiency in fungi ｩｾＭ n-öi weil established, but generally, vitamin deficiencies cause reductions in lipid production. When compared to cells grown in a pyridoxine-sufficient media, a 40°;'; reduction in lipid production occurs in Hanseniaspora vallbyensis grown in pyridoxine-deficient media.(80) Phytosphingosine content is lower in the deficientcells, while lipid phosphorus is unchanged. There is also evidence that pyridoxine influences the degree ofunsaturation. Nicotinic acid deficiency causes an increase in the total lipid content, while pantothenic acid deficiency decreases it. Klein and Lipmann(81) also reported that S.

1.5. Cellular Distribution of Lipids

27

cerev/SIae grown under pantothenate-deficient conditions contains less lipid. Suomalainen and Keranen( 82 1 reported that C 18 acid formation, particularly C I8 :1, is reduced, while C I6 acid accumulation is increased. The effect ofbiotin deficiency can be explained, at least in part, by the need for this co-factor in the synthesis of co-carboxylase, an enzyme involved in the chain lengthening process in fatty-acid biosynthesis (see Chapter 4). Since the first report that inositol is required for normal growth of certain yeast strains,(83 1 several studies have been conducted to determine its metabolic role. Johnston and Paltauf(8 4 1 have summarized the effects of inositol deficiency on Saccharornyces species: (1) growth retardation and cell aggregation, (2) reduction in respiration, (3) reduction in cytochrome and pyridine nucleotide content, accompanied by unbalanced growth and cell death, (4) alteration of metabolic behavior resuIting in abnormal release of acetaldehyde, acetone, glycerol, and other unidentified products, (5) accumulation of glucan in the cell wall, and (6) abnormally high lipid content. Lewin(85 1analyzed the lipid content of an inositol-requiringstrain of Saccharomyces carlsbergensis ATCC 9080. When compared to cells supplemented with inositol, the vitamin-deficient cells contained an average of 36 % more lipid, most ofwhich was nonphospholipid in nature. Challinor and Daniels( 86 1 also found that certain lipids accumulate in inositol-deficient S. cerevisiae and N eurospora crassa.( 87 1 Shafai and Lewin(881 compared lipid composition of inositol-deficient cells of S. carlbergensis with normal cells and found that the tri glyceride content is higher in the vitamin-deficient cells. Not only is the concentration of inositol-containing phosphatides low, but the principal phospholipids, phosphatidylcholine and phosphatidylethanolamine, are also significantly lower in the inositol-deficient cells when compared to normal cells. PaItauf and Johnston(89 1 reported similar results with respect to the triglycerides in vitamin-deficient cells, but with the exception of phosphatidylinositides, the concentrations of glycerophosphatides are unaItered. The exact role of inositol in fungal lipid metabolism, other than as a constituent of phosphatidylinositol, is uncertain.

1.5. CELLULAR DISTRIBUTION OF LIPIDS 1 .5.1. Cell Wall The structure and chemical composition of the fungal cell wall has been extensively studied, and the subject has been reviewed by BartnickiGarcia.( 901 Except during the developmental stages of a certain species, all fungi are enclosed by a characteristic cell wall which distinguishes them from all other organisms. AIthough both qualitative and quantitative changes

28

Introduction to Fungal Lipids

occur during fungal morphogenesis, the fungal cell wall is composed of 80 to 90 % polysaccharides with the remainder consisting of lipid and protein. In 1964, Dyke(91) was the first to show that the lipid found in the iso la ted cell walls of N adsonia elongata is a true constituent and not a cytoplasmic contaminant, yet the function of cell-wall lipid remains unknown. Notable differences are found in the hyphal, sporangiophore, and spore wall chemistry. Total cell-wall lipid abundances for several fungal species are given in Table 1.8. TABLE 1.8. Fungal Cell Wall Lipids Fungal Species

Lipid"

Reference

( ＧｪｾＩ＠ PHYCOMYCETES

Mucor rouxii (yeast) M. rouxii (hyphae) M. rouxii (sporangiophore) Pythium butleri P. myriotylum Pythium sp. PRL 2142 Apodachlya brachynema Dictyuchus sterile Saprolegnia ferax

5.7 7.8 4.8 12.0 8.0 8.2 3.1 4.7 5.0

90 90 144-145 152 152 146 146 146 146

ASCOMYCETES

Saccharomyces cererisiae (Baker's yeast) S. cerel'isiae (18.29)

S. cerevisiae S. cerevisiae (Baker's yeast) S.fragilis CECT 1207 S. cerevisiae (Baker's yeast) S. cerevisiae (Baker's yeast) S. carlsbergensis S. oviformis Candida albicans 806 C. albicans RM 806 C. albicans 582 C. albicans (yeast, blastospore) C. albicans (mycelial) C. albicans (juvenile) C. albicans (yeast) c. albicans (mycelial) Aspergillus nidulans (wild 13) A. nidulans A. nidulans (mutant, 13 I.OL.) Aspergillus sp. Cordyceps militaris

8.1 10.1 1.0 3.1 3.1-3.3 8.5 13.5 20) 19.2

0.9

t:J

...

ｾ＠

..,

§

So CD

S'

::s

S:c:: ...Q'

;;;'

ｾ＠

tQ

ｾ＠

78

Fatty Acids

blakesleeanus.ol ) ,t-Linolenic acid has been detected in a number of higherand lower-plant species and is the predominant isomer produced by higher animals. Based on the apparent differences in the mode of polyenoic fattyacid formation, Shaw(4) suggested that the lower fungi are of different phylogenetic origin from the higher (Ascomycetes and Basidiomycetes) fungi. However, since both isomers have been detected in the same Phycomycete species and since Je-linolenic acid has been detected in the imperfect Hyphomycete, Dactylaria ampulliforme,02) Sumner(13) suggested that this hypothesis is invalid. Bartnicki-Garcia(14) has discussed what appears to be valid phylogenetic relationships among the fungi based on other biochemical characters such as cell-wall chemistry, lysine biosynthesis, and sedimentation characteristics of tryptophan biosynthesis enzymes. Phycomycete fungi do, however, differ in their fatty-acid distributions, and these differences seem to follow taxonomic lines. Fatty acids of higher (Zygomycetes) and lower (Chytridomycetes and Oomycetes) Phycomycete fungi differ in their degree of unsaturation. The lower fungi appear to possess the greatest potential for long-chain polyunsaturated (> C I8 :3) fatty-acid synthesis. Although relatively few lower fungi have been examined, longchain polyenoic fatty acids were found in almost every species investigated. For example, the Chytridiales species, Dermocystidium sp., contains 24 % C 20 :2, 5 % C 20 :S (0)3), and lower relative concentrations of other long-chain polyunsaturated fatty acids. oS ) Fungal species belonging to the dass Oomycete also produce long-chain polyenoic fatty acids. Pythiaceous fungi, belonging to the order Peronosporales, produce the characteristic high relative abundances of C 16 , C I8 :1, and C 18 :2 which are accompanied by saturated and unsaturated C 20 and C 22 fatty acids. Shaw(16) has tentatively identified C 20 :3 (ß 8,11,14), C 20 :4 (ß S ,8,II,14), and C 22 :3 (ß 7 ,IO,13) as products of Pythium debaryanum. Other aquatic fungi have been examined for their fatty-acid composition, and each species analyzed possesses distinct fattyacid distributions. O 7) Rhizidiomyces apophysatus produce a large number of fatty acids ranging in carbon chain length from C 13 to C 20 and are characteristic in that they produce mono-, di-, and triunsaturated isomers of almost every chain length detected. Rhizophylyctis rosea contain 74 % oleic acid. The distribution of fatty acids in the Zygomycete fungi is less complex than the lower fungi. Sumner(13) suggested that higher Phycomycetes can be divided into two groups based on the unsaturated fatty-acid composition. The Mucoraceae and Thamnidaceae produce polyunsaturated fatty acids up to chain lengths of C I8 with 3 double bonds (JeC I8 :3 ), while the Choanephoraceae and Entomophoraceae are capable of producing fatty acids up to the C 20 :4 level. These fungi produce greater abundances of fatty acids ha ving chain lengths of C 20 and C 22 than other higher Phycomycetes. (16,18-20)

3.3. Distribution in the Fungi

79

Sumner et alY 1) analyzed several M ucorales fungi and reported that the quantitative distribution of fatty acids varies with the optimum growth temperature. The thermophilic fungi are characterized by large relative proportions OfC 18 :1 (ca. 50%), C 18 :2 (ca. 15%), and C 18 :3 (3-5%). Thermotolerant, mesophilic, and psychrophilic fungi produce C 18 :1 in relatively smaller amounts and C 18 :2 (15-24%) and C 18 :3 (8-19%) in higher amounts. Tyrren< I 9) analyzed 17 Entomophthora species and found the most characteristic features to be the presence of C 20 :4 and high relative abundances of C 14 . He found that these species could be separated into three groups according to their fatty-acid composition. The first group includes E. virulenta, E. musca, E. thaxterantia, E. conglomerata, E. tipula, and an Entomophthora sp. isolated from aphids. This group is characterized by 40% C I8 :1, 16-19 % C I6 :1, and 12-19 % C 20 :4 . The second group, which consists of E. muscae (different isolate than above), E. obscura, and two unidentified Entomophthora species, is characterized by high relative amounts of C I8 (22-34 %) and short-chain fatty acids (20 %) such as C IO and C 12 . The third group contains high relative amounts of C I4 (26-30%) and C 18 :1 (22-25 %), as weil as tri- and pentadecanoic acids. The saturated fatty acid content of this group is 50-60 % of the total fatty acids, 85 % of this amount being acids with chain lengths of less than 16 carbon atoms. Branched-chain fatty acids are rare in the fungi. They were first identified (by GLC-retention data) as fungal products in the imperfect fungi Pithomyces chartarum, Cylindrocarpon radicicola, and Stemphylium denderiticum.(22,23) Using GLC-mass spectrometric techniques, Tyrrell(24) identified 12-methyltridecanoic (iso C I4 ), 13-methyl-tetradecanoic (anteiso C I5 ), and 14-methylpentadecanoic (iso C 16 ) acids as 35 % of the total fatty acids, collectively, in Conidiobolus denaeosporus.

3.3.3. Ascomycetes and Deuteromycetes (Imperfect Fungi) The Ascomycetes are divided into two subclasses: (1) the Hemiascomycetes, wh ich include the yeast, yeastlike fungi, and certain parasitic mycelial forms, and are generally considered the more primative of this group, and (2) the Euascomycetes, wh ich include the mycelial forms and are considered as the more advanced of the true fungi. Both the Ascomycete and Deuteromycete fungi produce C 18 :3 (w 3 ) isomer rat her than the w 6 isomer produced in the Phycomycetes. The lower Ascomycetes generally have little or no long-chain polyenoic fatty acids but tend to accumulate monoenoic acids. Saccharomyces species contain less than 1 % C I8 :3 , while other yeast or yeastlike* fungi produce * Fungi that are generally regarded as yeast hut have no known sexual stage are included in this section.

80

Fatty Acids

this fatty acid in relative abundances ranging between 0 and 28 % of the total fatty acids (Table 3.5). The most distinguishing aspect of the fatty-acid distribution in Saccharomyces species is the unusually high relative concentrations of C 16 : 1 (palmitoleic acid), which reportedly range between 26 and 60 % of the total fatty-acid fractions. The readily extractable lipids usually do not contain fatty acids with chain lengths greater than C 18' * The yeastIike fungi, such as Candida and Torulopsis, do not accumulate C 16: 1 to the high levels found in the true yeasts. Rhodotorula species also have low C 16 : 1 and C 18 : 3 levels relative to the true yeasts. Taphrina deformans, a parasite of peach trees, is the only lower mycelial Ascomycete studied for its fatty-acid composition, and it contains over 50% C 18 : 1 (Table 3.6).(16) Taking into consideration the variation in culture conditions, isolates of the same species, and analytical techniques employed, there are, with a few exceptions, no apparent significant ditTerences in the fatty-acid distribution among the major Ascomycete taxa (Table 3.6). The general fatty-acid distribution seems to be consistent with that of most organisms, i.e., C 16 is the predominant saturated acid, and C 18 : 1 and C 18 : 2 are the predominant unsaturated fatty acids. The CI8:1/CI8:2 ratio varies depending on the species and growth conditions. Except in a few cases, these fungi accumulate relative1y low concentrations of C I8 : 3 . Fatty acids having chain lengths above C 18 are not abundant in this group. Mumma et al.(25) compared the fatty-acid composition of several thermophilic and mesophilic Ascomycete and imperfect fungi (Table 3.6). The general characteristics of the fatty-acid distributions in these organisms are consistent with other fungi of this group. Palmitic acid is the predominant saturated acid, and, in Chaetomium thermophile and Stilbella thermophila, this acid accumulates to between 40 and 60 % of the total fatty acids. Oleic is the most abundant fatty acid in most thermophilic fungi, while C 18 : 2 is predominant in the mesophiles.1t was conc1uded, however, that thermophily could not be explained by ditTerences in the fatty-acid composition when compared to c10sely re1ated mesophilic species. As noted above, branched-chain fatty acids are not widely distributed in fungi, but the iso-branched isomers of C 16 and C I8 were identified (by GLC retention times) in the mycelial lipids of Penicillium pu[villorum,(26) and small amounts of branched acids (presumably iso and anteiso) were detected in the lipid extracts of P. cyaneum.(27) Hydroxy and epoxy fatty acids have been identified in extracts of Claviceps species and are discussed in Section 3.5 ofthis chapter. * See next section of this chapter for the long-chain and hydroxy fatty acids of the yeast-cell envelope.

