No title

Advances in

MICROBIAL PHYSI0LOGY

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Advances in

MICROBIAL PHYSIOLOGY edited by

A. H. ROSE School of Biological Sciences Bath University England

J. GARETH MORRIS Department of Botany and Microbiology University College Wales Aberystwyth

Volume 17

1978

ACADEMIC PRESS London New York San Francisco A Subsidiary of Harcourt Brace Jouanouich, Publishers

ACADEMIC PRESS INC. (LONDON) LTD. 24/28 Oval Road London NW 1 United States Edition published by ACADEMIC PRESS LTD. 111 Fifth Avenue New York, New York 10003

Copyright 0 1978 by ACADEMIC PRESS INC. (LONDON) LTD.

'

All Rights Reserved

No part of this book may be reproduced in any form by photostat, microfilm, or any other means, without written permission from the publishers

Library of Congress Catalog Card Number: 67- 19850 ISBN: 0- 12-0277 17-4

Printed in Great Britain by William Clowes and Sons Limited London, Colchester and Beccles

Contributors to Volume 17 MARGARET M. ATTWO OD, Department o f Microbiology, University o f Shejjeld, England PATRICK J. BRENNAN, Department of Biochemistry, University College, Dublin, Ireland A. D. BROWN, Department ofBiology, University o f Wollongong, Wollongong, N.S.W. 2500 Australia BRUCE L. A. CARTER, Department o f Genetics, Trinity College, University $Dublin, Dublin 2, Ireland W. HARDER, Department o f Microbiology, The University o f Groningen, The Netherlands DOROTHY M. LOSEL, Department o f Botany, University o f Shefield, Shejjeld SIO 2TN,England A. D. WARTH, C.S.I.R.O. Division $Food Research, North Ryde, N . S . W . 21 I3 Australia

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Contents Molecular Structure of the Bacterial Spore by A . D. WARTH

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I1. Spore Morphology . . . 111. Exosporium and Appendages

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I . Introduction

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VI . VII . VIII . IX . X.

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A . Morphology . . . B.ChemicalComposition . Coats . . . . . . . . A . Morphology . . . . . . B . Chemical Composition and Structure . C. Biosynthesis . . . . . . D . FunctionofSporeCoats . . . . Cortexand GermCellWall . . . . A . Morphology . . . . . . B . ChemicalStructure . . . . . C . Lytic Enzynes . . . . . . D . Biosynthesis . . . . . . Core . . . . . . . . . A . Macromolecular Composition . . B . Low Molecular Weight Compounds . Ionic Composition of Spores . . . . Water Content and Physical State of the Core Mechanisms for the Dehydration of the Core Acknowledgement . . . . . . . . . . . . . References

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Physiology of Fungal Lipids: Selected Topics by PATRICK J. BRENNAN and DOROTHY M . LOSEL I . Introduction . . . . . . . I 1 . Location of Lipid in Fungal Cells . . . 111. Lipids and Fungal Membranes . . . . A . Membranes-General Considerations . B . Phospholipids-TypesandDistribution . C. Phosphoglycerides-PhysiologicalAspects D . Glycolipids . . . . . . . E. Sphingolipids . . . . . . IV . Biosynthesis of Fungal Lipids . . . . A . Phospholipids . . . . . . B . Glycolipids . . . . . . . C . Sphingolipids . . . . . .

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CONTENTS

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. V . Role of Lipids in Fungal Morphogenesis . A . Hormonal and Growth-Regulating Factors B . Lipid Reserves in Morphogenesis . . C. Lipid Reserves and Secondary Metabolites D . Lipid Metabolism in Morphogenesis . . VI . Role of Lipid in Fungus-Host Relationships . A . Fungal Associations with Plant Tissues . B . Fungi Associated with Insects . . . C . Fungi Pathogenic to Man and Animals . D . Discussion . . . . . . . References . . . . . . .

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113 114 121 130 133 140 140 155 159 163 164

Compatible Solutes and Extreme Water Stress in Eukaryotic Microorganisms by A . D. BROWN I . Introduction

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11. RoleofPolyhydric Alcohols

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A . General . . . . . . . B . Specific . . . . . . . . PhysiologyofXerotolerance . . . . A . XerotolerantYeasts . . . . . B . Xerophilic Yeasts . . . . . . C . Xerotolerant Fungi . . . . . D . Halophilic Algae . . . . . . E . IntermediateXerotolerance . . . Regulation of Compatible Solute Accumulation Summary . . . . . . . . Acknowledgements . . . . . . References . . . . . . . .

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181 184 184 188 197 198 212 215 215 223 223 237 239 239

The Yeast Nucleus by BRUCE L. A . CARTER I . Introduction . . . . . . . . . I1 . Nuclear Morphology . . . . . . . 111. Nuclear Division . . . . . . . . IV . Yeast Chromosomes . . . . . . . A . Introduction . . . . . . . . B . Histones . . . . . . . . . C. Sizeofyeast Chromosomes . . . . . D . Localization of Genes o n Chromosomes . . . . . V . Initiation of Nuclear DNA Synthesis . . . . . . . VI . Nuclear DNA Replciation VII . Nuclear Control Over Mitochondrial-DNA Replication . . . . . . VIII . Nuclear DNA Enzymes . A . DNA Polymerases . . . . . . . B . DNA-Dependent RNA Polymerases . . . C. Poly(A) Polymerases . . . . . . .

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244 245 247 249 249 251 254 256 260 266 271 271 271 273 280

CONTENTS

Expression of Nuclear Genes . . . . . . Expression of Yeast Genes in Eschen'chca coli . . . Integration of Growth and Nuclear/Cell Division . SomeTechnical Considerations . . . . . A . Isolation of Yeast Nuclei . . . . . . B. Isolation of a Nucleolar Fraction from Yeast Nuclei . . C . Isolation of Chromatin from Yeast Nuclei D . InhibitionofNuclear Functions . . . . E . DNA Estimation . . . . . . . F. Nuclear Staining . . . . . . . XI11 . Conclusions . . . . . . . . . XIV . Acknowledgements . . . . . . . . References . . . . . . . . . IX . X. XI . XI1 .

Biology. Physiology and Biochemistry of W . HARDER and MARGARET M . A-TTWOOD

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Hyphomicrobia by

I . Introduction . . . . . . . . . . . . I1. BiologyandPhysiologyofHyphomicrobia . . . . . . A . The Genus Hyphomicrobium . . . . . . . . B . Enrichment and Isolation . . . . . . . . . . . . . . . . . . . . . C. Nutrition D . Life Cycles and Pleomorphism . . . . . . . . E . Effect of Environment o n Morphology . . . . . . F. Ecology . . . . . . . . . . . . 111. Biochemistry of Hyphomicrobia . . . . . . . . A . Biochemistry of Growth on Reduced One-Carbon compounds . B . Biochemistry of Growth on Two-Carbon Compounds . . . C. Possible Role of Cytochromes . . . . . . . . D . Biochemical Basis for Restricted Methylotrophy in Hyphomicrobia IV . Conclusion . . . . . . . . . . . . . V . Acknowledgements . . . . . . . . . . . References . . . . . . . . . . . . . Author Index . . . . . . . . . . . . Subject Index . . . . . . . . . . . .

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Molecular Structure of the Bacterial Spore A. D. WARTH C.S.I.R.O. Division of Food Research, North Ryde, N.S. W. Australia 21 13

I. Introduction

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11. SporeMorphology

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111. Exosporium and Appendages . . A. Morphology . . . . . B. ChemicalComposition . . . IV. Coats . . . . . . . A. Morphology . . . . . . B. Chemical Composition and Structure . C. Biosynthesis . . . . . . D. FunctionofSporeCoats . . . V. Cortex and Germ Cell Wall . . . . A. Morphology . . . . . . B. Chemicalstructure . . . . C. LyticEnzymes. . . . . . D. Biosynthesis . . . . . . VI. Core . . . . . . . . A. Macromolecular composition . . B. Low Molecular Weight Compounds . V I I . Ionic Composition of Spores . VIII. Water Content and Physical State of the CoEe IX. Mechanisms for the Dehydration of the Core X. Acknowledgement. . . . . . References . . . . . . .

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I. Introduction

In the differentiation of a bacterial cell into a spore, a number of new morphological structures are formed. The cortex and germ cell wall are specialized adaptations of vegetative structures. Spore coats, 1

2

A. D. WARTH

exosporia and appendages are new structures embodying new classes of microbial products. Their novelty is made possible by the strategy of intracellular synthesis and assembly in the sacrificial sporangial cell. Within the spore cell, or core, unique constituents such as dipicolinic acid, and very different proportions of normal metabolites and electrolytes are found. Some of the intracellular macromolecules have spore-specific modifications, but in the main, a normal complement of enzymes, ribosomes and nucleic acids is present. The mature spore has very well known properties of resistance to heat, radiation, enzymes, disinfectants and other deleterious agents and an absence of endogenous metabolism. In this article, knowledge of the composition and structure of the spore cytoplasm and each of the spore integuments will be reviewed, and the contribution of each component to determining the essential properties of the spore will be considered. Of particular interest is the heat resistant and ametabolic state of spores. This appears to be mainly a consequence of a reduced water content in the core. The final section discusses the possible chemo-mechanical properties of the cortex, and examines models for its role in the dehydration of the core.

11. Spore Morphology

Spores of all species have the same basic structure. For example, the spore of Bacillus cereus (Fig. 1 ) has a central core (c) or protoplast, surrounded in turn by a plasma membrane (pm), germ cell wall (gcw), cortex (cx), coats (cts) and exosporium (ex.).The core, plasma membrane and germ cell wall constitute a condensed cell, which is contained within and is protected by the outer integuments. Much variation between species is found in the complexity of the coats. Even greater variation is found in structures external to the coats. An exosporium as seen in the spore of B . cereus (Fig. 1) is found in only a few species but more elaborate structures, termed appendages, are common among Clostridium spores (Rode, 197 1). Parasporal bodies of various forms are found in some Bacillus species, mostly insect pathogens. The best known of these are the large protein crystals formed by strains of B. thuringiensis. More detailed information on spore morphology is given in reviews of spore formation by Fitz-James and Young (1969)and Murrell(l967) and in a freeze etching study by Holt and Leadbetter (1969).

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

3

111. Exosporium and Appendages A. MORPHOLOGY

The exosporium of B. cereus consists of an outer layer of hair-like projections ( 2 5 nm thick), an intermediate layer (6 nm) and a basal layer (19 nm). The basal layer has several layers arranged in a hexagonally ordered lattice structure (Gerhardt and Ribi, 1964). Similar structures are found in B . fastidiosus (Holt and Leadbetter, 1969), some strains of B. megaterium (Beaman et al., 1972) and some Clostridium species (Samsonoff et al., 1970; Hoeniger and Headley, 1969). In some other strains and species, slightly different structures are seen (Hodgkiss et al., 1967; Mackey and Morris, 1972). The exosporium may be loosely fitting as in B . cereus or it may be tightly fitting or even integral with the coats. A recent observation of an exosporium in spores of B . subtilis revealed only after partial extraction of the coats, suggests that exosporia may be widespread occurrences but are often obscured by the dense outer coat (Sousa et al., 1976).

FIG. 1. Electron Micrograph of Spore of Bacillus cereus T showing: Core ( c ) ,plasma membrane ( P M ) , germ cell wall (ccw),cortex (cx), coats (CTS) and exosporium (EX).

A. D. WARTH

4

Appendages are loosely fitting structures of very diverse form that are commonly found on spores of Clostridium species. An excellent review is available (Rode, 197 1). Clostridium taeniosporurn spores have fifteen to twenty large ( 4 4 pm) ribbon-like appendages attached through a hook-like structure to the trunk which is continuous with the spore coats. An upper layer 9 nm thick overlies an electron-transparent layer of 3 nm. The main layer is about 100 nm thick and consists of multiple layers of 5 nm spherical subunits (Rode, 197 1). B. C H E M I C A L C O M P O S I T I O N

Chemical analyses are only available for exosporia of B. cereus T and appendages of C1. taeniosporum. Both are voluminous loosely fitting structures which are easily removed from the spore. Sonication was TABLE 1. Composition of the exosporium from Bacillus cereus T and the appendage from Clostridiumtaeniosporum. Exosporium

Appendage

%ofdry weight

Protein ( 17 amino acids) Neutral lipid Phospholipid Total P Neutral carbohydrate (as glucose) Glucose Glucosamine Rhamnose Dipicolinic acid Muramic Acid Diaminopimelic acid Ribose ~

~~

52.1 12.5 5.5 1.8 9.1 3.8 6.4 0.2

ND ND 0.7

79.7

10.4 3.7 4.9

+

ND ND ND

~~

N D indicates that none was detected. Data for B . cereus T from Matz et al. (19701, and for C . taemosporum tram Yolton etal. (1972).

used for isolation of appendages (Yolton et al., 1972) and passage through a needle valve under high pressure for exosporium (Gerhardt and Ribi, 1964). Estimates of the amount of exosporium range from 2% (Matz et al., 1970) to 10% (Gerhardt et al., 1972) of the spore dry weight.

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

5

In some aspects of their composition, the two preparations were similar (Table 1). In both, protein was the major component and both contained significant amounts of carbohydrate made up of glucose, rhamnose and glucosamine residues. Lipid composition was not reported for appendages but evidently it is much less than the 18% TABLE 2. Amino acid composition of exosporium appendages a n d coats. Amino Acid

Clostridiurn taeniosporum

Bacillus cereus Exosporium'

Coatb

AppendageC

CoatC

Moles/100 moles Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Pheny lalanine Lysine Histidine Arginine Cystine

+

13.5 6.8 6.2 12.0 4.2 6.8 5.5 6.8 0.9 5.8 7.5 6.2 4.5 3.1 2.1 6.5 1.2

11.3 7.2 6.1 8.2 4.9 11.3 7.8 6.8 5.1 7.2 4.7 5.4 4.5 2.5 3.8 3.4

14.9 9.8 6.5 7.5 6.8 11.9 5.6 7.6 0.6 5.4 4.1 4.6 3.0 5.0 0.6 2.8 2.7

13.8 4.6 5.2 10.4 4.1 10.6 5.7 5.0 2.0 4.4 6.0 4.6 3.6 9.7 1.7 4.9 3.6

References: a Matz et al. (1970); bAronson and Fitz-James (1968); 'Yolton et al. ( 1 9 7 2 ) .

found in exosporium. The amino-acid compositions of the exosporium and appendages are similar and resemble that of the spore coats (Table 2). Contamination with cell-wall material and ribonucleic acids was low, as shown by the small amounts of muramic acid, diaminopimelic acid and ribose detected. O n the other hand, some caution may be necessary with lipid analyses as lipid may be absorbed from the sporulation medium which contains lysed sporangia. The phospholipid in B. cereus T exosporium was almost entirely disphosphatidylglycerol (Matz et al., 1970). In other studies using whole spores of B. cereus T (Lang and Lundgren, 1970) and B . megaterium (Bertsch et al., 1969), diphosphatidylglycerol was found in a readily extractable form and could have originated in the exosporium. Exosporium lipid

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A. D. WARTH

was not significantly different from whole spore lipid in its fatty acid composition. Straight chain n-C,, and n-C,, fatty acids predominated, and branched-chain fatty acids were present in much lower amounts than is common among Bacillus species (Kaneda, 1967)or was found in vegetative cell membranes of B . cereus T (Beaman et al., 1974).Treatment of exosporia with phenol plus acetic acid, or with sodium dodecyl sulphate (SDS), solubilized components probably from the basal layer. The SDS extract contained 15%of the exosporium protein and consisted of spherical particles 11-44 nm in diameter, which on dialysis, spontaneously re-aggregated into sheets having a hexagonal lattice structure similar to that of the basal layer (Beaman et al., 197 1). This propensity for self assembly is consistent with the formation of the exosporium in the cytoplasm of the mother cell, apparently unaided by pre-existing cytoplasmic structures (Ohye and Murrell, 1973). Relative to exosporia, re-aggregated exosporia were enriched in lipid and contained 39%protein, 33%lipid and 12% carbohydrate. A component chemically related to the complex carbohydrate of the exosporia and appendage may be a common feature in spores, despite the difficulty with some species of recognizing exosporia in electron micrographs. Spores of a number of Bacillus species contained glucosamine in excess of the stoicheiometry required for peptidoglycan (Murrell and Warth, 1965) and a carbohydrate content of 1 to 5% is typical of spores (see Murrell, 1969). In spores ofB. subtilis and B . cereus, rhamnose, glucose and minor saccharides were present (Warth et al., 1963). The carbohydrate components could be associated with the delicate nap seen on exosporia or they may be present in the capsular material which very commonly engulfs spores. Walker (1969) looked for the location of carbohydrate in spores of B . cereus by oxidation of thin sections with periodate and staining with silver. Unexpectedly this procedure did not stain the exosporium, but did stain the developing cortex. However, the structure established for spore cortex peptidoglycan (Warth and Strominger, 1972) does not have periodate-sensitive bonds in the glycan chains, and other saccharides were not detected in cortex preparations. N o direct evidence exists as to the function of exosporia and appendages. Tipper and Gauthier (1972) suggested a function for the exosporium in controlling assembly of coat subunits during spore formation. Exosporia and appendages do not appear to be significant permeability barriers. Openings are present in appendages (Rode, 197 1) and in exosporia of Cl. pasteurianum (Mackey and Morris, 1972) and B. megaterium (Beaman et al., 1972). Solute uptake studies on B .

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

7

cereus did not distinguish an effect attributable to limited permeability of the exosporium (Gerhardt et al., 1972). Together with capsular material, and cell wall layers that in some species persist after sporangial lysis, exosporia and appendages usually form the outermost layer of the spore. As such they determine superficial properties such as adhesion, and antigenicity, which are of obvious ecological importance. I t is perhaps relevant that discrete exosporia appear to be very common in toxic o r pa thogenic species. IV. Coats A.

MORPHOLOGY

Spore coats show an interesting variety both in appearance and in complexity. Some examples are shown in Fig. 2. Three main types of layer can usually be distinguished in thin sections. The most distinctive is the middle layer which shows a very characteristic laminar pattern. This pattern is well developed in the spore coat of B . cougulans which has about seven lamellae spaced 5-7 nm apart. Beneath the laminated coat layer is a region of poorly structured material sometimes referred to as undercoat. Other less consolidated material, including possible remnants of the mother cell cytoplasm and the forespore membrane, may constitute the inner boundary of the coats. Outer coats particularly, vary in complexity between species. At one extreme, some species have heavily ridged and ornate coats (Bradley and Franklin, 1958; Murphy and Campbell, 1969).Thin sections show these to be of complex morphology (Fig. 2d; Holbert, 1960; Leadbetter and Holt, 1968). The spore of B . coagulans (Fig. 2b) has a simpler, thick, heavily staining layer, while that of B . cereus appears to lack an outer coat (Fig. 2a). Structures equivalent to the exosporium are integral with the coats in some species (Leadbetter and Holt, 1968; Beaman et ul., 1972). Freeze etching also reveals interesting details of coat structure (Holt and Leadbetter, 1969) but correlation with features seen in stained sections is not straightforward. A characteristic array of parallel fibrils, about 5 nm in diameter, is generally present and these probably correspond to a laminated layer. Often the fibrils wrap around much of the spore, but in spores of B . cereus they are present as smaller patches or domains. Underlying the fibrillar layer is a pitted layer. During sporulation, coats are formed in the mother cell cytoplasm and not on the cell or forespore membranes (Ohye and Murrell, 1973).The

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D. WARTH

morphogenesis of the spore coat and its morphological and chemical structure have been comprehensively reviewed recently by Aronson and Fitz-James (1976). B. CHEMICAL COMPOSITION A N D STRUCTURE

Spore coats consist very largely of structural protein. Smaller amounts of complex carbohydrate and lipid are generally also found and in some species quite large amounts of phosphorus. Coats can be prepared by mechanical disruption of spores, followed by extensive washing with buffers to remove soluble cytoplasmic components, and digestion with lysozyme to eliminate the cortex and germ cell wall. Such preparations comprise 30 to 60% of the spore dry weight and 40 to 80% of the spore protein. Electron microscopy shows the presence of the major morphological structures of the spore coats including the laminated inner coat, the more diverse and complex outer layers and, where present, the exosporium. Soluble, finely dispersed and protease-sensitive material can be lost. Cytoplasmic membranes are usually eliminated during the washing procedure, but the fate of the outer forespore membrane and the poorly structured material often seen between the cortex and the inner coats is not clear. In general, electron microscopy of thin sections does not give a clear indication of the chemical integrity of spore coats. Large proportions of the weight of the coats can often be extracted with little change in the appearance of the different coat layers and the presence of capsular material and close fitting exosporia is difficult to detect. Spore-coat components can therefore be lost if, during cleaning of the spores or spore coats, treatments with detergents, proteases, alkali or sonication are used, even though these treatments may not affect the viability, heat resistance and refractility of spores. Spore coats are substantially resistant to proteolytic enzymes and to a wide variety of chemical reagents. Part of the coat structure normally remains insoluble after all treatments short of severe hydrolysis or oxidation. The most useful agents for extracting coat components have been disulphide bond-breaking reducing agents, alkali, sodium dodecyl sulphate and urea. Despite the often striking differences in the morphological structure of the different spore coat layers, there appears to be remarkable uniformity in the extractable protein components of each layer. Unlike most other species, the spores of B . cereu5 T and B . megaterium KM have coats which can be almost entirely

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

9

FIG. 2. Electron micrograph showing structure of spore coats (a) Bacillus cereus T ib)

Bacillus coagulans ( c )Bacillus stearothermophilus id) Bacillus apiarius.

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A. D. WARTH

solubilized under mild conditions. Extraction of B. cereus T spore coats (Aronson and Fitz-James, 1968) or spores (Aronson and Horn, 1972) with dithioerythritol at pH 10.3 dissolved 82% of the coat protein with the concommitant disappearance of the inner or undercoat but not of the outer laminated layer. The extract contained only protein, which except for a lower cystine content closely resembled total coats in its amino-acid composition (Tables 2, 3). Further extraction in the presence of sodium dodecyl sulphate solubilized most of the remaining coat protein together with some polysaccharide, and eliminated the main structural features of the coats visible in the electron microscope. The residue, comprising less than 5% of the total protein, was mainly exosporium and contained lipid, protein and a small amount of carbohydrate. Both the dithioerythritol and sodium dodecyl sulphate extracts appeared to consist principally of the same polypeptide of about 12,000 molecular weight (Aronson and Horn, 1972). Apart from a small amount of high molecular-weight material which was attributed to aggregation, both extracts showed a single peak on gel electrophoresis, gel exclusion chromatography and sucrose gradient centrifugation. After dansylation and digestion of the extract with keratinase, three dansyl peptides were isolated. Their composition was consistent with a common amino terminal sequence for the polypeptide of: NH,-Ser-Gly-(Glu, Thr), in which the terminal serine residue was sometimes absent. Coat protein extracted from whole spores had mainly amino- terminal serine, whereas coat extracts, which presumably had suffered more exposure to peptidases, yielded more amino terminal glycine. The major extractable protein appears to be very similar in different species. With spores of B . subtilis, 85% of the coat was solubilized with sodium dodecyl sulphate and dithiothreitol and the major protein component had a molecular weight of 14,000. Serine was the major amino-terminal residue (Mitani and Kadota, 1976). Sodium dodecyl sulphate extracts of B . thiaminolyticus spore coats contained 53% of the protein and showed a single band of about 15,000 molecular weight on gel electrophoresis (Watabe et al., 1975). Urea plus mercaptoethanol extracts of spores of four Bacillus species and C1. bzfrmentans behaved identically on gel electrophoresis, showing a single major band. Their amino-acid compositions were very similar to each other and to the extract of B. cereus T spores (Table 3) and each showed partial antigenic homology with the crystal protein from sporangia of B . thuringzensis (Somerville et al., 1970). Similar results were obtained for a different

TABLE 3. Amino Acid Composition of Spore Coats and Crystal Proteins (For abbreviations see foot of table.) Bacillus thuringiensis alesti' berliner'

Bacillus cereu5 T a

Bacillus

Bacillus subtilis'

Banllus subtilis'

Bacillus subtilis"

Five species'

cereus

Fraction extracted

coats

coats

coats

coats

coats

spores

Extractant

50mMDTE 50mMME PH 10.5 PH 10.2

-

50 mM DTT 50 mM DTT 8 M urea, 1.6 M ME pH 10.5 pH 10.0 pH 8.5 -

Species

-

crystal

crystal

coats

8 M urea, pH 8.0

1.6M ME

z 0 E 0

;

Moles/100 moles

3

cn Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine 4 Cystine Tryptophan

11.3 6.8 5.8 7.9 5.3 11.6 8.3 7.0

-

5.4 7.9 5.0 5.3 4.3 2.5 3.8 1.2

-

13.1 6.7 6.2 10.4 5.3 10.1 8.4 7.9

1 .o

4.7 8.6 2.0 4.9 6.7 1.6 2.1 1.8

-

6.9 4.6 7.6 6.0 2.8 21.1 7.0 4.4

-

3.2 3.6 11.2 6.2 6.6 3.0 4.0

-

9.7 9.5 6.6 6.8 6.9 11.5 8.4 7.5

-

5.5 7.5 1.5 6.8 4.0 2.1 4.5 0.5

-

12.2 5.0 7.3 6.8 3.6 14.3 8.7 5.2 1.7 4.3 5.6 7.1 4.6 6.1

2.0 4.9 0.8 -

10.3-10.9 6.1-10.5 5.4-8.7 6.6-10.9 4.8-7.6 10.6-13.3 7.&12.0 7.4-8.0

-

10.6

9.1 6.8 8.5 6.4 11.6 9.3 7.7

12.4 6.3 7.3 12.0 4.8 7.7 5.4 8.1

-

-

5.8-8.2 6.8-8.8 1.2-3.2 3.6-5.5 3.5-5.2 1.6-2.1 1.4-3.8

6.6

6.0

7.8 2.1 4.7 4.1 1.9 2.7

8.3 4.8 5.2 3.2 2.2 6.4

-

-

-

11.2 6.2 6.1

12.7 4.2 8.8 7.2 6.7 1.9 5.0 8.3 3.3 4.1 4.6 2.2 5.0 0.9 1.4

-I p -I C p

m

$ -I

I

7 m

? v)

p

-o

Abbreviations: ME, 2 mercaptoethanol; DTE, dithioerythritol; DTT, dithiothreitol. References: "Aronson and Fin-James (1968); bSomerville and Pockett (1975); 'Spudich and Kornberg (1968); "Mitani and Kadota (1976); CSomervilleetal., (1970);fLecadetetal., (1972). a Data are given as moles of amino acid per 100 moles total amino acid recovered. Tryptophan, methionine and cystine analyses are often not reported. -..

12

A. D. WARTH

strain of B. thuringiensis by Lecadet et al. (1972). Some differences in amino-acid composition are apparent (Table 3) and in distinction to B. cereus T spore coat protein, phenylalanine was the principal amino-terminal amino acid of the extracted protein, although aminoterminal serine and methionine were found in intact coats. Lysine appeared to be the carboxyl-terminal residue. Peptide maps of keratinase digests of protein from spore coats of B. cereus T, B. subtilis 168 and B. megaterium KM gave very similar patterns of about twelve peptides (Aronson and Fitz-James, 1975) and coat proteins from B . cereus T and B . megaterium were interchangeable in in uitro reconstitution of spore coat layers (Aronson and Fitz-James, 197 1). A considerable amount of evidence has been obtained to indicate a close relationship between the protein of the parasporal crystal formed by strains of B. thuringiensis and the extractable protein of the spore coats. Amino-acid compositions are similar but not identical (Table 3), identical bands are obtained on gel electrophoresis, considerable antigenic homology exists, and both proteins are similar in the conditions required for solubilization and in their tendencies to reaggregate (Delafield et al., 1968; Somerville et al., 1968, 1970; Lecadet et al., 1972). Each contains the same amino and carboxyl terminal amino acids (Lecadet et al., 1972). Maps of the tryptic peptides from performic acid-oxidized protein of crystal and spore extract also were identical (Somerville et al., 1970). Spore extracts of several Bacillus species and acrystaliferous strains were all toxic to larvae of Lepidoptera species. Purification of the toxin from B . cereus strain 64a gave a protein of molecular weight approximately 32,000 very similar to B . thuringzensis crystal protein in composition (Table 4) but with a much lower specific toxic activity. Exosporium protein from B. cereus T also has similarities to this coat protein fraction, both in composition (Tables 2, 3) and in its tendency to re-aggregate (Beaman et al., 197 1). Morphologically the parasporal crystal seems to be formed on the developing exosporium (Somerville and James, 1970; Somerville, 197 1) and the crystal-specific antibody reacted with the inner layers of the spore coat and exosporium (Short et al., 1974) The foregoing work was based upon extraction of spores or coats with urea or sodium dodecyl sulphate in the presence of reducing agents. Kondo and Foster (1967) extracted spore coats of several Bacillus species with alkali. Mild sonication then dispersed a “paracrystal” fraction leaving a resistant residue of from 50 to 70% of the coat weight. Analyses o f these fractions from B . megaterium showed

TABLE 4. Amino acid composition of alkali-soluble proteins from spore coats

Amino acid

Species -.

Bacillus megatenurn

Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine 4 Cystine Tryptophan Ornithine

Alkali Extract'

Paracrystal fraction"

9.8 4.0 5.1 7.5 6.5 12.1 5.6 4.0 1.1 3.2 5.1 11.6 4.7 7.5 4.8 6.2 1.3

15.0 2.9 6.6 8.3 1.2 16.0 4.7 2.3 1.o 2.0 0.3 2.1 6.7 2.5 2.4 5.6 8.2 -

-

-

Residue" 11.0 5.1 6.1 10.0 4.8 6.2 4.7 3.8 3.5 5.8 0 4.9 5.4 18.4 3.1 4.5 1.9 -

Bacillus coagulans

Bacillus

Clostridium

cereus

sporogenes

Bacillus subtilis

Alkali extractb

Alkali extractb

Alkali extract

Alkali extractC

8.2 6.3 5.6 12.6 8.1 6.1 7.1 1.5 5.3 7.2 7.1 4.3 8.8 5.3 4.2 0 2.1

11.6 7.7 3.7 12.7 9.7 9.3 7 .O 1.5 5.5 8.0 3.5 4.2 6.3 2.6 5.4 0.3 0.9

10.0 7.1 5.3 11.1 10.2 8.7 8.1 1.8 4.5 9.3 4.0 5.0 7.6 3.6 3.1 0

8.1 3.4 4.3 8.1

0

References: a Kondo and Foster (1967).Coats were extracted with 0.06 N NaOH at 5 O O C . The paracrystalline fraction was solubilized by sonication. b Gould et al. (1970). Coats were treated with 7 M urea and 10%mercaptoethanol pH 2.8 before extraction with 0.1 N NaOH at 4°C. cWood (1972). Extracted as in Gould eta!. (1970).