3.4. Cellular and Extracellular Distributions

87

3.3.4. Basidiomycetes There are no apparent characteristics of the fatty-acid distributions of Basidiomycete fungi which may be used to distinguish them from the Ascomycetes and Deuteromycetes (Table 3.7). Like the other higher fungi, the Basidiomycetes produce a:-Iinolenic acid rather than the 0)6 isomer. Basidiomycetes are subdivided into two subclasses: (1) Homobasidiomycetes, which are saprophytic and include the mushrooms, puffballs, and related forms, and (2) Heterobasidiomycetes, which are parasitic and include the rusts and smuts. Shaw(28) compared the fatty-acid composition of fruiting bodies produced by three Homobasidiomycete species and found similar fatty-acid distributions except that C 20 :2 and C 20 :3 were present in Agaricus carnpestris and absent in Callybia sp. and Farnes sp. The long-chain polyunsaturated fatty acids are rarely reported for Basidiomycete fungi. Differences in the fatty-acid distribution between the stipe and pileus of Callybia sp. were found, and it was suggested that the distribution in the stipe may be intermediate in fatty-acid composition between that ofthe mycelia and pileus.(4) It was also suggested that, because of the differences in fatty-acid composition between the neutral and polar lipids, the overall fatty-acid distribution depends on the relative proportion of these lipids.(28) ResuIts obtained by Holtz and Schisler(29) on the fatty-acid analysis of neutral and polar lipids ofthe sporophores and mycelia of A. bisparus tend to support this hypothesis.

3.4. CELLULAR AND EXTRACELLULAR DISTRIBUTIONS 3.4.1. Mycelial and Yeast Cells 3.4.1.1. Cell Wall AIthough there are numerous reports on the chemistry and structure of the fungal cell wall, relatively little is known about their lipid composition. Over 40 species have been examined, and the total lipid abundances range between 0.5 and 15 % of the fungal cell wall (see Chapter 1). Because of the difficuIty in separating the plasmalemma and cell-wall components it is often difficuIt to distinguish the cell-wall lipid. Suomalainen et al. (30) and Nurminen et al. (31) found that, in addition to the ordinary cell wall, isolated cell envelopes of S. cerevisiae contain fragments of plasma membrane. Lipid constituents of the cell envelope include neutral lipids and glycerophosphatides. (32) These lipids are not constituents of the cell wall, but are contaminants from the attached membrane fragments. The lipids

(Y9OO) Candida albieansd (ATCC 10231) Candida lipolytiea e ./ Candida sp. #5 e ./ Candida scottii (AL25)"! Candida seottii (5AAP 2)'./ Candida petrophil/um' (SO-14) Debaryomyces hansenii' (NRRL Y-I448) Hanseniaspora valbyensis Lipomyces lipoferus (ATCC 10742) Pullularia pullulans' Rhodotorula glutinise •i Rhodotorula gracilis!" Rhodotorula graminis l Rhodotorula gracilism

Candida ulitis'

(NCYC 321)

Candida sp. 107" Candida ulitis b

Fungus

0.3

100 100 100 100 101

1.4

0.12 1.35 3.9 1.1

0.5

4.5 0.04 0.8

104 100 105 106 107

0.4

0.7

0.6

C I4

103 78

102

0.3

C ID _IJ

99

98

96 97

Ref.

1.1

0.7

tr

C14,1

0.4

tr

0.6

0.4

CIS

30.8 9 22.3 31.9 29.8

1.4 0.3 1.8

2.6

60.8 2.3

2.5

23.7 22.0 13.9

12 18 2 2 9.3

8.0

19 14 15 12 8.6

11.6

6.0

2 6.0

23 12.0 19.0

C16,1

C I6

0.47

4.2

tr

C I7

8.7 14 11.2 3.2 8.8

1.5 7.3

8.2

1 3 2 0.6

6.8

2.5

7

CIS

35.9

35

34 14

CIS,I

41.9 58 50.3 37.2 40.1

13.1 69.5

50.1

45 14 17 16 34.4

TAßlE 3.5. Fatty Acid Constituents of Yeast and Yeastlike Fungi

13.1 2 9.3 10.2 11.2

6.9

2.5

21 40 34 51 44.8

25.0

0.6 0 2.07 4.6 4.8

tr

1.5

0 11 28 17

9.6

12.0

27.1

27 37.3

24.0

CIS,J

C IS '2

0.27 4 0.35

3

C 2D

ｾ＠

ｾ＠

(')

:t.

"oe:

...

ｾ＠

tr 8.1 0.8 14.7

27.3 5.7 4.5 13.0

110 111 111 111 3.8+ -4.7 18.9 U

l.l

0.3

1.7

0.7

109

108

b

All acids not reported. , 45 Hr growth, batch growth. d Iso C I8 (1.8,%,), e Each species contained < 1 % each C 12 , Clz-monoene, C 14 , C I5 and C 17 . f C I7 -monoene. • Grown on glucose. C 17 ,1 (1.4 %). h 18: 4 or 20: 1 (3.1 %). i 20:2 (0.7%). j brC ls (7 %), brC I7 (3 %), nC 20 -monoene. k 11-5-270C. I C 20 and above (0.4 %). m C 12 and above (2.4 %). "2 % Glucose. °Other fatty acids 5.68 %. P Bottom fermenting. q 15: I (0.9 %), 16:2 (2.0 %), 16: 3 (1.9 %). , Semiaerobic. , 14: 2 (1.6 '%;J, 15: I (5.2 %), 15:2 (0.8 ,%,), 16: 2 (1.6,%,), 17: 1 (0.9 %). , Aerobic . • 15 : 1 (2.2 %), 17: 1 (0.7'%;).

aC 22 (2%), C 24 (1 %)-traces ofC II , C 12 , C 13 , CIS' C 17 , C I7 '3' C 20 ,3'

O

Saccharomyces cerevisiae (anaerobic Baker's yeast, A 2) Saccharomyces cerevisiae" (NCYC 712) Saccharomyces cerevisiae Brewer's yeast P •q Baker's yeast (A 2)'" Baker's yeast (A s)'·· 1.0 5.6 2.2

0.7

11.3 37.4 12.8 9.8

15.8

12.7

6.7 8.5 5.1 2.1

26.4 38.5 35.1 58.6

4.1 1.3

0.1 0.5 1.3

0.6

45.3

34.5

18.75 6.2 41.5 26.6

16.9

46.4

4.6 2.3 0.9

0.8 0.2

C)

:1:0.

ｾ＠

ｾ＠

a

＠ｾ o'

.

ｾＮ＠

tl

...

ｾ＠ Qj

!t

\')

Q1

ｾ＠

.

Cl..

ｾ＠

...

Qj

ｾ＠

CI)

ｾ＠

rose um

Humicola insolens! Humicula languinosa! Humico/a grisea J var. thermoidea M alhranchea pulchella! var. sulfurea M alhranchea pulchella

Humlcola nigrescens

Humicola brevisl'

Fusarium oxysporum Glomerella cingulata Humicola grisea t'

Fwwrium

Strain 43\ Ceratocystis Iagucearum d Chaetomium glohosum'· ChaelUmium thermophile! Cochliohalus miyaheanus' (Helminthosporium oryzae) Claviaps purpurea (M I) Claviaps purpurea (M 2 )h Claviceps purpurea (M ,)i Claviceps purpurea (33F\6. \/1) (33F\6,2/4)1 Taphrina deformans' Epico('('um nigrum Fwwrium solani f. phast'o/i

7.5 \\3

25

1.9

22

3.

11.4

tr tr \.I

1.3

26.2

4

3.2 2.5 5 7 \0.0 5.3 \6.6 5.8 1.5

5.7 5.9

5.4

3.3 5.\ 2.3

0.0 2.\ \8.7 \3.7 9.6 4.4 3

5.9 5.2

CI'

25

>4.\

0.5

2.2

tr

C I7

6.6 4.6 6.0

3.\

10.8

1.0 6.0 2.0

4.0 3.2 4.7 5.9

C I6 :!

3.6 \.I 4.5 2.2

\6 87 79 88 88 78 25 25 25 25 25 25

86 86 86 74

Ir

22

7.0 8.9 11.0 \0.\ \6.3 \2.0 30.6 57.8

22.2

40.\

C ln

23.2 24.\ 2\ \5 28.0 \7.5 11.0 43.7 \5.3 28.8 20.5 29.9 2\.4 28.8

3.0 1.0

1.\ 0.4

1.3

0.9

C l5

C 14 : 1

0.3 0.5

6.0 1.4

\.4

1.8

C I4

\9.5 200 22.7

8.2

>1.3 1.5

C\O_lJ

0.2 4.6 0.5

84 25 25 85

82 83

Cephalosporium diospyri

Ceratocystis (,(}f!ru!escens'

79 80 8\

Ref.

Aspergillus dauci Aspergillus niger" Cephlliosporium acremon;um"

Fungus

:t:. c.,

ｾ＠

a:

31.3

......ｾ＠ 50.7

tr

2.4

1.2

tr

ClO

35.0

\8.5 4.0 \2.2

1.2

2 4 1.0 8.2 3.8

1.3

7.2 \8.3 3.0

2.0 11.4

C 1R . J

26.6

3\.3 \2.7 7 34 2\.6 39.7 34.4 \9.8 33.9 41.3 34.3 31.8 8.6 28.5

32.4 \8.9 8.0

\7.6 46.\ \1.9 25.5 61.0 \0.\ 38.6 \7.\ 35.6 26.8 59

C 18 :2

ｾ＠

'
1.2

0.9 2.8

1.25.4

2.1 -I

1.9

1.4 0.9 5.7 6.2 D.4

tr

tr

-0.2

tr tr

2.1 3.4

-0.6

1.9 tr 1.0

1.8

26 27.8 23.8 12.2 15.4 18.9 25.2 34.4 10.0 13.7 19.9 23.9 14.6 18.2 30.4 28.4 17.0 19.5 42.5 -21 2.1 1.4 1.9 -3

1.2 2.3 tr 14.2 17.7

3.0 4.8

3.1

-I

1.6 0.8

1.4 2.1

tr

9.3 9.0 5.5 6.8 8.6 10.8 6.7 2.0 7.0 4.5 10.2 1.3 2.1 9.1 6.8 8.8 2.3 J3.7 -15

C,.

,4

" Grown on glucose at 25SC, C" (1.2%), C",. (1.0%). b Range for incubation time between 24-96 hr. , Range includes 3-46 days growth and surface and submerged cultures < C,. including C" (3.0- I 7.4 %), C ,O (2.4-20.4 %). d C" (1.2%), C 2I (I %), C 22 (2%), C" (1.6;',). " Mesophilic fungus. ! Thermophilie fungus. 'C 2 • (4.9%), (1.1 %). 'OH-C"" (5.9%). 'OH-C", (41.8%). j OH-C,., (40.1 %). • Small amounts of C" and C", I Major acids, grown in darkness. m Mutant strains. " Grown on glucose, C ,O ,. (4.2 %). " Range for growth of 11-20 days, C 20 " (0.3-4.2 %), C ,OA (OA-2.1 %), C 22 (0.5 2.0 %). p Isopalmitic (OA %), unknown (1.3 %), isostearic (1.0 %). q C 171 (-I %), C,O" (-0.3 %), C,o" (-0.2 %), C" (-I %), C (-2 %). 'Grown on glucose at 25.5"C, C,." (tr). , Range for rings of a culture for period of time.

StilheILa sp. StilheILa thermophila! Trichophyton rubrum I.p

Sporolriehum exile

Sepedonium ampuILosporum Sporotrichum thermophile!

Sderotium rolfsii Sclerotium hataticola"\

82 25 25 25 25 92

90 25 27

Penicillium chrysogenum n Penidllium chrysogenum Penicillium cyaneumO

Penicillium puLvillorum"P" PeniciILium ",ophi'

89

N eurospora crassa/,m

5.4 16.4 4.7 10.9 10.9 14.2 42.2 47.0 50.8 19.1 8.8 17.5 48.D 51.4 16.1 2.7 8.3 13.4 25.4 - 15

44.1 61.5 48.0 65.4 31.4 45.8 21.8 11.7 37.2 50.2 49.6 40.1 14.6 16.6 44.3 35.1 58.4 58.3 14.3 -45 5.0

1.0 13.2 0.5 0.1 0.4

6.0 2.8 4.0

2.6 14.1

-1.2

2.1

tr tr 0.2 0.3

4.5

ｾ＠

.,

.,

ｾＮ＠

ｾ＠

c::

5:

.

ｾ＠

i;j'

tl

Qi'

ｾ＠ｾ＠

ｾ＠

Q,

ｾ＠

Qi'

ｾ＠

c... ｾ＠

28

28 2.5

2.2

1.2

2.4 5.5

3.6 0.6 \.0 \.0

C lo 13

\.8

\.0

tr 0.7

0.8 2.2 0.4\.0 \.5 \.2

2.6 0.5 1.0 \.7 tr

C 14

1.4

\.8

\.0

1.7 1.3

\.6 0.5 \.0

C 14,1

\.1

1.7 tr

C I5

13.1

15.0

18.7 22.5

16.4 13.4 17.629.0 12,3 12.4

2\.4 2\.g 19.7 9.2 11.714.5

C I6

\.9

4.0

\.5 2.3

2,1

2.6 2.2 \.1\.6

2.6 4.3 3.0 2.5 tr3.2

C 16,1

1.2

C I7

1.7

3.0

9.6

\.8

5.6 7.3 6.212.6 12,6 3.1

9.6 3.7 7.3 0.2 2.43.9

CIS

6.3

5.0

42,8 20.4

4,6

23.1 22.3 12.333.0

12.3 2\.5 28.8 10.6 15.724.9

CIS,I

b

C 20 ,1 (4.2%). , Free acids C 6 (6.3 %), C s (4.4 %), CI U (\.3 %). dRange represents analysis of different rings within a culture occurring with time. 'C 20 ,1 (2.5 %). f Range given for fatty acid content between land 12 days of growth. , C I9 (tr), C 21 (tr), C 22 (1.7 ,%,,), C 23 (\.3 %), C 24 (19.1 %), C 25 (4.4 ,%,,), C 26 (46.6 %), C 27 (\.9 '%,), C 2S (23.9,%,,), C 30 (tr), C 32 (tr). 'C I7 ,1 (\.1 ,%,), C I9 (I 0;';), C 21 (I %), C 22 (24.8'%,), C 23 (1 O{), C 24 (26°{), C 25 (tr), C 26 (15.4°1,,). ; C 20 '3 (1.2 %), C 20 '2 (\.2 %), C 20 '3 (1.3 %).

°C'6'2 (\.9%).

(fruiting body)

Collybia sp.

(fruiting body)

Agaricus campestris;

(ATCC 12640)

Clitocyble illudens Coprinus comatus

95 78

94 28

F omes igniarius"h Fomes sp.

(fruiting body)

66 16 93

16 16 16 67 91

Ref.

Exobasidium vexans' Corticium solani Tricholoma nudium f

HOMOBASIDIOMYCETES

Rhizoctonia lamellifera Stilnum zacallo-xanthum O U stilago scitaminca b Tilletia controversa' Rhizoctonia solanid

HETEROBASIDIOMYCETES

Fungus

54.3

65.5

34.8 42.0

70,3

34.2 28.4 29.035.2

29.9 4\.4 32.3 66.2 54.466.6

C 18'2

TABlE 3.7. Fatty Acids of the Mycelia and Fruiting Bodies of Basidomycete Fungi

17.0

2.5

4.1

13.2 14.9 0.526.1

1.7

13.5 3.8

C IS '3

\.0

C 20

...... ｾ＠

C'l

:b

'


aeciospore vv

U. psoraleae

teliospore""

U. hedysari-obseuri

uredospore"

Uromyces phaseoli

teliospore"

Phyragmidium andersonii

aeciospore"

Peridermium stalactiJorme

aeciosporeqq

C. comandrae

aeciospore PP

Cronartium ribicola

aeciosporeoo

Frommea abtusa var. duchesneae

uredospore

Hemileia vastatrix""

aeciosporekk M elampsora lini uredospore" M. medosae uredosporemm

C. hasknessii

teliospore jj

Phragmidium speciosum

Fungus

60

59

60

60

60

60

60

60

60

0.9

0.5

1.1

0.1

1.0

1.3

0.7

0.5

0.3

0.2

0.6

1.1

1.5

0.9

0.1

19.9

15.8

25.4

6.6

19.7

21.6

21.6

16.3

5.0

32.0

0.6

60

1.3

60

11.1

1.1 0.4

10.4

C I6

0.6

CIS

17.7

C 14,1

0.8

0.4

C I4

60

60

Ref.

1.4

0.5

0.7

0.7

3.2

2.4

1.6

0.8

1.0

1.3

0.3

1.2

0.6

C 16,1

0.4

C I7

8.8

3.5

11.5

2.6

2.5

2.8

4.0

13.1

3.0

5.7

3.1

4.8

3.0

CIS

Fatty Acids (%)

TABLE 3.13--continued

40.9

7.5

32.7

20.1

8.0

6.8

5.9

9.6

2.0

7.0

2.2

6.9

14.7

CIS,I

5.2

40.9

8.3

23.0

5.0

6.2

8.0

4.2

7.0

6.7

2.0

5.2

20.1

C IS '2

15.6

27.7

17.4

43.9

10.9

12.1

19.1

9.7

1.0

7.8

3.0

12.1

42.3

C IS ,3

0.3

0.3

0.3

0.6

1.1

0.9

2.0

0.7

1.1

0.5

0.4

C 20

CII

t)

a:

:b

......ｾ＠ "
C 18 ) comprise over 50 % of the total acids in each species. Behenic (C 22 ) was found to be the most abundant acid in the conidia of S. humili, while a major portion of the saponifiable fraction from E. graminis was not identified. Fatty-acid distributions of conidia which are produced by the imperfect stages of saprophytic Ascomycetes are similar to that of the Phycomycete sporangiospores (Table 3.12).

3.5.1.3. B asidiomycetes The fatty-acid compositions of spores produced by over 40 Basidiomycete fungi have been reported (Table 3.13). Tulloch and Ledingham(59,60) analyzed the fatty-acid content of spores produced by numerous rust and smut fungi with the aim of establishing their chemotaxonomic importance. They found considerable variation among these species but concluded that differences between the various species and races are not sufficient to be taxonomically useful. This is also true for the spore forms (uredospores, aeciospores, and teliospores) from alternate hosts of wheat (Puccinia graminis tritici) and oak stern (P. graminis avenae) rusts. The fatty-acid distributions of the different spore forms are qualitatively similar with little quantitative variation. The teliospores produced by the oak stern rust contain higher abundances of 9,10-epoxy-octadecanoic acid (see below). The teliospores of both races contain 10% total lipid, while the uredospores and aeciospores range between 16 and 21 %. The similarities in the fattyacid distributions of the uredospores and aeciospores, harvested from their respective cereal hosts, and teliospores, harvested from barberry, suggest that the host plant has little inftuence on the fatty-acid composition of the fungal pathogen. It was also found that the fatty-acid composition of teliospores and basidiospores produced by Gymnosporangiumjuniperi-virginianae are very similar, but significant qualitative and quantitative differences were found in the aeciospores. Linolenic is the most abundant acid in the teliospores and basidiospores, while C I8 :2 , which is present in relative abundances of over 62 % of the total fatty acids in the aeciospores, is predominant.

3.5. Fatty Acids of Fungal Spores and Sclerotia

103

The distributions of fatty acids in rust and smut fungi are similar to those of other fungi. The predominant saturated fatty acid found in these spores is C 16 . The major unsaturated acids are C 18 :land C I8 :2 . Saturated and unsaturated acids with chain lengths greater than C I8 are often present. Fatty acids ofthe rust spores gene rally have a greater degree ofunsaturation, which is due to the unusually high relative abundances of C I8 :3 . The most characteristic feature of oils from the rust spores, however, is the presence of 9. lO-epoxy-cis-octadecanoic acid. which appears to be restricted to the Uredinales.* This acid varies in relative abundances between 4 and 78 ｾｉＢ＠ ofthe total fatty acids, depending on the species (Table 3.13), and seems to be found primarily in the trigylceride fraction of the spore oils. There seems to be an inverse relationship between the relative proportions of the epoxy acid and the CIS unsaturated acids. The 9.1O-epoxy-cis-octadecanoic acid has also been reported in extracts from several higher-plant species.(61,62) The fatty-acid composition of teliospores from several smut fungi, representing two families ofthe Ustilaginales, have been reported. Although the distributions are very similar, C 18 :2 is the major unsaturated acid of the Tilletiaceae family, and C 18 :1 is most often predominant in species of the Ustilaginaceae. Teliospores of the corn smut fungus Ustilago maydis have been studied by several investigators who found that C 18 :1 is the predominant fatty acid,(60,63,64) while Gunasekaran et al.(65) found that C 18 :1 and C 18 :2 are present in almost equal relative proportions. Weete(64) detected unusually high relative concentrations of C I5 (13.9 %) accompanied by low relative abundances ofC 13 , C I5 :1, C 17 , and C 17 :1 in the free fatty-acid fraction of U. maydis teliospores. In some smut species, unusually high relative proportions of C 16 :1 have been reported.(57,59) Approximately 60% of the total fatty acids from the oils of Urocystis agropyri spores consist of two unidentified compounds.(66) The culture of rust and smut fungi has been accomplished in some laboratories. but little attention has been given to the comparative chemistry of mycelial and spore materials. In one such study, however, Trione and Ching(67) compared the fatty-acid composition ofteliospores produced in the field by the dwarf bunt fungus and mycelia grown on synthetic media. They found the distribution of fatty acids to be qualitatively and quantitatively similar in both materials. These studies and others(64,68) have also shown that the fatty acids are not uniformly distributed throughout the spore. Palmitic acid was found in the highest relative abundances in the spore coat and C 18 :2 in the spore interior. Laseter and Valle(69) reported that the fatty acids are specifically distributed between the spore coat and interior of oat stern rust * A c's

epoxy acid has been suggested as a component of sclerotia of an unknown Claviceps species'

n (J)

:\

i,v

)

to,-s-" V,

(

AcylIransler'

ｨｾ＠

0

U1

ｾ＠

1I ' 1_ _ _ _ _ _ _ _ _

Cj

NADP)

s'c

00

CoA SH

) t

XDehydral,on

0')

,p'

'.s'

. )

n

'Al' n-o

,p

V/

FMNH 2

Reduclian 2

B ｾ＠

CO,

ｃｈＮｾ＠

0

0

OH 1

1

\, eH. HCV""" ./p'-. . / Ｇ｛ｾ＠

J'