8.2 8.3 4.4 3.4 1.1 3.2 4.1 12.9 2.8 14.6 4.4 8.9

-

3 rn

k 50

cn -I

W

i c W

rn

$ -I

I m

rn

E D cn I-

T

0 2 m

14

A. D. WARTH

some marked differences in amino-acid composition (Table 4). The paracrystal fraction clearly resembles sodium dodecyl sulphate and dithioerythritol extracts in composition and properties. It has high glycine, cystine and aspartic acid contents. Its colloidal suspension was cleared by sodium dodecyl sulphate to give a heterogeneous solution of high molecular-weight components which dissociated into small units in the presence of mercaptoethanol. Partial reduction of the paracrystalline fraction led to aggregation and precipitation. The alkali extract had quite a different amino-acid composition from the paracrystalline fraction, having a high tyrosine and low cystine content (Table 4). Other workers (Delafield et al., 1968; Gould et al., 1970; Wood, 1972; Somerville and Pockett, 1975) have examined alkali extracts of whole spores, often after prior treatment with urea and mercapto-ethanol. Extracts from a number of species showed a general similarity in amino acid composition to the alkali soluble fraction of B . megaterium coats (Table 4) and comprised from 1 to 6%of the weight of the spores. The protein from B. subtilis behaved as a single species on ion-exchange chromatography and gel filtration. Both very large and small molecular weight components were present but each appeared identical by immunological criteria. Treatment with sodium dodecyl sulphate, mercapto-ethanol and urea at 100°C effectively disaggregated the higher molecular weight components, and after sodium dodecyl sulphate gel electrophoresis only two bands were seen, corresponding to molecular weights of 10,000 and 56,000 (Wood, 1972). The resistant residue of B . megaterium spore coats after alkali treatment and sonication had a distinctive amino acid composition, high in lysine, aspartic acid and glutamic acid but deficient in cystine (Table 4). Most of the phosphorus of the coats was present in this fraction (Kondo and Foster, 1967). Although in the case of spores of B . cereus, some B . megaterium strains and some Clostridium species, a major part of the coat protein can be solubilized, this is not typical, and with spores of many other species a substantial resistant residue remains. Possibly this is correlated with the presence of coat layers external to the laminated coat, including possible equivalents of the exosporium. Minor coat components such as lipid, complex carbohydrate, hexosamine and phosphate (Warth et al., 1963) are reminiscent ofexosporia (Table 1) and the superficial naplike morphological structure in some species is also similar (Holt and Leadbetter, 1969). The phosphorus content of spore coats differs considerably between species (Murrell, 1969). However, some of the higher values may have resulted from

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

15

contamination of the coat preparation with inorganic phosphate precipitates. Serine phosphate linkages have been demonstrated in B . subtilis spore coats (Sano et al., 1975; Kondo et al., 1975) and galactosamine phosphate in those ofB. megaterium (Kondo et al., 1975). Earlier reports of a phosphomuramic acid polymer appear to be incorrect. Other unusual chemical features of spore coats include taurine and a very high glutamic acid content in the ridged coats of B. breuis 636 spores (Warth et al., 1963). Contrary to a previous report, bacitracin does not appear to be a component of B. lichen$ormis spore coats (Marschke and Bernlohr, 1970). The reported presence of t--(aspartyl)lysine links in the spore coats of B. sphaerzcus (Tipper and Gauthier, 1972) points to an unusual type of cross-link between peptide chains in this species. Other types of cross link such as the t.-(y-glutamyl)-lysine found in keratins from the medulla (Pisano et al., 1968) have not yet been reported in spore coats. Spore coats have a number of similarities with the keratins, including X-ray diffraction patterns (Kadota et al., 19651, extraction properties and resistance to enzymes. Both are complex morphological structures containing a number of structural proteins including some rich in cystine. A number of specific differences and similarities are evident but a detailed comparison does not seem warranted at present as knowledge of coat structure is still fragmentary and keratins consist of several groups of' proteins with diverse properties and compositions (Bradbury, 1973). Compared with the efforts expended on the study of structural proteins from higher vertebrates, spore coat chemistry has so far received trivial attention. Progress had been made, especially in the B. cereus system, to a point where questions can be more clearly stated and the major practical problems are apparent. In many ways, the properties and the problems such as solubilization techniques, tendencies to aggregate, heterogeneity and complex morphology are similar to those of the keratins but hopefully will prove somewhat less complex. In particular, the use of stable alkylated or oxidized derivatives of the reduced proteins has proved essential in the keratin studies but has been little used in spore coat investigations and may help resolve questions of polydispersity of molecular weight and heterogeneity. Many of the spore-coat studies have not described precautions taken to prevent oxidation of thiol groups during manipulation of reduced protein, nor has the extent of reduction or denaturation achieved been apparent in some cases. These conditions together with the need sometimes to work near the lower limit of p H value for solubility, are conducive to aggregation and molecular weight heterogeneity. Despite these diffi-

16

A. D. WARTH

culties it is clear that a degree of uniformity exists in the major polypeptides within and, to a lesser extent, between species. Whether there exist a few small polypeptides o r families of closely related polypeptides, will require genetic analysis o r sequencing studies lor resolution. Investigation of the chemical linkages and polypeptides in the resistant fraction is understandably more difficult and may require much work to develop suitable chemical o r enzymatic techniques. Unfortunately this resistant fraction is often a major part of the coat. The major challenge lies in describing the processes involved in the assembly of polypeptides into the morphological structures of the spore coats. C. BIOSYNTHESIS

Construction of spore coats is a major activity of the mother cell during spore formation. Two aspects of coat formation can be distinguished; first, biosynthesis of coat polypeptides, and second, aggregation, modifications, additions and re-arrangements involved in the assembly of the morphological structures. Spore-coat structures first appear in electron micrographs after the forespore has contracted and concommitant with cortex formation and the beginning of refractility (e.g. see Fig. 4). Completion of coat formation is a very late event, occurring at the same time as full refractility and heat resistance (Fitz-James and Young, 1969). Different coat components may be synthesized at different times. In B. subtilis, incorporation of phenylalanine into coat precursors was a late event, occurring maximally at the time of appearance of the morphological structures, but the fraction soluble in sodium dodecylsulphate with dithiothreitol was labelled slightly earlier than the resistant fraction (Spudich and Kornberg, 1969). Serine-phosphate linkages were also synthesized late in spore formation (Sano et al., 1975). In contrast, the alkali-soluble coat protein was synthesized at a uniform rate from very early in spore formation (Wood, 1972). In B . cereus, unlike B . subtilis, incorporation of amino acids was continuous from an early stage, suggesting the accumulation of precursor proteins (Aronson and Fitz-James, 1968; Aronson and Horn, 1969). Cystine incorporation, however, was much greater late in spore formation Winter, 1959; Aronson and Fitz-James, 1968). Some cystine appears to be incorporated directly into outer coats of B. cereus by disulphide interchange reaction with coat protein thiols o r disulphides. Reduction of coats released free cystine, predominantly from the sodium

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

17

dodecyl sulphate-soluble outer coat fraction. About one mole per mole of coat polypeptide was released (Cheng et al., 1973). Furthermore, the in vitro reconstitution of spore coats as judged by antigenic (Horn et al., 1973) or morphological criteria (Aronson and Fitz-James, 197 l ) , was facilitated by cystine. On the other hand Setlow and Kornberg (1969) could not show Cys-S-protein links in B . megaterum spore coats. Much of the cystine for coat formation comes from a pool of' reduced glutathione. Limitation of cysteine availability results in lysozyme sensitive spores, presumably defective in outer coat formation. Achievement of this cysteine deprivation required the use of mutants defective in both cysteine synthesis and glutathione reductase (Cheng et al., 1973). Since, in B. cerem T, the major polypeptide ofboth the inner and outer coats appears to be the same (Aronson and Horn, 19721, the major difference in their morphological structure must be caused by packing and conformational differences, perhaps mediated through disulphide-sulphydryl interchanges between polypeptides and some cysteine. Minor components may also be important. The presence of minor proteins up to 20% of the total coat protein has not been excluded, but the occurrence of very cystine-rich proteins such as are found in keratins has not been reported. D . FUNCTION O F S P O R E COATS

Coats have no significant role in the heat, and ultraviolet radiation resistance mechanisms of the spore. Disruption of coat structure by mutation (Cassier and Ryter, 197 1 ; Aronson and Fitz-James, 19751, inhibition of synthesis (Fitz-James and Young, 1969) or extraction of coat proteins (Gould et al., 1970; Somerville et al., 1970; Aronson and Fitz-James, 1971; Wood, 1972; Vary, 1973) generally gives spores which are heat and radiation resistant and which retain their calcium and dipicolinic acid (DPA) contents and their refractility. Such spores are, however, made sensitive to lysozyme and octanol and may show differences in their response to germinants. These latter properties suggest a role for the spore coat as a protective permeability barrier preventing access of lysozyme to the sensitive cortical peptidoglycan. In conjunction with the exosporium, the coats doubtless function to protect the interior parts of the spore from a wide range of deleterious substances, particularly surfactants and enzymes. Their own extreme resistance to enzymes and other agents may well be due to the complementary resistance of the various components, hence their morpho-

18

A. D. WARTH

logical and chemical complexity. Much of their stability appears to result horn hydrophobic interactions and covalent cross-linking. Coats also provide obvious mechanical protection and, together with other superficial layers, determine the physical properties of the spore surface which affects binding and dispersal from surfaces. Spores have quite unusual surface behaviour as exemplified by their tendencies to form films on glasslair surfaces and their hydrophobic behaviour in two phase systems (Sacks and Alderton, 1961). V. Cortex and Germ Cell Wall A. MORPHOLOGY

The cortex in mature spores appears as a featureless, electrontransparent zone between the core and the coats. When the spore germinates, the cortex loses its refractiveness to staining and a fibrous network can be seen. At its inner surface is a more dense layer which develops into the cell wall of the emergent cell while the cortex lyses. The inner layer has been termed cortical membrane, primordial cell wall and germ cell wall. O n disruption of the spore and in preparations of the spore integuments, the cortex swells greatly, revealing a fibrous network which is now readily stained by heavy metals. Under conditions where the cortex is swollen, its inner surface and the germ cell wall become folded; see figures in Warth et al., (1963);Murrell and Warth (1965) and Fitz-James and Young (1969). The folding appears to result from anisotropic swelling of the cortex. Swelling occurs along the radial axis but no extension of the surface dimensions is evident. Thus, in a fragment of a spherical shell, expansion is directed inwards with distortion of the inner surface. B. CHEMICAL STRUCTURE

Treatment of disrupted spores with lysozyme dissolves the cortex and usually the germ cell wall. Structural determination of the digestion products from B. subtilis spores indicated the structure shown in Fig. 3 (Warth and Strominger, 1972). Essentially identical results were obtained with spores of B . sphaericus (Tipper and Gauthier, 1972), B. cereus, B. megaterium, B . stearothevnophilus and Cl. sporogenes (Warth and Strominger, 19721, and spores of eight other Bacillus species (A. D. Warth, unpublished observations) appear also to have the same struc-

35% r

47%

A

V

18%

A

f

I /

h

1

in

0, H

0

NHAc H-C-CH,

co

3.

L-Ala

5-

c-c i

I

II

H O Muramic d-lactam

0

J. L-Ala

D-Glu J.V IIl-Dpill

3. D-Ala FIG. 3. Repeating units of peptidoglycan from spores of Bacillus subtilzs. The relative frequency of each unit is shown as a pri~cc~ntage. The sequence ofthe units is not random. Muramic lactam units tend to alternate with the other units. O n average 19% ol tliv tetrapeptides were linked through their D-alanine carboxyl group to the €-amino group of diaminopimelic acid ( D i m )ol'another-peptide side chain (from Warth and Strominger, 1972).

-I

I

m

20

A. D. WARTH

ture. The structure is related to type I peptidoglycan (Ghuysen, 1968) which is common in vegetative cell walls, but some modifications unique to spores are present. The most striking of these is that 45-60% of the muramic acid residues in the glycan chains lack both a peptide and an N-acetyl substituent and instead form an internal amide muramic lactam”. In a further 18% of the muramyl residues, the peptide side chain is curtailed to a single L-alanine residue. Compared with vegetative cell-wall peptidoglycans, the degree of cross-linking is very low. Only one peptide in five initiates a cross-link and only 35% of the muramyl residues bear a peptide, thus giving about one peptide crosslink to every sixteen residues in the glycan chain. End-group determinations indicate an average glycan chain length of 80 to 100 saccharides which is longer than is common in vegetative cells. Other polymers found in the cell wall, such as teichoic acid, have not been detected in spores (Chin et al., 1968; Warth and Strominger, 1972). Perhaps the most significant structural feature is the uniformity of structure of the spore peptidoglycan compared with the species variability of cell walls. Walls of B . subtilzs differed from the spore cortex in having amidated diaminopimelic acid residues and in lacking the carboxyl terminal D-alanine from the peptide (Warth and Strominger, 1971). In B . sphaericus, the change in structure is even more radical, with the diaminopimelic acid in the spores being replaced by lysine in the cell wall and a D-isoasparaginyl residue being incorporated in the peptide cross link (Hungerer and Tipper, 1969). I t is likely that other variations in peptidoglycan structure exist in the vegetative cell walls of spore-forming species, but the structure of the cortex does not appear to vary. This conservation and specificity of structure must imply an important function for the cortex, common perhaps to all species. Although the sequence along the glycan chain of the various muramic acid substituents is not known, the relative yields of the products of lysozyme digestion suggest that a degree of regularity exists, with lactams alternating with peptide or alanine substituents (Warth and Strominger, 1972).This conclusion has been confirmed by a study of the kinetics of formation of lysozyme digestion products and by a non-enzymic degradation procedure employing alkaline hydrolysis and nitrous acid, which specifically breaks the glycan chain at muramic lactam residues (A. D. Warth, unpublished results). Direct analyses of purified germ cell walls have not been reported. A number of observations suggest that it is a rudimentary form of cell (6

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

21

wall lacking some of the distinctive features of the vegetative cell wall, but it probably does not have the unique structural features of cortical peptidoglycan. Germ cell wall is formed earlier than the cortex and the morphology of their formation strongly suggests that the germ cell wall is synthesized by the forespore cytoplasm and membrane, whilst the cortex is synthesized by the mother-cell cytoplasm and outer forespore membrane. Bacillus sphaericus spores contain peptide side chains of both cell and spore types (Tipper and Gauthier, 1972).During spore formation, diaminopimelate ligase, a n enzyme specifically required for synthesis of spore-type peptides, was found only in the mother cell (Tipper and Linnett, 1976). I t was formed after most of the germ cell wall had been laid down (Holt et al., 1975), whereas L-lysine ligase which is required for cell wall type peptides was present at highest specific activity in the forespore. I n B . cereus, the germ cell wall shares the vegetative cell wall property of lysozyme resistance, which is probably occasioned by the absence of N-acetyl substituents from some glucosamine residues in both cells walls (Araki et al., 197 1 ) and spores (Warth, 1968). O n the other hand, amidated peptides characteristic of B. subtilis walls were not present in spore peptidoglycan (Warth and Strominger, 1972). Teichoic acid and other major cell-wall polymers have not been detected chemically in spores of B . subtilis (Chin et al., 1968; Warth and Strominger, 1972) o r other species (Warth, 1965, 1968). Electron micrographs show the germ cell wall as a simple layer adjoining the cortex and d o not suggest the presence of any of the more complex features of cell walls of some species, although cell-wall antigens were detected in the germ cell wall of B . cereus (Walker, 1970). A characteristic cell-wall layer of protein subunits in B . polymyxa was formed immediately after germination (Murray et al., 1970). Cross linking of peptides appears to be greater in the germ cell wall than in the cortex, as is suggested by its less expanded appearance. Peptides from cell walls of germinated B . megaterium were more cross-linked than total spore peptidoglycan (Cleveland and Gilvarg, 1975). The lysozyme-resistant fraction of B . cereus and B . subtilis spore peptidoglycan was mainly germ cell wall and had more peptides and peptide cross-links than cortical peptidoglycan (Warth, 1968, and unpublished results). Muramic lactam appears to be confined largely o r entirely to the cortex. It is formed late during spore formation at the same time as dipicolinic acid (Wickus et al., 1972; Imae and Strominger, 1976a, b). Peptidoglycans containing muramic lactam residues can be solubilized after mild alkaline hydrolysis by treatment with nitrous acid which

22

A. D. WARTH

breaks the glycan chains specifically at the muramic lactam residue (A. D. Warth, unpublished results). This method removed the cortex from B . cereus and B. subtilis spores leaving the germ cell wall apparently unaltered. C . LYTIC ENZYMES

In addition to peptidoglycan, the cortex and germ cell wall contain a number of degradative enzymes. On germination or even simply on disruption of the spore, the enzymes become active. The cortex structure is more or less completely solubilized whereas the germ cell wall is stretched by the swelling cell but persists and becomes the cell wall of the young cell. In disrupted spores both the cortex and the germ cell wall often autolyse but, in some species, germ cell wall may persist. The principal lytic activities present in B . subtilis, B . cereus and B . megaterium spores are endo-N-acetylglucosaminidase, which hydrolyses glycosidic links in the glycan chain of the peptidoglycan, and N-acetylmuramyl- L-alanine amidase which cleaves peptide side chains from the glycan chains (Warth, 1972; Hsieh and Vary, 1975). Each of these enzymes has only a limited action during germination or autolysis, cleaving only a few of the glycosidic and amide bonds present. The products are large peptidoglycan fragments, which in B . megaterium had a molecular weight of 15,300 and appeared relatively monodiserse (Record and Grinstead, 1954) and a few small peptides. Spores of B . cereus also contain D-alanine carboxypeptidase and N-acetylglucosamine deacetylase activities (Warth, 1972). These two enzymes are not themselves lytic but could modify the germ cell-wall structure in such a way that it approaches that of the vegetative cell wall and may modulate the action of the lytic enzymes. For example, removal of acetyl groups from N-acetylglucosamine residues makes the peptidoglycan resistant to lysozyme (Araki et al., 1971). In B. cereus, the lytic enzymes are readily extractable (Strange and Dark, 1957) but, in most other species, they are bound to spore structures. The spore lytic enzymes have not been separated and the particular properties and function of each enzyme determined. All studies using spore lytic enzyme have used this mixture of enzymes, and the relative participation of each will depend upon the conditions of pH value, cation concentration and substrate used. Great caution is therefore necessary in comparing work from different laboratories or involving quantitative measurements of activity.

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

23

Two major hnctions for the spore lytic enzymes can be envisaged, namely lysis of the cortex and decrease in the rigidity of the germ cell wall to facilitate swelling of the young cell. In addition, the normal complement of cell-wall enzymes necessary for growth may be present. N-Acetylglucosaminidase appears to be the main enzyme associated with cortex lysis. Its action would be essential to break the long glycan chains of the cortical peptidoglycan to the size of' the fragments produced. A spore enzyme preparation from B . cereus lysed isolated cortices with the formation of reducing groups but not of amino groups (Gould and King, 1969). The optimum pH value of 5 to 6 found for autolysis in B. subtilis and B. coagulans (Warth, 1965, 1972) is typical of bacterial cell-wall N-acetylglucosaminidases (Berkeley et al., 1973). Lytic activity in a B. cereus spore extract was stimulated by Co2+ at an optimal pH of 7.8 (Strange and Dark, 1957). This suggests the presence of the muramyl- L-alanine amidase with like properties which was purified from autolysing sporangia of B. thuringiensisby Kingan and Ensign ( 1968). A proper comparison of the spore enzymes with those from vegetative cell walls will require a more thorough characterization of the individual spore enzymes. Both amidase (Herbold and Glaser, 1975) and N-acetylglucosaminidase (Berkeley et al., 1973)have been purified from B. subtilis walls. A mutant lacking N-acetylglucosaminidase apparently was capable of both sporulation and germination (Ortiz, 1974) but a mutant with a temperature-sensitive amidase (Fan and Beckman, 1973) has not yet been tested for spore related functions. D. B I O S Y N T H E S I S

The existence of differences in structure between spore and vegetative peptidoglycans has made a study of peptidoglycan synthesis during spore formation an attractive system for gaining insight into control mechanisms operating during the cellular differentiation process. Interesting questions arise as to the respective roles of the forespore and forespore membrane on the one hand, and the mother cell and the outer forespore membrane on the other. Vinter (1963) showed that peptidoglycan synthesis as indicated by diaminopimelic acid incorporation was maximal at two periods during spore formation of B. cereus. The first period corresponded to the formation of the germ cell wall and the second was co-incident with cortex synthesis and the development of refractility. In B. megaterium,

24

A. D. WARTH

Pitel and Gilvarg (197 1 ) failed to find diaminopimelate incorporation until engulfment of the forespore was complete. Subsequent incorporation was not clearly separated into two maxima. Muramic lactam synthesis in B. cereus T and B. megaterium occurred only during the later phase (Wickus et al., 1972 ) . Penicillin-binding capacity, taken as a measure of transpeptidase activity and possibly other reactions involved in biosynthesis and crosslinking of peptidoglycan, also showed maxima during these two periods (Lawrence et al., 1971). Mutants blocked at different stages of'spore formation failed to express either the second or both maxima in binding (Rogolsky et al., 1973). Anwar et al. ( 1974) separated five binding components from B . subtilis cells, but found no new components or changes in the relative proportions during spore formation. Possible pathways for the formation of muramic lactam and the N acetylmuramyl- L-alanine units in the cortex have been discussed (Warth, 1968; Tipper and Gauthier, 1972). So far, neither muramic lactam synthesis nor any of the postulated enzymic activities have been reported in cell free preparations. Sporulating cells of B . sphaericus (Guinand et al., 1974) and of B . subtilis (Guinand et al., 1976) contained a particulate y-glutamyl diaminopimelate peptidase. Bacillus subtilis also had muramyl-L-alanine amidase activity. The relevance of these enzymes to cortex formation is not known. Other functions of these enzymes could be sporangial lysis or post-germinative modification of the cell wall. Tipper and his colleagues have studied the activity of the enzymes involved in synthesis of the UDP-N-acetyl muramyl peptide precursors of peptidoglycan in B . sphaericus. Most of the enzymes are common to both vegetative cell and spore peptidoglycan, and were synthesized during two periods preceding the two peaks of biosynthetic activity. Diaminopimelate-ligase is specifically required for addition of diaminopimelic acid to the precursor and its activity does not appear until just before cortex synthesis (Tipper and Pratt, 1970) and is confined to the mother cell (Tipper and Linnett, 1976) thus providing strong evidence for synthesis of the cortex by the outer forespore membrane under the control of the mother cell cytoplasm. This is to be expected since, at this stage, the forespore cytoplasm is condensed and is unlikely to be active metabolically and the outer forespore membrane is proximal to the cortex. The lysine-adding enzyme is present throughout all stages of spore formation. It is found also, along with the other enzymes necessary for formation of vegetative type pre-

MOLECULAR STRUCTURE

OF THE BACTERIAL SPORE

25

cursor, at a relatively high specific activity in the mature spore. It seems probable that these enzymes participate in synthesis of the germ cell wall and later, after germination, of new vegetative cell wall. VI. Core A.

MACROMOLECULAR COMPOSITION

In terms of its macromolecular constitutents, the core is a relatively normal cell. Many of the enzymic activities of vegetative cells are found in spore extracts. Most spore enzymes which have been studied had very similar properties to their vegetative counterparts, and it is probable that both are specified by common genes (Kornberg et a/., 1968; Sadoff, 1969).Adenylate kinase (Spudich and Kornberg, 1969)deoxyribonucleic acid polymerases (Falaschi and Kornberg, 1966; Terano et al., 19751, inorganic pyrophosphatase (Tono and Kornberg 1967a, b), lysyl- tRNA synthetase (Steinberg, 1974) and ribonucleic acid polymerase core enzyme (Ben-Ze’ev et al., 1975) from cells and spores have been studied in detail and no important differences in kinetic or molecular properties have been found. Other enzymes, such as aldolase (Sadoff, 1969), glucose 6-phosphate dehydrogenase (Ujita and Kimura, 1975)and purine nucleoside phosphorylase (Gilpin and Sadoff, 197 11, are similar but have significant differences in properties suggesting specific modification of the spore form. In some cases, spore enzymes have been modified by serine protease action either in vivo or during extraction; in others, the nature of the modifications are unknown. Ribosomes from B. subtilis (Bishop et al., 1969) and B . megaterium (Chambon et al., 1968) spores are similar to vegetative ribosomes in physical properties and protein-synthesizing activity but, in B. cereus spores, some ribosomal proteins were missing, causing defective subunits and poor synthetic activity (Kobayashi, 1973). Hybridization studies with B. subtilis DNA (Bishop and Doi, 1968; Edge11 et al., 1975) indicate that both spore and vegetative rRNAs are transcribed from the same genes. Messenger-RNA competitive with vegetative and sporulation messenger has also been found in spores (Jeng and Doi, 1974). The tRNA complement of spores is similar to that of vegetative cells. Several new types have been detected in sporulating cells and some of these were also found in spores (Lazzarini, 1966;Jeng and Doi, 1975). DNA from spores of B. megaterium (Chambon et al., 19681, B . subtilis (Sakakibara et al., 1969)and B . cereus (Tsuji et al., 1975)had properties

26

A. D. WARTH

not significantly different from vegetative DNA. Unlike spore protein and RNA, which are largely synthesized during spore formation, DNA is incorporated into the spore by partition of the parent cell DNA. Bacillus subtilis contained a single complete genome (Callister and Wake, 1976), but other species may have multiple copies (Fitz-James and Young, 1959).The properties of spore and cell nucleic acids have been reviewed by Doi (1969). The lipid composition of spores is very similar to that of vegetative cells. The major phospholipids of both cells and spores of B . polymyxa (Matches et al., 19641, B. megaterium (Bertsch et al., 1969)and B . cereus (Lang and Lundgren, 1970) were phosphatidylglycerol, diphosphatidylglycerol and phosphatidylethanolamine. Bacillus megaterium also has a glycosaminylphosphatidylglycerol. Disruption or hydrolysis of spores is necessary for complete extraction of lipid. The readily extractable fraction in B. cereus and B. megaterium had a high diphosphatidylglycerol content and probably came from the exosporium and other peripheral structures. Lipid from Bacillus species is characterized by a very high proportion of branched-chain fatty acids (Kaneda, 1967). In B. megaterium, CI5branched-chain isomers comprised 70%of the total fatty acids. Spores contained relatively more of the C,, branched-chain isomers than cells but the proportion depended upon the amino-acid composition of the sporulation medium. Phospholipids accounted for two-thirds of the fatty acids of cells but only one third of those of the spore. (Scandella and Kornberg, 1969). Bacillus thuringzensis spore lipid contained more is0 and less anteiso isomers than cell lipid, whereas straight-chain fatty acids remained constant at 10 to 11% of the total (Bulla et al., 1975). Bacillus spores have quite low lipid contents, but the data are not reliable (Murrell, 1969). Clostridial spores on the other hand contained 13 to 38% lipids. In two species of thermophilic clostridia, the lipid content was 13.5%and 16.3%and was nearly all firmly bound in the spore. Normal saturated and monounsaturated fatty acids from C,, to CLB, and a hydroxystearic acid were the main components (Pheil and Ordal, 1966). Setlow (1974, 1975a, b) has recently discovered a group ofbasic, low molecular-weight proteins present in the core of B. megaterium spores. These proteins may constitute as much as 30 to 50% of the protein in the core. Their main function is as a reserve material for germination as they are very sensitive to proteolytic enzymes and are rapidly degraded on germination. Amino-acid analysis show a very high proportion of polar amino acids, and cystine and tryptophan were absent.

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

27

Most of the glutamic and aspartic acid residues must be amidated as the isoelectric points were high (pH 9.8). N o enzymic function has been ascribed to these proteins but in vitro they complexed with DNA, raising its melting temperature. Other storage polymers have not been conclusively demonstrated in mature spores. B. megaterium accumulated /3-hydroxybutyrate (Slepecky and Law, 1961) at the end of exponential growth and C. butyricum formed an intracellular polysaccharide (Bergcre et al., 1975), but these reserves are utilized during spore formation. B.

LOW MOLECULAR WEIGHT C O M P O U N D S

In contrast to the unexceptional nature of the macromolecules of the spore core, the composition of small molecules in the spore differs dramatically fi-om that of vegetative cells. Localization of small molecules presents special problems in a structure with the morphological complexity but small size of a bacterial spore. The ionic composition of the spore is dominated by dipicolinate anions and calcium cations. After many years of uncertainty it now seems clear that the dipicolinate and much of the calcium are located in the core and not the cortex. Leanz and Gilvarg (1973) studied the attenuation of /3 particles emitted from tritium labelled dipicolinate and labelled components in the coats, cortex and core. A central location, in the core, was clearly indicated. Some excellent ultraviolet photomicrographs of sporulating B. subtilis (Wyckoff and Ter Louw, 193 1) also provide convincing evidence for a core or core plus cortex location. Arguments in support of a cortical location have been based on the similar time of syntheses of dipicolinate and the cortex, the large cortical space, and the dependance of dipicolinate retention on the cortex. A cortical location seems most unlikely, however, as the cortex shows no significant capacity to bind dipicolinic acid in vitro. In coat-deficient spores, the cortex is accessible to lysozyme but the spores retain dipicolinic acid. The coats may even be extracted (Aronson and Fitz-James, 1971) leaving no visible permeability barrier outside the cortex, and yet the spores still retain dipicolinic acid. Calcium, as the dominant cation, must be associated with at least some of the dipicolinate. In several species, the calcium and dipicolinic acid are present in nearly equivalent amounts (Murrell and Warth, 1965) but significant departures from 1 : 1 stoicheiometry implies that not all the dipicolinic acid or the calcium need be associated as the 1 : 1 chelate. Attempts to locate cal-

28

A. D. WARTH

cium by electron probe X-ray micro-analysis (Scherrer and Gerhardt, 1972) and by micro-incineration (Knaysi, 1965) showed calcium throughout the spore. Calcium was concentrated in the core, but significant amounts could well be present also in the cortex and coats. Indirect evidence for the location of calcium dipicolinate in the core comes from the peculiar property of spores, discovered by Robinow (1953), of exploding when treated with strong acid. Electron microscopy (Robinow, 1953; Fitz-James and Young, 1969) shows the core contents extruded through a break in the membrane, cortex and usually the coats. Only strong acids which have a soluble calcium salt will cause the reaction and only spores containing calcium dipicolinate will explode (A. D. Warth, unpublished results). I t would appear that the acid entering the core dissociates calcium dipicolinate forming a high concentration of calcium ions. If the membrane or cortex retains a low permeability to calcium ions, a transient osmotic force would be generated. If the calcium dipicolinate were located in the cortex the coats, but not the membrane, and cortex would be disrupted . The content of free amino acids was similar in spores and cells, but in spores consisted very largely of glutamic acid, arginine and lysine with very low levels of some of the other amino acids (Pfennig, 1957; Lee and Ordal, 1963; Nelson and Kornberg, 1970a). Spermidine was the predominant polyamine in B. megaterium (Setlow, 1974). Lesser amounts of putrescine and spermine were sometimes found. The total ribonucleotide pool in B. megaterium spores was somewhat less than in cells, and consisted mainly of the monophosphates and some diphosphates. Some high-energy compounds such as ribonucleotide triphosphates, reduced nicotinamide nucleotides and sugar phosphates were present at very low levels (Setlow and Kornberg, 1970; Setlow, 1973). On the other hand, a relatively large amount of 3-phosphoglyceric acid was present in B . megaterium and other Bacillus species. This was utilized during the first few minutes of germination to generate and maintain ATP levels (Nelson and Kornberg, 1970b; Setlow, 1975~).In addition to high levels of dipicolinic, glutamic and phosphoglyceric acids, B. subtilis but not B . cereus or B. megaterium spores contained a large amount of sulpholactic acid (Bonsen et al., 1969; Wood, 1971).Among the 14 strains of Bacillus listed inTable 6, sulpholactic acid was present in seven including B. subtilis, B. licheniformis and B . brewis.

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

29

The unique nature of the molecular environment of the vital macromolecules in the spore core is strikingly evident from the quantitative composition of the soluble fraction of spores (Table 5 ) . Protein and nucleic acids comprise only 50 to 60% of the dry weight, the remainder being principally dipicolinate, and other low molecular-weight anions and cations. The amount of enzymic protein is probably even less than that shown, at least in B . megaterium, as 30 to 50% of the protein is a basic low molecular-weight fraction that serves as a storage polymer and is probably of some importance itself in the physical structure of the core (Setlow, 1975b). Evidently the enzymes and other sensitive macromolecules are in a solvent containing a very high concentration of electrolytes. The properties of the solvent phase and its effect upon the activity and heat stability of the macromolecules depends very critically on the amount of water present. Evidence relating to the water content and physical state of the core is discussed later. VII. Ionic Composition of Spores

Divalent inorganic cations are of particular importance in the ionic composition of spores and have major effects on spore formation, heat resistance, and dormancy. Compared with vegetative bacteria, spores have very high calcium and manganese contents but often contain less TABLE 5. Composition of the Soluble Fraction of Spores Component

Bacillus megateriuma

Bacillus subtilis a

Bacillus subtilisb

% of spore dry weight

Dipicolinic acid Phosphoglyceric acid Sulpholactic acid Glutamic acid Arginine Ribonucleic acid Deoxyribonucleic acid Protein Inorganic cations

9.8 0.9 0 0.4 0.2 5.2 0.9 14.8 2.9

8.8 0.8

10 1

1.3 0.5 1.4

3-6

3.3 0.6 8.8 2.6

1

-

1

4 8 3.6

'Unpublished data of A. D. Warth Bacillus megaterium QM B 155 1 , Bacillus subtilis Porton strain. 'Data from Nelson et al.. (19691.'

A. D. WARTH

30

TABLE 6. Content of Ions in Spores

Bacillus cereus

Bacillus subtilis

Bacillus

SPP.

,u equiv./gm dry weight

Calcium Magnesium Manganese Potassium Sodium Polyamines Lysine Histidine Arginine Peptidoglycan" Total cations

1893 300 56 54 20 22 3 14 126 210 48 3043

1832 173 54 140 30 16 287 129 164 77 2902

780-1900 93-526 48-253 40-640 10-70 10-32 200-400 50-220 130-270 28-8 1 2050-3600

Dipicolinic acid Phosphate estersb Peptidoglycan" Sulpholactic acid Glutamic acid' Aspartic acid Amide

1890 3 16 195 < 10 590 500 -450

1513 293 317 <10 599 42 1 -503

940-1 890 201-815 115-350 0-160 460-1020 340-600 - 3 8 0 - 4 70

Total anions

304 1

2640

1990-3580

Bacillus cereus var. mycoides FRR B689, Bacillus subtilis strain FRR B692 and twelve other strains of seven Bacillus species were grown on 0.15% yeast extract, 0.15% beef extract, 0.5% peptone, 0.01 M K,HPO,, 1.5% agar, pH 7.2 plus spore salts (Warth, 1968). "Peptidoglycan cations = 0.8 x diaminopimelic acid, anions = 2.8 x diaminopimelic acid + 0.18 x muramic acid. Phosphate esters x total P. Phosphomonoesters will increase the anionic charge slightly. 'Excluding glutamic acid from peptidoglycan. (Unpublished data ofA. D. Warth).

potassium and magnesium (Curran et al., 1943; Slepecky and Foster, 1959; Crosby et al., 1971; Table 6). Calcium is accumulated preferentially to other divalent cations and is necessary in approximately stoicheiometric amounts to dipicolinic acid for dipicolinic acid synthesis (Black et al., 1960).The metal-ion content of spores depends to some extent upon the metal composition of the sporulation medium and on the spore-cleaning treatments. Precipitates of complex phosphates and other inorganic salts are a particular problem because of the common practice of adding supplements of metal salts to phos-

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

31

phate buffered media, and the typical rise in pH value during spore formation. Centrifugation is often ineffective in removing these precipitates and sonication followed by density-gradient centrifugation, two-phase treatment (Sacks and Alderton, 1961) or acid washing may be necessary. Each"of these treatments will also modify the ionic composition of the spore by cation exchange with sites on the spore surface. A variety of divalent metal cations including Mn2+, Ni2+, Zn2+ (Slepecky and Foster, 1959)Sr2+and Ba2+(Pelcher et al., 1963; Foerster and Foster, 1966)are accumulated during spore formation and, when added in unnaturally high concentrations to the medium, lower the calcium and dipicolinic acid content and the heat resistance of the spores. As expected Sr2+ replaced Ca2+with the least abnormality in spore properties. Specific transport systems in sporulating cells have been described for Ca2+(Eisenstadt and Silver, 1972), Mn2+, K+ and Mg2+(Scribner et al., 1975). As the location of soluble ions cannot easily be demonstrated, some perspective on the composition and relative importance of ions in the main spore compartments can be gained by consideration of the total ionic composition of spores (Table 6). Fixed ions are mainly aminoacid side chains of proteins, phosphate esters and peptidoglycan. Protein is present both in the core and coats and possibly also in the cortex. Analyses of coat proteins (Table 3) indicate that a major part of the protein ionic groups in Table 6 are from the spore coats. Phosphate esters in the core are present principally as RNA, DNA and phosphoglyceric acid. A variable amount of the total phosphorus, up to 75% depending on the species, is found in the coats. Peptidoglycan, which occurs exclusively in the cortex and germ cell wall, has total anionic groups of 115-350 pe/g which is only 20 to 30% of the fixed anions in the spore, or 5 to 1 1% of the total anions. Titration of spores to pH 4 and back titration with Ca2+has profound effects upon heat resistance and other spore properties (Alderton and Snell, 1963). In two strains of 121.perfringens, 220 and 480 pe of H'/g were found at p H 4 (Ando, 1975). Spores of B. megaterium pre-treated at pH 4 were able to bind 144 p e of Ca2'/g and isolated coats, possibly including some cortex, bound 220 pe/g (Rode and Foster, 1966). These values are consistent with a significant fraction of the potential ionogenic groups in the superficial layers of the spore being involved in this exchange reaction, but do not indicate to what relative extent the coats and cortex are involved.

32

A. D. WARTH

When the ionic complement known to be present in peripheral spore layers is subtracted from the total, the overwhelming dominance of dipicolinic acid and calcium in the core becomes more evident. N o matter how the minor cations are distributed, a major fraction of the dipicolinate must be associated with Ca2+. Furthermore, as mentioned by Nelson et al. ( 1969) the other anionic molecules glutamic acid, phosphoglyceric acid, and sulpholactic acid have a structural resemblance to dipicolinic acid. The commonly observed approximate equivalence of the calcium and dipicolinic acid contents can be seen to be a consequence of the cation specificity of the spore and the dominating level of dipicolinic acid present. I t does not imply that the Ca2+and dipicolinic acid occur entirely as the 1 : 1 chelate. More likely, a variety of ionic and chelating interactions occur in the core.

VIII. Water Content and Physical State of the Core

The physical state of enzymes and low molecular-weight solutes depends very critically on the amount of water present in the core. In common with most other dormant cell forms, spores have a lower water content than vegetative forms as evidenced by their high refractility and density. The refractive index of spores, as measured by interference microscopy, is close to 1.55 (Ross and Billing, 1957; Leman, 1973) comparable with that of dry protein. Although the refractive increment for spore constituents is not known with accuracy at high solids concentrations, the result indicates a very low water content of probably less than 20%. On the other hand, direct measurement of water content in B . cereus T spores in centrifuged pellets gave a value of 67% (Black and Gerhardt, 1962). Marshall and Murrell (1970) found values of 42 to 79% depending on the species and the assumed interstitial space. The disparity between the two techniques is much too large to be ignored. Much of the differences is doubtlessly due to the heterogeneity in water distribution within the spore. Interference microscopy averages the refractive index linearly through the spore and consequently -underweights the contribution of superficial layers to the spore volume. Furthermore, superficial layers which are extended and of high hydration, e.g. capsular material and some exosporia, may be overlooked by microscopy, whereas volume measurements are particularly sensitive to them. Spores have buoyant densi-

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

33

ties in Urografin solutions close to 1.34 g/ml (Dean and Douthit, 1974; Hsieh and Vary, 1975).This is similar to that of proteins and indicates that that part of the spore which is not permeated by the Urografin is very tightly packed and cannot contain substantial amounts of water. Of more critical concern than the total water content of the spore is its disposition among the various parts. The most refractile part of the spore appears to correspond to the core, suggesting that the core has a tighter packing with less free water and the superficial layers a more hydrated structure. Buoyant density measurements of B . cereus spores, which showed a small increase in density after extraction of the coats (Aronson and Fitz-James, 197 1) support this conclusion, as do lightscattering measurements of dry spores. These have been interpreted as showing a dense inner zone of refractive index (n = 1.54) and a less dense “coat” layer 25-85 nm thick (n = 1.46; Wyatt, 1975). Conclusions about the hydration of the cortex are very difficult to make with the present data. Electron microscopy shows a volume of 13 to 60% of the spore (Could and Dring, 1975a) yet peptidoglycan amounts to only 9 to 15%of the spore weight (Warth, 1968) suggesting a very hydrated structure. As the cortex can be sensitive to lysozyme in vzuo, it is probably permeable to smaller solutes such as Urografin and hence will not greatly affect the buoyant density. Any significant amount of water present in the cortex will strengthen the evidence supporting a tightly packed core. Spores differ from vegetative cells and disrupted spores in having a lower water content at high water activity (Neihof et al., 1967; Maeda et al., 1968; Marshall and Murrell, 1970).Similar or greater amounts of water are absorbed at intermediate water activity showing that hydrophilic sites are present. The lower uptake at high water activity implies that solvation and osmotic pressures are being constrained by mechanical forces either intramolecular in origin or caused by tension in spore integuments. Indirect evidence for a relatively dry and condensed core comes from several physical studies. Similar photoproducts, differing from the normal thymine dimers, were formed by ultraviolet irradiation of spores and of DNA dried in the presence of calcium dipicolinate or salts (Donnellan and Setlow, 1965). Dielectric studies by Carstensen and Marquis (1975) show very low conductances at high frequencies indicating that the high concentrations of electrolytes present in spores are not free to diffuse appreciably. Dielectric loss measurements at a higher frequency ( 10 GHz) suggest that any water in the spore is bound (Ballario etal., 1975; Maeda etal., 1968).

34

A. 0.WARTH

Several spectrographic techniques have been used to study the bonding of dipicolinic acid in the spore. Ultraviolet-absorption studies are consistent with the presence of calcium dipicolinate chelate but not of substantial amounts of acid or simple salts or esters (Bailey et al., 1965). Electron paramagnetic resonance shows that manganese is largely in a chelated form in spores in contrast to the ionic state found in cells (Windle and Sacks, 1963). Infrared spectra are not very informative but the Raman spectrum clearly shows that dipicolinic acid is not present as the tridentate calcium dipicolinate chelate nor as a simple salt or the free acid. Most of the dipicolinic acid (DPA) is bound in a form in which the carboxyls are polarized to an extent intermediate between DPA2- and H,DPA (Woodruff' et al., 1974). More work with model compounds involving calcium ions, dipicolinic acid, proteins and amino acids is urgently needed to enable the spectrographic evidence to be interpreted more fully. The available data points to a core structure in which the low molecular-weight substances are tightly packed in with the vital cell constituents, probably to the exclusion of a Auid aqueous phase. The Raman spectral data may indicate that a specific pattern of bonding exists between dipicolinic acid and other constituents, but X-ray scattering shows little sign of extensive ordering (Kadota and Iijima, 1965). More likely, components are concentrated such that a variety of polar hydrogen bonding and aromatic ring stacking interactions occur. As is evidenced by the rapid release of calcium ions and dipicolinic acid on germination or disruption, these bonds are stable only by virtue of the extreme concentration present and in effect replace a fluid solvent, water, with an amorphous immobile one largely consisting of calcium dipicolinate and structured water. At this time, there appears to be no need for special hypotheses about any interactions or structures unique to calcium dipicolinate or spores. Such a structure for the core may readily fulfil the requirements for heat resistance and metabolic inactivity. Proteins (Altman and Benson, 1960) can be extremely heat stable when heated at an appropriate low water content. The presence of an amorphous phase of suitable low molecular-weight polar compounds may make the water content much less critical by providing a space-filling medium with properties suitable for the maintenance of native enzyme structures, and may lessen unfavourable interactions between cell constituents. I t may also facilitate germination by allowing the core to hydrate by the replacement of the dipicolinic acid phase with water without the immediate necessity for swelling. Sur-

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

35

viva1 of bacterial cells during freeze drying and storage is greatly enhanced by such compounds (Orndorff-andMacKenzie, 1973). Glutamate is particularly effective and Salmonellae dried in glutamate can survive many hours at 100°C (Annear, 1964). Protective compounds require at least three polar groups and high solubility. High solubility would not be an advantage in vivo as the small size of the cell and the presence of proteins would inhibit crystallization, and a high osmotic activity would make concentration of the core contents more difficult. All the major spore electrolytes, dipicolinic acid, glutamic acid, phosphoglyceric acid, sulpholactic acid and arginine appear suitable for this function and the divalent cations have clear advantages in the formation o f a matrix of low osmotic activity. In keeping with this role of making the resistant state less critical is the recent isolation by Hanson et al. (1972) of mutants of B . cereus T whose spores lack dipicolinic acid but are fully heat resistant. However, on storage in water, heat resistance was soon lost. Within the normal range, heat resistance is not correlated with DPA content (Murrell and Warth, 1965) but in all other circumstances heat resistance requires the presence of dipicolinic acid. The ubiquitous distribution of calcium ions and dipicolinic acid in all endospores investigated from several genera implies that they have uniquely suitable properties. IX. Mechanisms for the Dehydration of the Core

A relatively dehydrated core containing high concentrations of hydrophilic solutes requires a mechanical constraint to the osmotic and swelling pressure. To some extent, this will be provided by the various intramolecular bonds between dipicolinic acid, calcium ions and other constituents. The remainder appears as the pressure exerted by the integuments to oppose solvent uptake. Pressure could be applied to the core in several ways. The simplest, first postulated by Lewis et al. ( 19601, is a contraction of the cortex. Conversely, the cortex might be expanded against an inextensible coat (Alderton and Snell, 1964; Gould and Dring, 1974, 1975a, b). A third and plausible model that has received little discussion since it was first suggested by Alderton and Snell (1963) is for anisotropic swelling of the cortex in the radial direction. In this case, the outer layers of the cortex would be under tension. The germ cell wall and spore cytoplasmic membranes may also make limited contributions, especially during the latter part of stage IV of spore formation where the core is partly condensed and

36

A. D. WARTH

phase white but before full refractility and completion of cortex and coat formation. It is also necessary to consider the chemical mechanisms by which these forces might be generated during spore formation. The cortex is a loosely crosslinked polyanionic structural polymer; as such, its state of swelling or contraction depends upon the type and concentration of cations present. Dehydration of the core could be controlled during spore formation by changes in the cation content of the cortex (Lewis et al., 1960). Alternatively, the swelling properties of the cortex could be modified during spore formation by hydrolysis of peptide side chains (Could and Dring, 1975b). What is the experimental support for these models? Firstly, it seems clear that the cortex is indispensible for heat resistance, is involved in the final stages of dehydration of the core, and is necessary for maintenance of the dehydrated heat-resistant state. Inhibition of cortex synthesis by antibiotics Winter, 1959; Fitz-James and Young, 1969) or by limitation of precursors (Freese et al., 1970; Imae and Strominger, 1976a, b) produces spores that are either unstable and lyse, or that lack heat resistance and full refractility. Hydrolysis of the cortex with lysozyme or other lytic agents causes loss of refractility and heat resistance. During spore formation, the transition from phase whiteness to full refractility and the acquisition of heat resistance occurs at about the same time as completion of cortex synthesis Winter, 1959; Wickus et al., 1972; Imae and Strominger, 1976~). In diaminopimelate auxotrophs of B. sphaericus, cortex synthesis is dependent on the supply of diaminopimelate. Spores containing approximately 20% of the maximum amount of cortex are refractile and accumulate DPA, but more than 90% of the cortex is necessary for heat resistance (Imae and Strominger, 1976b). Likewise, in a mutant of B . cereus, small amounts of cortex are sufficient for calcium dipicolinate accumulation and refractility (Pearce and Fitz-James, 197 1). Among spores of several species, the degree of heat resistance was correlated with the cortex content (Murrell and Warth, 1965). The cortex is associated mainly with the final stage of dehydration of the core and the accumulation of calcium dipicolinate, and does not decrease the volume of the core by more than a few per cent. Before the cortex is formed, the contents of the core are already condensed and the volume of the core has been reduced by 50% from its maximum at Stage IV (Fig. 4, D. Ohye, unpublished results). Large spores sometimes appear phase white at this stage. This preliminary concentration may occur through quite different mechanisms from those operating later.

MOLECULAR STRUCTURE

OF THE BACTERJAL SPORE

37

FIG. 4. Electron micrographs of sporulating cells of Bacillus cereus T. (a) After 6 h of spore formation showing the forespore protoplast at its maximum volume. ( b ) 1 h later, the volume is reduced 5 I%, the cytoplasm is more tightly packed and the DNA has a fibrous structure and peripheral location. The cortex is .just beginning to form (results of D. F. Ohye.).

Spore coats do not seem to be required for refractility o r heat resistance (see Section IV-D). Electron micrographs of coatless spores (FitzJames and Young, 1969; Aronson and Fitz-James, 197 1 ; Cassier and Ryter, 197 1) show no tendency for the cortex to swell outwards. The core retains its condensed appearance, the spores retain full refractility and the buoyant density is slightly greater. Similar results have been obtained in this laboratory. These properties are quite inconsistent with a simple expanded cortex model (Gould and Dring, 1975b) which requires an intact and strong coat to contain the swelling pressure. A contracted cortex also seems inconsistent with electron micrographic evidence. The large volume of the cortex, compared with the peptidoglycan content, suggests an expanded rather than a contracted

38

A. D. WARTH

structure (Gould and Dring, 1974, 1975a). In mature spores, no structure can be seen in the cortical region unless thin sections are stained. Then the structure appears expanded, and there is usually no suggestion of a tightly contracted cortex or a void between it and the coats. In disrupted or germinating spores and in spore integument preparations, the cortex stains more readily and always appears very expanded. With disrupted spores and fragments of integuments, the expansion is always inwards, irrespective of the presence of attached coats, and causes folding of the inner cortical layers and the germ cell wall (see figures in Warth et al., 1963; Fitz-James and Young, 1969). This picture is indicative of anisotropic expansion of the cortex with maximum expansion along the radial axis. Electron microscopy could be misleading, but dehydration and treatment of the cortex with heavy-metal cations would be likely to give artefactual shrinkage but not swelling. Overall, the electron microscopy data appear inconsistent with a cortex that is simply contracted around the core or expanded against the coats. They do, however, support the anistropic cortex model in which pressure is applied to the core by swelling along the radial axis. Of the two mechanisms which suggest themselves for the origin of the swelling pressure in the cortex, the ion-exchange model for several reasons seems the less important. In coat-defective spores, and probably in normal spores also, the cortex can almost certainly equilibrate with external electrolytes. Such spores do not lose refractility over a much wider range of ionic strengths than would fully expand or contract common ion exchange polymers including cell walls (Ou and Marquis, 1970) and lysozyme-digested spore peptidoglycan (Baillie and Murrell, 1974). Heat resistance of spores can be affected by manipulation of the ionic content of the spore (Alderton and Snell, 1963, 1964) and spores with weakened coats are sensitive to heating in 2MCaC1, (Gould and Dring, 197513).I t is not certain that these treatments had affected the cortex and not damaged some other essential site such as the cell membrane. Nevertheless, heat resistance is no doubt a very sensitive indicator of the hydration of the core and interactions between cations and the fixed anions of the cortex must affect to a limited extent the pressure applied to the core. Swelling cannot depend only upon a low ionic-strength and higher ionic strengths d o not substantially reverse it. Electron microscope studies on the cortex of acid-popped spores of B . stearothermophilus (D. F. Ohye, unpublished results) support this conclusion. The cortex retains

MOLECULAR STRUCTURE OF THE BACTERIAL SPORE

39

its characteristic inward swelling (Fig. 1 in Murrell and Warth, 1965) after treatment at several pH values and at electrolyte concentrations up to saturated calcium chloride solution. The chemical structure of the cortex and the morphology of its formation suggest the following model. Cortex formation begins with synthesis by the outer forespore membrane of a layer of peptidoglycan. At this stage, the core cytoplasm is already concentrated (Fig. 4) but the forespore is not refractile and lacks dipicolinic acid. It is likely that the cortex peptidoglycan is synthesized initially, in the same way as cell walls, with a full complement of peptide side chains and a moderate degree of peptide crosslinking. Removal of peptide crosslinks by hydrolysis and transamidation to form muramic lactam, Lalanine and tetrapeptide subunits will allow the structure to swell. If the mechanical constraint is less along the radial axis, as, for example, if the long glycan chains are preferentially oriented parallel to the surface, the swelling pressure is directed inwards. Further layers are built up at the outer surface until the cortex meets the coats which are forming in the sporangial cytoplasm. Control of the relative rates of synthesis and hydrolysis would enable a structure to be built with uniform stress in each layer. Only a small decrease in the volume of the core is seen during cortex formation (D. F. Ohye, unpublished results). At the same time as cortex is formed, calcium dipicolinate is synthesized and taken into the core, where it will displace a significant volume of water. In normal spores, this may make a significant contribution to achieving the degree of dehydration necessary for full heat resistance. Mutant spores unable to make dipicolinic acid are usually not fully heat resistant (Zytkovicz and Halvorson, 1972; Imae and S trominger , 197 6a). Anisotropic behaviour is common to all structures which have oriented molecular arrangements. Recognition of this property in the spore cortex has led to the resolution of some apparent contradictions. I t is now obvious that swelling and contraction cannot be used without specifying the direction. Radial expansion alone in a spherical shell will exert pressure inwards but expansion parallel with the surface has the opposite effect. Effects such as the re-establishment of refractility in freshly germinated spores by treatment with polycations (Cislavska et al., 1970)can be interpreted as an electrostatic interaction bridging gaps in the outer layers of the cortex and causing contraction of the surface. Other data, interpreted as favouring an expanded cortex (Gould and Dring, 197513)are likewise consistent with this model.

40

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X. Acknowledgements I wish to thank Mr. D. F. Ohye for the electron micrographs and for many helpful discussions on this topic.

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Physiology of Fungal Lipids: Selected Topics PATRICK J . BRENNAN" Department of Biochemistry. University College. Dublin. Dublin 4. Ireland and

DOROTHY M . LOSEL Department of Botany. University of Sheffield. Sheffield. SlO 2TN. England I . Introduction . . . . . . . I1 . Location of Lipid in Fungal Cells . . . 111. Lipids and Fungal Membranes . . . A . Membranes-General Considerations . B . Phospholipids-Types and Distribution . C. Phosphoglycerides-Physiological Aspects D . Glycolipids . . . . . . E . Sphingolipids . . . . . . . . . IV Biosynthesis of Fungal Lipids . A . Phospholipids . . . . . . B. Glycolipids . . . . . . . . . . . . C. Sphingolipids . V. Role of Lipids in Fungal Morphogenesis . A . HormonalandGrowth-RegulatingFactors B . Lipid Reserves in Morphogenesis . . C . LipidReservesandSecondary Metabolites D . Lipid Metabolism in Morphogenesis . VI . RoleofLipid inFungus-Host Relationships . A . Fungal Associationswith PlantTissues .

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B. Fungi Associated with Insects . . C. Fungi Pathogenic to Man and Aninials D . Discussion . . . . . . . . . . . References .

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155 159 163 164

I. Introduction

The physiology of fungal lipids can be defined as the relation of lipids to the process of life in fungi. In necessarily restricting our coverage of this vast topic, we have been governed by recent developments in a narrower functional context. In particular, we have emphasized areas which have not been reviewed in the several recent major reviews o n fungal lipids (Weete, 1974; Erwin, 1973; Brennan et al., 1975). For instance, the involvement of lipids in host-fungal parasite relationships has not previously been treated in the secondary literature. A review on fungal glycolipids is also timely since it now appears that glycolipids of higher plant and fungal membranes assume the physical role of the glycoproteins in mammalian cell membranes, and there has been much recent work on the chemistry of fungal glycolipids. Other areas which have seen substantial recent progress and which are reviewed are: the involvement of lipids in morphogenesis, a field in which correlation of biochemical data and ultrastructural studies has been valuable; the role of polyprenols in mannan synthesis; the metabolism of phosphoglycerides and their interrelationship in fungal membranes; and the physiology and spherosomes and lipid inclusion bodies. A discussion of the structure, distribution and biosynthesis of simple fungal lipids, such as fatty acids, sterols and hydrocarbons, has been deliberately avoided since these areas have been thoroughly explored in other reviews. 11. Location of Lipid in Fungal Cells

Besides the essential presence of lipids as membrane components of the endoplasmic reticulum, plasmalemma and organelles, a variety of lipid-containing inclusions occur in plant cells, usually becoming most prominent in reproductive organs and spores. Lipids are also found as cell-wall components and o n the surfaces of spores (Table 7 , p. 122) and various aerial structures or as extracellular lipid clinging to mycelium or yeast cells in certain conditions. Some of the information recorded on the location of fungal lipid is summarized in Table 1. The usually low proportion of lipid in walls may be greatly exceeded in

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

49

certain specialized structures. An outstanding example is the sporangiophore of Phycomyces sp., a single cell which can grow to a height of over 10 cm, with a wall lipid content of 25% (Kreger, 1954) and an electron-dense surface cuticle (Peat and Banbury, 1967). Differences in lipid composition between walls of yeast cells, which grow by budding, and those of filamentous hyphae are of interest in relation to the dimorphism which is characteristic of some of the fungi pathogenic to man and animals. In the well known dimorphic growth of Mucor rowcii, 2% lipid is found in the hyphal-phase walls and 0.8% in walls of the yeast form (Bartnicki-Garcia and Nickerson, 1962). Lipid inclusions are often abundant within vegetative mycelium but, although these have been the subject of much biochemical investigation, their ultrastructural distribution within vegetative hyphae has received less attention than that in reproductive organs and has not been related to other morphological and taxonomic features. Reproductive structures show some characteristic patterns of lipid distribution in different taxonomic groups of fungi. This aspect will be discussed more fully in Section V (p. 113). Although aspects of morphological organization and life cycle used in fungal taxonomy often correspond very satisfactorily with wall chemistry (Bartnicki-Garcia, 1968, 1970) and other chemotaxonomic characters such as fatty-acyl composition (Shaw, 1966; Erwin, 1973) or carotenoid content (Valadon, 19761, little use has been made of chemotaxonomy in identification of fungi. A computer programme for cluster analysis has recently been applied to data on fatty-acyl content in taxonomic studies by Dart et d.(1976a, b). The various detailed investigations which have been made of the fine structure of vegetative hyphae have been mainly concerned with cellwall formation and distribution of mitochondria, ribosomes, dictyosomes and vacuoles in relation to regions of growth. Relatively few have shown a particular interest in lipid physiology, but the published electron micrographs can often provide much information on the involvement of lipids in vegetative growth. The clear pictures of Grove et al. (1970)of the hyphal tip region of Pythium sp. (Fig. 1) show an apical zone with many vesicles, extending back three to five microns from the hyphal tip. Behind this a subapical, non-vacuolated zone, richly supplied with mitochondria and ribosomes, showed occasional lipid droplets. Grove et al. (1970) mention the absence of lipid for several hundred micrometres behind this, i.e. well into the zone of vacuolation, which began 50 to 100 pm behind the apex. In the more

TABLE 1 . Distribution of lipid in fungal structures Vegetative cell wall Saprolegniaferax Phytophthora cinnamomi Mucor ruuxii (hyphae) Mucor rouxii (yeast) Candida albicans (hyphae) Candida albicans (yeast) Aspergillus nidulans Aspergrllus nidulam Geotnchum candidum Agaricus bisporus Cell membranes Saccharomyces cereuisiat, Candida utilzs Cytoplasm of vegetative cells Pythium ultimum

Aphanomyces eutiches Antheridial hyphae Saprolegniaferax Oogonium Bremia lactucae

Phytophthora capsin Gametangia Allomyces arbuscula Zoosporangia Allomyces spp.

Sajnolegniaf e r m Phytophthwa infestam

Ln

0

5%of dry weight 2.4%of dry weight 2% of dry weight 0.8%of dry weight 4.5%ofdryweight 1.8% of dry weight 10.5%of dry weight 4.6%of dry weight 8%of dry weight 1.5%of dry weight

Seitsma el al. (1969) Bartnicki-Garcia (1969) Bartnicki-Garcia and Nickerson (1962) Bartnicki-Garcia and Nickerson (1962) Bianchi (1967) Bianchi (1967) Bull (1970) Zonnenveld ( 197 1) Seitsma and Woutern (1971) Michalenko et al. (1976)

40%of dry weight ofprotoplast membranes

Hunter and Rose ( 197 1)

Some lipid bodies in subapical zone, abundant in mature regions Lipid bodies in mature hyphae. Lipid in “striated inclusions” in vacuoles at sporulation

Grove et al. (1970)

Large amounts of lipid bodies

Gay et al. ( 197 1 )

Oosphere with large lipid drops; lipid-rich periplasm. Large lipid drop in oospore Large lipid bodies during oospore delimitation

Tommerup ct al. (3974)

5 5:

Bartnicki-Garcia and Hemmes ( 1976)

rn r

Hoch and Mitchell (1972)

m

a rn z z D z D z

0

P

Crowns of lipid droplets around all nuclei Carotenoid in male gametangium Lipopolysaccharide and lipid-containingfibrous material in plugs of zoosporangial discharge papillae (30%of dry weight) Dense bodies with banded appearance Myelin-like lipoidal inclusion in vacuoles

P

c

Youatt (1976) Gay and Greenwood (1966) Elsner et al. (1970)

u)

LUUIlJULCJ

Bhtocladiella emersonii

Lipid sac attached to large mitochondrim

Blasfocladiella emersonii Blastocladiella emersonii Allomyces sp. Phytophthora parasitica

“Side body”4ipid sac complex 4.8 pg per spore Lipid bodies in periphery of nuclear cap Lipid droplets in cytoplasm; “liposomes” (lamellar inclusions in vacuoles) Lipid bodies in cytoplasm

Bimpongand Hickman (1975)

Relatively low lipid content Numerous lipid bodies coated with ferritin

Gunasekaran et al. (1972) Grove ( 1976)

Lipid droplets in mature zygospores

Hawker and Beckett (197 1)

Lipid in large inclusion bodies Abundant lipid droplets in rnacroconidia

Campbell (1971) Marchant (1966)

Large lipid inclusions

Stevenson and Becker (1972)

n C

Hyphae with few vacuoles and high lipid content

Mercer et al. (1975)

t)

Lipid-rich cells of inner wall layers

Jackson and Wheeler (19741

Lipid granules and microbodies in outer layers of perithecium wall

Hohl and Streit (1975)

Large lipid drops

Lowry and Sussman (1968) Illingworth et al. (1973)

Oval lipid bodies, 0.8 prn in diameter

Williams and Ledingham (1964)

Lipid drops in spores and aecial peridium

Lose1 and Lewis (1974)

Lipid droplets in developing basidiospores Lipid bodies and microbodies in mature basidiospores Lipid throughout spore-wall layers Several lipid drops

Vogel and Weaver (1972) Greuter and Rast (1975) Rast and Hollenstein (1977) Heintz and Niederpreum (1970)

Large lipid bodies, closely associated with “unidentified organelles” possibly glyoxysomes

Allen et al. (1971)

Phytophthora palmivora Sporangiospores Rhizopus arrhizus Phycomycs blakesleeanus Zy gospores Rhizopus sexualis Conidia Aspergillusfumigatus Fusarium culmorum Chlamydospores Fusarium oxysporum Sclerotia Colletotrichum lindemuthianurn Cleistothecia Sphaerotheca mors-uvae Perithecia Neurospora lanceolata Ascospores Neurospora tetrasperma Saccharomyces cerevisim Urediospores Puccinia graminis Aeciospores Pwcinia poarum Basidiospores Agaricus bisporus Agaricus bisporus Agaricus bisporus Coprinus lagopus Teliospores Tilletia caries

Lessie and Lovett (1972) Cantino and Truesdell (1970) Subercropp and Cantino (1973) Hill (1969) Hemmes and Hohl(197 1)

W

5

z

P

vl

P

-I

: 0 vl

52 P. J. BRENNAN AND D. M. LOSEL

FIG. l(b). Subapical non-vacuolated zone, densely supplied with ribosomes. Some densely staining lipid bodies are also present. M mitochondria, ER endoplasmic reticulum, N indicates nucleus, L lipid, R ribosomes, D dictyosomes, V vesicles. cn w

54

P. J. BRENNAN AND D. M. LOSEL

FIG. I(c). Zone of' vacuolation about 100 pm behind the apex. Lipid bodies are absent. V vesicles, VA vacuoles, D dictyosomes, N nucleus, M mitochondria.

mature regions, the abundant vacuoles and lipid bodies occupy substantial and almost equal volumes of the hypha, leaving relatively little cell volume occupied by organelles and ground cytoplasm. The electron-microscope pictures correspond well to the observations of the same workers on living hyphal tips by interference microscopy. In hyphae with a growth rate of 1 mm per hour, mitochondria were mainly lacking from the apex, but were seen moving in and out of the apical zone. More than a hundred microns behind this, the region of vacuolation could be clearly identified. Considerable confusion of terminology attends the description of lipid-containing organelles of fungi. Obvious storage deposits of lipid tend to be referred to as lipid bodies, fat bodies, osmiophilic globules and granules. These vary greatly in size with age of the cell, condition, species and nutritional balance of the medium. Bracker (19671, in his review of ultrastructure in fungi, described lipid droplets as occurring in mature or aged cells. He noted the frequent appearance of an unstained clear centre, surrounded by a dark cortical zone, which reacted

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

55

FIG. l(d). Mature hypha 4-5 mm behind the tip, with large, irregularly shaped lipid bodies in close contact with the mitochondria and vesicles. W wall, L lipid, VA vacuoles, M mitochondria, V vesicles, N nucleus.

with the fixative, and a bounding membrane-like structure lacking the usual tripartite organization of intracellular membranes. The name “spherosome” is well established for lipid bodies of seeds and various plant tissues, described by Matile (19751, in his monograph on the lytic compartment of plant cells, as osmotically inactive carbon reserves, which are transformed on germination into sugars.