../

, 1 NHI 1

1 1

1

ｾｓｈ＠

1

0

CH NH CH. NH CH.' '-.", '-. .,/ " " , '-.", ,""SH 0 / - , CO CHo CO eH. r CH 3 CH 3 I 1 1

20.15A

• I 1

1

1

I

Fig, 4.5, (a) Hypothetical model of 4'-phosphopantetheine function in yeast fatty-acid synthetase complex,(20) (b) Chemical nature of"peripheral" and "central" thiol groups.(32)

supporting evidence that the "central" SH group is that of the 4'-phosphopantetheine portion of the ACP molecule. The next reaction is that of condensation between the acetyl and malonyl moieties to form acetoacetyl-SENZ at the "central" thiol group. The remaining reduction and dehydration reactions of the fatty-acid biosynthesis cycle occur with the respective substrates bound at the "central" thiol group. A hypothetical model has been proposed which explains the function of 4'-phosphopantetheine as a portion of ACP and an origin ofthe "central" thiol group. This model is iIIustrated in Fig. 4.5. 4'-Phosphopantetheine is envisioned as forming a flexible arm, 20 A in length, extending radially from the center of the muItienzyme complex. The flexible arm allows optimal

4.2. Biosynthesis

0' Saturated Fatty Acids

123

juxtaposition of the substrates and active sites of the enzymes for the reactions to occur. 4'-Phosphopantetheine is fixed in place by the apoprotein portion of the ACP molecule bound at the center of the synthetase complex. When the second reduction is complete, the substrate may be transferred to the "peripheral" thiol group for further elongation reactions beginning with condensation with malonyl-Sc-ENZ (Fig. 4.4). The cycle repeats until the preferred fatty-acid chain length is formed. For example, the cycle is repeated six times in the formation of palmitic acid. It is important to note that the subtrates acetate, malonate, and all intermediates of fatty-acid synthesis are covalently bound to the enzyme complex. No intermediates are released until the endproducts, palmitate and stearate, are produced. A third transfer reaction then takes place to remove the fatty-acid product and release the synthetase complex to accept acetate and malonate molecules to continue the process. The fatty-acid product is transferred to coenzyme A. Why are fatty acids with chain lengths of 16 and 18 carbon atoms the principal products of fatty-acid synthtases? Under normal conditions, neither sharter chain fatty acids nar longer ones are released from the complex. Schweizer et al.(33) demonstrated that the selective formation of C I6 and C I8 is not due to the acyl transferase present in the yeast synthetase, since no preference for chain lengths ranging from C 2 to C 20 can be shown far this enzyme. However, this phenomenon may be related to the mechanism of the transfer and condensing reactions, the relative rates of these reactions, and the relative affinities of the transfer-binding sites for the intermediates and products of fatty-acid biosynthesis. Before chain elongation can continue, the acyl group must be transferred from the "central" to the "peripheraI" thiol group so that an incoming malonate can combine at the "central" position. In order for the fatty-acid product to leave the complex, it must first be transferred to the transferase-binding site and then to CoA. Schweizer et al.(33) also pointed out that the "central" position is spatially separated from the active center of the transferase until acyl chain-Iengths OfC I6 and C I8 are reached. With chain elongation, the acyl group becomes increasingly Iipophilic, which may cause conformational changes in the quarternary structure of the fatty-acid synthetase, and becomes available for the transferase. Furthermore, Sumper et al.(32) suggested that the probability of a covalently enzyme-bound acyl group forming a product rather than continuing in the chain-e1ongation process may be chain-Iength dependent. They obtained experimental evidence to support their proposal that the growing acyl chain interacts with the enzyme protein only after a chain length of C 13 is reached, and this interaction changes the relative velocities of the transferring and condensing enzymes in favor of product formation by an energy increment of -0.9 kcal per methylene.

724

Fatty Acid Metabolism

4.2.5. Control of Fatty Acid Synthesis The factors which may regulate the terminal transfer reaction and, hence, the nature of the endproducts of fatty-acid synthesis were discussed above. Numerous studies have shown that several metabolic factors inftuence the rate of fatty-acid synthesis, but it is not known exact1y how this process is regulated in nature. The control of fatty-acid synthesis has been included in reviews by Stumpf,(34) Vagelos,(35) Lynen,(14,20) and Kumar et al.(19) It has been shown that intermediates in the glycolytic and tricarboxylic acid pathways stimulate fatty-acid synthesis in cell-free preparations from several sources. White and Klein(36,37) screened over 35 compounds and found that the rate of fatty-acid synthesis by cell-free preparations of S. cerevisiae is stimulated by citrate, isocitrate, glucose-6-phosphate, fructose1,6-diphosphate, and L-IX-glycerophosphate. Stimulation by these compounds is attributed to the activation of acetyl-CoA carboxylase, wh ich is not an integral component of the multienzyme complex, but a soluble enzyme. The activity of this enzyme may, however, be associated with the membrane elements of the cell, since it can be stimulated by phospholipids.(38) In animals, the activation mediated by these compounds is due to aggregation of the component parts of the enzyme.(39) Activation in yeast preparations seems to occur by a different mechanism, since no differences in the sedimentation behavior of the enzyme are noted in the presence or absence of IXglycerophosphate and citrate.(40) The carboxylation of acetyl-CoA is considered the rate-limiting step of fatty-acid synthesis in animals, and the yeast, Lipomyces lipo!er,(41) but this does not appear to be the ca se for other yeasts(42) and certain higher plants(43) since no activation by these compounds is observed. In his review on fatty-acid synthesis, Kumar et al.(19) compared the reaction rates of the component enzymes of fatty-acid synthetases from several sources and showed that the acetyl CoA: ACP transferase, and possibly the palmityl transferase, reactions are rate-limiting. On the other hand, Lynen(44) proposed that the condensation reaction is the rate-limiting step in yeast. This is supported by the results obtained by Schweizer and Bolling,(45) who compared the component enzyme and total fatty-acid synthetase activities of a wild-type yeast (Table 4.3). They reported that the "condensing enzyme" has the lowest specific activity, which is followed by the dehydrase. These investigators also isolated and characterized a mutant of S. cerevisiae, defective in saturated fatty-acid synthesis, which requires C 14 , C 16 , C 18 , or C 18 :1 for growth. A comparison between the fatty-acid synthetases from this mutant and a wild-type yeast reveals that the respective component enzyme activities of each synthetase are almost identical, except that the "condensing enzyme" is completely inactive in the mutant (Table

4.2. Biosynthesis

0' Saturated Fatty Acids

125

TABLE 4.3. Component Enzyme and Fatty-Acid Synthetase Activities of Wild-Type and FAS14 Mutant Yeast (45) Specific Activity· Reaction Wild-type ( x 2180, mat 1 - IJ() Malonyl transfer Acetyl transfer Condensation First reduction Dehydration Second reduction Palmityl transfer Fatty-acid synthetase

11,000 32 0.36 5,000 3.3 33,000 370 2,050

FAS-15

10,500 33 b ｾ＠

4,800 3.9 32,500 370 ｾ＠

b

• Activities are expressed as units/mg; a unit is defined as the turnover of I J1mole of substrate per minute. b No detectable activity.

4.3). Meyer and Schweizer(46) isolated a mutant of S. cereUlszae wh ich requires saturated fatty acids (C 14 or C 16 ) for growth at 33°C but grows normally at 22°C. Again, the limiting step was found to be the "condensing enzyme." It was shown that enzymatic activity ofthe thermolabile component is related to whether the synthetase complex is synthesized at higher or lower temperatures, rat her than the incubation temperatures for the completed complex. It was concluded that the altered protein is stabilized by its incorporation into the structural framework of the multienzyme complex. It is weil known that long-chain thiolesters of coenzyme A bind nonspecifically to proteins and cause enzyme inhibition by a detergentlike action.(47.48) Inhibition of several enzymes such as glucose-6-phosphate dehydrogenase(58,59) and phosphatidic acid phosphatase(50) by acyl thiolesters has been reported. Also, inhibition of fatty-acid synthesis by the longchain thiolesters has been reported far pigeon liver,(51) rat liver,(52) and yeast(36) preparations. Palmityl-CoA was shown to inhibit the citratecondensing enzyme(48) and the rate of acetyl- and malonyl-CoA condensation by rat brain preparations,(53) while acyl-CoA derivatives competitively inhibit acetyl-CoA carboxylase activity with respect to activation by citrate.(54) Lust and Lynen(55) reported experimental evidence that long- chain acyl-CoA derivatives may be of importance in the regulation of de novo fatty-acid biosynthesis by yeast. They found that long-chain acyl-CoA thiolesters (C 12 to C 18 ) competitively inhibit the fatty-acid synthetase of yeast with respect to malonyl-CoA. The inhibition can be partially reversed

126

Fatty Acid Metabolism

by serum albumin. The competitive inhibition is chain-Iength dependent (C 14 > C I6 > C I8 > C 12 ). Noncompetitive inhibition of acetyl-CoA and NADPH was also noted. The regulation of fatty-acid biosynthesis is probably due, not to any single factor, but to a combination of factors controlled by the dynamic balance of intermediary metabolism. For example, the availability of substrates, cofactors, and cations may participate in this regulation. A reduction in NADPH levels resuIts in the production of ß-ketoacyl- and O!,ß-unsaturated acyl-CoA endproducts rather than the normal saturated fatty acids. Cations, such as magnesium,(36.37) stimulate fatty-acid synthesis, and their availability may exert a rate-Iimiting control in this process.

4.2.6. Fatty Acid Elongation The de nova synthesis described above is the principal source of saturated fatty acids in all biological systems. The synthetases, which catalyze reactions leading to the formation of these lipids, produce chain-Iengths no longer than 18 carbon atoms. However, as noted in Chapter 3, fatty acids with chain lengths greater than 18 carbon atoms are present in fungi, as weil as in plants and animals. The elongation of fatty acids has been shown experimentally in both animals(56.57) and plants.(58.59) Two types of fatty-acid eIongations have been reported; one involving the condensation of acetyl-CoA with a preformed acyl-CoA, and the other involving condensation of malonyl-CoA. Mooney and Barron(60) extracted a soluble mitochondrial fatty-acid elongation system from mammals which could eIongate acyl-CoA thiole sters of chain lengths ranging from C 4 to C 20 . It was shown that inner mitochondrial membranes contain enzymes which synthesize C I4 to C I8 acids (de nova or with a medium chain-Iength primer), and the outer membranes contain enzymes which catalyze the elongation of medium- and long-chain fatty acids.(61.62) In this elongating system, acetyl-CoA is the C 2 donor, and NADH and NADPH are required for maximum activity. They also reported evidence that 3-hydroxy and fl2 fatty acids are intermediates in the elongating process.(63) They proposed that the unfavorable energetics for the addition of C 2 units, via acetyl-CoA, is overcome by the rapid reduction by which the 3-hydroxy acid is produced. No evidence of a keto acid intermediate was found. The second type of fatty-acid elongation is associated with the microsomal fraction of the cell. Malonyl-CoA is the C 2 donor and NADPH is the electron donor.(64.65) Fatty acids containing chain lengths of C IO to C 16 , C I6 :1, C I8 :1, C I8 :2, and C 18 :3 serve as substrates for this system. Intermediates are reported to be 3-keto, 3-hydroxyl, and fl2-unsaturated fatty acids. A

4.3. Biosynthesis of Unsaturated Fatty Acids

127

microsomal fatty-acid elongating system was not detected in S. cerevisiae, but a mutant of this species, deficient in fatty-acid synthetase, is capable of elongating dietary fatty acids of medium-chain length (C 13 to C 17 ) by 2 carbons or more.(66) The extent of elongation is dependent on the fatty-acid chain length added to the growth medium, and no fatty acids containing greater than 18 carbons are produced. The contribution of this elongation pathway to the total fatty-acid synthesis by the cell is probably insignificant.(45) The fact remains, however, that very long-chain fatty acids are present in yeast. Fu1co(67) reported that these acids (C 20 to C 26 ) are not present in Candida utilis and S. cerevisiae grown on lipid-free media but are detected when [3HJ labeled C 22 is added. He found a chain elongation pathway in C. utilis that is specific for chain lengths of C 20 to C 24 . When these fatty acids are added to the medium, C 26 is the principal product of the elongation enzymes, and, when C 21 or C 23 are added, C 25 and C 27 are the principal products. This chain-length specificity explains why very longchain fatty acids are not detected in the yeast grown on lipid-free medium.