56

P. J. BRENNAN AND D. M. LOSEL

More than one type or organelle seems to fit the description ofspherosome. Matile (1975) quotes the observations of Schaffner (1974) that Saccharomyces cerevisiae spherosomes differ from those of higher plants in containing phospholipids, sterols and _sterol esters and a smaller proportion of triglycerides, suggesting that they may function as storage sites for membrane lipids. Matile and his coworkers appear, on the whole, to equate “spherosome” and “lipid body”. The observations of Schwarzenbach (1971a, b) on the development of spherosomes from endoplasmic reticulum in Ricinus sp. explain the characteristic “half-unit” membrane. Schwarzenbach described deposition of triglyceride in the lipid middle layer of the double membrane of an endoplasmic reticulum cisterna, causing the half-membrane layer adjacent to the cytoplasm to swell out, bounding the spherical body of lipid, which continued to accumulate, even after the young spherosome had budded off from the rest of the endoplasmic reticulum. The inner membrane layer remained as a core still visible in the developing spherosomes. Wilson et al. (1970) and Armentrout et al. (1976) regard spherosomes as lysosomes, as defined by Gahan (19671, i.e. single-membranebounded organelles containing more than one hydrolytic enzyme. By cytochemical reactions, Wilson et al. ( 1970) demonstrated activity of acid phosphatase, aryl sulphatase (the natural substrate of which is still unknown) and deoxyribonuclease in spherosomes of several species of fungi. Such spherosomes are most abundant in the hyphal tip cells and aggregate also in dead and dying cells. The observations of movement of spherosomes in parasitic hyphae of the mycotrophic zygomycete, Piptocephalis virginiana, into the haustorium formed in the hypha of the host fungus, Monotypha microspora, (Armentrout and Wilson, 1969) indicated the involvement of spherosomes in hostparasite interactions. Wilson et al. ( 1970) suggested that enzymes from the parasite’s spherosomes were transferred across the haustorium membrane, through the sheath matrix surrounding the haustorium, into the host cytoplasm. Acid phosphatase was demonstrated on the surface of the haustorium. Host spherosomes seemed to break down on penetration of the host cell by the haustorium and could be involved in autolysis of the host cell. Wilson et al. (1970) also found lysosomes implicated in deliquescence of Ceratocystis sp. and breakdown of extraspore contents of the ascus and the ascus membrane, before discharge of the ascospores from the perithecium. Fusion of lysosomes with vacuoles was observed, apparently forming auto-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

57

phagic vacuoles. The same authors suggested that lysosomes may be involved in release of sexual and asexual spores, anastomoses, rapid +is of some basidiomycete fructifications and release of certain pathogens, e.g. Plasmodiophora sp. from host tissue. Sorokin ( 1967 ) similarly equated spherosomes and lysosomes and distinguished them from oil droplets which contain mainly neutral lipid. Spherosomes had mainly phospholipid and very little neutral lipid, thus corresponding to the Sacch. cereuisiae spherosomes described by Schaffner (1974). Sorokin (1967) pointed to the need for further study of the role of lysosomes or spherosomes in lipid synthesis. The developmental sequence in spherosomes observed by Schwarzenbach is consistent with the opinion of Frey-Wyssling et al. (1963) that spherosomes may sometimes be precursors of oil droplets. Thus, differences in relative proportions of membrane lipids and neutral lipids, on which Sorokin ( 1967) distinguished spherosomes from oil droplets, may simply represent stages of development and increasing accumulation of neutral lipid within the original spherosome. That lipid may be transferred to vacuoles for storage, or prior to its mobilization, is indicated by plates published in Matile’s (1975)monograph, which show, for example, a large lipid droplet being engulfed by a vacuole in Sacch. cereuisiae and some comparable pictures form lipid-storing plant tissue. Lipases for mobilization of reserve lipid are associated with spherosomes of various plant tissues. As Matile ( 1975) points out, by lipase action lipophilic neutral fats are hydrolysed to fatty acids, which can be released from the oil droplets. Enzymes for /?-oxidation of fatty acids and glyoxylate-cycle enzymes, to metabolize the resulting acetate units, are contained in organelles variously termed microbodies or glyoxysomes. Beevers ( 1969) described glyoxysomes as membrane-bound organelles where enzymes of /?-oxidation and the glyoxylate cycle are localized. Matile ( 1975) includes electron micrographs showing a glyoxysome surrounded by oil-droplets in seeds, and suggests that lipase determines the flow rates of fatty acids from spherosomes to glyoxysomes. An equally striking configuration appears in electron micrographs of mushroom spores (Greuter and Rast, 1975), in which lipid bodies are in intimate contact with mitochondria and microbodies. In the electron micrographs of Grove et al. (1970)of the hyphal tip of Pythium (Fig. la, p. 5 5 ) , mitochondria appear in close contact with lipid bodies, although this is not commented on and microbodies are not labelled.

58

P. J. BRENNAN AND

D. M. LOSEL

This recalls also the close proximity of lipid bodies, mitochondria and small vacuoles in the sporangiophore of Phycomyces species (Peat and Banbury, 1967)and the intimate association of lipid sac and giant mitochondrion in zoospores of Blastocladiella emersonii (Cantino and Truesdell, 197 0). In asexual spores of Phycomyces blakesleeanus and certain other zygomycetes, lipid bodies, which disappear in the first stage of germination, where shown by Grove (1976) to be surrounded by ferritin, apparently forming a paracrystalline monolayer rigid enough to give the drops an angular profile. Grove (1976)reports the suggestion of David and Esterbrook ( 197 1) that the ferritin functions as a storage form of iron required early in germination. Membrane systems of fungal cells contain 30 to 50% lipid (Weete, 19741, the major component of which is phospholipid, accompanied by sterols, sterol esters and some neutral lipids. The phospholipids of fungal membranes will be discussed in Section I11 (p. 65). Various osmiophilic inclusions recorded in electron micrographs of fungal structures show a spiral or whorled arrangement, and have been referred to as myelin figures or dismissed as artefacts. There is evidence that these may sometimes represent membrane material, either in the course of synthesis or breakdown. More than one such system was noted by Beck and Greenawalt (1976) in Neurospora sp. Whorled structures, which occurred in the cytoplasm during germination and at other times, appeared to be derived from membranes. Different whorled structures, appearing during germination, corresponded in number to the level of carbohydrates available, suggesting membrane synthesis. The lipid nature of some osmiophilic inclusions found in mitochondria was confirmed by their removal by 90% acetone. That they were present in living cells and not merely artefacts of fixation was demonstrated by their occurrence in freeze-fractured preparations. Beck and Greenawalt (1976) suggested that these inclusions were sites of membrane synthesis. Similarly, other membrane complexes, such as those seen within the basal region of haustoria of Uromyces appendiculatus by Hardwick et al. (19711, may represent stages of membrane synthesis. The relative ease o f identification of lipid structures and the convenient manipulation of physical and nutrient growth conditions permitted by their hyphal organization, make fungi an ideal system for the productive correlation o f structural and biochemical aspects of lipid physiology, in growth and differentiation.

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

111.

A.

59

Lipids and Fungal Membranes

M EM B R A N ES-GENERAL

CONS ID ERATI ONS

The primary functions of membranes are to provide a permeability barrier and sites for active transport of ions and facilitated diffusion of hydrophilic molecules. In animal cells, the plasma membrane also provides antigenic sites, attachment and recognition factors for other cells and for foreign substances. In many eukaryotes, the plasma membrane mediates the triggering of cellular responses to external influences, such as hormones and antigens. In fungi and higher plants, it is probable that the plasma membrane provides attachment sites for cell-wall components. In eukaryotic cells, such as fungi, there are usually a variety of cytoplasmic organelles with membranes, and the individual metabolic activities of many of these organelles are due to membrane-associated proteins. For instance, enzymes of the tricarboxylic acid cyclE are partially located in the mitochondrial inner membrane and those of the respiratory chain and oxidative phosphorylation system form membrane-bound multi-enzyme complexes, located in the mitochondrial inner membrane. Other relevant activities are those in the plasma membrane which are responsible for the synthesis and secretion of cell-wall precursors. Membranes, whatever their source, consist of mostly lipid, protein and some water. The lipid is a mixture in which phosphoglycerides usually predominate. However, in some membranes, particularly those of mycoplasmas, chloroplasts and some Gram-positive bacteria, glycolipids may predominate. Both membranous phospholipids and glycolipids are amphipathic substances, having polar head groups and long hydrophobic tails, and are suited for formation of a lipid bilayer. The early classical model of cell membranes (Danielli and Davson, 1934) represented the bilayer as a symmetrical structure. Recent evidence suggests otherwise. In the red-blood cell, the membrane carbohydrates, proteins and lipids are all unequally distributed between the two sides of the plasma membrane (Bretscher, 1974). Similarly, the membrane proteins, phospholipids and carbohydrates of influenza virions are also asymmetrically distributed (Tsai and Lenard, 1975). I t appears that the plasma membranes of milk-fat globules, erythrocytes and influenza virions are similar in that the

60

P. J. BRENNAN AND D.

M. L ~ S E L

choline-containing phospholipids, phosphatidylcholine and sphingomyelin, are concentrated in the outer half of the lipid bilayer and ethanolamine- and serine-containing phospholipids are mostly contained in the inner half (Patton and Keenan, 1975; Tsai and Lenard, 1975). Since influenza virions acquire their lipid coat by budding from the plasma membrane of a variety of hosts, it is reasonable to suggest that asymmetric phospholipid distribution may be a general feature of the structure of animal plasma membranes. Whether plasma-membrane asymmetry also applies to the phospholipids of other eukaryotic cells is not yet known. In animal cells, most of the glycosphingolipid carbohydrate is associated with the outer plasma membrane (Nicolson and Singer, 1974). The glycophosphosphingolipids of fungal cell surfaces may be situated likewise (see p. 96). New plasma membrane of eukaryotic cells is considered to arise from the fusion of secretory vesicle membranes with old plasma membrane. According to the membrane-flow hypothesis, the choline-containing phospholipids, glycolipid, glycoproteins and certain proteins are located in the inner surfaces of Golgi apparatus and secretary vesicle membranes and, with fusion, these become the outer plasma membrane. Sterols are typical of eukaryotic membranes. Animal cell membranes, such as rat-liver plasma membrane, contain 6 7 % cholesterol. The average sterol content of fungal organisms ranges from 0.7 to 1.O% of the dry tissue. This represents 5-6% of the membrane mass. The most common sterol is ergosterol. The chemistry, distribution and biosynthesis of fungal sterols have been thoroughly reviewed (Weete, 1973; Weete and Laseter, 1974; Nes, 1974). Nevertheless, there have been few contributions to an interpretation of the molecular role of fungal membrane sterols. Some clues can be derived from a comparison with other eukaryotic systems and synthetic bilayers. When cholesterol is introduced into a bilayer, the plate-like steroid rings interact with and partly immobilize these regions of the hydrocarbon chain closest to the polar head group (leaving the rest of the paraffin chain flexible). This prevents crystallization of the fatty acids (Chapman, 1973). A result of this is a more condensed and thicker phospholipid bilayer. Furthermore, the permeability rates of anions, cations, and non-electrolytes are considerably lowered. in the presence of cholesterol. Thus, in high concentrations, cholesterol abolishes temperature-induced phase transitions; it has the dual effect of preventing formation of crystalline gel areas, while inhibiting the overall

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

61

flexing motions of the hydrocarbon chains. In this way, it imparts an intermediate fluid condition to the interior of the bilayer. Therefore, in general terms, cholesterol controls the fluidity of the hydrocarbon chains of phospholipids, providing a more stable structure over a wide temperature range. This stability is accompanied by a general decrease in permeability and electrical conductivity and an increase in thickness and electrical capacitance. Hamilton-Miller ( 1974) compared the protecting effects of free sterols, sterol esters and sterol glucosides on leakage of electrolytes through membranes, and found that sterol esters and glucosides were inactive. Apparently only free sterols are involved in controlling membrane stability and membrane permeability, indicating the importance of the hydroxyl group at C-3 of the ring. As expected, sterols can also have significant effects on membrane proteins, for lowering the fluidity of membrane can inhibit the activity of certain enzymes that require a critical fluid phospholipid environment. Like some other membrane constituents, cholesterol is also apparently asymmetrically distributed across the plane of the erythrocyte’s plasma membrane (Fisher, 1975). Membrane splitting by freeze-fracture, and chemical analysis of the outer and inner halves, showed a two-to-one distribution of cholesterol in favour of the outer half. The polyene antibiotics (e.g. nystatin, filipin) inhibit growth of many fungi. These complex with the sterols present in the plasma membrane, thereby making it leaky, with the loss of ions, sugars and amino acids. The mode of action of polyene antibiotics on fungal sterols has been well reviewed (Hamilton-Miller, 1974). In animal cells, a cholesterol to phospholipid ratio of one is characteristic of plasma membranes with a high degree of purity. The observation of Weinstein et al. (1969) that surface membranes of animal cells possessing a high cholesterol-phospholipid ratio may also have a high glycolipid content, if applicable to fungi, would indicate that their glycolipids are associated with the plasma membrane and are possibly involved in cell-wall synthesis and attachment. I t is difficult to obtain reliable figures for the carbohydrate content of fungal plasma membrane since protoplasts are usually obtained after enzymic digestion involving glucosidase activity. However, within this constraint, Hunter and Rose ( 197 1 ) reported figures ranging between 3 and 6% for the carbohydrate content of the plasma membrane from various yeasts. The functional nature of this carbohydrate

Q,

N

P

TABLE 2. Phospholipids of fungi

L Acidic Phosphoglycerides

Neutral (Zwitterionic)Phospholipids

W

m

_.

rn

0

0

(i) Phosphatidic acid (PA)

II

z z

II

CH,O-C-R,

I

o

R,-C-OCH

I II CH,O P-OI

0

( a ) Phosphatidylethanolainirie IPE)

II

DG-P-0-CH,-CH,-NH,

+

I

P

D

z 0

0-

P

0

0

0

II

(ii) Phosphatidylserine (PSI

II

I

0-

0

0

(iii) Phosphatidylglycerol ( P C )

v)

“DG-P-0-CH,-CH-COO-( b ) Ptio~phdt~dy~lnonolrlethylt.thdno~DG-P-O-CH,-CH,-NH,(CH,) + I I driiiiir ( P M M E ) 0N H:

II

DG-P-0-

CH,

I

HOCH

I

CH,OH

(c)

Phosphatidyldimethylethanolarnine (PDME)

DG-P-O-CH,-CH,II

I

0-

NH(CH,),

rn

r

0

(iv) Diphosphatidylglycerol (DiPG)

II

0

DC-P-0-CH,

I

0-

I HOC H I

DG-P-o-cH,-CH,-~-(CH,), II (d) Phosphatidylcholine (PC)

0

I

II

0-

CH,O-P-0-DG

I

0 0

iv) Phosphatidylmyo-inositol (PI)

DG-P-o

I 0

OH H O

4

With di- and triphospho-inositides, the hydroxyl groups at positions C-4 and C-5 are esterified with phosphate to give 1-phosphatidylmyo-inositol-4-phosphate (diphospho-inositide, DPI) and 1-phosphatidylmyo-inositol-4,5-diphosphate itriphospho-inositide, TPI) “DG refers to the diacylglycerol moiety of the phosphoglycerides.

(c) Splringomyelin ISM,

cera~nylphosphorylcholit~e) 0

I1 CH,-(CH,),,-CH=CH-CH-CH-CH~-O-P-O-CH,CH,-N-~CH,~,

I

OH

I NH I c=o I R

I 0-

+

-n

C

z 0

E

nD

v

cn

cn W

64

P. J. BRENNAN AND D. M. LOSEL

is not known but some, or perhaps all, of it is due to membraneous proteoglycans and glycosphingolipids. Proteins are the predominant components of all eukaryotic membranes, comprising 41-54% of the mass of most membranes, compared to 26-32% for phospholipid. These facts are the basis of recent models of membrane structure which are apparently of general applicability. The model receiving greatest attention is that proposed by Singer and Nicolson (1972).The basic premise is that the majority of membrane protein is in helical conformation, and can assume a shape such that hydrophilic amino acids (and the glycosylated peptides in the case of plasma membranes) are in contact with the exterior, and the apolar amino acids are embedded in the paraffin area of the bilayer. The membrane is a mosaic of alternating globular protein and phospholipid bilayer in a dynamic state of flux whereby there is a constant mobility of the components in a lateral direction. Two types of membrane proteins are present, namely integral and peripheral. The integral proteins, constituting the bulk of membrane proteins, cannot be readily separated from their surrounding phospholipids. These are largely globular in nature and are not spread out as monolayers. The peripheral proteins may not be directly relevant to the structure of the membrane; they are held to one of the surfaces of the membrane by weakly non-covalent interaction and are dissociated by an increase in ionic strength or use of chelating agents and, when isolated, are usually lipid-free. With fungi, it is doubtful if there are periphereal proteins on the outer surface of the plasma membrane, in view of the presence of a cell wall. Preparation of yeast protoplasts, usually the first step towards isolating plasma membrane, has been reviewed by Kuo and Yamamoto (1975). Most procedures use lytic enzymes present in gut juice of the snail Helix pomatia, which has disadvantages under certain circumstances, as already mentioned. Similarly the review of Wiley (1974) on the isolation of sphaeroplasts, the consequent cytoplasmic membrane vesicles and other subcellular structures from yeasts and filamentous fungi, describes procedures which rely entirely on digestive enzymes. The mechanical procedure described by Nurminen et al. ( 1976) seems effective and simple for the isolation of yeast plasma-membrane fragments. Whole cell homogenates were subjected to zonal centrifugation on sucrose and iso-osmotic Ficoll, yielding plasma-membrane fragments of different densities. The methods developed by Scarborough ( 1975)for the isolation of plasma membranes are also worthy

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

65

of particular mention. They have been applied to a cell wall-less mutant of Neurospora crassa but should be applicable to many types of' fungal sphaeroplasts. Plasma membranes were stabilized against fragmentation and vesiculation by treatment of intact cells with concanavallin A. Ghosts were isolated by low-speed centrifugation and converted to vesicles by removal of the concanavallin A. Unfortunately these were contaminated by significant amounts of nonmembrane carbohydrate (about 14% of the membrane mass). However, the molar ratio of sterols to phospholipids was about l . 3 and the membranes contained 60% of cellular sterols. Hereward ( 1974) described the isolation and features of rough membranes of Schizosaccharomyces pombe, and Smith ( 1972) described the isolation of Golgi apparatus from the same organism. Schatz and Kovac (1974) described the isolation and characterization of respiratory-deficient mitochondria1 precursors (promitochondria) from Sacch. cereuisiae grown anaerobically on a fermentable carbon source. The promitochondria were devoid of cytochromes a, as, b, c and ubiquinone, and contained low levels of ergosterol and unusual fatty acids. B . PH

o s P H o L I P I D S-TY'P

Es AND D I STRIBUTI

oN

The structures of all of the major phospholipids found in fungi are shown in Table 2, and the distribution of phosphoglycerides in selected fungi is shown in Table 3. I t is obvious that phosphatidylcholine, phosphatidylethanolamine, phosphatidylserine, phosphatidylinositol and diphosphatidylglycerol are the major fungal phosphoglycerides. Phosphatidylglycerol, common in bacteria and chloroplasts, is rare in fungal extracts. I t seems from these patterns that fungi exhibit a conformity of pattern reminiscent of the organ similarity of phospholipids from different animals. However, it is possible that insufficient numbers of the higher and lower fungi have been examined to detect a distinctive pattern. Jack (19661, in a brief examination, conclude-d that the pattern of major phosphoglycerides present in yeasts applied to other fungi, although considerable variation in the relative amounts of the individual species was found. Mumma et al. (197 1) quantitatively compared the phosphoglycerides of a thermophilic deuteromycete, Humicola grisea var. thermoidea, with that of other fungi, and concluded that phosphatidylcholine and phosphatidylethanolamine were the major phosphoglycerides in basidiomycetes, deuteromycetes and phycomycetes.

m m

TABLE 3. Phosphoglyceride distribution i n selected f u n g i Phosphoglyceride (percent of total phospholipid) (for key, see foot of table)

Organism

PC

PE

PS

PI

DPG

PG

PA

Phytophthora parasitica Pythium ultimum Glomus mossez Phycomyces blakesleeanus Schizosaccharomyces pombe Saccharomyces cereuisiae * Saccharomyces cereuisiae Saccharomyces cereuisiae Saccharomyces cereuisiae Saccharomyces cerevisiae

41.3 33.7 15.9 31.2 44.9 46.4 45.2 42.0 27.0 55.0

20.3 24.1 34.1 41.0 19.4 14.9 16.7 22.4 29.0 20.0

0.6 0 17.3 11.9 5.7 9.5 9.3 7.3

6.6 10.7

2.6

2.4 11.6' 19.0

1.6 10.7

Saccharomyces cereuisiae Saccharomyces cereuisiae Saccharomyces cereuisiae Saccharomyces marrianus Saccharomy cesfragilis Saccharomyces pastorianus Saccharomyces telluris (themotolerant) Candida macedoniensis Candida lipolytica (rnesophile) Candida sp. (psychrophile) Candida lipolytica (rnesophile) Candida parapspsilosis (themotolerant)

42.0 42.4 44.0 42.0 30.5 31.4 40

35.0 18.3 24.0 22.0 24.4 24.3 25

11.0 19.0 9.1 7.6 2

9.0 22.6 21.0 17.0 22.8 16.2 14

41.0 36.0 44.0 42 40

26.0 29.0 25.0 32 26

17.0 17.0 16.0 2 4

16.0 14.0 7.0 9 14

DMPE

References

P 0 OMYCETES ZYGOMYCETES HEM IASCOMY CETES

3.4 17.6 12.7 19.8' 22.5 23.0e 15.0'

1.8

13.7 2.6 7.5 5.3d 2.2 <5

2.4 1.6 2.5 1.4

1.6

<5

12.0 7.1

5.2 1.9 10

I .0 1.4 2


2

Hendrix and Rouser (1976) Bowmanand Mumma 11967) Cooper arid Losel(1978) Hendrix and Rouser (1976) White and Hawthorne (1970) 0.2 Get2 et al. (19701 Deierkauf and Booij (1968) 2.2 Letters (1966) Longley et al. (1968) Suornalaineri arid Nurminen (1970) Vignais etal. (1970) Trevelyan (1966) Graf el al. ( 19681 Grafetal. (1968) Hendrix and Rouser (1976) Hendrix and Rouser ( 1976) Arthur and Watson (1976) Grafet al. (19681 Kates and Baxter (1962) Kates and Baxter (1962) Arthur and Watson (1976) Arthurand Watson (1976)

1

n

rn 2 2

4

D

2 0

0

5 r

0:

cn

rn

r

EUASCOMYCETES

BASIDIOMYCETES

DEUTEROMYCETES

Candida sloofri (thermophilic) Torulopsis bouzna (thermophilic) Trigonopsis uariabilis Brettanomyces truxellensis Hansenula anomala Endomycopsis selenospora Neurospora crassa Neurospora massa Arthroderma uncinatum Tricholoma nudum Agaricus bisporus Agaricus bisporw Schizophyllum commune Polyporus uersicolor Uromycesphaseoli (uredospores) Uromycesphaseoli (germinated uredospores) Bullera alba Leucosporidiumfrigidum (psychrophile) Humicola gnsea var. thermodiae

30 30 51.0 41.0 53.0 46.0 44.4

20 20 18.0 21.0 23.0 23.0 24.3 47.6 23.8 40.1 21.0 59.0 26.0 3 9 . d 13.8 47.81 50.2 42.1 30.3 46.6 25.1 54.4 27.2

8 7

10.5

11.7d

13.7 11.5 7.9

4.3 3.4 6.1

2.7 3.3 2.8

2.8 3.4 1.6

Arthurand Watson 11976) Arthur and Watson (1976) Grafetal. (1968) Graf et al. (19681 Graf et al. ( 1968) Grafetal. (1968) Hendrix and Rouser (19761 Kushwahaetal. (1976) KishandJack(1974) Leegwater et al. (1962) O’Sullivan and Lose1 (197 1 ) HoltzandSchisler(1971) Hendrix and Rouser (1976) Hendrixand Rouser (1976) Hoppe and Heitefuss (1974b)

0.6

1.9

Hoppe and Heitefuss (1974b)

8

2

Graf et al. (1968) Arthur and Watson (19761

12

2 16.0 16.0 15.0

14.0 5.5 4.8 15.6 8.0 12.3

20 6 25 15.0 22.0 9.0 17.0 9.9 5.3 4.8 8.1 9.7

52.5

32.4

8.0

4.6

54.0

30.0 43

8.0 6

8.0

27 33.0

18.3

3.3

12.8

7

3.9

2.8 19

4.9 7.0

28.1

Mumma et al. ( 197 1)

PC indicates phosphatidylcholine, PE phosphatidylethanolamine, PS phosphatidylserine, PI phosphatidylinositol, DPG diphosphatidylglycerol, PG phosphatidylglycerol, PA phosphatidic acid, and DMPE dimethylphosphatidylethanolamine. ‘Contains lysophosphatidylethanolamine.*Late stationary-phase cells. ‘Contains PG. dContains PA. PI + PS. See text for discussion. Raju et al. (1976) recently examined the lipids of a number of thermophilic fungi ( o n the basis that their ability to survive and grow at high temperatures may be due to intrinsic heat stability of their membrane components). In Thermoascus aurantiacus, Humicola lanuginosa, Malbranchea pulchella var. sulfurea and Absidia ramosa, there was considerable variation in the levels of the various phosphoglycerides: PC (15.9-47%), PE(23.4-67%), PS (9.3-17.6%) and PI ( 1.9-1 1.9%). DPG occurred in quantity only in Humicola lanuginosa and Malbranchea pulchella var. sulfurea. PA was present as a minor component u n l y in MalbrancheapulchellaVal.. sulfurea and Absidia rumosa and I S apparently not a characteristic lipid ot therniophilic lungi

68

P. J. BRENNAN AND D. M. LOSEL

I t appears from what is known that the bulk of fungal phosphoglycerides are phosphatidylcholine and phosphatidylethanolamine, which is a reasonably consistent eukaryotic trend. Values for phosphatidylcholine range from 35-50% ofwhole-cell phosphoglycerides. However, Byrne and Brennan (1975) could not detect phosphatidylcholine in the mycelial or sporophore stages of Agaricus bisporus. Moreover, Mlodecki et al. (1972) were unable to detect phosphatidylcholine in fresh and dried mushrooms of species Leccinum scabrum, Leccinum aurantiacum and A. bisporus. It seems that the phospholipid composition of some eukaryotes is subject to genetic and dietary variations, and the prokaryotic trait of the virtual absence of phosphatidylcholine and phosphatidylinositol is sometimes exhibited by fungi. Merdinger ( 1969) was unable to detect phosphatidylcholine or phosphatidylinositol in the ascomycete Pullularia pullulans in studies extending over several years. In this fungus, phosphatidylethanolamine and phosphatidylserine were the principal phospholipids. Phosphatidylethanolamine has been observed in all fungi examined, comprising 14-35% of total phosphoglycerides. Unlike in prokaryotes, phosphatidylserine is a prominent fungal phosphoglyceride, its content ranging from 4-20%. The level of phosphatidylinositol in fungi is also very variable, ranging from 8 to 23%. Phosphatidylinositol 4-phosphate and phosphatidylinositol 4,5-diphosphate have been recognized in small amounts in yeasts (Lester and Steiner, 1968; Prottey et al., 1970).The relative amounts of phosphatidylinositol : diphospho-inositide : triphospho-inositide in Sacch. cerevisiae is 140 :4 : 1 (Lester and Steiner, 1968). Only in a few cases have the fatty acids of individual phosphoglycerides been examined; 1,2-dipalmitoleyl-sn-gl~cerol-3phos@orylcholine has been identified as the major species in members of the Saccharomycetaceae family, Sacch. cereuisiae (Hanahan and Jayko, 19521, Sacch. carlsbergensis (Shafai and Lewin, 1968), Hunseniaspora ualbyensis (Haskell and Snell, 19651, and in a pseudosaccharomycete Candida sp. (Kates and Baxter, 1962). DeBell and Jack (1975) have shown stereospecific distribution of fatty acids in the phosphoglycerides from the mycelium and sporangiospores of Phycomyces blakesleeanus. In phosphatidylcholine and phosphatidylethanolamine, 85%of the fatty acids on sn-1 are C,6,0,C i s : ?and C18.3. The sn-2 position contains 98% unsaturated fatty acids, mostly C,8:2and C1x:3.In phosphatidylcholine fi-om Lipomyces lipoferus, the sn-2 position was 26.7% more

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

69

unsaturated than sn-1 , and in phosphatidylethanolamine sn-2 was 10.8%more unsaturated than sn-1 (Haley and Jack, 1974). The fattyacid content of phosphoglycerides as a class has been extensively examined (Weete, 1974). As with phosphoglycerides from other biological sources, the predominant saturated acid is palmitic. However, unlike animal and higher plant sources in which oleic acid is generally the major unsaturated fatty acid of phosphoglycerides, in many yeasts palmitoleic acid occupies this position. This applies in particular to yeast phosphatidylcholine and phosphatidylethanolamine since, in phosphatidylserine and phosphatidylinositol from the same sources, much of the palmitoleic acid is replaced by palmitic acid. For instance, the phosphatidylserine species of Lipomyces starkeyi is almost en tirely 1 -palmitoyl-2 -0ley1-sn -glycero- 3 -p hosp horylserine ( Suzuki and Hasegawa, 1974). C.

P H 0 S P H 0 G L Y C E RI D E S - P H Y

S I OLO G I CA L A S P E C T S

I t is now fairly certain that all of the phosphoglycerides of fungi are associated with plasma or cytoplasmic membranes. Northcote and Horne (1952) reported that yeast cell-wall lipids were mostly triglycerides with little phosphoglyceride. The substantial amounts of phosphoglycerides in cell walls, reported by Nurminen and Suomalainen (197 l ) , are attributable to fragments of plasma membrane. Longley et al. (1968) carried out an extensive investigation of the protoplast membrane of Sacch. cereuisiae. These were very similar in phospholipid composition to whole-cell extracts, and contained only slightly more phosphatidylethanolamine, phosphatidylinositol and phosphatidylserine and slightly less phosphatidylcholine. Membranes accounted for 13.2% of the weight of dry yeast cell and contained 39% lipid, 49%protein, 6% sterol, 4.6%carbohydrate and about 7% RNA. The principal structural lipids were a range of phosphoglycerides comprising between 15 and 25% of the membrane. These lipids were associated with proteins as complex lipoproteins, forming the functional membrane structure. Analysis of protoplast membranes of Candida utzlzs yielded similar results, namely 38.4%protein, 40.4% lipid, 5.5% carbohydrate and 1 . 1 % RNA (Mendoza and Villanueva, 1967). And again the analyses of Boulton (1965) on the plasma membranes of Sacch. cereuisiae support these values, and the general contention that practically all fungal phosphoglyceride belongs to membranes.

70

P. J. BRENNAN AND D. M. LOSEL

More recently, a novel approach adopted by some workers has led to new insight into the role of fungal-membrane phosphoglycerides. In this, the nature of the polar head groups was altered by various means, the reason being that the electrostatic field extending into the aqueous environment from the lipid bilayer determines some important membrane phenomena; for example, the specificity and activity of enzymes can be profoundly influenced by the surface charge and ionic environment of the phospholipid bilayer, as can the action of certain hormones, vitamins and drugs. Bangham ( 1961 ) suggested that an important feature of biological membranes is their low surface charge density, and this is achieved by a partial negation of the charge on anionic phospholipids by non-ionic and zwitterionic lipids. The idea that this membrane charge is conserved under extenuating conditions is now becoming accepted. To test this idea, attempts have been made to alter the phospholipids of several micro-organisms by genetic and dietary means. Crocken and Nyc (1964) and Hubbard and Brady (1975) took advantage of the large number of available mutants of Neurosporu crussa defective in phospholipid synthesis to explore the consequences of variations in phospholipid head groups. Crocken and Nyc ( 1964)first established the principle that the phospholipid composition of this fungus could vary within broad limits without lethal damage to cell membranes. Hubbard and Brody (1975) later established the details of these limits. They used four auxotrophic mutants and a wild-type strain. The mutants were: chol-1 (defective in phosphatidylethanolamine methylation); chol-2 (defective in phosphatidylmonomethylethanolamine methylation) : inos (defective in a phosphatase for myo-inositol 1-phosphate); and a double mutant chol-I : chol-2 (see Section IT, p. 102 for details on biosynthesis of phosphatidylcholine). When these mutants were grown with enough supplement to support growth, they exhibited bizarre phospholipid patterns. By an appropriate choice of mutant and supplement, it was possible to vary the relative level of every phospholipid in the organism, with the exception of diphosphatidylglycerol. The maximum ranges reported for the zwitterionic species were : phosphatidylcholine (0.9-53.1%), phosphatidylmonomethylethanolamine (0.0-55.5%), phosphatidyldimethylethanolamine (0.0-53.9%) and phosphatidylethanolamine (9.8-43.3%). For the anionic species, the ranges were phosphatidylserine (1.7-10.4%) and phosphatidylinositol (3.6-25.196). Despite this wide variation in the relative proportions of the individual

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

71

phospholipid species, five quantities remained constant: that of diphosphatidylglycerol, total phospholipid (50 pmoles phospholipid P/g lyophylized mycelium), total content of zwitterionic species, total amount of anionic .species, and the ratio of zwitterionic to anionic totals. One observation was that, when the cellular content of phosphatidylinositol in the inos strain was lowered to less than half of its normal level, the deficiency was compensated by an equimolar increase in the level of the anionic phosphatidylserine. Paltauf and Johnson (1970) had noted a similar increase of phosphatidylserine in inositoldeficient cells of Sacch. carlsbergensis. These results suggest the existence of an internal compensatory biosynthetic mechanism which maintains a fairly constant contribution by the phospholipid components to the overall membrane charge. In addition, the results suggest that i t is possible to manipulate extensively the phospholipid content of a eukaryote without drastic variation in growth conditions. Hubbard and Brody (1975), commenting on these results, state that the use of the mutants provides a different approach for in uivo and in vitro investigation of the effects of altered membrane structure in nuclear function, protein conformation, respiratory efficiency, transport and membrane-related processes. Minnikin and colleagues (Minnikin et al., 1972a, b, 1974) used a nutritional approach to arrive at related conclusions. However, they mostly used bacilli and pseudomonads, although some of the conclusions seem to apply to a yeast and a mould. They noted that, in certain bacilli under conditions of phosphate-starved batch culture and phosphate-limited chemostat culture, phosphatidylethanolamine was partially replaced by diglucosyldiglyceride. In Bacillus cereus T, anionic phospholipids under conditions of phosphorus limitation were partially replaced by anionic glycolipids (mostly glucuronosyl diglyceride) and, in the case of Bacillus subtilis 3610, by an anionic pep t id ol i pid . With various pseudomonads limited in their supply of nutrient phosphate, analogous changes took place. In Pseudomonas diminuta, the anionic phosphoglycerides were replaced with anionic glycolipids. In PS.Juorescens, the zwitterionic phosphatidylethanolamine was replaced by an ornithine amide. These results seem to indicate that neutral glycolipids of the diglycosyldiglyceride type and zwitterionic ornithine amides are interchangable to various degrees with the zwitterionic phospholipids. Similarly, anionic glycolipids containing hexuronic acids may replace anionic phospholipids. The latter results are more

72

P. J. BRENNAN AND D. M. LOSEL

akin to those obtained with the N . crassa mutants, although neither Hubbard and Brody (1975) nor Crocken and Nyc ( 1964)examined the status of the newly found glycolipids of N . crassa (see Table 6, p. 86). The results of Minnikin and his colleagues further support the contention that phosphorus-free polar lipids of bacterial membranes have the same basic membrane function as phospholipids, and that the net charge on the lipid head groups remains fairly constant despite serious alterations in the nutrient composition, i.e. roughly equal proportions of acidic and neutral (or zwitterionic) polar lipids prevail. Johnson et al. (1973) reported that, in phosphate-limited chemostat cultures of Candida utilis, the major phosphoglycerides were largely replaced by three unidentified glycolipids (two of which were probably simple glycosylceramides). A less dramatic effect was observed in phosphorus- depleted batch - cultured A . niger ; monoglucosy loxyfatty acid became the dominant polar lipid class, but there was no evidence for a compensating zwitterionic phospholipid (Laine et al., 1972). Nevertheless, the notion seems to be established that the presence of phosphoglycerides as major components of at least some fungal membranes is not obligatory provided that other polar lipids are present. Phospholipids are intimately involved in the functional organization of a number of complex enzyme systems, most notably the mitochondrial electron- transport chain. Moreover, several individual enzymes have been shown to require absolutely one or more species of phospholipid for activity (Finean et al., 1974). Most of these systems have been studied in mammals. However, there are several examples of this type of interaction in fungi. In particular, the involvement of phospholipid in yeast mitochondria1 function has been examined. As with animal systems, yeast diphosphatidylglycerol has almost exclusively been associated with mitochondria and the amount present is closely correlated with the state of development of the yeast mitochondrial membrane (Jakovcic et al., 197 1). Mangnall and Getz (1973) report that, following the transfer of glucose-grown cells to a nonfermentable medium, the respiratory competence increased more than three fold and this was accompanied by a doubling of the diphosphatidylglycerol content. In the case of the “petite” (respiratorydeficient) strains of yeast, the diphosphatidylglycerol content of mitochondria dropped from 15.6%of total phospholipid to 9.1% and this was accompanied by a corresponding increase in phosphatidylinositol Oakovcic et al., 197 1). Moreover, respiratory-deficient mitochondria (promitochondria) obtained from anaerobically grown Sacch. cerewisiae

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

73

have less diphosphatidylglycerol and more phosphatidylinositol as well as alterations in the other phospholipids (Paltauf and Schatz, 1969). Arthur and Watson ( 1976) showed that the respiratory-deficient Candida slooJji, was completely devoid of diphosphatidylglycerol. Mangall and Getz (1973) suggest that diphosphatidylglycerol is an important component of the “scaffold” membrane of the inner mitochondrial membrane. Presumably, it is into this area of the inner mitochondrial membrane that respiratory and phosphorylative components are inserted during the transition of promitochondria to functional mitochondria. Phospholipids have also been involved in membrane transport of‘ phosphate and arsenate in fungi (Cerbon, 1969, 1970).Arsenate enters a constitutive system normally reserved for phosphate, which i t inactivates, and transport ceases. Phosphatidylinositol in particular has been implicated in this phenomenon. In arsenate-adapted cells, there was a large increase in phosphatidylinositol content, whereas synthesis of the other usual phosphoglycerides was decreased. One suggested explanation was that arsenate combines with newly synthesized inositol (perhaps by substituting for the phosphate group) thereby facilitating arsenate uptake. Phosphatidylcholine has been implicated in lysine unptake by N . c r a m (Sherr, 1969) since a phosphatidylcholine-deficient mutant had a decreased capacity for lysine uptake. Hydrolysis of phosphatidylglycerolphosphate to phosphatidylglycerol in yeast has been considered (Deierkauf and Booij, 1968) to be important for the active transport of D-glycerol and amino acids. Polyphospho-inositides may be involved in transport of protons o r other cations across yeast membranes. The rapid turnover of the phosphomonoester groups of di- (DPI) and triphospho-inositides (TPI) in brain is probably due to the combined action of phosphatases and kinases, and this rapid turnover is thought to be a significant aspect of their function. The discovery in fungi of di- and triphospho-inositides with a rapid metabolic turnover (Steiner and Lester, 1972) suggested that polyphospho-inositides may have a general biological role, not solely restricted to nerve tissue. Talwalkar and Lester ( 1973) showed that the adenylate energy charge of the yeast cell was a big factor in determining the concentration of the polyphospho-inositides. When a respiratory-deficient mutant strain of Sacch. cerevisiae was transferred from complete growth medium to buffer, there was an immediate (within one min) and large drop in the levels of DPI, TPI, ATP and

74

P. J. BRENNAN AND D. M. LOSEL

adenylate charge (ATP + iADP)/(ATP+ ADP + AMP). In contrast, the major cellular phospho-inositides, phosphatidylinositol and mannosyldi-inositoldiphosphorylceramide (CerP,I,M) were unchanged. A similar starvation experiment on the parental yeast strain showed a qualitatively similar pattern. When glucose was supplied to the starved respiratory-deficient cells, a rapid increase in the concentrations of ATP, DPI and TPI ensued. Thus, there was a clear-cut correlation between the adenylate energy charge and the concentrations of’ the polyphospho-inositides. In explaining the biological implications of the turnover of the phosphomonoester groups of the phosphoinositides, Talwalkar and Lester ( 1973) considered both stoicheiometric and catalytic mechanisms. Binding and release of Ca2+, or transport of other ions, may be stoicheiometrically coupled to one turn of the cycle (PI-DPI and TPI-PI) with some of the energy charge released from ATP being conserved for driving this sequence. In addition, polyphospho-inositides could have a catalytic role. Located as they are in membranes, their phosphate groups could combine with cations to produce molecular species of lowered net charge, thereby facilitating the diffusion of such ions across the membrane.

D.

GLYCOLIPIDS

The nitrogen-free glycolipids of fungi can be divided into two broad structural classes, namely hydroxy acid glycosides and acylated sugars or polyols. They can also be subdivided on a functional basis; many appear to be membrane associated whereas others, in particular the sophorosides and ustilagic acids, are largely secondary metabolites. 1. Hydroxy Acid Glycosides The major glycosides of yeasts, moulds, and some basidiomycetes, are listed in Table 4. Glycosides corresponding to the sophorosides from C. bogoriensis and T . upicola have not been isolated from the cell; they are all extracellular products (Stodola et ul., 1967).They are therefore dealt with in Section V (p. 131). Laine et al. (1972) described a major lipid from A . niger as a monoglucosyloxyoctadecenoic acid. A related compound from A . bisporus (Byme and Brennan, 197 5 ) appeared to be a monoglucosyloxyhexadecenoic acid or its methyl ester. The assignment of these structures was based on the following information: (i) a glucose to hydroxyfatty

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

75

acid ratio of about unity; (ii) the presence of a carboxylic function, as shown by infrared spectroscopy; (iii) thin-layer chromatographic properties similar to those of synthetic monoglucosyloxyoctadecenoic acid; (iv) sphingosine bases were apparently absent; (v) the presence of 2-hydroxyfatty acids, as shown by gas-liquid chromatography-mass spectroscopy. However, more recently in trying to isolate more of this material from A . bisporus, only a glucosylceramide was found (P. J . Brennan and J. Madden, unpublished results). The monoglucosyloxyfatty acid of A. niger is apparently cell associated; it is absent from the growth medium of A. niger (Laine et al., 1972). Results reported by Byrne and Brennan (1974) implied a membraneous role for this compound. Aspergillus niger grown batch-wise in a fermentor, or grown shaken for long periods in a phosphate-deficient medium, produced the monoglucosyloxyfatty acid which was the dominant polar lipid since there was virtually no phospholipid in the mycelium. However, when the mycelium was grown in the presence of high concentrations of phosphate, phosphoglycerides were the predominant polar lipids. The results seem to suggest that the glycolipid was retained in membranes under conditions in which phosphoglycerides apparently were not synthesized o r were degraded. Since there were no observable microscopic differences between phospholipid-rich and phospholipid-poor mycelium, and obvious cellular disruption had not taken place, it appeared that the glycolipid could assume the role of phosphoglycerides as structural components of the membranes. These interpretations are compatible with the view that glycolipids and phosholipids have complementary and supporting functions in the lipid bilayer of biological membranes. The observation that monoglucosyloxyoctadecenoic acid of A . niger is apparently capable of supporting membrane functions under conditions in which the cellular phospholipid complement is depleted may have further physiological implications. Aspergilli are noted for their ability to survive under conditions of very low external pH value, the high acidity of the medium being a result of their own metabolic processes. In baker’s yeast, the increase in concentration of hydrogen ions in the medium has been shown to be coupled with the intracellular accumulation of potassium ions (Riemersma, 1964). The membrane ATPases responsible for such ionic exchanges processes in higher organisms show an absolute requirement for phospholipid (Uesugi et al., 197 1 ; Kyte, 197 1). This phospholipid requirement has been made the basis of molecular theories of ion exchange in which roles for the

TABLE 4. Glycolipids of fungi Glycolipid

Source

HYDROXYACID GLYCOSIDES 13-((2’-0-P-o-glucopyranosyl-P-o-glucopyranosy1)oxy) docosanoic Candida bvgvriensis acid 6’6” -diacetate (Ac,Glc,HDA) 13-~~2’-0-~-~-glucopyranosyl-~-~-glycopyanosyl~oxy~ docosanoic Candida bogoriensis acid 6’-monoacetate (AcClc,HDA) 13-((2’-0-p-D-glucopyranosyl-P-o-glucopyranosy1)oxy)docosanoic Candida bvgoriensis acid (Glc,HDA) 1 7 -((2‘-O-~-o-glucopyranosy~-~-D-glucopyranosyl)oxy)octadecanoic Torulvpsis apicola acid 6’6” diacetate 1 7-((2’-0-/3-~-glucopyrranosyl-/?o-g1ucopyranosyl)oxy)octadecenoic Tvrulopsis apicvla acid 6’6” diacetate Tvrulvpsis apicda, 1 7 - L-((2’-O-~-o-glucopyanosyl-~-o-glycopyranosyl)oxy) octaTorulvpsis gropengensis “ decanoic acid 1,4”-lactone 6’6” diacetate (a sophoroside lactone) Tvrulvpsis apicvla 17 - ~ - ( ( 2 ’0-pD-glucopyranosyl-p- D-glucopyranosyl)oxy)octadecanoic acid 1,4”-lactone 6” monoacetate (a sophoroside lactone) Aspergdlus niger 2’-glucosyloxyoctadec-trans-3 -enoic acid USTILAGIC ACIDS Glucoustilic acid A is the 2”-acylated P-cellobiosyl residue glycosidically Ustilago maydis attached to ustilic acid A : OH

I

HOH,C-CH-(CH,),,-COOH

Reference

P Tulloch et al. (1968b)

L

Tulloch et al. (1968b)

W n rn

Esders and Light (1972a) Gorin et al. ( 196 1 ) Tulloch et al. (1962) Gorin et al. ( 1961) Tulloch et al. (1962) Tulloch et al. (1968a) Jones (1967) Tulloch et al. ( 1968a) Laine et al. (1972) P. J . Brennan and F. Gunstone (unpublished data) Reindel(1930) Lemieux et al. (1953) Stodola et al. (1967)

z

z

P D

z 0

P

T r

0 v)

r rn

Glucoustilic acid B is the Z"-acylated p-cellobiosyl residue glycosidically attached to ustilic acid B: OH OH

I

Ustilago nudu

Bhattacharjee et al. (1970)

I

HOH,C-CH-(CH,),,-CH-COOH The acyl groups at position 2" of the cellobiosyl residue are acetate, ~-3-hydroxyhexanoate,L-3- hydroxyoctanoate and hexanoate

ACYLATED SUGAR ALCOHOLS AND SUGARS Rhodotorula graminis 6-0-(3-o-hydroxyhexadecanoyl)o-mannitol. Also esters of o-arabitol and xylitol with 3-D-hydroxyoctadecanoicacid Rhodotorula graminis Mannitol and perititol esters of3-o-hydroxyhexadecanoicand 3- o-hydroxyoctadecanoic acids. One molecule of the acid is attached to each polyol and most of the remaining hydroxyl groups, including that o n the fatty acid, are acetylated Saccharmyes cerevisiae Mono- andpolyacylglucoses, acyltrehalosk Agaricus birporus (mycelium) Pullularia pulluluns ~ - 0 - ~ ~ , ~ , ~ , 6 - t e t r a - ~ - a c y ~ - ~ - ~ - m a n n o p y r a n o s y ~ ~ - D -Ustilago e r y ~ r muydis ito~ Fatty acids are from C,, to C,,, with C,, predominant GLYCOSYLDIGLYCERIDES 1+2 Glucosyl-a- or -Glucosyl-a-(3-1,Z-sn-diglyceride) 1+6 1+2 Galactosyl-p- or -Galactosyl-/3-(3-1,Z-sn-diglyceride) 1-6 The principal fatty acids are palmitic and oleic acid Mono- and diglycosyldiglycerides

Aspergtllus niger

Tulloch and Spencer (1964) Tulloch and Spencer ( 1964)

Brennan et al. (1970) Bryne and Brennan (1975) Merdinger et al. (1968) Boothroyd et al. (1956) Fluharty and O'Brien (1969)

P. J. Brennan and J . A. Hackett

(unpublished observations)

Blastocladiella emersonii

Mills and Cantino (1974)

'Jones (1967) at one time considered that the hydroxyl group at C-4 ofthe relevant glucose was not esterified,but that an acetate residue was present the hydroxyl group at C-3 and that the fatty-acid carboxyl group was linked at the hydroxyl at C-6. However, this impression was corrected by Tulloch kt al. (1968a).. at

78

P. J. BRENNAN AND D. M. LOSEL

acidic phospholipids as transient ion binders were proposed (Schatzmann, 1962; Ansell and Hawthorne, 1964). I t is possible that the monoglucosyloxyoctadecenoic acid, an acidic glycolipid, has a similar function. A molecular hypothesis of ion-exchange involvement might make use of the existence of lactonized forms within this glycolipid class (Table 4, p. 76). Delactonization at the cell surface, potassium salt formation, transmembrane transport of the complex by means of an energy-linked process, followed by internal lactonization would have the net effect of K+-H+ exchange, intracellularly generated hydrogen ions being neutralized by potassium hydroxide.

2. Acylated Sugars and Sugar Alcohols Haskins et al. (1955) and Boothroyd et al. (1956) partially characterized an oil from the corn-smut fungus, Ustilago maydis, as an acetylated disaccharide, containing both erythritol and mannose. However, when Fluharty and O'Brien (1969) re-examined U. maydis they could not detect extracellular glycolipid production. They found instead that the glycolipid was cell-associated, whence they isolated the material and established the complete structure (Table 5). Since the extracellular product, when observed, is apparently structurally identical with the intracellular glycolipid, it appears that minor differences in growth and culture conditions, or strain selection, can affect the location of these glycolipids. Similar considerations may apply to the yeast sophorosides, or alternatively their secretion may be due tolack of control on their synthesis ; intracellular sophoroside is detectable in C. bogoriensis several days before glycolipid excretion commences (Esders and Light, 1972b). I t is not known if these glycolipids, when cellassociated, are membrane-bound or part of the numerous oil droplets which can be readily observed in such fungi. Glycosyldiglycerides are readily identifiable as lipid components of membranes; they are prominent in Gram-positive bacteria, chloroplasts and some animal tissues (Sastry, 1974). Until recently, it was not clear if glycosyldiglycerides are present in fungi. The report by Baraud et al. ( 19701, which claimed discovery of a monogalactosyldiglyceride in yeast, suffered from inadequate characterization of the putative glycolipid. The evidence that mono- and diglycosyldiglycerides are major components (16%of total lipids) in the zoospores of the water mould, Blastocladiella emersonii, is more impressive, although identification was

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

79

based largely on comparative thin-layer chromatography (Mills and Cantino, 1974). However, an extensive survey of the major glycolipids of baker's yeast (Tyorinoja et al., 1974) seemed to conclude that the majority of alkali-labile compounds with t.1.c. properties indicative of glycosyldiglycerides were in fact acylsugars (Table 5). P. J . Brennan (unpublished results) isolated minute quantities ( 8 mg from 4 g of extractable lipid) of an alkali-labile compound from A . niger which he identified as a mixture of diglucosyl- and digalactosyldiglycerides (Table 4, p. 77). While this was apparently the first report of diglycosyldiglycerides in fungi, in which the presumptive compounds have been adequately characterized, the paucity of the amounts detracts from the significance of the observation.

3. Polyprenol-Containing Glycolipids

Polyprenols, their esters and their phosphates, occur in the membranes of prokaryotes and eukaryotes in extremely small amounts. In prokaryotes, they are usually the C,,-undecaprenol analogues, while in eukaryotes they usually contain from 14 to 20 isoprene units. Much of the polyprenol phosphates are linked via either a phosphate or a pyrophosphate bridge to sugar. These activated sugar derivatives are hydrophobic, and their synthesis is catalysed by enzymes associated with the cell membrane. In bacteria, it is clear that the function of these carrier lipids is to convert water-soluble activated sugars to hydrophobic molecules that can glycosylate macromolecules within the hydrophobic environment of the membrane (Horecker, 1972 ; Lennarz and Scher, 1972). In eukaryotic systems, their function seems to be solely concerned with glycosylation of protein (Lennarz, 1975).In fact, the dolichylphosphates in some eukaryotes, including fungi, are apparently capable of' donating oligosaccharides from 7 to 20 glycose units long for cell-surface glycoprotein synthesis (Behrens et al., 197 1 ; Lucas et al., 1975; Lehle and Tanner, 1975). To date, polyprenols of fungi have been implicated in transfer of mannose or N-acetylglucosamine residues only. In Aspergillus fumigatus, the lipophilic acceptors are a family of hexahydropolyprenol phosphates containing between 18 and 20 isoprene units in each molecule (Stone et al., 1967; Stone and Hemming, 1967). In A . niger, the active polyprenols are a family of exomethylenehexahydroprenol phosphate; other polyprenol phosphates, such as ficaprenol or cetyl alcohol, were inactive (Barr and

OD

0

P

c

TABLE 5 . Free l o n g - c h a i n bases, ceramides, p h o s p h o r y l a t e d and acetylated ceramides of fungi

m

Class of sphingolipid

Composition

Fungal source

Reference

n

rn

Z

Free bases

C,,. and C,,-Phytosphingosine

Free ceramides’

Mostly amides of C I Sor CYO phytosphingosine and 2-hydroxyhexa- o r 2-hydroxyoctacosanoic, or 2,3-dihydroxyfatty acids. Also C,, and C,, sphingosine and dihydrosphingosine with 2-hydroxyfatty acids ranging from C,, to C,,, in addition to some non-hydroxy saturated and unsaturated fatty acids

Candidu intermedia Phycomyces blahesleeanus Saccharomyces cereuisiae Candida utilis Penicillium notatum (Q1751 Aspergillus sydoun Aspergillus niger Fusarium lini Amanita muscuna Amanita rubescens Agaricus bisporus

Free anhydroceramide

and Amides of 1,4-anhydrophytosphingosine its C,, homologue, in equal amounts. Major fatty acid is 2-hydroxyhexacosanoic. Also small amounts of C,,-C,, 2-hydroxyfatty acids

Clitocybe tabescens Hypholoma fasiculure Marasmius scorodonius Saccharomyces cereuisiae

Kimura et al. (1974) Weiss et al. (1973) Weinert etal. (1973) Stanarev and Kates (1963) Oda (1952) Bohonos and Peterson (1943) Wagner and Fiegert ( 1969) Hackett and Brennan (1976) Weiss and Stiller (1972) Zellner (191 1); Weiss and Stiller (1972) Weiss and Stiller (1972) Weiss and Stiller (1972) Hackett and Brennan (1976) Prostenik and Cosovic (19741 Zellner ( 19 I 1 ) Froschl and Zellner (1928) Kishimoto et al. (1974)

z

P P0 P

3 r

0 v)

rn r

9

Ceramide phosphate

Tetra-acetylphytosphingosine

Phosphate groups attached to position I of Saccharomyes cereuiszae C,,-phytosphingosine. The acyl group is principally 2,3-dihydroxyhexacosanoicacid. Also present are all fatty acids in the C22-C27 saturated 2,3-dihydroxy range; the C,,-C,, saturated 2-hydroxyl range; the C,,-C,, saturated and mono-unsaturated range C,,-Phytosphingosine in which all of the Hansenula c$mi hydroxyl groups and the amino group are acetylated : OCOCH, OCOCH, NHCOCH, I I I CH,-(CH,),,-CH -CH CH-CH,OCOCH,

Oda and Kamiya (1958) Hoshi etal. (1973)

Stodola and Wickerham (1960)

-u

5

v,

0

~

Triacetylphytosphingosine

C,,-Dihydrosphingosine in which all of the hydroxyl groups and the amino groups are acetylated : PCOCH, YHCOCH, I

CH,-(CHJ,,-CH

Hansenula ciferri

Stodola el al. (19621 L

z

t,

> i

1 .

NH-CH,OCOCH, -

1 71 0

“CIS-Sphingosine, sphing-4-enine, ~-erythrol,3-dihydroxy-2-amino-trans-4-octadecene, CH,-(CH,),,-CH=CH-CH-CH-CH,OH; C,,-Dihydro-

I

I

cn

rn

O H NH, r sphingosine, sphinganine, o-erythm-1,3-dih~droxy-2-amino-octadecane; C,,-Phytosphingosine, 4-~-hydroxysphinganine,o-ribo-1,3,4-trihydroxy-Z- rn 0 amino-octadecane. u b Ceramide, cerebrin, cer, a N-acyl derivative of a sphingosine-type base.

4 +

z

0

cn

82

P. J. BRENNAN AND D. M. LOSEL

Hemming, 1972). In Sacch. cerevisiae, the polyprenolmannose acceptor consists of a family of dolichols with from 14 to 18 isoprene units (Jung and Tanner, 19 7 31. Letoublon et al. ( 1973) characterized the particulate mannosyl transferase enzyme from A . niger responsible for transfer of mannose from GDP-mannose to the polyprenol phosphate. The ensuing polyprenol-phosphate mannose is apparently involved in the biosynthesis of aspergillus mannan (Barr and Hemming, 1972); that these mannan units are attached to protein can be inferred from the results of Letoublon and Got (1974). Similar results were obtained with the yeasts, Hansenula holstii and Sacch. cereuisiae (Bretthauer et al., 1973; Babczinski and Tanner, 1973). In addition, it was shown that the products of mannose transfer from polyprenol phosphate were mostly glycopeptides with mannose linked to serine or threonine (Bretthauer and Tray, 1974; Bretthauer and Wu, 1975; Sharma et al., 19741, although there was evidence for linkage to other amino-acids. It appears that these mannosyl- 0-serine (threonine) linkages are part of the cell-wall mannan-protein complex of yeasts. Sharma et al. (1974) and Lehle and Tanner (1974) made the important distinction that the dolicholmonophosphate is involved only in transfer to an appropriate amino acid of the yeast mannan-protein. N o lipid intermediate takes part in mannosyl-transfer reactions to mannosyl groups, in which case GDP-mannose is used directly. Therefore, the sequence depicted in Fig. 2 A probably applies for biosynthesis of yeast and aspergillus mannan-protein. Moreover, from several lines of evidence reported by Gold and Hahn ( 1976), it appears that a mannosylphosphorylpolyisoprenol is an obligatory intermediate in the in uiuo mannosylation of particulate protein in N. crassa. Recently, Lehle and Tanner (1975) reported that incubation of a membrane fraction from Sacch. cerevisiae with UDP[14C]-N-acetylglucosamine catalysed transfer of N-acetylglucosamine to endogenous lipid as well as to a methanol-insoluble polymer. The lipid fraction was subdivided into three components by thin-layer chromatography. Two were identified as dolicho1pyrophosphate-Nacetylglucosamine and dolicholpyrophosphate-di-N-acetylchitobiose. Radioactivity was also transferred to a lipid containing two mannose residues and a di-N-acetylchitobiose (i.e. a tetrasaccharide). In view of evidence (Lennarz, 1975) for pre-assembly of oligosaccharide chains of certain animal glycoproteins on a polyprenol carrier prior to their transfer to the nascent polypeptide, it seems that

nGD P- Man

A # Mannosyl- 0 serine-peptide

Polypeptide with free hydroxyl group of L-serine (threonine)

N-acetylglucosaminylasparaginyl-peptide

Polypeptide with free amino group of L-asparagine

NH,

+

dolicholypyrophosphate

I

(GlcNAc),-(Man), Polypeptide with free amino group of L-asparagine

-c

Mannan-glycoprotein of cell walls

Mannan-glycoprotein of cell walls

nGDP-Man N H -( GlcN Ac)2 - Man2

di-mannosyl-di-N-acrtylchitobiosylasparaginyl peptide

4

c

NH -( GlcNAc),(Man)n+2

Mannan glycoprotein of cell walls

u !?

8

r rn

9 mi

0

g

n

FIG. 2. Postulated biosynthesis of yeast mannan demonstrating the involvement of polyprenol and nucleotide sugars Yeast mannan is a covalently linked polysaccharide-protein complex. Some of the mannose is attached to the polypeptide chain as short oligosaccharides, glycosidically linked to serine and threonine (Ballou, 1974). Biosynthesis of these segments is represented in A. However, the majority of mannose is attached as polysaccharide chains with perhaps 150 o r more mannose units linked via N-acetylo-glucosamine to asparagine (Sentandreu and Northcote, 1968). Possible routes for biosynthesis of these segments are demonstrated in B and C. Ballou (1974)pointed out that little is known about this type of linkage because the attachment of mannose to glucosamine and the number of glucosamine units at the linkage point are uncertain.

i/j

84

P.-J.