4.3. BIOSYNTHESIS OF UNSATURATED FATTY ACIDS Lipids produced by most biological systems generally contain a high see Chapter 3) which is due to the monodegree of unsaturation ＨｾＯｭｯｬ･Ｌ＠ and polyenoic fatty acids present in their structures. Monounsaturated fatty acids are produced by virtually allliving organisms, and, with the exception of bacteria, polyunsaturated fatty acids are also widely distributed in nature. Monoenoic fatty acids generally have the cis configuration. In most organand oleic [C I8 :1 ＨｾＹＩｊ＠ are the most common isms, palmitoleic [C I6 :1 ＨｾＹＩｊ＠ monoenes, but other positional isomers are produced by certain groups of organisms. Linoleic [C I8 :2 ＨｾＹＮＱＲＩｊＬ＠ a-linolenic [C I8 :3 ＨｾＹＮＱＲＵＩｊ＠ and Alinolenic [C I8 :3 ＨｾＶＮＹＱＲＩｊ＠ are the common polyenes (see Chapter 3).

4.3.1.

Monounsaturated Fatty Acids

4.3.1.1. Aerobic Pathways

Two principal aerobic pathways of monounsaturated fatty-acid biosynthesis are found in nature. One pathway appears to be restricted to animals, fungi, and certain microorganisms and is specific for thiole sters of coenzyme A. The other pathway is restricted to plants and is specific for stearyl-ACP.

128

Fatty Acid Metabolism

The desaturase system found in animals, fungi, and bacteria appears to be an aggregate of enzymes associated with the microsomal fraction of the cell.(68-71) The desaturase system from hen liver contains three components: (1) a desaturase, (2) NADH cytochrome b s , and (3) cytochrome b s .(72,73) Thiolesters (CoA) of CUi and C I8 are desaturated by this system to their respective A9 monoenes. Reduced NADP+ and molecular oxygen are required in the desaturation process: 02' NADPH

Stearyl-CoA - - - - - - - - - - - . . Oleyl-CoA Microsomal enzymes

The oxygen dependency of these reactions explains why unsaturated fatty acids (oleate or linoleate) are required for S. cerevisiae grown under anaerobic conditions. (68) The desaturase enzyme system from the mycelial fungus, N eurospora crassa, was reported to be similar to that ofthe yeast and animals.(74) StearylCoA is rapidly converted to its A9 unsaturated isomer by enzymes of the microsomal fraction from this fungus. Both NADPH and NADH are effective electron donors for this system, but there is a pronounced preference for the latter. It was also shown that the acyl-CoA substrates are tightly, but not covalently bound to the microsomes, and that they are available for several competing reactions: (1) nYcleotide-Oz-dependent desaturation, (2) glycerophosphatide formation, (3) triacylglyceride formation, and (4) deacylation. Bennett and Quackenbush(7S) found a similar type of desaturase system in Penicillium chrysogenium. The soluble plant desaturase systems are associated with the chloroplast and catalyze the following reaction : 02. NADPH

Stearyl-ACP -------.. Oleyl-ACP Ferredoxin

Plant desaturases are specific for C 18 ACP thiolesters. Fatty acids with hydrocarbon chains of less than 18 carbon atoms must und ergo chain elongation prior to desaturation. As in animals, plant (Euglena gracilis) desaturase systems are composed of three components, adesaturase, NADPH oxidase, and ferredoxin.(76-78) There is a large volume of information concerning the biosynthesis of monounsaturated fatty acids, but the exact mechanisms of these dehydrogenation processes are not weIl established. This subject has been reviewed by Bloch(79) who pointed out that there is no analogous chemical or biochemical reaction in nature which removes hydrogen atoms from the nonactivated center of an aliphatic chain. The desaturase is highly specific for introducing a double bond in the 9, 10 position of fatty-acid thiole sters with 18 or less carbon atoms.

4.3. Biosynthesis of Unsaturated Fatty Acids

129

The absolute specificity of desaturases (regardless of chain length) for introducing 11 9 double bonds probably arises from the enzyme binding the activated carboxyl end ofthe substrate, wh ich allows the 9 and 10 methylene groups to be properly positioned at the active center.(80.81) A second 11 9 desaturase, specific for short-chain fatty acids, may be present in some species.(82) However, Schul tz and Lynen(82) reported a yeast system that converts decanoyl-CoA to 11 9-decenoyl-CoA, with C 16 :1 (11 9 ) and C 18 :1 (11 9 ) being the major monoenes produced by this organism. They suggested that the appearance of certain monoenes in a species is not governed by the specificity of the desaturase, but by the specificity of the fatty-acid synthetase which provides the acyl-thiolester substrates. Other than binding the acyl substrates, the requirement for thiolesters cannot be readily explained. The thiolester does little to '"activate" the 9,10 positions (C-H bond scattering). However, Richards and Hendrickson(87) proposed that the hydrocarbon chain may not be fully extended in the enzyme-substrate complex but may assurne a pseudoannular conformation, allowing the reacting carbon atoms to approach the thiolester. They also suggested that the oxygen may attack the sulfur and not a carbon atom, resulting in aperthiolester that would provide an "active" oxygen for withdrawl of hydrogens from the 9, 10 methylene groups. The requirement of molecular oxygen and NADPH suggests that a mixed function oxygenase (see Chapter 2) may be involved in the desaturation process. This would involve a two-step process in which the saturated fatty acid would be hydroxylated and then dehydrated to form the corresponding olefin. Several studies have been conducted to test this mechanism, but no hydroxylated intermediates were detected,(78) and radioactively labeled hydroxy acids (ACP or CoA derivatives of 9- or lü-hydroxystearic acid) were not converted to the corresponding monoenes.(78,83) Although the mixed-function oxygenase system does not appear to be operative, an electron-transport chain is involved. Based primarily on studies with the soluble enzymes of Euglena gracilis,(78) Bloch(79) summarized the current information on the flow of electrons in the desaturation process (Fig. 4.6). As noted above, the soluble desaturase enzymes from E. gracilis can be fractionated into three catalytic proteins. One fraction contains flavin and oxidizes NADPH, and another fraction contains a nonheme pro tein (ferredoxin). Both make up the electron-transport chain for reducing molecular oxygen. A third fraction contains the desaturase, wh ich is unstable and inactive in the absence of the other two fractions. The next question on the mechanism of fatty-acid desaturation that arises is concerned with the stereochemistry of hydrogen removal. This was first investigated by Schroepfer and Bloch(84) who grew the bacterium, Cornebacterium diphtheriae, in the presence of chemically synthesized,

Fatty Acid Metabolism

130

ｎａｏｐｈｾ＠

ｆｐｾｲｦ［Ｌ＠

\(oesalurase )(RCH:CHR +H 20

ｆｌａｖｏｐｒｾ＠ NAOP

ｾ＠ FPH 2

NAOPH OXidase

Fe+++ Prolein Ferredoxin

( 02

Desalurase ＢＧｏｾＯ＠ \

ｒｃｈＲＭｾ＠

H20

Fig. 4.6. Electron-transport chain far oxygen activation in the formation of monounsaturated fally acids P9 )

stereospecifically tritium-labeled stearic acids: D- and L-[9- 3HJ- and D- and L-[1O- 3H] stearic acids. They showed that the D-9- and D-lO-tritium atoms are lost and the L-9- and L-lO-trition atoms are retained in the formation of oleic acid from the stereospecifically labeled C 18 . The same stereospecificity has been shown in Chlorella vulgaris and others.(86) Morris(86) showed that, as in the bacterial system, the two hydrogens removed from stearic acid in its conversion by a mammalian and algal systems have the cis configuration. AIthough the stereospecificity of hydrogen removal in the biosynthesis of oleic acid is established and seems to be uniform in microbes, plants, and animals, the mechanism of hydrogen removal remains uncertain. Schroepfer and Bloch(84) reported that in C. diphtheriae the desaturation of stearic acid involves a stepwise removal of hydrogen. The D-9-hydrogen atom is the first to be removed, and this is the rate-limiting step. Morris(86) on the other hand, reported that, in algal (Chlorella) and mammalian systems, the formation of oleate involves a simuItaneous concerted removal of hydrogen from stearate. This eliminates the requirement of an oxygenated intermediate in fatty-acid desa tura tion. 4.3.1.2.

Anaerobic Pathway

Monounsaturated fatty acids can also be produced via a nonoxidative pathway, rather than the direct oxygen-requiring desaturation routes described above. This anaerobic pathway seems to be restricted to the bacteria and has been studied extensively in Clostridia species and Eseheriehia eoli by Goldfine and Bloch,(88) Scheuerbrandt et al.,(89) and Lennarz et al.(134) In this pathway, saturated fatty-acid formation proceeds as usual until the 10 carbon level is reached. At this point, the enzyme, ß-hydroxydecanoyl thioester dehydrase, catalyzes the dehydration and double-bond isomerization reactions which lead to the formation of eis-L\3-decenoyl-ACP (ß, ,1.) and trans-L\2-decenoyl-ACP (IX, ß). The trans isomer is the normal intermediate in saturated fatty-acid biosynthesis and undergoes chain elongation to the preferred saturated product. The cis isomer, however, cannot be

131

4.3. Biosynthesis of Unsaturated Fatty Acids

I

CH 3(CH2)5 CH - CHCH2COSR

o

+3C,

CH3(CH2)5CH-CH(CH2)7COSR 9

I

.1 - C I6

+C,

CH3(CH2)5CH - CH (CH 21g COSR

LX I -

I

CH3(CH2)5CH2CH -CHCOSR

NAOPH

CH3(CH2)5CH2CH2CH2COSR

t

polmilole

I

sIeorale

C I8

Fig. 4.7. Pathway of saturated and an aerobic monounsaturated fatty-acid biosynthesis in E. co/i. Reactions catalyzed by 3-hydroxydecanoylthioester dehydrase are indicated by a P9 )

reduced by enoyl-ACP reductase, but instead, continues in the elongatio r , reactions of fatty-acid synthesis as an unsaturated intermediate. The unsaturated intermediates remain bound to the synthetase as ACP, as in the formation of saturated fatty acids. This pathways explains the origin of the /).7, /).9, and /).11 position al isomers detected in bacteria. One of the most common monounsaturated fatty acids in bacteria is vaccenic acid [C I8 : 1 (/).11c)], which is shown in Fig. 4.7.

4.3.2. Polyunsaturated Fatty Acids As noted in Chapter 3, there are three families of unsaturated fatty acids, w 3 , w 6 , and w 9 , which are classified according to the number ofmethylene groups between the terminal methyl group and the first double bond toward the carboxyl end of the molecule. Generally, polyunsaturated fatty acids have the cis configuration and are oft he methylene-interrupted type (see Chapter 3). Polyunsaturated fatty-acid biosynthesis has been studied in organisms from various taxonomie groups including fungi, protozoa, slime molds, algae, higher plants, and animals. (90.91) In these systems, including Candida utilis(92) and Penicillium chrysogenum,(75) polyunsaturated fatty-acid formation

Fatty Acid Metabolism

132

UJ9-Family

C'S -

w 6 -Family

C'S" (öge )

1

1+c

2

C20 '2 (öSe." e)

1

Fig. 4.8. Principal pathways of polyunsaturated fatty-acid biosynthesis in plants and animals. 1911

proceeds via sequential desaturations of C 18 :1 (.'1 9) and chain elongation. The double bonds are introduced into the molecule in one of two ways. In plants, desaturation occurs between the original central (.'1 9) double bond and the terminal methyl group and is referred to as w 3 type desaturation (Fig. 4.8). In this pathway, C 18 :3 (.'1 9. 12 ,15) or ｾＭｬｩｮｯ･｡ｴ＠ is produced via C 18 :2 (.'1 9,12) or linoleate. Certain algae desaturate palmitate to the respective C 16 unsaturated acids via the w 3 pathway.(93) Higher animals cannot produce polyunsaturated fatty acids by terminal methyl-directed desaturation and, hence, are incapable of synthesizing C 18 :2 (.'1 9,12). The .'1 6,9 isomer is the C 18 diene produced by these organisms by carboxyl-directed desaturation. w 9 Fatty acids, such as C 20 :2 (.'1 8,11) and C 20 :3 (.'1 5,8,11), are produced by chain elongation and further desaturation in the same manner. The C 18 triene produced by animals is C 18 :3 (.'1 6,9,12) or ),linolenate, which is formed via dietary C 18 :2 (.'1 9,12). Certain lower plants and phycomyceteous fungi also produce A-linolenate. The w 6 family is produced by animals and lower organisms by carboxyl-directed desaturation and chain elongation. Of particular importance is arachidonic acid [C 20 :4 (.'15.8.11.14)J, which is a common constituent of animals, lower plants, protozoans, and a few fungi (see Chapter 3). This tetraene may be produced by two pathways. The pathway predominant in animals and certain algae involves the chain elongation to C 20 :3 (.'1 8. 11 ,14) followed by carboxyl-directed desaturation. In the slime mold, Physarum polycephalum,(95) and certain protozoans, C 18 :2 (.'1 9,12) first undergoes chain elongation to C 20 :2 (.'1 11. 14) which is desaturated to C 20 :4 (.'1 5,8.11,14) via C 20 :3 (.'1 8. 11 ,14). Generally, the ｾＭｬｩｮｯ･｡ｴ＠ pathway is characteristic of plants and the A-linolenate pathway is characteristic of animals, but the lower forms of life may contain characteristics of both pathways.