BRENNAN AND D. M. LOSEL

Tanner’s results point to a similar phenomenon existing in eukaryotic micro - organisms. Ballou ( 19741, in discussing yeast mannans and their biosynthesis, envisaged a mechanism in which protein or short polypeptides are assembled; manno-oligosaccharides are then built on the serine and threonine units, and the longer polysaccharide chains are formed by addition first of N-acetylglucosamine to asparagine followed by stepwise addition of mannose units. From Tanner’s results, it appears that part of the longer chains are preformed on the carrier polyprenol before donation to the asparagine residue. Since it is not yet clear how many glucosamine units are at the linkage point, the relative importance of the two procedures for glycosylation is not known. Schemes by which biosynthetic and structural observations on yeast mannan can be correlated are summarized in Fig. 2. Letoublon and Got (1974) suggested that polyprenolphosphomannose is the form in which active mannose is transported across the plasma membrane for cell-wall biosynthesis. A difficulty in the Roseman (1974) hypothesis of cellular adhesion, extended to slime moulds in Section E (p. 98), is to explain how the sugar nucleotides can pass through the permeability barrier of the cell membrane into the extracellular area. However, if the active sugar is lipid-linked, then it should readily diffuse through the plasma membrane. E.

SPHINGOLIPIDS

Long-chain sphingosine-type bases are found in fungal extracts in the form of glycophosphosphingolipids, phosphosphingolipids, glycosylceramides, ceramides, acylated long-chain bases or sometimes in the free form. A small amount of an anhydrocerebrin has been obtained from baker’s yeast without the use of hydrolytic procedures (Table 5, p. 80). Whether it occurs in nature or is an artifact of isolation is not known. Previously we (Brennan et al., 1975) suggested that the bulk of fungal sphingolipids will prove to be glycosphingolipids of the type found in higher plants or animals. In animals tissues, the sugar residues are in direct glycosidic conjugation with the primary hydroxyl group of the N-acetylated sphingosine-type base (ceramide). In higher plants, the direct glycosidic bond is seen only in the simple monoglycosylceramides (cerebrosides).The remainder of plant glycosphingolipids have a phospho-inositol bridging the ceramide and

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

85

glycosyl moieties. Both types of glycosphingolipids have recently been found in fungi and have often been given the generic name of mycosphingolipids. 1. Free Long- Chain Bases and Ceramides

There have been several isolated reports of the presence of free bases in fungi. For instance, in Hansenula cferri, about 3% of’ the phytosphingosine occurs in the free form; the remainder is present as the tetra-acetylphytosphingosine. Probably the best authenticated report is that from Kimura et al. (1974; Table 5 , p. 80). They isolated substantial quantities (60 mg from 120 g of dried cells; 2-3% of total lipid) of a ninhydrin-positive substance from Candida intermedia grown on glucose. The compound corresponded to authentic C,,-phytosphingosine, particularly with regard to the infrared and nuclear magnetic resonance spectra, and the production of serine after periodate oxidation and hydrolysis. However, gas-liquid chromatography-mass spectroscopy of this material showed that 10%of it was the C,,-phytosphingosine. I t is noteworthy that this yeast was devoid of free ceramides of tetra-acetylphytosphingosine. Hence it appears that the organism may be deficient in the enzymes responsible for the acetylation of phytosphingosine. Until recently, it was considered that the major class of fungal sphingolipids were free ceramides (cerebrins). Table 5 (p. 80) list the fungi from which these have been isolated, and their probable structures. In some of these cases, the ceramides comprise a very large proportion of the total cell weight; in several of the moulds, mushrooms, phycomycetes and Fungi Imperfecti, they account for 0.2-0.3% of the dry tissue. In Amanita muscaria, ceramides may amount to about 3% of the dry sporophore. On the question of the distribution, total levels, and structures of fungal ceramides, the reports of Weiss and colleagues (1972, 1973) are the most comprehensive. The unprecedented abundance of ceramides in fungal extracts led to the suggestion (Brennan et al., 1975) that they might result from alkaline degradation of complex glycosphingolipids; most of the authors listed in Table 5 (p. 80) used an alkali-stable lipid fraction as a source of ceramides. I t was conceivable that treatment of lipid extracts with alkali cleaved the phosphodiester bridge of glycophosphosphingolipids resulting in the release of some free ceramides. In fact, Steiner et al. (1969) demonstrated degradation of mannosyldi-inositoldiphosphorylceramide(CerP,I,M)

TABLE 6. Complex sphingolipids of fungi Probable structure

Trivial name and abbreviation

Composition of ceramide

Source

Reference

0

II

Cer-( l+j-O-P-O-inositol

Inositolphosphorylceramide (CerPII

I

0-

Phytosphingosine and hydmxy Cz6: tatty acid

Saccharomyces cerevisiae

Smith and Lester (19741

P

0

II

L

Cer-( 1-4-O-P-O-inositol

Inositolphosphorylceramide ( CerPl]

I 0-

Phytosphingosine and C z 6 : tatty ac1d

Saccharomyces cerevisiae

Smith and Lester (19741

2

II

Inositolphosphorylceramide ( Cer PI I

Cer-( l-W-P-O-inositol

I 0-

Phytosphingosine and Saccharomyces dihydroxy Cz6: fatty acid cerevisiae

Smith and Lester (19741

.

Cer-( I+I-O-P-O-inositol

.

I

Inositolphosphorylcerarnide (CerPI)'

Not determined

Neuro$ora crassa Lester e l al. (19741

Inositolphosphorylceramide ( Cer PI Ib

Not determined

Aspergillus n i p

Di-inositoldiphosphorylteramide (CerP~Ipl

Phytosphingosine and hydroxy Cz*: tatty acid

Neurospora crassa Lester et al. ( 19741

00

II

Cer-( l+l-O-P-O-inositol

I

0-

II

Cer-0-P-0

I

Hackett and Brennan(l977)

0

II

-inos-0-P-0

O-Na+

I

> 2 0

P T

0

0

rn 2 2

P

0

I1

33

-inos

0-Na'

0

0

0-

0-

II II Cer-0-P-0-inos-(mann)-O-P-O-inos I 1

Mannosyldi-inositoldiphosphoryl- Not determined' ceramide (CerP,I,M)

Saccharomyces cereuisiae

Steiner et al. ( 1969); Steiner and Lester (1972)

Mannosylinositolphosphorylceramide (CerPIM)

C,*- and Cz0Phytosphingosine. Mixture of 2-hydroxyand non-hydroxysaturated and unsaturated fatty acids

Saccharomy ces cereuisiae

Wagner and andZofcsik (1966a. b)

Mannosylinositolphosphorylceramide (CerPIM)

C I8 -Dihydrosphingosine, 2-hydroxy- and nonhydroxy C,, fatty acids

Saccharomyces cereuisiae

Mannosylinositolphosphoryceramide (CerPIM)

C 1 8-Sphingosine, 2-hydroxy C,,-C,, fatty acids

Agaricus bisporus

0

II

Cer-0-P-0-inos-mann

I

0

W

< 0,

and Candida utilis

0

II

Cer-0-P-0-inos-mann

I

0Cer- 1'-phosphory- l-inos-(Ztl)-a-O-mann

GalactosylmannosylinositolNot determined phosphorylceramide (CerPIMGal)

Aspergtllus niger

P. J. Brennan (unpublished results)

Brennan and Roe (19751 Roe (1976)

v)

-I

? 0 v)

0

II

Cer-0-P-0 -inos- (rnann-gaI-glcId

1

0-

Glucosylgalactosylmannosylinositolphosphorylceramide (CerPIMGalClc)

Not determined

Aspergillus niger

Roe (1976)

TABLE 6.-cont. Probable structure

Trivial name and abbreviation

03 03

Composition of ceramide

Source

Rehence

0

I

Cer-0-P-0-inos-(mann,-gal,)

Trigalactosyldimannosylinositolphosphorylceramide (CerPIM,Gal,)

Not determined

Aspergillus nzger

Cer-(l’)phosphoryl-(l)inos(6cl)a-o-GlcUA Fucosyltrigalactosylglucuronosyl2 inositolphosphorylceramide (CerPI(GlcUA)Gal,Fuc)

Not determined

Agaricus bisporus

Large variety ofphytosphingosines. Mostly 2-hydroxy C , 6 : ,fatty acid Mostly C,,-sphingosine and C , , tatty acid Mostly C,, phytosphingosine and 2-hydroxy C,, fatty acid Mostly phytosp hingosines. In Amanita muscaria the major bases are o f n 22 : O and i 21 : O types. In Amanita rubescens they are mostly i 19 : 0 and i 20 :O. In Agaricus bisporus the principal bases are of the n 18 :0 and i 2 1 : 0 variety. In all sources 2-hydroxy fatty acids constitute 40-60% of the total fatty acids

Phycomyces blakesleeanus

Weiss et al. (1973)

Hansenula cferri Fusarium lint

Kaufmann et al. (1971) Weiss et al. (1973)

I 0-

t

Byme and Brcnnan (1976) (unpublished results) P. J. Brennan and J. A. Hackett, (unpublished

1

a - o - G a l ( 2 cl)a-D-GaUZ+ I ) a - o - G a l ( 2 c l)a-L-Fucc C e r-(l’tl)gl u c o s e

Glucosylceramide (Cer-Glc)

Cer4 1 ’+ 1)glucose

Glucosylceramide (Cer-Glc)

Cer-(l’+-l)glucose

Glucosylceramide (Cer-Glc)

...

,

Cer-( 1’+- 1)glucose

Glucosylceramide (Cer-Glc)

z Z

B

z B

z

P

5 r

Amanita muscaria Weiss and Stiller Amanita rubescens (1972) Agaricus bisporusf Prostenik and Clttucybe /abuscuns@ C o s ~ \ . l C( 1974)

rn

r

Cer-( 1‘C1)-galactose

Galactosylceramide (Cer-Gal)

Cer-(glucose-galactose-galactosegalactose)

Monoglucosyltrigalactosyl ceramide (Cer-(glc-gal,))

Mostly C,,-, C,,-dihydro- Saccharomyces cereuisiae sphingosine, C l x sphingosine, and 2Candida utilis hydioxy Ci6: tdtty acid Cis-sphingosine. C,,-di- Aspergillus nigerf hydrosphingosine and 2hvdroxv-Clu I fattv .” . , acid. Phytos;hiiigosine and 2- Neurospora crassa hydroxy C,,: fatty acid

Wagner and Zofcsik ( 1 966a) Wagner and Zofcsik (1966hi Wagner and Fiegert, (1969) -u

Lester etal. (1974)

5

“Neurospora c~assaseems to contain three CerPIs. bAspergillus niger seems to contain two CerPIs. ‘Tyorinoja et al. (1974)seem to have also isolated this sphingolipid from Saccharomyces cereviszae. If so, the major long-chain base is C,,-phytosphingosine and, from a previous publication (Nurminen and Suomalainen, 197 11, the principal fatty acids are 2-hydroxy-C,,: and non-hydroxy C,,: o. *Tentative structures. ‘This is a generalized structure. Two spingolipid preparations were examined, each containing at least two glycophosphosphingolipids. These differed in the presence or absence of o-glucuronic acid and L-fucose. ’P. J. Brennan and J. Madden (unpublished results) also found g1ucos;lceramide in Agaricus bisporus and Aspergillus niger. N o galactosylceramide was found in either sperics. B O f the two glucocei-ebi-osides obtained from Clitocybe tabescens, one is a glucosylceramide containing some 2-hydroxystearic acid, but with a predominance 01 heptadecanoic and decanoic acids. The other cerebrosides is a N-acilphytosphingosylglucosidecontaining mostly 2-hydroxystearic acid. There is tenuous evidence for a P-glycosidic link in these cerebrosides.

0 c)

<

% -n

C

z

G)

r U

9 v)

rn r rn

! U

s -I

0

rJY

m (9

90

P. J. BRENNAN AND D. M. LOSEL

to mannosylinositolphosphorylceramide (CerPIM)under alkaline conditions which were mild compared with those used in much of the earlier work. However, while extensive treatment of fungal lipids with alkali probably increases the free ceramide content, this alone does not account for their prevalance; lipid extracts of Aspergillus niger and Agaricus bisporus obtained with neutral solvents contain substantial amounts of free ceramides with non-hydroxylated, monohydroxylated and dihydroxylated fatty acids (Hackett and Brennan, 1976). I t therefore seems that many of these free ceramides are products of' fungal autolysis. This view was also expressed by Hoshi et al. (1973) who quoted as favourable evidence the inability of Oda and Kamiya (1958) to isolate free ceramides from intact baker's yeast. However, Oda and Kamiya (1958)did isolate a free ceramide phosphate (Table 5, p. 81) in fairly substantial quantities (0.024%of fresh yeast), and the yield of this increased after brief autolysis of the yeast. Presumably free ceramides arise after further autolysis. Since the probable source of these free ceramides has emerged only in recent times, nothing is known of the enzymology of their catabolism. However, it is clear that they are the end products of the autodegradation of complex membranous glycophosphosphingolipids. 2 Simple Glycosylceramides ( Cerebrosides) There are reports of substantial quantities of glucosyl- and galactosylceramides in moulds, yeasts and basidiomycetes (Table 6). Some of the very early investigators attempted to isolate cerebrosides from species such Lycoperdon bovista (Landsiedel and Bamberger, 1905), Amanita muscaria (Zellner, 19 11 ; Rosenthal, 19221, Polyporus pinocola (Hartmann and Zellner, 1928) and Marasmius scorodonius (Froschl and Zellner, 1928). The products were inipure and their characterizations incomplete by present-day standards, and it is not certain that sphingolipids were in fact being examined. The first substantial record of' the existence of a simple glycosylceramide in fungi came from Wagner and his associates. They reported the presence of galactosylceramides in Sacch. cerevisiae, C.utilis (Wagner and Zofcsik, 1966a,b) and A . niger (Wagner and Fiegert, 1969; Table 6). The glycolipid from A . niger yielded both C,, spingosine and C,, dihydrosphingosine, and over 80% of the fatty acid of the ceramide moiety was 2-hydroxyoctadecenoic acid. The galactosylceramide from C. utilis had C,, and C,, dihydrosphingosine, CIS sphingosine and

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

91

principally 2-hydroxystearic acid. In the strain of A . niger used by Wagner, the galactosylceramide was apparently a major lipid since it was obtained in the crystalline form following t.1.c. of an entire chloroform - methano 1 extract. Kaufmann et al. ( 197 1) described in detail a monoglucosylceramide from Hansenula czferri. Apparently this organism did not contain any of the galactosylceramide present in Sacch. cereuisiae and C. utilis. Again, phytosphingosine was not a major constituent; the long-chain bases corresponded to unsaturated C,, and C,, types. The fatty acids (Table 6 ) were also unusual in that 2-hydroxy acids were relatively minor. In view of these reports, it is surprising that Tyorinoja et al. (1974) in an extensive survey of the major glycolipid of baker’s yeast did not report any glycosylceramides. Weiss and Stiller (1972), in particular, and Prostenik and Cosovic ( 1974) have examined in detail the glucosylceramides of a number of different agaric species (Table 6 ) . A point about the work of Weiss and Stiller (1972) is that it gives the impression that phytosphingosines are the sole long-chain bases in agaric cerebrosides. As one would expect, this apparently is not the case. Prostenik and Cosovic ( 1974) isolated and partially characterized the cerebrosides from Clitocybe tabescens Scop, and showed the presence of a variety of long-chain bases. I t is obvious from their work and that of Weiss and Stiller (1972) that the cerebrosides in all of the basidiomycetes which were examined are glucosylceramides. 3 . Polyglycosylcerarnides

The question has often been raised as to whether fungi and higher plants contain polyglycosylceramides similar to the globosides, cytolipin, ganglioside or blood-group glycosphingolipids of animal cells. Lester and his colleagues have now partially answered this question. They (Lester et al., 1974) have provided convincing evidence for a galactosyl-galactosy1-galactosyl-glucosylceramidein Neurosporu crussu. While examining this organism for glycophosphosphingolipids, they prepared a “sphingolipid concentrate” by extracting selective lipids with ethanol-diethyl ether-pyridine and precipitating sphingolipids with acetic acid. Column chromatography on porous silica-gel beads, pretreated with sodium hydroxide, showed a major glycolipid which was devoid of phosphorus. Methanolysis followed by gas-liquid chromatography of the trimethylsilyl ethers showed only galactose

92

P. J. BRENNAN AND D. M. LOSEL

(1.93 pmoles per mg of glycolipid) and glucose (0.65 pmole per mg). Equivalent amounts of fatty acid and long-chain base were found, and these appeared to be largely 2-hydroxytetracosanoic acid and C phytosphingosine. The glycolipid was stable to mild alkaline methanolysis indicating that the fatty acid was amide linked. This and other evidence suggested the structure given in Table 6. Lester et al. (1974) reported evidence for this glycolipid in other strains of Neurospora. However, from other work, it appears that similar glycolipids are not prominent in other moulds, yeasts or mushrooms. 4. Phosphosphingolipids and Glycophosphos~hingolipids As mentioned previously, first clues to the existence of complex sphingolipids in fungi came from the high amounts of free ceramide observed in lipid extracts; it was assumed that these were degradation products of more complex types. The extensive work of Carter and his associates (Carter et al., 1969) showing the presence of a unique array of glycophosphosphingolipids (phytosphingolipidsi in higher plants suggested that related substances may exist in fungi. Wagner and Zofcsik ( 1966a, b) were probably the first seriously to examine a fungus for such substances. They isolated a sphingolipid containing phosphorus, myo-inositol, and D-mannose from Sacch. cereuisiae and C . utilis, and suggested the structure, mannosylmono-inositolmonophosphorylceramide (CerPIM; Table 6). It seemed, at one stage, that CerPIM was solely an alkaline degradation product of a more complex sphingolipid obtained from Sacch. cereuisiae by Lester's group (Steiner et al., 1969). Analytical and degradation studies on this compound suggested a mannosyldiinositoldiphosphorylceramide (CerP,I,M) structure (Table 6). This is evidently a major constituent of the yeast strain employed by Lester since it represented 27.9% of the lipid-soluble inositol and 8-9% of the lipid-phosphorus. Of particular note was the susceptibility of' this compound to mild alkaline hydrolysis, the products of which had the properties of CerPIM and phospho-inositol. Since Wagner and Zofcsik (1966a) had isolated CerPIM after treatment of the crude lipid with aqueous alkali for 24 h, it appeared likely that CerPIM was solely a cleavage product of CerP,I,M. However, this now appears unlikely for a number of reasons. In a subsequent examination of a sphingolipid concentrate which had not been exposed to alkali, Smith and Lester ( 1974) isolated and characterized significant quantities of CerPIM.

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

93

While the natural occurrence of CerPIM in Sacch. cerevisiae was thus clearly shown, the levels were low. It constituted only 1.95% of lipidinositol and 0.7%of lipid phosphorus. Further evidence for the natural occurrence of CerPIM comes from unpublished work of P. J. Brennan and his colleagues. A CerPIM was isolated from Agaricus bisporus and its complete structure established (Table 6). It represents 5 0 4 0 % of the total sphingolipid mixture, and it is a major phospholipid of theagaric. Although alkali was used in its preparation, the mildness of the condition precluded degradation of a phosphodiester linkage. Other evidence for the presence of CerPIM and CerP,I,M in yeast comes from interpreting results reported by Tanner (1968, 1969) in a manner different from his own explanation. He found that ['4Clmyoinositol was incorporated by Sacch. cerevisiae into three water-soluble components (called X, Y and Z). Component X was identified as O-aD-mannopyranosylmyoinositol(IM),and Component Z was tentatively identified as p ho sphoryl- 0 - a - D - mannopyranosyl my0 inositol (PIM ) . Component Y was a mixture of phosphatidylinositol and natural glycerylphosphorylinositol (GPI). In addition, two lipids, lipids I and 11, were labelled. Kinetic studies indicated that the water-soluble IM, PI, PIM, and GPI, and lipid I arose as natural degradation products of lipid 11. In particular, Tanner (1968) showed that, when cells pulsed with ['4Clinositol were chased with non-radioactive inositol, all components, except lipid 11 increased in radioactivity and 87% of this increase could be accounted for by the decrease in lipid I1 (63%)and in free inositol (24%). Over 91% of lipid I1 was phosphatidylinositol. However, the remaining 8.4% could not be de-acylated and, on dephosphorylation by ammonolysis, it produced inositol and inositolmannose, evidence which may indicate that the compound was CerP,I,M, i.e. 0

0

II

Cer- 0-P-0

I

0-

Ceramide

I1

-inosi tol(mannose)-0-P-0

I

I 0-

-inositol

IO%NH,OH; 16OoC; 18 h

+ 2Pi + inositol-mannose + inositol

An examination of lipid I (Tanner, 1968) indicated that a minority (40%) of i t was lysophosphatidylinositol. The majority on dephos-

94

P. J. BRENNAN AND D. M. LOSEL

phorylation yielded inositol-mannose and inositol. I t is known that, under these dephosphorylation conditions, some cleavage of glycosidic groups takes place (Khuller and Brennan, 1972). It is possible, therefore, that the bulk of lipid I is CerPIM, in which case CerPIM would be a natural degradation product of CerP,I,M. Such an explanation seems to provide the simplest correlation of Tanner’s results with the evidence from Lester that CerP,I,M is the predominant sphingolipid in yeast. I t is more difficult to explain Tanner’s results if it is assumed that lipid I1 contains CerPIM. Angus and Lester ( 1972) conducted essentially similar experiments to those of Tanner, but the products were homogenous and well characterized. They appeared to presume that Tanner’s lipid I was CerP,I,M, but received its label from phosphatidylinositol during the chase. In addition, Tanner’s experiments, by providing an explanation for the origin of natural IM and PIM, also demonstrate the origin of some of the free ceramides of Sacch. cereuisiae. Incidentally, natural IM was recently isolated from Sacch. cereuisiae and its structure established as 6-O-a-~-mannopyranosylmyoinositol (Wells et a/., 1974). If this is a catabolite of CerPIM or CerP,I,M from Sacch. cereuisiae, it implies that these have the mannose unit glycosidically attached at C-6 of the inositol ring unlike CerPIM from A . btsporus which employs the hydroxyl at C-2 (Table 6). Further evidence for CerPIM/CerP,I,M in fungi was provided by Nurminen and Suomalainen (197 1). They obtained a sphingolipid from preparations of cell envelopes isolated from a commercial strain of Sacch. cereuisiae. Quantitative analysis of the sphingolipid showed that it contained 1.14 moles of mannose per mole of phosphorus. Therefore, it appeared to be CerPIM. However, Smith and Lester (19741, commenting on this work, observed that the sphingolipid was prepared according to Steiner et al. (1969) and that this procedure could yield a fraction containing both CerPIM and CerP,I,M. In any case they (Tyorinoja et al., 1974) later claimed that their sphingolipid was CerP,I,M based on the evidence that its chromatographic mobility was similar to that reported by Steiner and Lester (1972). Trevelyan ( 1968) also provided evidence for inositol-mannose-containing sphingolipids in Sacch. cereuisiae. A biosynthetic precursor of CerPIM and CerP,I,M should be inositolphosphorylceramide (CerPI). Smith and Lester (1974) isolated and characterized three members of this type of sphingolipid from

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

95

Sacch. cereuisiae (Table 6). Lester et al. (1974) recognized at least three members in Neurospora crassa, and Hackett and Brennan (1977)recognized at least one in A . niger (Table 6 ) . Yet another major non-glycosylated phosphosphingolipid was isolated by Lester et al. (1974) from N . crassa. This was shown to have the general structure: ceramide-(P-inositol),(CerP,I,) (Table 6) and i t was obviously the major sphingolipid in this fungus, representing 4060% of lipid-soluble inositol, its level exceeding that of phosphatidylinositol. This sphingolipid does not seem to be a biosynthetic precursor of CerP,I,M since the mannose of the latter appears to be internal in a linear chain (cf. Brennan et al., 1975). A pulse-chase experiment with [H31-inositol(Lester et al., 1974) suggests that CerP,I, may be an anabolic end product, similar to CerP,I,M. The possibility that fungi may contain glycophosphosphingolipids of the type associated with higher plants with a substantial oligosaccharide segment was investigated by Brennan and his colleagues (Byrne and Brennan, 197 6 ; Roe, 1976 ; and unpublished results). Agaricus bisporus contains at least four distinct glycophosphosphingolipids. These were partially purified by “dry” silica-gel H column chromatography. The structures shown in Table 6 were suggested by gas-liquid chromatographic analysis of the products of mild acid hydrolysis, enzymic degradation, and periodate oxidation. These workers also examined the glycophosphosphingolipids of A. niger. This organism contains at least eight distinct glycosphingolipids, some of which were tentatively characterized (Table 6). Prostenik’s group (Cosovic et al., 1974) described an unusual glycophosphosphingolipid from the agaric Clitocybe tabescens Scop. The fungus was extracted by the Folch procedure. From the upper aqueous phase, a water-soluble nitrogen- and phosphorus-containing fraction was obtained which chromatographed similarly to the phytosphingolipid from peanuts. Acid hydrolysis of the sphingolipid yielded mannose, glucuronic acid, glucosamine and at least two unidentified compounds. However, the authors did not detect inositol. An issue was made of the absence of inositol since it could not mediate the attachment of the ceramide-phosphate to the oligosaccharide moiety. However, it seems more likely that the hydrolysis conditions were not sufficiently strong to hydrolyse the phosphodiester link and release inositol. If this is the case, the glycolipid corresponds closely to some of those from higher plants (Carter et al., 1969; Kaul and Lester, 1975).

96

P. J. BRENNAN AND D. M. LOSEL

5 . Functional Features of Sphingolipids

The combined use of inositol starvation and inositol-requiring mutants of N . c r a s ~ ayielded interesting results on the molecular role of phosphatidylinositol and the inositol-containing sphingolipids. It had been proposed that autolysis and death resulting from the withholding of inositol from such mutants was due to release of proteases which, under normal circumstances, are encased in a lipo-inositol-rich lysosome membrane (Matile, 1966). With the absence of exogenous inositol this membrane could rupture and release the proteases. However Lester et al. (1974) have noted that, when growth of an inositolrequiring mutant of N . craJsa is limited by inositol starvation, there is a continual breakdown of phosphatidylinositol and synthesis of Cer(PI),. Hence, the lethal effects of inositol deprivation on inositolless mutants may be due either to decreased levels of membranous phosphatidylinositol or abnormally high amounts of the inositolcontaining sphingolipids. Therefore, in some inositol-requiring mutants, the effect of the abberation may be akin to that seen in diseases known as sphingolipidoses. The cellular location of fungal sphingolipids is not known. Nurminen and Suomalainen ( 197 1) obtained CerP,I,M, and the longchain 2-hydroxyfatty acids associated with sphingolipids, from wholecell envelopes of S u c h . cerevisiae, and concluded that CerP,I,M was present in both the cell wall and plasma membrane. In addition, free ceramides, which are probably autolysis products of glycophosphosphingolipids, were present in the cell-envelope fraction (Tyorinoja et al., 1974). However, the cellular fractions were such that the possibility of contamination of cell wall with membrane material could not be excluded. Since glycophosphosphingolipids have been found only in eukaryotes bearing a cell wall, it has been suggested that they may play a role in the production and function of the cell-wall material. Furthermore, the effects of inositol deprivation (Ghosh et al., 1960; Power and Challinor, 1969) and inositol antagonists (Deshusses et al., 1969; Deshusses, 1974) on inositol-requiring yeasts suggest a role for inositol in metabolism of cell-wall polysaccharides. Since practically all of the yeast’s cellular inositol is in the form of various phosphoinositides, these seem to be implicated in cell-wall formation. Further use of specific fungal growth inhibitors, such as deoxyglucose and isomytilitol, antagonists for glucose/mannose and myoinositol respec-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

97

tively, should be particularly informative. Both of these are incorporated into fungal phospho-inositides at a fairly normal rate (Steiner and Lester, 1972; Deshusses, 19741, so they may not yield much biosynthetic information. However, the abnormal lipids s o synthesized are apparently not able to participate in normal functioning of the cell, probably due to abnormal cell-wall formation as evidenced by bizarre growth. Hence, these antagonists should have important uses in determining the physiological role of the phospho-inositides. Several Gram-positive bacteria contain a variety of phosphoglycolipids (Shaw, 1975) not unlike the simpler of the glycophosphosphingolipids in physicochemical properties. They both usually have Iong lipophilic hydrocarbon residues, phosphodiester group: and hydrophilic sugar residues of varying numbers. Also, throughout Gram-positive species of bacteria are found lipoteichoic acids which are polymers of glycerol phosphate in which phosphodiester bonds link the 1 and 3 positions of adjacent glycerol moieties (Baddiley, 1972). The polymer, consisting of up to 35 residues of glycerophosphate, contains a variable number of disaccharide and alanine residues bonded to the secondary alcohol functions of the glycerol phosphate units. In several Gram-positive bacteria, the disaccharide is similar to that which occurs in the phosphoglycolipids and glycolipids of the same organism. Wicken and Knox (1970) found-that some of the glycolipids were attached to membrane teichoic acid by a phosphodiester bridge (lipoteichoic acids). Those lipoteichoic acids are situated on the outside of the cytoplasmic membrane and are thought to occur in all Gram-positive bacteria. The glycerolphosphate polymer of lipoteichoic acid is visualized as extending from the membrane into the cell wall. Considering the highly hydrophilic nature of the long gly.cero1 phosphate polymer and the hydrophobic environment of the cell membrane, which contains all of the lipid of Gram-positive bacteria, it is reasonable to assume that the polymer is anchored to the membrane by glycolipid. Pieringer and Ganfield ( 1975) constructed spacefilling models of phosphatidylkojibiosyl diglyceride (which contains four long-chain fatty acids instead of the usual two) and the associated polyglycerolphosphate polymer. They concluded that the non-polar end offered a hydrophobic region capable of interacting with, and being imbedded in, the non-polar region of the cell membrane. Its polar “head” is strongly hydrophilic due to the anionic phosphate groups and the hydroxyl groups of the glucose residues. In addition to

98

P. J. BRENNAN AND D. M. LOSEL

interacting with the more polar molecules above the surface of the membrane, the hydroxyl groups provide a chemically reactive site for coupling the long glycerol phosphate polymer to the lipid anchor. Such a cell-surface anchoring role may apply to the glycophosphosphingolipids of fungi. Not enough is yet known about fungal cell walls to suggest an equivalent for lipoteichoic acids. Powell et al. (1974) have shown that micrococci are apparently the only Grampositive bacteria without membrane lipoteichoic acid, but instead possess membranous mannan with properties analogous to the lipoteichoic acids of other Gram-positive bacteria. Yeasts, aspergilli and other fungi contain mannan or related cell-surface polysaccharide. If the analogy between mannan and lipoteichoic acids can be extended to fungi, then perhaps the glycophosphosphingolipids may have functions similar to the phosphoglycolipids. Indeed, there is convincing evidence that mannan is a genuine component of yeast plasma membrane (Nurminen et al., 1976). The nature of some of the sugar residue in fungal glycophosphosphingolipids suggests that they may also be involved in cellular recognition and association. For instance, glucuronic acid is directly involved in mediating re-aggregation of Microciona prolfera (Turner and Burger, 1973); mannose is the major constituent of the sexual agglutination factor released by Hansenula wingei (Yen and Ballou, 1973); mannose and other sugars are also apparently implicated in the agglutination of gametes from Chlamydomonas (McLean and Bosmann, 1975), Glycophosphosphingolipids have been found in Mucor rouxii and Saprolegniaferax (P. J. Brennan, unpublished results). There is as yet no evidence as to whether glycosphingolipids are involved in fusion of gametangia or sexual organs in these fungi, or in the cellular slime moulds. There is, however, considerable speculation and some evidence that glycosphingolipids are involved in the cellular aggregation process in Dictyostelium discoideum. This cellular slime mould provides an excellent system for studying aspects of eukaryotic differentiation, particularly cellular aggregation. In the presence of a food supply, D . discoideum multiplies and remains as individual cells. Upon starvation, the homogeneous population of individual cells becomes mutually adhesive, aggregate to form multicellular organisms comparaable to an animal tissue, transform into at least two distinctly different cell types, and mature as viable spore cells or supporting vacuolated stalk cells (Loomis, 1975). McMahon (1973) has proposed a model for amoe-

PHYSIOLOGY

OF FUNGAL LIPIDS: SELECTED TOPICS

99

boid aggregation, which in some respects is not unlike that suggested by Roseman (1974)for intercellular contact in higher animals. He suggested that there are “contact-sensing” molecules on the surface of cells that regulate the internal concentration of‘cyclic AMP which provides positional value. The “contact-sensing’’ molecules are activated by interaction with complementary molecules on adjacent cells. The chemical composition of these cohesion factors has been examined, to some extent. Some of these are glycoproteins (Frazier et al., 1975). However, Wilhelms et al. ( 1974) have implicated glycosphingolipids in amoeba aggregation. A phenol-water extract from aggregationcompetent cells yielded two different antigens ( I and 11). Antigen I was assumed to be a glycosphingolipid since it contained fatty acid, phytosphingosine, phosphorus, but apparently no inositol. Surprisingly, it also contained ethanolamine and at least 19 sugar residues. There was no indication that antigen I was homogeneous. It is possibly an aggregate of relatively simple glycosphingolipids and a lipid analogous to the lipophosphonoglycan isolated from Acantharnoeba castellanii (Korn et al., 1974; Dearborn et al., 1976) which accounts for about 31% ofthe mass of the plasma membrane of this Soil amoeba. Approximately 26% of the compound is made up of neutral sugars (glucose, mannose, galactose, xylose; 5 : 4 : 1 : 1); 3.3% are glucosamine and galactosamine, 10% are aminophosphonates, 3.2% is phosphate and 14% are fatty acids. Antigen I from a non-aggregating mutant of D. discoideum contained markedly less fucose than the aggregation-competent cells. The results may suggest an absence of the “glycosyl-extension response” in aggregating amoebae. This term, coined by Hakomori et al. (1972), has arisen from the observation that the amount and sugar complexity of glycosphingolipids increases when animal cell cultures reach confluency, and this increase has been attributed to enhanced glycosyl extension of precursor lipids. This observation, combined with the evidence that the glycosphingolipids of animal cells are primarily localized in plasma membranes, has led to the suggestion by Hakomori et al. (1972) that, in growing cells, a certain proportion of glycosphingolipids of animal cells are primarily localized in plasma membranes, has led to the suggestion by Hakomori et al. (1972)that, in growing cells, a certain proportion of glycosphingolipids and glycoproteins are arranged in a complementary order to similar structures on an apposing cell. With confluency, the carbohydrate chains are linked together through complementary structures. Roseman ( 1974) suggests that the surfaces also contain glycosyltransferaseswhich would

100

P. J. BRENNAN AND D. M. L ~ S E L

then be responsible for both glycosyl extension and intercellular linkages at confluency. This concept of the function of glycosphingolipids on animal-cell surfaces may be directly applicable to aggregation in the Acrasiales. In the case of the Eumycotina, glycophosphosphingolipids, while providing an attachment point for cell-wall constituents to the membrane, may also provide aggregation sites or sexual agglutination factors. From the work quoted above, it is obvious that the glycophosphingolipid structure does not occur in animals; the phospho-inositol bridge between the ceramide and oligosaccharide moieties is specific to fungi and higher plants. Hence, these lipids are logical targets for antifungal chemotherapy. In addition, there is a case for examining these components in the fungi responsible for cutaneous and systemic mycoses; they may be amenable to vaccine development or be of use in developing specific immunological diagnostic methods. IV. Biosynthesis of Fungal Lipids

Biosynthesis of fungal sterols has been recently reviewed by Goodwin (1973)and Weete (1974). Erwin(1973)andWeete(1974) have reviewed most aspects of fungal fatty-acid biosynthesis, and the reviews by Harwood (1975) and Stumpf (1975) are also relevant to fungi. Erwin's ( 1973) review is particularly comprehensive in its treatment of unsaturated fatty-acid synthesis. Weete ( 1974) dealt with the latest information on hydroxyfatty-acid synthesis. In this connection, the evidence for a novel 2-hydroxy fatty acid in a glycolipid from A . niger (Laine et al., 1972; P. J. Brennan and F. D. Gunstone, unpublished results) raises some interesting biosynthetic problems. Aspergillus niger contains a glycolipid which is thought to be an acidic monoglucoside of 2-hydroxyoctadecenoic acid (Laine et al., 1972).The double bond was recognized as having trans configuration, but was not specifically located. More recently P. J. Brennan and F. D. Gunstone (unpublished observations) subjected this fatty acid to Von Rudloff oxidation with potassium permanganate and iodate. The oxidation product, after esterification, contained one major component, namely methyl pentadecanoate. This was indicative of' A3 unsaturation. The trans configuration of the double bond was also confirmed by a sharp infrared signal at 965 an-'.2-Hydroxy acids are constituents of various fungal lipids in particular triglycerides and sphingolipids (Weete, 1974 and Table 6, p. 87). Such acids are often long-chain (above C,,), and mainly saturated. In addition, acids with A3 unsatura-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

101

tion, which is always trans, are rare but known. The majority of unsaturated fatty acids have the cis configuration. The Z-hydroxyoctadec-trans-3-enoic acid, containing these unusual features, was previously unrecognized. While the biosynthesis of this fatty acid has not been investigated, thoughts o n it are relevant to the subject matter of this review (Fig. 3). Octadecanoyl- CoA

CH,-( CH,) ,,-CH2-CH2-CH2-CO-CoA

I CH,-(CH2)H-CH,

Desaturase

(0,) 0 ctadec- cis- 2 -enoyl- CoA

-C=C-CO-CoA

I I

H H

H

I

Oxygenase

I

CH,-(CH,) ,,-CH-CH-CH-CO-COA

\ /

(0,)

Octadec-2,3 epoxyenoyl-CoA

YH+

I H

I

CH,-( CH 2) ,,--C=CH-CO-CoA

I I

2 - H ydroxyoctadec-trans- 3 -enoyl-CoA

H OH FIG. 3. Possible biosynthesis of the novel 2-hydroxyoctadec-trans-3-enoic acid of Aspergtllus niger

Stearyl-CoA is the most likely immediate precursor. Direct ahydroxylation of it, as occurs in the case of a-hydroxypalmitate (Morris, 19701, seems unlikely since the product would have to undergo desaturation to yield a AS trans product and all of the known eukaryotic desaturase systems yield a cis-unsaturated product (Erwin, 1973). Another slight possibility isthat Bloch’s (1969) anaerobic pathway for fatty-acid synthesis could be utilized, followed by a-hydroxylation. However, while the enzyme P-hydroxydecanoyl thioester dehydrase could yield an octadec-3-enoic acid, it would have the cis con-

P. J. BRENNAN AND D. M. LOSEL

102

figuration. Similarly, while the products of de now fatty-acid synthesis are trans, the unsaturation is A2. The most likely explanation (Fig. 3) is that stearyl-CoA undergoes specific oxidative desaturation. Such desaturation would be stereospecific, with D-hydrogens removed from both carbons to yield the cis bond. Perhaps the mechanism for introduction of conjugated unsaturation (Gunstone, 1967) is then utilized. In many conjugated polyunsaturated fatty acids, the feature: -R-CH=CH-CH-R-

I

OH

is evident. A point about the scheme in Fig. 3 is that introduction of the P,y-trans double bond and the a-OH group is closely connected. A.

PHOSPHOLIPIDS

The subject of fungal phosphoglyceride synthesis and the closely related triacylglycerol synthesis has been exhaustively reviewed by Hunter and Rose (1971), Mangnall and Getz (19731, Weete (19741, Brennan et al. (1975) and Rattray et al. (1975). From all of these and other reviews (Kates and Marshall, 1975; van Den Bosch, 1974; Gatt and Barenholz, 1973),it appears that there exist in nature four types of pathways for phospholipid synthesis:(a) CDP- Diglyceride pathway : phosphatidylinositol

f

/y

inosi t o l d

phosphatidylserine

-

phosphatidylethanolamine

phosphatidylglycerolphosphate glycerol diphosphatidylglycerol

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

103

( b ) CDP- Base pathway: CDP-choline + diglyceride + phosphatidylcholine + CMP CD P - ethanolamine + diglyceride +. phosphatidylethanolamine + CMP CDP-choline + ceramide + sphingomyelin + CMP (c) Modajicationpathway, e.g.: phosphatidylserine -+ phosphatidylethanolamine + CO phosphatidylethanolamine + S-adenosylmethionine -+ mono- and dimethylphosphatidylethanolamineand phosphatidylcholine phosphatidylglycerophosphate + phosphatidylglycerol + Pi phosphatidylinositol + ATP di- and triphosphoinositides (d) Exchangeltransferpathways: phosphatidylcholine + ceramide -+ sphingomyelin + diglyceride phosphatidylglycerol + phosphatidylglycerol diphosphatidylglycerol + glycerol phosphatidylcholine + ethanolamine -+ phosphatidylethanolamine + choline phosphatidylserine + ethanolamine -+ phosphatidylethanolamine + serine phosphatidylethanolamine + serine 4 phosphatidylserine + ethano lamine phosphatidylethanolamine + choline ---t phosphatidylcholine + ethano lamine -+

-

Insufficient work has been done to determine the extent or relative importance of these pathways in fungi. However Syeiner, M. R. and Lester (1972) showed that CDP-diglyceride could give rise to phosphatidylserine, phosphatidylinositol, phosphatidylglycerophosphate and diphosphatidylglycerol in Sacch. cereuisiae. The possibility that deoxy-CDP-diglyceride is involved in any of the fungal liponucleotide-requiring reactions, as happens in E . coli (Raetz and Kennedy, 1973), has been discounted by Schneider and Kennedy ( 1976). They isolated the liponucleotide fraction from Sacch. cereuisiae and showed that it was all CDP-diglyceride, constituting about 0.08% of the total glycerides of the yeast. Phosphatidylethanolamine apparently arises mostly by the CDP-ethanolamine pathway, but more of it comes by decarboxylation of phosphatidylserine. Phosphatidylcholine, on the other hand, seems to be formed mostly by methyla-

104

P. J. BRENNAN AND D. M. LOSEL

tion of phosphatidylethanolamine (Waechter et al., 1969; Steiner and Lester, 19691,but some of it is obtained via CDP-choline. Many of these observations also apply to Neurospora crassa, particularly the two pathways for phosphatidylcholine synthesis (Crocken and Nyc, 1964; Scarborough and Nyc, 1967a, b ; Sherr and Byk, 197 1). In their studies with Neurospora sp., Scarborough and Nyc( 1967a)found that microsomes from a mutant of this organism were deficient in phosphatidylethanolamine: N-methyltransferase activity, while microsomes from a second mutant could not catalyse methylation of either phosphatidylmonomethylethanolamine or phosphatidyldimethylethanolamine. This finding, and other work (Scarborough and Nyc, 1967b),was important in showing that, in N. crassa and probably other organisms, a single enzyme converts phosphatidylethanolamine to phosphatidylcholine. The CDP-diglyceride-dependent synthesis of phosphatidylserine and its decarboxylation to phosphatidylethanolamine have been demonstrated in a particulate subfraction of N . crassa (Sherr and Byk, 197 1). Regulation of phosphatidylcholine synthesis by yeast is related to the free choline available to the cells (Waechter et al., 1969; Steiner and Lester, 1969). Presence of choline in the growth medium repressed synthesis of enzymes involved in the methylation pathway, thereby presumably ensuring the effectiveness of the CDPcholine pathway. There is some evidence that not all fungal phosphatidylinositol arises by the CDP-diglyceride pathway. The existence of an “activated” phospho-inositol was mooted by Steiner and Lester (1972); a CDP-inositol or inositol 1 : 2 cyclic phosphate or inositol exchange from one of the sphingolipids were possibilities. White and Hawthorne (1970) seemed also to imply the existence of a second phosphatidylinositol synthetic pathway since they clearly could not demonstrate the CDP-diglyceride pathway in Schizosacch. pornbe or Sacch. cerevisiae. They unsuccessfully explored several other reasonable alternatives, such as exchange of the ethanolamine of phosphatidylethanolamine for free inositol. They also investigated the reversal of a phospholipase D-type action, i.e. condensation of phosphatidic acid and inositol. However, Deshusses ( 1974) suggested that this additional pathway in Schizosacch. pombe and N. crassa may involve donation of the phosphatidyl moiety of phosphatidylglycerolphosphate to free inositol, i.e. :

phosphatidylglycerolphosphate + inositol -+ glycerolphosphate

phosphatidylinositol

+

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

105

This is related to the phosphatidylserine pathway described by Borkenhagen et a/.(1961). The evidence for such a route for phosphatidylinositol synthesis was based on the observation that no activation of synthesis took place when CDP-diglyceride was added to the incubation mixture. Of several substances tested on the enzyme from both fungi, only a-glycerophosphate enhanced incorporation suggesting that the direct precursor of fungal phosphatidylinositides was possibly phosphatidylglycerolphosphate. Some of the workers investigating the possibility of a CDPdiglyceride-independent pathway for phosphatidylinositol synthesis in fungi seem to be unaware of the observations of Paulus and Kennedy (1960). They reported that inositol, in the presence of Mn2+,could be incorporated by liver microsomes into phosphatidylinositol in the absence of added CDP-diglyceride, and suggested that this occurred by an exchange reaction with endogenous phosphatidylinositol and that the exchange reaction and the CDP-diglyceride reaction were part of the same enzyme systern : CDP-diglyceride + enzyme gphosphatidyl-enzyme

+ CMP phosphatidyl-enzyme + inositolephosphatidylinositol + enzyme Biosynthesis of the polyphospho-inositides in yeasts is assumed to be similar to the process in animals, i.e. catalysis by ATP-Mg*+requiring kinases which convert phosphatidylinositol to di- and triphospho-inositides. Talwalkar and Lester ( 1974) demonstrated that conversion of phosphatidylinositol to diphospho-inositide was catalysed by a soluble fraction of a yeast cell homogenate. However, Wheeler et al. ( 1972)regard this activity as membrane-associated which is more in keeping with the role of the enzyme in maintaining the cell’s “adenylate charge” (Talwalker and Lester, 1973). Fungal disphosphatidylglycerol is known to arise by the CDPdiglyceride pathway. The possibility that fungal diphosphatidylglycerol may also be synthesized by the prokaryotic pathway, involving interaction of two molecules of phosphatidylglycerol, has not been examined. Synthesis of diphosphatidylglycerol in yeasts is associated only with mitochondria; other phosphoglycerides are synthesized primarily by microsomal fractions (Cobon et al., 1974). Some of the early steps in phosphoglyceride and acylglyceride synthesis in fungi have received considerable attention (Fig. 41, for instance, acylation of a-glycerophosphate by yeast extracts (Kuhn and

P. J. BRENNAN AND D. M. LOSEL

106

CH,OH

CH,OH

.

1 CHOH

sn-~lycerol-3phosphate

I c=o I

I

CH,O-POS-

k

Mono-acyl-sn-

Dihydroxyacetonephosphate

CH ,-P 0 ; Fatty acyl CoA-1

CH,OCOR NADP' NADPH glycerol - 3 I CHOH phosphate (lysophosphatidic I acid) CH,O-PO;-

+ H+

, -

CH,OCOR

&=o I

Mono-acyl dihydroxyacetonephosphate

CH,O-PO:-

Fatty acyl-CoA

Phosphatidic acid

CH,OCOR 1 CHOCOR

CTP

A

CDP-diglyceride

I CH,O-PO;-

/\

Phosphatase a,P-Diglyceride

k

Fattyacyl CoA

Triglycerides

i

Phosphoglycerides CDP-Bases

1 Phosp hoglycerides

FIG. 4. Precursor role of glycerophosphate in fungal glyceride synthesis

Lynen, 1965; Johnston and Paltauf, 1970; White and Hawthorne, 1970) and acylation of dihydroxyacetone phosphate (Johnston and Paltauf, 1970). The mechanism of synthesis of CDP-diglyceride by a yeast particulate fraction (Hutchison and Cronan, 1968) and rnitochondria (Mangnall and Getz, 197 1) as shown in Fig. 4 is apparently questioned by Steiner and Lester ( 197 2) who provided some evidence that an endogenous lipid other than phosphatidic acid was the precursor of the diglyceride moiety. There is conflicting evidence for some of the exchangehransfer-type reactions for synthesis of some phosphoglycerides in certain fungi (White and Hawthorne, 1970; Deshusses, 1974). However, there is no reason why some of these reactions should not exist. Ullrnan and Radin (1974) consider that, in mouse liver microsomes, the transfer

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

107

method is the major and perhaps the only means of synthesizing sphingomyelin. Such a “transfer” pathway seems to apply to synthesis of other sphingolipids in fungi (see p. 109). B.

GLYCOLIPIDS

Biosynthesis by fungal extracts of both types of hydroxy-acid glycosides, diglycosyldiglycerides, sterol glucosides and polyprenol phosphate mannose has been examined. Jarvis and Johnson ( 1949) described an extracellular glycolipid, L-rhamnosyl ( 1- 3)-~-rhamnosylhydroxydecanoylhydroxydecanoate,which was secreted into growth medium by Pseudomonas aeruginosa. Biosynthesis was shown (Burger et al., 1963)to proceed as follows: 2 p-hydroxydecanoyl-CoA

-

p-hydroxydecanoyl- p- hydroxydecanoate /

/p-

TDP- L- rhamnose

L-rhamnosyl- p- hydroxydecanoylTDP- L-rhamnose

hydroxydecanoate

4

L-rhamnosyl-L-rhamnosyl- P-hydroxydecanoylp- hydroxydecanoate Since the sophorosides are also hydroxyfatty acid glycosides, and are primarily extracellular in location, Esders and Light (197213)sought a related pathway for synthesis of these in Candida bogoriensis, and established the sequence: ,B-hydroxydocosanoicacid + UDP-glucose

(1)

glucosylhydroxydocosanoic acid ( G l c H D y UDP-glucose glucosyl-glucosylhydroxydocosanoicacid 2 acetyl-CoA (Glc,HDA) diacetyldiglucosylhydroxydocosanoic acid (Ac,Glc,HDA)

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P. J. BRENNAN AND D. M. LOSEL

Esders and Light ( 197213) purified 20-30 fold the glycosyltransferase activities (reactions 1 and 2) but found that they maintained a constant ratio throughout the purification steps and were apparently one enzyme. The transferases were specific for UDP-glucose. Of several hydroxyfatty acids tested, P-hydroxydocosanoic acid was the most effective initial substrate. The methyl ester of GlcHDA was more effective for subsequent glycosylation than was the free-acid form. Acetyltransferase(s), which catalysed incorporation of the acetyl group from acetyl-CoA, was also found in crude extracts of C. bogoriensis. The authors tentatively concluded that the other sophorosides of C. bogoriensis (AcGlc,HDA and Glc,HDA) were formed by de-acylation of Ac,Glc,HDA. P. J . Brennan and J. A. Hackett (unpublished observations) demonstrated the existence in A. niger of a UDP-glucose: 2-hydroxyoctadectrans-3-enoic acid glucosyltransferase, apparently catalysing the following reaction: UD P-glucose + 2-hydroxyoctadec-trans- 3-enoic acid monoglucosyloxyoctadec-trans-3-enoic acid

-

+ UDP

The properties of the glucosyltransferase responsible for biosynthesis of the monoglucosyloxyfatty acid are in many respects similar to those of the glucosyltransferases in C. bogoriensis. UDP-Glucose was the glucose donor in both systems; other glucose nucleotides were inactive. Both enzymes were associated with particulate cellular fractions. Both systems exhibited closely similar pH-value and temperature optima, inactivation temperature and linear incorporation of glucose over several hours. The principal difference between the two systems was in their relative specificity for the aglycone glucose acceptors. In C. bogoriensis, various hydroxy C,, fatty acids were active as glucose acceptors yielding sophoroside-like materials: The glucbsyltransferase of A. niger shows singular specificity for the natural fatty acid; other 2-hydroxyfatty acids were inactive in this system. Biosynthesis o f glycosyldiglycerides in Gram-positive bacteria has been reviewed by Shaw (1975) and Sastry (1974). while synthesis in higher plants has been ably reviewed by Rosenberg (1973), and by Mudd and Garcia (1975)in addition to Sastry (1975).Taking as a guide the reaction sequence already identified in several bacterial and plant sources, P. J. Brennan and J. A. Hackett (unpublished observations) showed that microsomal preparations of A. niger catalysed incorporation from UDP-[6-3Hl-glucose into an alkali-labile lipid, chromato-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

109

graphically similar to the diglycosyldiglyceride mixture previously obtained from the same organism. Typically, 1% incorporation per mg of protein was observed. I t thus appeared that the diglucosyldiglyceride of A . niger was formed by the following reaction sequence:

-

diglyceride + UDP-glucose 3 monoglucosyldiglyceride + UDP diglucosyldiglyceride monoglucosyldiglyceride + UDP-glucose UDP

+

Adenosine diphosphate-glucose and GDP-glucose were inactive as glucose donors under conditions where glucose incorporation from UDP-glucose was optimal. The biosynthetic reaction was relatively insensitive to divalent cations and dithiothreitol. The bptimum pH value for the enzyme(s) lay in the range 7-7.5, while optimal activity was observed at 25OC. The incorporation of label from UDP-[6-3H]glucose into diglycosyldiglyceridevaried linearly with incubation time up to 3 h. Several publications have suggested that various fungi contain sterol glycosides. However, the structural evidence was never convincing. Esders and Light (1972c), from biosynthesis studies, proved the existence of such a glycolipid in C. bogoriensis. Crude extracts of sonicated C. bogoriensis catalysed incorporation of radio-activity from UDP[*4C1-glucoseinto a lipid other than the sophorosides. The lipid acceptor for the labelled glucose was identified as ergosterol, although cholesterol was also active. Therefore the following reaction was demonstrated : ergosterol

+ UDP-glucose-+ergosterolglucoside + UDP

It appears that the product is present in tangible amounts, but attempts to isolate it were thwarted by its instability. Hence clear-cut chemical characterization was not effected. The established features of polyprenolphosphomannose synthesis are dealt with in the context of their function (p. 83). C.

SPHINGOLIPIDS

Some aspects of the biosynthesis of fungal sphingolipids have been examined in considerable detail. The knowledge that sphingosine is derived from palmityl-CoA and serine via a pyridoxal phosphatelinked enzymes was obtained by Snell and his colleagues using Hanseniaspora ualbyensis and Hansenula cferri. Weete ( 197 4) has extensively reviewed this topic.

P. J. BRENNAN AND D. M. L ~ S E L

110

CH,(CH,),,COOH ATP, CoASH

+ PPi

$AMP

CH&CH,),pCO-SCoA

CH,(CH * "-C-CH-CHZOH 11

I

1

0 NHZ

CHs(CHz)l,-CH-CH-CH20H

I\

1';

4 CH,CO-SCoA /

/

,'

I

I

OHlNH, I,, 5 \, ,,CH&CHz)ZlCH-CO-SCoA \

\

/

I

'

OH CH,( CH ,) ,,-CH-CH-CH

I

I 1 O H NH

I

I I

w

CHS(CHz)l4-CH-

I

OCOCH,

2~

H

I c=o

CH-CH,OCOCH,

1

I

CH-OH

I

NHCOCH,

(?&)Zl

I

Phosphatidylinositol

9,

Diacylglycerol

CH,( CH z) ,-CH-CH-CH

I

I

O H NH

,-0-P-

0

II

I

0 -1nositol

0

I

C=O

I

CHOH

I

(CH,),I

I

CH, FIG. 5 . Summary of the only sphingolipid-synthesizing steps established for fungi

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

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Barenholz and Gatt (1969, 1972) described a microsomal enzyme from H . cfewi, namely long-chain base-acetyl-CoA acetyltransferase, which catalysed acetylation of sphingosine, dihydrosphingosine, and phytosphingosine, both at the amino group (N-acetylation)and at the hydroxyl groups (0-acetylation). The enzyme also further acetylated the N-acetyl derivative of these bases. However, it did not act on the N-octanoyl- or N-palmitoyl derivatives of sphingosine or dihydrosphingosine nor on psychosine ( 1-0-galactosyl-sphingosine). It was not determined whether the enzymic preparation contains two separate enzymes, one for N-acetylation and one for 0-acetylation, or whether one enzyme is responsible for both activities. Barenholz et al. (1973) showed that this enzyme was present in the microsomes of high and intermediate producers of extracellular acetylated sphingosines but was practically absent from the microsomes of low producers. The five enzymic steps involved in the conversion of serine and palmitate to acetylated dihydroshingosine were examined in high and low producers (Fig. 5). Overall synthesis of dihydrosphingosine was greater in the high producer. However, it was the condensing enzyme, and not the reductase, which showed a big increase (5-10 times) in high producers as compared with low producers; activities of the reductase were about equal. In addition, the activity of the long-chain base-acetyl-CoA acetyl-transferase was at least 30 times greater in the high producers, whereas that of the palmityl thiokinase was about equal in high and low producers. These results demonstrated the existence of two enzymes whose activities are greater in the high producer, namely the condensing enzyme and the acetylating enzyme. The genetic aspects leading to such a dual metabolic lesion are unknown. In addition, Barenholz et al. ( 1973) concluded from examining the relative rates of four of the enzymes that condensation of palmityl-CoA and serine is 1. Long-chain fatty-acyl synthetase (palmityl thiokinase). 2. 3-Ketodihydrosphingosine synthetase (condensing enzyme). Krisangkura and Sweeley ( 1976) recently found that synthesis of the 3-ketodihydrosphingosine in Hunsenula czJerri involves the replacement of the a-hydrogen atom and the carboxyl group of serine by a proton from the medium and a palmitoyl group, rather than a previously proposed mechanism in which the a-hydrogen of serine is retained. 3. 3-Ketodihydrosphingosinereductase (reductase). 4. Long-chain base-acetyl- CoA acetyltransferase (acetyltransferase). 5 . Long-chain base-acyl-CoA acyltransferase (acyltransferase). 6. Ceramidephosphatidylinositolphosphoinositol transferase (phosphatidylinositol exchange).

112

P. J. BRENNAN AND D. M. LOSEL

the rate-limiting step in biosynthesis of sphingosine at least in this organism. Direct glycosylation of ceramides to yield monoglycosylceramides has not been examined in fungi, although it is well studied in animal tissues (Basu et d.,1968). A small amount of work on biosynthesis of the glycophosphosphingolipids has been conducted by Lester and Brennan and their colleagues. A likely pathway is “exchange/transfer” involving phosphatidylinositol and ceramide, in a fashion similar to synthesis of sphingomyelin although, with the circumstantial evidence for other forms of “activated” phospho-inositol, this is not certain. Angus and Lester ( 1972) showed that, when Sacch. cerevisiae was uniformly labelled with both [32PlPi and [2-3Hlinositol and transferred to unlabelled medium, the major yeast glycophosphosphingolipid, CerP,I,M, accumulaated both 3H and 32Pat a rate suggesting that phosphatidylinositol (PI) served as the phospho-inositol precursor and that the bond between phosphorus and inositol was not broken during the conversion. However, these experiments did not rule out the possibility that glycerylphosphorylinositol, of which there is considerable amounts in yeast, or some other form of phospho-inositol arising from PI, was the direct precursor. In order directly to test the capacity of phosphatidylinositol to donate its phospho-inositol moiety in an in vitro system, Hackett and Brennan (1977) incubated microsomal extracts of A . nzger with [32Plphosphatidyl-[3Hlinositol. Both labels were convefted into watersoluble materials which were characterized as [32PlPiand [3Hlinositol. I t was concluded that any biosynthetic activity present in the membrane fraction was masked by breakdown of the added substrate. However, incubation of soluble protein fractions of A . niger with [32Plphosphatidyl-[3Hlinositol resulted in label incorporation into an alkali-stable lipid fraction, with preservation of the original 32P :3H count ratio. This activity was considerably enhanced by addition of A . niger ceramide (containing 2-hydroxy fatty acids); 2,3-dihydroxy fatty acid-containing ceramides inhibited activity. The label incorporation was low (typically 5% per mg protein). Thin-layer chromatography of the labelled lipid extract alongside labelled inositol phosphorylceramide (CerPI) previously obtained from A . nzger (Table 6, p. 186) showed that the labelled lipid extract cochromatographed with CerPI. The variation in the extent of the biosynthetic reaction with incubation time was studied. In common with other complex lipid biosynthetic reactions in A . nzger, maximum label incorporation into

PHYSIOLOGY

OF FUNGAL LIPIDS: SELECTED TOPICS

113

CerPI was obtained after about 4 h incubation. This evidence supports the idea that CerPI biosynthesis proceeds as follows : ceramide (2-hydroxyfattyacid-containing) + phosphatidylinositolinositol phosphorylceramide + diacylglycerol. Incidentally, these observations also provide evidence that the phosphate-inositol linkage in the intact complex glycophosphoshingolipids was via the 1-hydroxyl of inositol, since the inositolphosphate moiety of phosphatidylinositol is so constructed. To elucidate the biosynthesis of glycophosphosphingolipids, fungi other than A . niger will have to be employed, since its major sphingolipid is CerPI. Saccharomyces c e k i s i a e should be ideal for exploring the synthesis of compounds with the CerPJ, moiety, and some of the basidiomycetes should be useful for examining CerPI-oligosaccharide synthesis. Alternatively, the use of appropriate mutants could be invaluable. Such mutants, lacking one or more enzymes in a biosynthetic sequence, accumulate relatively large quantities of the intermediate immediately proximal to the lesion, when grown in unsupplemented medium. In particular, a number of well characterized mutant strains of Neurospora sp. with defects in phospholipid synthesis (Hubbard and Brody, 1975; Crocken and Nyc, 1964) and a requirement for inositol (Mishra and Tatum, 1973)have been described. At one stage, Rizza et al. (1970) suggested the existence of CDPceramide in Bacteroides melanogenicus, and its possible role in the biosynthesis of phosphosphingolipids in this bacterium. Moreover, the demonstration by Schneider and Kennedy ( 1973) that the enzyme diacylglycerol kinase of E . coli readily phosphorylates ceramide lent some support to the suggestion. However, Schneider and Kennedy (1976)could find no evidence for CDP-ceramide as an intermediate in the biosynthesis of phosphosphingolipids in yeast. They synthesized CDP-ceramide labelled with L3H1in the cytidine moiety and [32Plin the phosphoceramide portion, and showed that the liponucleotide fraction from Sacch. cerevisiae was devoid of such a compound.

V. Role of Lipids in Fungal Morphogenesis

The almost universal occurrence of lipid bodies in fungal reproductive organs, spores, gametangia and gametes has been recorded in light- and electron-microscopic studies and in biochemical assays.

P. J. BRENNAN AND D. M. LOSEL

114

Comprehensive reviews of such observations and of investigations of the role of lipids in sporulation and germination have been provided by Weber and Hess (1974), and Hess and Weber (1974, 19761, who have gathered together a wealth of previously rather scattered information. In the period since these reviews, further reports on these topics have continued to appear. Many aspects of the lipid physiology of fungal differentiation involve secondary metabolism. The close relationship between differentiation in fungi and secondary metabolism has been stressed at various times (Bu'Lock, 1967; Smith and Berry, 1974). This can also be seen in the complex tissues of higher plants, but some aspects are amenable to more direct investigation in the simpler situation of fungal morphogenesis. Smith and Galbraith (197 1) point out that fungal differentiation occurs in essentially endogenous, self-sufficient systems which have to a greater or lesser extent cut themselves off from the environment, and that some degree of nutritional deficiency may be involved. Like secondary metabolism, differentiation characterisically occurs in older parts of colonies, not in actively growing marginal hyphae which are exploiting fresh medium. Lipid components with a special role in development of fungal spores, fructifications and resting or resistant structures include hormonal and growth-regulating factors, membrane lipids of sporulating or reproductive structures and lipid reserves which provide carbon and energy sources for motility of zoospores, dormancy of asexual or sexual spores and germination. In addition, protective hydrocarbons accumulate on the surfaces of some reproductive or resting structures. Comprehensive accounts of' these have been given by Weete (1972, 1974). A.