4.3. Biosynthesis of Unsaturated Fatty Acids

133

The mechanisms of polyene formation are thought to be analogous to those of monoene biosynthesis. The coenzyme-A thiolester of oleate is the principal substrate for the desaturase and molecular oxygen and NADH or NADPH are the required cofactors. Meyer and Bloch(70) reported that polyene biosynthesis may require additional cofactors, since the desaturase from Candida ulitis requires an unknown soluble factor for activity. In most organisms, including the yeast C. utilis and N eurospora crassa,(96) desaturase activity is associated with the particulate (lOO.OOO-g) fractions of the cel\. In Chlorella, the conversion of linoleate to linolenate was observed in the chloroplast fraction, but, as in the yeast system, soluble activating enzymes are required for activity.(97) Based on experimental evidence, it has been proposed that certain glycolipids and glycerophosphatides may be involved in polyunsaturated fatty-acid biosynthesis. This proposal is based on studies with Chlorella.(98- 101) Euglena,' I 02) and N eurospora crassa.(96) As described above, aerobic monounsaturated fatty-acid biosynthesis pathways involve the acyl thiolesters of CoA or ACP as the initial ｳｵ｢ｴｲ｡ｾＮ＠ In these systems, however, [14C] oleate is rapidly incorporated into more polar lipids, particularly phosphatidyIcholine and monogalactosyldiacylglyceride, and then is converted to its diunsaturated isomer. Based on these studies, Erwin(90) outlined the possible pathways of polyunsaturated fatty-acid biosynthesis

Fig. 4.9. Possible pathways of polyunsaturated fatty-acid biosynthesis in Chlorella and Euglena. (91 ) MG DG, monogalactosyldiacylglyceride: PC, phosphatidylcholine: heavy arrows indicate a possible pathway in fungi (Neurospora crassa)!95 I

134

Fatty Acid Metabolism

as they occur in Chlorella and Euglena (Fig. 4.9). According to Baker et al.(95) stearyl-CoA is rapidly converted to oleyl-CoA which is subsequently incorporated, almost as rapidly, into an oleyl phosphatide by microsomes from N. crassa.ln the presence ofNADH or NADPH, the oleyl phosphatide is converted to a linoleyl phosphatide by the same fraction. Thus, polyunsaturated fatty-acid formation by fungi (N. crassa) may follow the pathway involving phosphatidylcholine, as shown in Fig. 4.9.

4.4. BIOSYNTHESIS OF UNUSUAL FATTY ACIDS As defined in Chapter 3, unusual fatty acids are those wh ich are generally limited in distribution to certain taxonomic groups and have some aspect of their structure that varies from the straight-chain, saturated or unsaturated (methylene interrupted) monocarboxylic acids. This section is concerned primarily with unusual fatty acids produced by fungi and their biosynthesis by these organisms. This includes the acids containing nonmethylene interrupted ethylenic bonds, acetylenic acids, and the substituted acids containing hydroxyl, epoxy, and methyl groups.

4.4.1. Nonmethylene I nterrupted Ethylenic Acids Polyenoic fatty acids containing nonmethylene interrupted ethylenic bonds may be of two types (see Chapter 3): (1) a conjugated system in which two double bonds are separated only by a single co valent bond between two carbon atoms, and (2) a system in which two double bonds are separated by two or more methylene groups. Fatty acids with conjugated double bond systems have not been reported as fungal products, but the ｾＵ＠ fattyacids reported for the slime mold, Dictyostelium discoideum, are ofthe latter type of non methylene interrupted systemY03,I04) The ｾＵ＠ fatty acids are also products of certain higher plants such as Pinus spY05) and certain bacteria.(I06-108) Fulco(I05-107) reported that the formation of ｾＵ＠ fatty acids in certain bacteria is similar to the formation of ｾ＠ 9 fatty acids; molecular oxygen and iron are required. The formation of ｾ＠ 5 or other fatty acids with double bonds in unusual positions has not been sufficiently investigated but Hitchcock and Nichols(91) suggested that they arise through the action of normal or abnormal desaturases followed by modification ofthe molecule.

4.4.2. Acetylenic Acids Fatty acids containing acetylenic bonds are considered secondary metabolites, and their distribution was not considered in Chapter 3. There

4.4. Biosynthesis of Unusual Fatty Acids

135

has been a large number of searches for these compounds in plants and fungi which have produced the identities of over 400 natural acetylenic compounds.(109) About a fifth ofthese compounds were isolated from fungi, most of which were Basidiomycetes. Mono, di, and triacetylenes occur in fungi ranging from C 6 to C I8 in chain length. In fungi, C 9 and C IO acetylenes are most common. Most acetylenes do not occur as the simple acid, but may contain various oxygenated functional groups, i.e., hydroxyl, epoxy, aldehyde, or a second carboxyl group. The bio genesis of natural acetylenes is not weIl established, but has been discussed by Turner(108) and reviewed by Bu'locky09) The biosynthesis of acetylenes will not be discussed here in detail; only the formation of the simple monoacetylene crepenynic acid [C I8 :2 (,19c, 12a)] will be outlined (Fig. 4,10), Bu'lock and Smith(11 0) showed that [1O_ 14 C] oleic acid is converted to [1O_ 14 C] crepenynic acid by Tricholoma grammopodium and showed that the proposed intermediate, dehydrocrepenynic acid [C I8 :3 (,19c,12a.14c)], is present in polyacetylene-producing fungi. Polyacetylenes are apparently CH 3(CH 2116 COOH Stearic Acid

1-2H CH 3(CH 217CH. CH (CH 217COOH Oleic Acid

!-2H CH 3(CH 214 CH ·CHCH 2 CH. CH(CH 217 COOH Linoleic Acid !-2H ｃｈＳＨｾＱＴ＠

C= CCH 2CH. CH(CH 217COOH crepernic Acid

CH3(CH 212 CH·CHC=CCH 2 CH aCH (CH21-rC00H

!-nH POLYACETYLENES

1

B - oxidation

Cg - CI6 Polyocetylenes

Fig. 4.10. Possible biosynthesis of crepenynic acid and other natural acetylenes 010 )

136

Fatty Acid MBtabolism

produced through successive dehydrogenations (ane --+ ene --+ yne) at saturated portions of crepenynic acid and its dehydrogenated products. The common shorter chain acetylenes may be formed by ß-oxidation (Fig. 4.10).

4.4.3. Substituted Acids 4.4.3.1.

Hydroxy Acids

Several hydroxy fatty acids have been identified as fungal products and inc1ude a 12-hydroxy-oleic acid, 17-hydroxy-palmitic and stearic acids, 2hydroxy-palmitic and hexacosanic acids, and 9,1O-dihydroxy-stearic acid (see Chapter 3). The most extensively investigated of these hydroxy acids is 0-12hydroxy-il 9 -octadecenoic acid (ricinoleic acid). There are two major natural sources of this fatty acid: castor plant (Ricinus communis) and the immature sc1erotia or mycelia ofthe ergot fungus, Claviceps purpurea. The biosynthesis of ricinoleic acid occurs by distinct pathways in each of the two systems and has been reviewed by Morris.(86) In the higher plant system, ricinoleic acid is produced by the direct hydroxylation of oleic acid in which a single hydrogen atom having the o-configuration is replaced.{1l2) Molecular oxygen and NADPH are the required cofactors(113) (Fig. 4.10). In the fungal system, ricinoleic acid is produced via the hydration of linoleic acid, which does not require molecular oxygen (Fig. 4.11). As discussed in Chapter 3, ricinoleic acid does not exist as a free acid, but is esterified to glycerol, and the hydroxyl function is further esterified to a long-chain fatty acid (estolides). Morris(86) suggested that, instead of a simple hydration reaction for the formation of ricinoleate, long-chain fatty acids may be bound direct1y across ERGOT FUNGUS

CASTOR PLANT CH 3 (CHz17CH • CH (CH z 17COOH

CH3(CHZ14CH = CH CHzCH = CH (CHzl.,GOOH LINOLEIC ACID

OLEIC ACID

Hydroxylotion (Oz,NADPHl

(01

T""'

OH CH 3 (CH z14 - CH z - eH -CHz -CH = CH -(CH z17COO H RICINOLEIC ACID

Fig. 4.11. Biosynthesis of ricinoleic acid by Ricinus communis (castor bean) and Claviceps purpurea (ergot fungus). (a) See reference (112). (b) See reference (114).

4.4. Biosynthesis of Unusual Fatty Acids

137

the double bond of linoleate, resuIting in the production of the estolide structure. Extracellular glycolipids produced by Torulopsis species contain 17-Lhydroxy C 18 acids bound glycosidically to the disaccharide sophorose. The hydroxy acids are formed first and then are linked to the sophorose. During this conversion. the hydroxyl oxygen atom of the fatty acid is retained. In this yeast. hydroxylation occurs primarily at the penuItimate carbon atom (w - I). but placement of the hydroxyl groups (w or w - 1) depends on the fatty-acid chain length and degree of unsaturation.(116) Heinz et alY 14) showed that the hydroxyl oxygen atom is derived from molecular oxygen rather than water. They also showed, through thc use of specifically labeled pentadeuterostearic acid. that L-17-hydroxy-stearic acid is formed via the direct hydroxylation of stearate and an unsaturated intermediate is not involved. Using D- and L-[3H] stearic acid, they showed that the L-tritium atom is displaced by the hydroxyl group. and the configuration is

16.18- Pentadeuterostearic Acid

retained. In a cell-free system from a Torulopsis sp., Heinz et al.(116) showed that hydroxylation activity is found in 48,000 g particulate fraction and requires molecular oxygen and NADPH. This system has the characteristics of a mixed-function oxidase. Mixed-function oxidases were discussed in detail in Chapter 2 in the section on the oxidation of hydrocarbons. In the yeast Candida utilis and Saccharomyces cerevisiae, very longchain 2-hydroxy (a-hydroxy) fatty acids are associated with the cell envelope and represent part of the cerebroside structure of sphingolipids. The ahydroxy compounds are formed during the a-oxidation (see section 4.5.1) of fatty acids. a-Hydroxy fatty acids are also found in higher plants as part of the cerebroside fraction,(118) and their biosynthesis has been included in a review by Morris.(86) Hydroxy acids from plant, animal, and fungal tissues have the D-configuration. In plants, 2-hydroxy-palmitate is formed by the direct hydroxylation of palmitate, rat her than via a a,ß-unsaturated intermediate. The :x-hydroxylase system also has the same stereochemical characteristics as mixed-function oxidases.(87) The mechanisms of a-hydroxylation have not been investigated in fungi. However, und er the conditions described for fatty-acid chain elongation in Candida utilis (see section 4.2 of this chapter), Fulco(67) reported that an enzyme is present that converts hexacosanoic acid (C 26 ) to 2-hydroxy hexacosanoic acid. The enzyme shows a strong preference for the C 26 . The :x-hydroxylation is associated with the :x-decarboxylation pathway in this yeast (see section 4.5.1).

138

Fatty Acid Metabolism

9,10-Dihydroxystearic acid has been detected in certain fungi, and is discussed in the following section.

4.4.3.2. Epoxy Acids Cis-9,1O-epoxyoctadecanoic (9,lO-epoxystearic acid) acid is a major fatty-acid constituent of rust spores (see Chapter 3) and is formed during sporulation on infected tissues. Knoche(118) reported that [14C] labeled acetate, stearate, and oleate are incorporated into the epoxy acid by Puccinia graminis. Light has little inftuence on epoxide formation,(118) and molecular oxygen is required.o 19) Based on the relative rates of incorporation of stearate and oleate, a tentative pathway of9,1O-epoxystearic acid biosynthesis is shown in Fig. 4.12. Although [14C] labeled oleic acid is rapidly converted to the corresponding epoxide, the immediate substrate for the epoxide forming enzyme is uncertain. Since over 95 % of the epoxide resides in the triacylglyceride and glycerophosphatide fractions of the rust uredospore

d. ＢＧｾ＠

F,.,

r'

B;""It,,,;,

CH 3 (CH 2 )16 COOH

I

5teoric Acid Oesoturotion (02)

CH3(CH2)-rCH. ｃｈＨｾＩＷｏ＠ Oleic Acid

_______ 'Oleyl-POLAR or NEUTRAL LlPIO"

,0, ｃｈＺｐｩｾｲ［ＭＨＲＩｏ＠

_ __

ｃｈｪｾＩＷ＠

9.IO-Epoxysteoric Acid

•

I

"Epoxy-POLAR or NEUTRAL LlPIO"

I H,O 9H «H

-CH (CH2lrCOOH 9.IO-Oihydroxysteoric Acid

Fig. 4.12. Tentative pathway of cis-9.IO-epoxyoctadecanoic acid biosynthesis and its degradation to 9,1 O-dihydroxyoctadecanoic acid.

4.4. Biosynthesis of Unusual Fatty Acids

139

oils, it has been proposed that the substrate for epoxidation enzymes may be the complex acyl lipids rather than the free acids or their thiole sters. Oleate may be rapidly incorporated into the complex lipids and then converted to the epoxide (Fig. 4.12). This is analogous to the proposed pathways of oleate desaturation (section 4.3.2 of this chapter) and hydrocarbon biosynthesis in bacteria (see Chapter 2). Cis-9,lO-epoxyoctadecanoic acid of P. graminis has the L-configuration(87.120) and is hydrated to threo-9,lO-dihydroxy stearic acid when the uredospores are incubated in water.(121-123) The dihydroxy acid has the L-9, D-I0 configuration, and, thus, the hydroxyl function at the lO-position originates from water and the oxygen function of the 9-hydroxyl originates from the epoxide(87) (Fig. 4.12). 4.4.3.3.

Methyl Branched-Chain Acids

Substituted fatty acids with a methyl group present in the hydrocarbon chain (Cf. to UJ - 1) are common in bacteria and animals and are produced by some plant species. Based on the few reports available, methyl-substituted fatty acids appear to be sparsely distributed in fungi. A methyl group may be incorporated into the fatty-acid molecule during de noro synthesis or alkylation may occur after completion of the fatty-acid chain. The biosynthesis of iso-(w - 1) and anteiso-(w - 2) branched-chain fatty acids, as intermediates in the formation of aliphatic hydrocarbons, were discussed in Chapter 2. 2-Methyl- and 3-methyl-butyrate (from leucine and isoleucine, respectively) may substitute for acetate during fatty-acid synthesis and may be elongated to the respective branched-chain acids. Methyl branches mayaIso occur at locations other than the iso and anteiso positions and may arise by one of two known mechanisms. Methyl branches may arise by the incorporation of propionate via methyl malonate, rather than malonate, during the chain elongation of fatty-acid synthesis. Placement of the methyl branch depends on the length of the intermediate fatty-acid chain that condenses with methylmalonate and the degree of subsequent chain elongation. Multiple methyl branches may arise if more than one propionate molecule enters the fatty-acid synthetase. Methyl branches mayaIso be introduced into the fatty-acid chain through the alkylation of a preformed unsaturated fatty acid. The mechanism of fatty-acid alkylation is similar to that described in Chapter 6 for the alkylation of ergosterol. For example, a soluble enzyme system has been isolated from the bacterium, Mycobacterium ph/ei, which catalyzes the alkylation of glycerophosphatide-bound oleate to lO-methylenestearate, which is subsequently reduced to 10-methylstearate,o 24) S-adenosylmethionine is the methyl donor in this reaction.

140

Fatty Acid Metabolism

4.5. FATIV ACID DEGRADATION 4.5.1. a-Oxidation The process offatty-acid degradation by iX-oxidation involves the oxidative decarboxylation of an acid in which CO 2 and an acid containing one less carbon atom are produced. The 2- or iX-carbon atom of the original acid (n) becomes the carboxyl carbon of the newly formed fatty acid (n - 1). This pathway is widely distributed in nature and was first studied in germinating peanut cotyledons(125) and then in leaf(126) and mammalian tissues.(127) iX-Oxidation has been included in a review by Stumpf(127) and was discussed by Hitchcock and Nichols.(91) The exact mechanisms of fatty-acid degradation by iX-oxidation are not weIl established. Although they have several properties in common, the !X-oxidation systems differ among the various tissues studied. In germinating peanut cotyledons, the !X-oxidation system contains a peroxidase, which is involved in the decarboxylation of a fatty acid to CO2 and an aldehyde containing one less carbon atom, and an aldehyde dehydrogenase requiring NAD+, which converts the aldehyde to the corresponding acid.(129) Hydrogen peroxide is required and may be genera ted through the action of glycolic acid oxidase on its substrate. The enzymes which catalyze the following reactions in germinating cotyledons are found in the mitochondrial, microsomal, and supernatant fractions: CO 2

RCH 2COOH

...J

RCHO ----. RCOOH ----. etc.

In contrast, the iX-oxidation system from leaf tissues has, as its first stable intermediate, an !X-hydroxy acid and requires molecular oxygen rather than hydrogen peroxide. NAD+ is also required by this system. The !X-oxidation enzymes from leaf tissues are believed to catalyze the following reactions : CO 2 RCH 2COOH ----. L-RCHOHCOOH

..4

RCOOH ----. etc.

Apparently both the D- and L-hydroxy isomers are formed du ring the oxidation process, but it has been shown that only the L-isomer is converted to the corresponding acid. The D-isomer tends to accumulate and appears to be incorporated into the sphingolipids. !X-Oxidation of fatty acids by mammalian systems differs from the two plant pathways. An iX-oxidation system solubilized from rat brain microsomes requires ATP, NAD+, molecular oxygen, and a ferrous ion.o 30) A

4.5. Fatty Acid Degradation

141

keto acid is believed to be an intermediate in this IX-oxidation pathway:

RCHzCOOH

ｾ＠

RCHOHCOOH

ｾ＠

CO z

RCOCOOH

--4

RCOOH ｾ＠

etc.

The catabolism of fatty acids by IX-oxidation has not been extensively investigated in the fungi. Fulco(67) has shown that long-ch&in fatty acids are decarboxylated by the yeast, Candida utilis. This system catalyzes the oxidative decarboxylation of IX-hydroxy acids ranging in chain lengths between C I8 and C 26 . Maximum activity occurs with IX-OH C I8 , but decreases with increasing chain length. Aldehydes containing one less carbon atom than the IX-hydroxy acid have been detected and are presumed to be intermediates in the decarboxylation of fatty acids by this yeast. Thus, IX-oxidation of fatty acids may occur in fungi via pathway similar to that of the germinating seeds.

4.5.2. ß-Oxidation The ß-oxidation pathway offatty-acid degradation is widely distributed in nature and is the principal pathway offatty-acid catabolism. This pathway involves the stepwise removal of C z units from a pre-existing activated fatty acid to acetyl-CoA and an acyl-CoA containing two carbons less. It is called ß-oxidation because the 3- or ß-carbon atom is oxygenated during the process. The enzymes of ß-oxidation are restricted to the mitochondria in animals, where the pathway is linked with the energy metabolism of the cel!. This is also true for plants, and, in addition, ß-oxidation enzymes are present in the glyoxysomes in which fatty-acid degradation is associated with the conversion of lipid to carbohydrate via the glyoxylate cycJe. Unlike the IX-oxidation pathway, the initial substrate for the enzymatic degradation of fatty acids via ß-oxidation is the thiolester of coenzyme A. Three activating enzymes have been identified, each with a specificity for a different range of fatty-acid chain lengths. Acetyl-CoA synthetase (discussed in section 4.2.1.1 of this chapter) catalyzes the activation of acetate and propionate. Two activation enzymes have been identified wh ich catalyze the activation of fatty acids with chain lengths ranging from C 4 to C IZ and from C 8 to C I8 . Activating enzymes are not restricted to the mitochondria, but are found in most fractions of the cel!. As is the case with acetyl-CoA (see section 4.2.1.1), the mitochondrial membrane is impermeable to the longchain acyl thiolesters. An enzyme has been identified that catalyzes the transfer of an acyl moiety of acyl-CoA to carnitine, which can traverse the

742

Fatty Acid Metabolism

ACYL LIPIDS

I

R CH 2 (CH2l NCH2CH 2 COOH AMP,PPi

i

Acyl Dehydrogenose ｒｃｈＲＨｾｎ｟ｏｯａＧ･｣ｹｉ＠

C g

ｒｃｈＲＨｾｎ＠

NADH

CoA, ATP, Mq++

•

ｒｃｾＨｈＬＮ｣Ｒｏｯａ＠

3-Ketoocyl Thiolose

1

/l

Thiokinase

_

CH 3 COCOA CoA

FAD) FADH 2

tI

- CH 2COCoA

•

R ｃｈｾＲｬｎＮ＠

CH COCoA

EnOYI

3 - Hydroxyocyl Dehydrogenose

)

Hydrotose

OH RCH2(CH2l NCH-*CH 2 COCOA

Fig. 4.13. ß-Oxidation cycle.

mitochondrial membrane. The reverse reaction occurs in the mitochondria: Acyl-CoA

+ Carnitine ｾ＠

Acylcarnitine

+ CoA

The ß-oxidation cycle consists of four reactions which may be repeated until the fatty acid is completely oxidized to C 2 units (Fig. 4.13). The first reaction of this cycle is catalyzed by the ftavoprotein enzyme, acyl-CoA dehydrogenase, and involves an IX,ß-dehydrogenation of the acyl-CoA substrate. Three acyl dehydrogenases, with different chain-length specificities, have been identified. Each is a ftavoprotein and contains 2 moles of FAD per mole of protein. The reduced ftavoprotein produced in this reaction is oxidized by a ftavoprotein ofthe electron-transport system. The trans-IX,ßunsaturated (trans-2) acyl-CoA produced in this reaction is converted to L( + )-ß-hydroxyacyl-CoA by an enoyl-CoA hydratase (crotonase). This enzyme has a broad chain-length specificity (C 4 to ClS)' The third reaction of the ß-oxidation cycle is catalyzed by L-3-hydroxyacyl-CoA dehydrogenase and involves the conversion of L( + )-ß-hydroxyacyl-CoA to the corresponding 3-keto acyl thiole ster. This enzyme is highly specific for the Lconfiguration and NAD+. In the fourth reaction, an acyl-CoA transacetylase

4.5. Fatty Acid Degradation

143

(acetyl-CoA transacetylase, or thiolase) catalyzes the cleavage of the 2,3 carbon-carbon bond, and coenzyme A displaces an acetyl-CoA moiety. The single thiolase catalyzes two reactions: o 0 C o 11

11

(1) RCH 2C-CH 2CCoA

+

o

HS-E

11

(2) RCH 2C-S-E + CoA

11

RCH 2C-S-E ｾ＠

11

+ CH 2C-CoA

ｾ＠

0 11

RCHCCoA + HS-E

A yeast thiolase which has a molecular weight of 170,000 and is cornposed of 4 subunits with molecular weights of about 42,000, has been identified. The products of this fourth reaction are acetyl-CoA and an acyl thiolester containing two less carbon atoms. The acyl chain may be recycled through the ß-oxidation cycle until the fatty acid is completely oxidized into C 2 units. Saturated fatty acids undergo these reactions during their degradation via ß-oxidation. The oxidation of unsaturated fatty acids follows this same general pathway, but because ofthe specificities of certain enzymes, additional reactions are necessary. F or example, the degradation of linoleic acid (cis-d 9, cis-d !2-octadecadienoic acid) proceeds normally through the cycle du ring the removal of the first three C 2 units. The product after three turns through the cycle is ciS-d 3, ciS-d 6 -dodecadienoyl-CoA, which contains a cis-ß) (cis d 3) double bond. This bond is not attacked by enoyl hydratase but is converted to the trans-rx,ß(d 2 ) isomer, fungi > animals. In animals, lanosterol is the first cyclic intermediate in the formation of 4-desmethyl sterols. It is now generally accepted that cycloartenol fills that role in plant systems. Lanosterol is also the first cyclic intermediate in the formation of 4-desmethyl sterols by fungi. Gibbons et al. (164) recently reported the resuIts of an investigation on the comparison of the utilization of lanosterol and cycloartenol by a mammalian (rat Iiver) and a higher plant (corn) system to determine whether certain inferences into the evolutionary origins of the major groups represented could be made. They reported that the plant systems can convert radioactively labeled lanosterol and cycloartenol into the predominant 4-desmethyl sterols equally weil, which suggests that plants do not discriminate between the two compounds. Conversely, when similar experiments are carried out with rat liver homogenates, only lanosterol is significantly converted to 4-desmethyl sterols, which indicates that in mammals one or more of the enzymes in this pathway cannot accept cycloartenol. Definite conclusions concerning the evolutionary origin of

202

Sterol Biosynthesis

photosynthetic and nonphotosynthetic organisms cannot be made from this data, but when combined with the fact that lanosterol is the natural sterol precursor in fungi, this data is interpreted as supporting the hypothesis of a partly or completely different origin for these groups. It will be meaningful to determine whether, and to what extent, cycloartenol can be converted to ergosterol or other sterols by fungi. In complex polycyclic organic molecules, such as the sterols, asymmetrie carbon atoms are present, and certain groups at these centers can be arranged in one of two possible steric configurations. Of particular interest is the asymmetrie carbon atom created by alkylation ofthe sterol side chain, which occurs in both plant and fungal systems. Patterson(159) suggested the steric configuration of the Cl or C 2 groups at the C-24 position may have phylogenetic implications. Table 6.1 points out that sterols containing alkyl groups having the 24R (a) configuration are characteristic of higher plants and certain algae, while the 24S (ß) configuration is characteristic of some green algae. The few fungal sterols examined for their C-24 alkyl steric structure have the 24S configuration. The C-20 of the sterol side chain is also an asymmetrie carbon atom and mayaiso be ofphylogenetic significance. Most naturally occurring sterols have .he 20R configuration, although in most studies the steric configuration at this position is assumed, and experimental evidence is rarely given for its confirrnation. Sterols with the 20S configuration have been identified as natural products and include sargasterol (C-20 epimer of fucosterol) and haliclonasterol (C-20 epimer of campesterol), which were isolated from marine brown and green algae, respectively. Sterols with the 20S configuration have not been reported from fungal sourees. The predominant sterols of animals, plants, and fungi have ｾＵ｟ｃＲＷＧ＠ ｾ＠ 5-C 29 , and ｾ＠ 7 -C 28 structures, respectively. As was po in ted out before, the basic steps leading to the formation of major 4-desmethyl sterols is the same in each of these major taxonomie groups. Investigations with stereospecifically labeled sterol precursors have shown that these reactions may differ stereochemically among the major groups of organisms (Table 6.1). For example, the ｾＲ＠ double bond is characteristic of both plant and fungal sterols, but it is not found in the sterols of higher animals. In the biosynthesis of ergosterol by the alga, Ochromonas danica, the 23-pro-Rhydrogen is lost in the formation of this bond, while the oPI?osite is true in the fungus, Asperigillus jumigatus. Similar stereochemical differences are noted in the reactions occurring during the nuc1ear double-bond shifts ＨｾＸ＠ ｾ＠ ｾＵＩ＠ in animals, plants, and fungi. It is weIl documented that the 7ßhydrogen is lost du ring the ｾ＠ 8 ｾ＠ ｾ＠ 7 conversion in representative plant and animal systems, while the 7a-hydrogen is lost in this reaction by yeast. The ｾＷ＠ ｾ＠ ｾＵＮＷ＠ conversion in animal and fungal systems (A. jumigatus) appears to have the same stereochemistry, that is, the cis elimination of the

6.7. References

203

6a-hydrogen. No information has been reported on the stereochemistry of this reaction as it occurs in representative plant systems. Likewise, no information is available for comparison on the ,17 saturation in the ,15,7 -+ ,15 conversion in plant and fungal systems, while in mammalian systems this reaction involves the addition of a hydride ion to the 7a position and a proton to the 8ß position. This discussion on the comparative aspects of the composition and biosynthesis of sterols by representative plant, animal, and fungal systems has been included only for the purpose of pointing out its potential value in a phylogenetic consideration. and not to make inferences into relationships among these groups. Too few representative species of important taxa such as the algae. protozoans. lower animals. plants. and other fungi have been examined in this area to warrant such speculations.

6.7. REFERENCES I. T. w. Goodwin. Blochem. J. 123: 293 (1971). 2. L. J. Goad. in Terpenaids In Plants. (J. B. Pridham. ed.). Academic Press. New York (1967). 3. E. Heftmann and E. Moseltig. Blochemlstry 0/ Steraids. Reinhold Publishing Corp., New York (1960). 4. J. O. Frantz and G. J. Schroepfer. Ann. Ra. Biochem. 36:691 (1967). 5. G. Popjak and J. W. Cornforth, Blochem. J. 101:553 (1966). 6. K. Bloch, Sc/ence 150: 19 (1965). 7. R. C. Ottke, E. L. Tatum, I. Zakin, and K. Bloch, J. Blol. Chem. 189:429 (1951). 8. G. E. W. Wolstenholme and M. O'Connor (eds.) Ciba Foundatlon Symposium on the Blosynthesls 0/ Terpenes and Sterals, Little, Brown and Co., Boston (1959). 9. J. W. Cornforth. 1. Lipid Res. 1:3 (1959). 10. G. Popjak and J. W. Cornforth, in Advances in Enzymology. Vol. 22. p. 281 (F. F. Nord. ed.), Interscience. (1960). 11. K. Bloch, Harvey Lectures 48:68 (1952). 12. R. B. Clayton, Quart. Ret·. 19: 168 (1965). 13. K. Folkers. C. H. Skunk, B. O. Linn, F. M. Robinson, P. E. Wittreich, J. W. Huff. J. L. Gilfillan, and H. R. Skeggs, in Biosynthesis 0/ Terpenes and Sterals, Little, Brown and Co., Boston ( 1959). 14. J. D. Brodie, G. Wasson, and J. W. Porter, J. Biol. Chem. 238: 1294 (1963). 15. B. H. Amdur, H. C. Rilling, and K. Bloch, J. Amer. Chem. Soc. 79:2646 (1957). 16. T. T. Tchen, J. Amer. Chem. Soc. 79:6344 (1957). 17. T. T. Tchen, J. Blol. Chem. 233: 1100 (1958). 18. K. Bloch. S. Chaykin, A. H. Phillips, and A. DeWaard, J. Biol. Chem. 234:2595 (1959). 19. B. W. Agranoff, H. Eggerer, U. Henning, and F. Lynen, J. Blol. Chem. 235:326 (1960). 20. J. W. Cornforth, K. Clifford, R. Mallaby, and G. T. Phillips, Proc. Roy. Soc. (Landon) Sero B 182:277 (1972). 21. G. Popjak, Blochem. Soc. London Symp. 29: 17 (1970). 22. G. Popjak and J. W. Cornforth, Biochem. J. 101:553 (1966). 23. J. W. Cornforth, R. H. Cornforth, C. Donninger, and G. Popjak, Proc. Roy. Soc. (Landon) Sero B 163:492 (1966).

204

Sterol Biosynthesis

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CHAPTER 7

Acylglycerides, G Iycosylglycerides, and Simple Esters

7.1. INTRODUCTION Acylglycerols (glycerides) are esters of fatty acids and glycerol and are the primary constituents of natural fats (so lids) and oils (Iiquids). This class of lipids includes monoacyl, diacyl, and triacyl esters; the latter group is generally found in the highest relative proportions. Since fatty acids are the major source of energy in Iiving systems, yielding over two times more calories per gram upon oxidation than either carbohydrates or proteins, triacylglycerols represent the most efficient form of energy storage. In fungi, triacylglycerols are the major constituents of oil droplets suspended in the mycelial or spore cytoplasm, but the relative abundances depends on the cuIture conditions and stage of growth. These lipids mayaIso be minor constituents of membranes and cell walls of fungi. As noted before, lipids containing carbohydrate moieties are not treated as a separate lipid class (glycolipids) in this text, but are included with the appropriate lipid class having other structural features in common. This chapter deals, in part, with the glycosyldiacylglycerides which are particularly common in plants and bacteria. This chapter also includes the simple esters. These lipids are the methyl and ethyl esters of long-chain fatty acids and the fatty-acid esters of sterols. These simple esters have been reported as fungal products, but few studies have been concerned with their occurrence and biosynthesis in these orgamsms. 209

210

Acylglycerides. Glycosylglycerides. lind Simple Esters

7.2. ACYLGLYCERIDES 7.2.1. Nomenclature and Structure

° 11

H2 2,3diacylglyceride > monoacylglyceride. The stimulation is higher in the early stages of the reaction, but decreases in the later stages. Stimulation by bile salts is apparently not due to enzyme activation, but to the initial increase in absorbed enzyme. Decreases in the rate ofhydrolysis are due to the salt which forms micelIes with the monoglyceride and fatty-acid products of the initial reaction, which interfere with the approach of the enzyme.(34) The rate oflipolysis catalyzed by M ucur lipase is also influenced by the degree of unsaturation in the acylglycerides.(34) Rates of hydrolysis of lipids containing C 18 : 1 and C 18 : 2 are relatively high, but the rate ofthe trilinolenin (containing 3 linolenic acid molecules) degradation is reduced by 75 %. This may be due to the change in the configuration of the fatty-acid hydrocarbon chain caused by the double bonds. In C 18 : 3' the terminal methyl group is close to the carboxyl group (Fig. 7.5), which may interfere by steric hindrance with the enzyme-substrate association. The most extensively investigated lipase of fungal origin is that produced by Rhizopus arrhizus. The first of these studies were conducted by Laboureur and Labrousse(64-66) who reported that the Rhizopus lipase has properties similar to pancreatic lipase but it is extracellular and more stable. U pon purification, the lipase (Lipase I) produced by R. arrhizus is slowly

224

Acylglycerides, Glycosylglycerides, and Simple Esters

GOOR H

H

H

H

H H

GOOR GH 3 H

(bI

(cl

Fig. 7.5. Structures of oleic (a), linoleic (b), and linolenic (c) acids.

converted in the cold to another form (Lipase 11), which is almost as active, more cationic, and even more stable.(67) Lipase I is a glycoprotein having a molecular weight of about 43,000 and contains ＱＳｾＴ＠ molecules of mannose, 2 molecules of hexosamine, and a single N-terminal aspartic acid (or asparagine) residue. It consists of two apparently noncovalently linked portions, a glycoprotein of 8500 molecular weight, and the enzyme. The glycopeptide does not playa role in the catalytic function of the enzyme, since, upon storage, the glycopeptide separates and Lipase I is converted to the second active form, Lipase 11. The role of the glycopeptide is not known, but it may facilitate passage ofthe lipase through the plasma membrane.(68) The lipase produced by C. paralipolytica can be modified by dialysis and separated from an unknown substance which absorbs at 260 nm by Sephadex G-75 column chromatography.(53-56) It has been shown that the Rhizopus lipase has the same positional specificity as the pancreatic lipase, that is, the 1- and 3-acyl groups are preferentially attacked. (25 ) Through the use of 1,3-dipalmityl-2-[9,103H] oleylglycerol, it was shown that acyl migration occurs under the conditions optimum for Rhizopus lipase activity, and the specificity for the 1- and 3positions is practically absolute. Rhizopus lipase can also hydrolyze chylomicrons.(69,70)

7.2.4.3. Stereospecilic Analyses 01 Acylglycerides As was discussed in a previous section of this chapter, there are a large number of possible molecular species of triacylglycerides which may occur

7.2. Acylglycerides

225

in nature. It was also shown that the acyl groups are generally not randomly distributed among the three hydroxyl groups of glycerol, but the fatty acids may be specifically distributed according to chain length or the degree of unsaturation. Hydrolysis of triacylglycerides, followed by gas chromatographic analysis, can only give information concerning the qualitative and quantitative fatty-acid content and not their specific positions on the glycerol molecule. However, methods for the stereospecific analysis oftriacylglycerides have been recently developed, and the structures of individual molecular species can be determined. These methods employ thin-layer (silver-ion argentation) and gas-liquid chromatography combined with the reactions of lipases (with known specifi