HORMONAL A N D GROWTH-REGULATING FACTORS

Sussman (1976), in a review of activators of germination, has provided lists of fungal spores activated by detergents, organic acids or lipids, and of membrane-located enzymes that require phospholipids or detergents for activity. He emphasizes the physiological importance of membrane permeability changes, due to heat or lipophilic compounds, which affect diffusion across membranes separating enzymes and substrates. Of the many fungal and plant metabolites regulating growth and reproduction of fungi, a striking proportion are lipids, particularly terpenoids. These include the three fully character-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

115

ized fungal sex hormones (Machlis, 19721, sirenin, antheridiol and trisporic acids. 1. Fungal Sex Hormones

a. Sirenin secreted by female gametes of the Chytridiomycete water mould, Allomyces, attracts the motile male gametes. First discovered by Machlis (19581, chemically characterized by Rapoport (Machlis et al., 1966)and later synthesized (Plattner and Rapoport, 197 11, sirenin is an oxygenated sesquiterpene. b. Antheridiol, the sex hormone of the oomycete water mould, Achlya bisexualis, the subject of the classical investigation of Raper from 1939 to 1950 (Raper, 19521, isolated by McMorris and Barksdale (1967),has the carbon skeleton of stigmasterol. Antheridiol was shown by Barksdale (1963) to act both as Raper’s “hormone A”, which is secreted by vegetative hyphae of the female thallus and induces formation of antheridial branches, and as L‘hormone C”, by which the oogonial initials on the female thallus attract the antheridial branches and induce differentiation of antheridia. Antheridiol is effective at a concentration as low as lo-’“ M with receptive strains. The factors diffusing from the antheridial hyphae which induce development of the oogonia (“hormone B”) and initiate cleavage of the oogonia into female gametes or oospheres (“hormone D” of Raper) have not yet been identified. Studies on the Achlya hormones have been reviewed by Barksdale (1969). c. Trisporic acids stimulate formation of zygophores in zygomycetes of the order Mucorales, and are produced only when both (+) and (-1 strains are present in the same culture. Zygophores are hyphae with a high carotene content which grow towards and fuse with zygophores from a colony of compatible mating type. The trisporic acids were originally recognized as the fraction from chloroform extracts of mated cultures of Blakeslea trispora which increase @-caroteneproduction by single-strain cultures. They are C,, carboxylic acids which are formed from p-carotene, but virtually nothing is known oftheir mode of action. Addition of trisporic acid induced an 80-fold increase in @carotene production (Thomas and Goodwin, 1967).The physiology of differentiation in the Mucorales and the biosynthesis of trisporic acids by cleavage of p-carotene via retinal are reviewed by Gooday (1973). This review includes a brief report on experiments showing that, during sexual reproduction, carotene can be oxidatively polymerized

116

P. J. BRENNAN AND D.

M. LOSEL

to give the sporopollenin of the zygospore wall, the highly resistant hydrocarbon better known from the protective exine of pollen grains of seed plants. d. Other reproductive hormones. Controversy still surrounds the other sexual hormones for which evidence has been obtained in the Mucorales. Volatile factors have been implicated (Hepden and Hawker, 1961) and sex-specific agglutinins may be involved in the rapid and inseparable fusion of gametangia (Hawker and Gooday, 19671. The cellular slime moulds respond to morphogenetic factors which may be in some ways comparable to the above hormones. Aggregation of Acyrostelium leptosomum is stimulated by sterols and alkaloids (Hostak and Raper, 1960). Studies on yeast sexual hormones by Takao et al. (1970)have shown that cells of Sacch. cereviseae secrete a steroid resembling, but not identical to, testosterone and estradiol, compounds which induce expansion of cells of “a” and “a” mating type.

2 . Short-Chain Fatty Acids Short-chain fatty acids have been implicated in dormancy-breaking and stimulation of germination in spores of Phycomyces sp. (Robbins et al., 19421, rust urediospores (Reisener et al., 1961)and basidiospores of Agaricus bisporus (Losel, 1967 ; O’Sullivan and Losel, 197 1 ; Rast and Stauble, 1970). The occurrence and activity of these were reviewed briefly by Brennan et al. ( 1975) and included in the review by Weber and Hess (1974). Low concentrations of nonanoic acid in the atmosphere over spore cultures were stimulatory to germination of Aspergillus niger spores (O’Sullivan and Losel, 197 11, while the corresponding aldehyde nonanal was reported to overcome self-inhibition of Puccinia graminis urediospores (Allen, 1958) and stimulated vegetative growth of Pestalotia rhododendri (Norrman, 19681 and certain basidiomycetes (Fries, 1961). Medium chain-length fatty acids have more frequently been associated with inhibitory effects on fungi, particularly at low pH values. Examples of this include leakage of cell materials and inhibition of endogenous respiration of Boletus variegatus by low concentrations of C, to C,, fatty acids (Pedersen, 19701, which was attributed to interaction with lipophilic components of membranes, and the sporostatic activity of nonanoic acid against Cunninghamella elegans, Fusarium

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

117

oxysporum, A . niger and other filamentous fungi (Garrett and Robinson, 1969). The inhibited spores germinated subsequently on transfer to fresh medium without nonanoic acid, which was shown to be one of the metabolites produced by various sporing fungi which prevented germination of spores on their parent culture (Robinson and Park, 1966). The well known fungistatic activity of undecylenic acid against several dermatophytes was shown to be dependent on the concentration of undissociated acid (Prince, 1959). So far, these shorter-chain fatty acids have been relatively little studied and are recorded from few fungal sources, e.g. yeasts (Stevens, 19601, mushroom mycelium, sporophores and spores (Rast and Stauble, 1970; Stauble and Rast, 1971), and Boletus variegatus (Krupa and Fries, 197 1). The usual techniques for analysis of longer-chain fatty acids tend not to detect the volatile fatty acids. Further investigation is, however, likely to show short-chain fatty acids to be widely distributed fungal metabolites, as was demonstrated by Garrett and Robinson (1969)for non-anoic acid. It has recently been shown, using electron-capture gas-liquid chromatography, that up to 6.1% of mycelial dry weight of six soil fungi ( Asperigillus niger, Cephalosporium sp., Cladosporium herbarum, Fusarium sp. and Penicillium frequentens) is composed of hydroxybutyrates (Nuti et al., 1975). If the cultures are shaken, the amounts of hydroxybutyrates rise to remarkable levels. ALpha-Hydroxybutyricacid was present in large quantities in all of the micromycetes examined, and the p- and y-hydroxybutyrates occurred in smaller amounts. Apparently these are not polymerized, unlike Sacch. cerevisiae which accumulates p-polyhydroxybutyrates. Previous work by Nuti and his colleagues suggests that hydroxybutyrates enter into fatty acid metabolism in microfungi, and serve either as metabolic intermediates or reserve material. 3. Growth-Regulating Terpenoids

a. Carotenoids and Photomorphogenesis. The interest long attached to close association of carotenoid production with formation of reproductive structures and with photoresponses was intensifed when the trisporic acids, first recognized as factors stimulating carotene production in Blakeslea trispora (Caglioti et al., 1964), were found to play a fundamental role in sexual conjugation of various members of the Mucorales. Carotenoids are abundant in many fungal structures

118

P. J. BRENNAN AND D. M. LOSEL

showing phototrophic growth, e.g. sporangiophores of species of Pilobolus and Phycomyces, and in fructifications of various higher fungi which require light for proper development. In Aspergillus giganteus, light induces carotenogenesis, conidiophore extension and orientation of the tall conidiophores (Trinci and Banbury, 1969).The apical 240 pm of the developing conidiophore is positively phototropic towards light of wavelength less than 520 nm. This light triggers a chain reaction which maintains growth and carotenogenesis. The stimulatory effects of light on sporulation and production of cleistothecia in Aspergillus species are reviewed by Smith and Anderson ( 1973). Diphenylamine treatment of Neurospora crassa represses both production of conidia and carotenogenesis (Youatt et al., 1971). Whether the striking yellow and red pigments, widely distributed in apothecia of discomycetes, and the oil drops of the asci have special functions in formation of apothecia and phototropic orientation and discharge of the asci does not appear to have received much attention. The recent observations of Webster (1976)might indicate a short-wave light response in production of the bright yellow apothecia of Pezizella ericae. Carlile and Friend ( 1956) showed that apothecium formation could occur in polyene-inhibited cultures of Pyronema omphalodes, and concluded that the carotenoids which are responsible for orange pigmentation of normal apothecia were not essential for the photomorphogenetic development of these fructifications. Ingold and Marshall ( 1963) found that “puffing” of asci in apothecia of Ascobolus sp. was induced by light of wavelengths 400 to 460 but not at 500 to 750 nm. The slime moulds, another group of organisms often having bright yellow and orange pigments in their fructifications or even in the vegetative plasmodium, have been studied by several workers in relation to the characteristic photoresponses shown in their fruiting behaviour (Daniel and Rusch, 1962; Gray, 1953; Wolf, 1959). Gray ( 1953) demonstrated the sensitivity of photoreceptor pigments of Physarum polycephalum to wavelengths of 436 nm. Sauer ( 1973), in his review of differentiation in Physarum sp., summarized work on the yellow pigment, a conjugated polyene, which may be a photoreceptor. Its action spectrum showed peaks at 330 to 540 nm and 630 to 7 13 nm. Green light of 540 to 620 nm wavelength could reverse the stimulatory effects of the active wavelengths.

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

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In spite of the above examples of correlation with photomorphogenesis, the role of the often abundant carotenoids of fungi remains obscure, since there is strong evidence for flavonoids being the photoreceptors in some of these responses, e.g. in development of sporangiophores of Pilobolus (Page, 1956).The overlapping of action spectra of flavonoids and carotenes makes it difficult to resolve the question of whether carotenoids may have a direct role in photoresponses or whether they protect photoreceptor systems against photo-oxidation, as postulated for carotenes associated with photosynthetic systems in plants and bacteria. Some basidiomycetes produce sporophores with carotenoid pigmentation, but their possible involvement in photomorphogenesis has not been investigated, although the carotenoids of various basidiomycetes have been studied from the point of view of taxonomy. In a recent study of the carotenoid content of Cantherellus infundibulformis, Valadon and Mummery (1975)isolated the first epoxycarotenoid to be found in a fungus, together with ,!land &carotene and translycopene. Since this compound has the same absorption spectrum as neurosporene, Valadon and Mummery (1975) pointed out that previous identifications of “neurosporene” in other fungi may have to be reexamined. Changes in carotene production during development have been demonstrated in the ubiquitous cellulose-decomposing soil chytrid Rhizophyctis rosea (Davies, 196 l), young cultures of which appear red-pink in colour, due to formation of lycopene, while mature cultures produce, in addition, P-carotene and appear orange in colour. Here, too, the possible physiological and ecological significance of the pigments has not been explored. The more intensively studied chytrid, Allomyces javanicum, is characterized by the marked carotenoid content of the antheridium. Turian ( 1962) showed that, even when carotenoid synthesis was 95%inhibited, male gametes were still liberated which, in spite of decreased motility, fused with the female gametes to give normal zygotes. Similarly, in the related Blastocladiella sp., the significance of the abundant carotene in resting sporangia is not clear. Carotene biosynthesis has been investigated in cell-free systems by Chichester’s group, using extracts of Phycomyces blakesleeanus (Yamamoto et al., 1961; Yokoyama et al., 1962; Lee and Chichester, 1969). Bramley and Davis (1975), working with two mutants of P. blakesleeanus, namely, a red form producing lycopene and a yellow mutant accumulating only P-carotene, demonstrated active synthesis

120

P. J. BRENNAN AND D. M. L ~ S E L

of these carotenoids from ['4C]-labelled mevalonic acid, even in the absence of light and oxygen. Maximum formation of lycopene took place in dark, anaerobic conditions, but @-carotene production was stimulated by light. Carotenoid synthesis in wild-type P. blakesleeanus appeared more responsive to stimulation by light. The rate of formation of @-carotenein the yellow mutant and of lycopene in the red mutant, as well as the yield of total lipid, paralleled the growth curves. Bramley and Davis (1975) suggested that such production of carotenoid throughout the active growth phase should not be considered as secondary metabolism. This sort of situation is, however, often encountered in non-synchronous cultures of filamentous fungi, where maturing and static stages of mycelium are accompanied by marginal h p h a e which are still actively growing. Bramley and Davis (1975)suggested that removal of /?-carotene in lipid droplets at later stages of growth might prevent feedback control and allow high levels of carotene to accumulate. Reviews of fungal carotenoids have been published by Valadon (1968) and several surveys of carotenoids in particular groups of fungi have been carried out by the Service de Phytochemie and Phytophysiologie in Lyon, France, e.g. by Arpin (1968)for discomycetes and by Fiasson (1968) for basidiomycetes. Singh (1975), investigating the storage materials of two aquatic hyphomycetes, namely Lemonniera aquatica and Articulospora tetracladia, found that their lipid reserve contained a high proportion of carotenoids. b. Sterol Growth Factors. Since the occurrence and function of fungal sterols have been thoroughly reviewed by Weete (19741, only certain aspects will be discussed here. Hess and Weber (1974) have provided a detailed survey of available information on the role of sterols in each of the different fungal classes, particularly certain oomycetes which lack the ability to synthesize sterols. The part played by sterols in fungal morphogenesis has also been discussed by Smith and Berry (1974). Previously, Hendrix ( 1970) had reviewed the role of sterols in growth and reproduction of fungi, and had pointed out that members of the Pythiaceae were unable to synthesize sterols which they required for asexual and sexual reproduction, although they could grow vegetatively without sterols. Some of these species must obtain their sterols by parasitic growth on living plant tissue. Hendrix (1975) studied uptake of cholesterol and sitosterol by various isolates of a range of species of Pythium and Phytophthora. Differences were found in their cholesterol uptake and ability to form sterol esters or a more polar metabolite, but there appeared to be no correlation between reproduction and sterol

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metabolism. Hendrix ( 1975) concluded that, under these conditions, any steroid hormone which might be produced would have been masked by other metabolites or by sterol autoxidation. Sterols, too, have been implicated in photomorphogenetic responses and pigmentation. Light induces formation of sterols and sporulation in Rhiropus arrhizus (Weete et al., 1973). Hendrix (1964) showed that both light and sterols were required for sporangia and zoospore production in Phytophthora sp. Ergosterol in the presence of light was necessary for development of yellow pigmentation, conidiophores and conidia in Phymatotrichum omnivorum (Baniecki and Bloss, 1969). A similar involvement of ergosterol was found in induction of conidiation in Stemphylium solani by ultraviolet radiation (Sproston and Setlow, 1968) and sporulation was prevented on medium containing a sterol inhibitor, even in the presence of ultraviolet radiation. Marked changes in contents of sterols and fatty acids have been observed during sporophore formation in Agaricus bisporus (Holtz and Schisler, 1972) where sterol becomes the principal neutral-lipid component although it is absent from vegetative mycelium. Sterol formation by rust fungi and by fungi involved in insect symbioses will be discussed in Section VI (p. 140). N o information appears to be available on sterols of powdery mildews and other obligate biotrophs.

8.

LIPID RESERVES I N M O R P H O G E N E S I S

Lipids are present on spore surfaces, in fungal spore walls (Bartnicki-Garcia, 19731, as membrane components and as nutrient reserves within the spores (Table 1, p. 50). Hess and Weber (1974) discussed the involvement of lipids in the physiology of fungal sporulation. Their review (Weber and Hess, 1974) and the reports of the Second International Fungal Spore Symposium (Weber and Hess, 1976) give a full account of ultrastructural and biochemical observations of many workers on the lipids of dormant and germinating spores, of the turnover of different lipids during germination, and of the metabolism of fatty acids and acetate units and their incorporation into various cell materials, including lipids. The occurrence of surface lipids, which presumably have protective and waterproofing properties, has been investigated by various workers (Table 7 ) and reviewed by Weete (1974). The presence of surface lipid on conidia of such leaf-surface fungi as species of Alternuria

P. J. BRENNAN AND D. M. LdSEL

122

TABLE 7. Some records of surface lipids on fungal spores Lipid on spore surface Rhizopw stoloniJer Neurospora crassa Botrytisfabae Alternaria tenuis Pithomyces chartarum Tilletia spp. Puccinia striqormis (urediospores)

Lipid-free surface Mucor rouxii Nectria galligena Erysiphe graminis Erysiphe cichoracearum Penicillium expansum Verticillium alboatrum Sphaerotheca mcularis

Reference Fisher et al. (1972) Fisher et aL(1972) Fisher et al. (1972) Fisher et al. (1972) Fisher et al. (1972) Fisher et al. (1972) McKeen et al. (1966) Bertaud et al. (1963) Weete et al. (1963) Jackson et al. (1973)

and Botrytis, rust urediospores and teliospores of smut fungi is not unexpected, but it is more difficult to explain the absence of surface lipid from spores of powdery mildews and Penicillium spp. The freeze-etch replica of an Erysiphe graminis conidium (Hess, in Weete, 1974) showed a wax-like surface pattern. Unwettable spore coats of soil fungi such as Penicillium spp. have been claimed to prevent washing down of spores in soils, so that they remain in upper horizons of the soil profile, in atmospheres of composition favourable for growth. Practically all fungal spores appear to contain lipid reserves which are mobilized to varying extents during germination. Particularly high levels are found in some cases, e.g. 27% in ascospores of Neurospora tetrasperma (Lingappa and Sussman, 1959) and 35% in teliospores of Tilletia controuersa (Trione and Ching, 19 7 1 ) . A single large lipid globule occupying the bulk of the oospore is a familiar feature of oomycetes and even provides a reliable taxonomic criterion, e.g. in the genus Achlya (Johnson, 1956). Among the relatively few fungal spores stated to have a low lipid content are asexual spores of zygomycetes (Weber and Hess, 19741, Fusarium spp. (Rambo and Bean, 19691, Phymatotrichum omniuorum (Gunasekaran et al., 197 4) and Alternuria spp. (Campbell, 1968). Some records of utilization and synthesis of lipids by fungal spores are listed in Table 8 . Zoospores of the lower fungi can often remain active for long periods without external nutrients, using lipid reserves during the freeswimming phase. In electron-microscope studies of zoospores of Phyto-

TABLE 8 . Changes in lipid during germination of fungal structures Decreasing Myxomycete spores Arcyria cinerea Dictyostelium discoideum Encysted zoospores Phytophthora palmivora Sporangiospores Rhzopus arrhizus (early germination, neutral lipids) . Cunninghamella elegans Conidia Aspergillus nidulans Aspergillusfumigatus Penicilliumoxalicum Elysi$he graminis hordei Sphaerotheca macularis Veerticiliumalboatrum Fusan'um solani Urediospores Puccinia graminis Urumycesphaseoli (first 20 minutes) Cronartiumfusforme Smut spores Ustilago muydis (neutral lipid and phospholipid) Basidiospores Ooidia

Increasing

Rhizopus arrhizw (later germination, polar lipid)

Reference Mims ( 197 1 ) Cotter et al. ( 1 969a)

V

Bimpongand Hickman (1975)

V,

Gunasekaran et al. (1972)

c,

I

<

g

Hawker et al. (1970) -n

Shepherd (1957) Campbell (197 1 ) Raj et al. (1970) Yanagita and Kogane (1963) Gottlieb and Ramachandran (1960) McKeen (1970) Mitchell and McKeen (1970) Walker and Thornberry ( 19 7 1 ) Cochrane et al. (1963)

C

0 -I

Urmycesphaseoli(sterols) Melampsora lini (lipid phosphate)

Staples and Wynn (1965) Langenbach and Knoche ( 19 7 1) Laseter et al. (1973) Lin et al. (1972) Jackson and Frear (1968)

Ustilago muydis (sterols, phospholipid, diglyceride)

Davidoff ( 1964)

Schizophyllum commune Coprinus lagopus

Aitken and Niederpreum (1970) Heintz and Niederpreum (1970)

AspergillusJlauus (phospholipid) Aspergtllus niger Penicillium atrouenetum (lipid doubles)

Uromycesphaseoli (phospholipid 5-10 hours)

z

c,

F

1

I? cn 0

;;I

: 0 v)

124

P. J. BRENNAN AND D. M. LOSEL

phthora parasiticu, Hemmes and Hohl ( 197 1) observed lipid droplets in the cytoplasm and vacuoles containing lamellar inclusions, which they designated liposomes. Bimpong and Hickman ( 1975) demonstrated the presence of lipid in lipid bodies in Phytophthora palmivoru zoospores, along with an unknown material in “crystalline vesicles”. The bulk of the cytoplasm was occupied by lipid bodies, “crystalline vesicles” and granular vesicles containing protein. Bimpong ( 1975) showed that acyl glycerides and free fatty acids provide the major source of energy during a six-hour motile period of these zoospores and a subsequent two-hour germination period of the cysts. Carbohydrates and protein decreased slightly in content during the motile period but increased significantly during germination. Glycogen provided the major reserves in sporangia of P. uythroseptica, but Bimpong ( 1975) comments that the lipids in zoospores provide a more efficient energy source, stored in more concentrated form than glycogen. In Koch’s studies on chytrid zoospores, summarized by Webster (19701, a lipoid body was the most conspicuous feature within the zoospore, its position and size varying from genus to genus. The blepharoblast at the base of the active flagellum seems to be attached to the lipoid body in some cases by a disk-like structure, which Chambers et al. (1967) suggested may be a photoreceptor sensitive to light concentrated by the lens effect of the lipoid body. Here, too, swimming may be prolonged for some hours, and appears to be at the expense d stored lipid. Zoospores of the closely related Blastocladiales also have abundant lipid in the periphery of the prominent nuclear cap of Allomyces sp. (Hill, 1969) and in the lipid sac, adjacent to the giant mitochondrion, of Blastocladiella zoospores. In their detailed review of metabolism, activity and morphogenesis of chytridiomycete spores, Cantino and Mills (1976) interpret this “side body” of Blastocladiella emrsonii zoospores as a complex symphiomicrobody consisting of microbodies and lipid bodies. Lipid appears also to be involved in the formation of zoospores of Synchytrium sp., the chytrid causing the destructive wart disease of potato. In resting spores of Synchytrium endobioticum, induced to germinate by leachate from susceptible plants, Sharma and Cammack ( 1976) observed a large lipid globule in the resting spore before the extrusion of a thin-walled hyaline vesicle, into which the spore contents flowed before differentiating into a single sporangium containing zoospores.

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

125

The changes in lipid metabolism accompanying ascus and ascospore morphogenesis in Saccharomyces cerevisiae were reviewed by Tingle et al. (1973). Lipid increases markedly up to the meiotic divisions and early differentiation of the ascospores, then again during spore maturation. Illingworth et al. (1973) found the lipid content of yeast ascospores increased to four times that of vegetative stationary-phase cells, due mainly to rapid synthesis of triacylglycerols and sterol esters and, to a lesser extent, phospholipid. The triacylglycerols and sterol esters appeared to be located in low-density structures similar to those found in liver cells. Henry and Halvorson (1973) reported that both haploid and diploid yeast cells synthesized phospholipid as well as neutral lipid in the first 12 hours on acetate-containing sporulation medium, the diploid cells accumulating a higher proportion of neutral lipid. In the second phase of lipid synthesis, from the 20th to the 25th hour, [l4Clacetate incorporation was mainly into neutral lipid. This second phase of synthesis occurred only in diploid sporulating cells, co-incident with differentiation of ascospore membranes and spore walls, into which some of the lipid material may have been incorporated. Since the phospholipid synthesized during yeast ascus differentiation was more than estimated to be required for membrane formation in the ascospores, Illingworth et al. (1973) suggested some may be present in membranes enclosing newly synthesized vesicles. These vesicles, which were electron- transparent in permanganate-fixed material, became arranged in the “prospore” wall region during ascospore development, possibly associated with deposition of wall materials. It was suggested that the increased proportion of unsaturated fatty acids found at this stage might give greater mobility to membranes of vesicles involved in transport. In the relatively few observations which have been made on the various types of complex fructifications of the higher fungi, lipid reserves have been found not only in spores but also in accessory structures. Abundant lipid is seen in cells of the wall structures surrounding the pycnium and aecium of the rust Puccinia poarum imbedded in Tussdago leaves (Lose1 and Lewis, 1974). Mature cleistocarps of the powdery mildew, Sphaerotheca mom-uvae, had three or four layers of thin-walled cells, within the outer wall layers, with contents giving a strong fat reaction with Sudan IV (Jackson and Wheeler, 1974). During differentiation of ascospores within the ascus, lipid, initially present throughout the cytoplasm, became organized into one or two lipid

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P. J. BRENNAN AND D. M. LOSEL

bodies. Substantial amounts of fat were present in undischarged spores but distributed in very small bodies throughout the cytoplasm of discharged spores. Lipid was probably not directly involved in the sporedischarge mechanism since the swelling of the ascus, which bursts the cleistothecium allowing discharge of the ascospores, is associated with a decrease in the glycogen content. Neurospora spp. reproduce asexually by pink, carotenoid-containing conidia and sexually by ascospores arising from fertilization of a female organ, the ascogonium, by microconidia from another culture of compatible mating type. The ascogonia are normally ensheathed in dark, melanized hyphae, forming protoperithecia which, after fertilization, develop into the dark, flask-shaped perithecia, enclosing the developing asci and ascospores. Control of these developmental stages, which has been investigated by various workers, was reviewed by Turian ( 1969). Growth on acetate-containing medium induces production of conidia without protoperithecia (Turian, 196la, b). Formation of protoperithecia requires operation of the tricarboxylic-acid cycle (Turian, 1962) and involves tyrosine metabolism and melanin formation. On acetate-containing medium, Neurospora spp. formed ascogonia, which were rich in orange lipid drops (Turian, 19751, but which were unable to proceed with the development of perithecia and accompanying formation of melanin-type pigment. The proportion of bound to free lipid always remained higher in differentiating cells. The perithecium of Neurospora lanceolata had two outermost layers richly provided with lipid granules and other storage materials (Hohl and Streit, 1975) and also containing microbodies, some of which were involved in lipid degradation. A possible role of the perithecial-wall tissue in storing food materials for future use, besides the obvious function of protection of the asci, was suggested. During the development of the asci, numerous small lipid inclusions, which initially fill the cytoplasm, fuse and become concentrated around the evenly spaced nuclei within the confines of the future spores. Holh and Streit (1975) considered that this was brought about by a highly fenestrated " ascus vesicle", which sweeps lipid granules, and possibly other components, towards the nuclei. Parts of the cytoplasm of the mature asci were found filled with granular material, possibly glycogen, which might function in ascospore discharge by providing a high concentration of osmotically active material within the ascus.

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

127

Observations on functionally analogous asexual structures show similar lipid reserves. The acervulus stroma produced by Colletotrichum lindemuthianum in host tissues consists of hyphae with few vacuoles and a high content of lipid (Mercer et al., 1975). The conidia contain globules of neutral lipid as well as a subspherical inclusion with denser contents bound by a single membrane, which was thought to resemble “phospholipid vesicles”. Both types of inclusion were present in germ tubes and young hyphae where the large lipid droplets were less densely stained than in ungerminated spores. The presence of smaller amounts of lipid in the dense cytoplasm of the appressorium formed at the point of penetration of the host may be associated with meeting the requirements for energy and materials for invading the host tissues. In sclerotia of Colletotrichum coccodes (Campbell and Griffiths, 19741, conversion of dried sclerotia into acervuli involves a remarkable formation of secondary hyphae from areas of activity within individual sclerotial cells. The energy for this transformation and the materials for growth of the secondary hyphae, through intervening sclerotial tissue to the exterior, were considered to come from lipid reserves in the dormant sclerotial cells. Cytological and ultrastructural investigations of morphogenesis in basidiomycete fructifications have not so far been particularly concerned with the occurrence of lipids, although significant differences in lipid content between vegetative mycelium and fructifications and within the fructification have been noted. Increased amounts of linoleic acid and phospholipid were found in sporophores of Tricholoma sp. (Leegwater et al., 1962) and the fatty-acid composition changed from stipe to pileus in various basidiomycetes, the fatty-acid composition of the stipe being intermediate between that of the mycelium and that of the pileus or cap (Shaw, 1966). The ratio of neutral lipid to phospholipid in fruiting bodies varies between 1 :2 and 2 : 1 (Shaw, 1966) but, in vegetative mycelium from a 12 day-old culture of Tricholoma nudum, this ratio rose to 27 : 1. The very high linoleic acid content of the polar-lipid fraction of Agaricus bisporus sporophores (Holtz and Schisler, 197 1) and the stimulation of mushroom production by oleic acid and linoleic acid (Wardle and Schisler, 19691, are compatible with the suggestion of Hughes (1962) that linoleic acid may have a special role in the fruiting mechanism of A. bisporus. Lehrian et al. (1976)observed that addition of low concentrations of either linoleic acid or sodium acetate to complex media

128

P. J. BRENNAN AND D. M. LOSEL

stimulated growth of A . bisporus by 30%.The similar patterns of incorporation of I4C from either sodium 1,2-[l4C1-acetateor [ 1-14Cl-linoleic acid led him to propose that linoleate is degraded to acetate before being incorporated into mycelial lipid. These workers suggested that sodium acetate acts as a micronutrient in stimulating a pathway leading to synthesis of linoleate. Electron-microscope observations on the development of the basidia of A . bisporus (Thielke, 19671 showed that lipid drops appeared in the basidia following nuclear fusion and meiosis. Vogel and Weaver (1972) found the cytoplasm of basidial cells densely filled with ribosomes, glycogen and lipid drops, with numerous mitochondria in the apical portion. Lipid droplets surrounded by glycogen granules were first to enter the spores, followed by cytoplasm with ribosomes and mitochondria. As the spore reached its maximum size, its attachment to the sterigma became blocked with lipid droplets and glycogen. With separation from the sterigma, the spore wall became very thick, compressing the inner constituents. The dominant feature of the cytoplasm was a group of lipid droplets in the inner region. The mitochondria were greatly compressed and rarely visualized. Successive workers have commented on the impermeability of this remarkably thick wall of the mushroom spore towards fixatives and other reagents required for electron microscopy (Manocha, 1965; O’Sullivan, 1969). Greuter and Rast (1975)obtained very fine electron micrographs after slightly cracking the spores with glass beads in a-cell mill. Large lipid bodies which occupied much of the cytoplasm were closely associated with organelles having a uniform matrix bounded by a single unit membrane, which were interpreted as glyoxysomes, and with mitochondria and glycogen deposits. This assemblage suggests an efficient system for metabolism of the spore reserves in germination. Besides acting as energy and carbon sources, a special function has been proposed for the lipids of powdery mildew conidia, well known for their ability to germinate under conditions of low atmospheric humidity. McKeen (1970) suggested that spore lipids could contribute to the water requirements of the germinating spores, since oxidation of hydrogen in respiration of lipids could yield considerable amounts of water. A detailed examination of the lipid reserves of Erysiphe graminis by combined gas-liquid chromatography-mass spectrometry (Johnson el al., 1976) identified spore hydrocarbons, naturally occurring methyl esters of fatty acids and free fatty acids. I t was thought that the fattyacid esters could be parts of membrane-lipid complexes or could be

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

129

energy reserves in equilibrium with the free fatty acids. This group of workers considered that the hydrocarbons and the unusually high proportion of fatty acids of chain length greater than C,, might be related to the ability of the powdery mildew spores to retain relatively large amounts of water in dry atmospheres. Besides the many reports of the utilization of lipid materials by spores during dormancy and germination, a smaller number of studies have recorded increasing levels of lipid (Table 8, p. 123)at least after the initial stages of germination. Particularly active synthesis of lipids during germination has been found in certain basidiomycetes, e.g. Schizophyllum commune shows a three-fold increase in total lipids in 12-h germlings on glucose-asparagine broth (Aitken and Niederpreum, 1970) and there is a three-fold increase during germination of oidia of Coprinus lagopus (Heintz and Niederpreum, 1970). In Schizophyllum commune, both phase-contrast and electron-microscope examination showed many lipid vacuoles in germlings and more abundant endoplasmic reticulum than in dormant spores. In germinating spores of Penicillium atroveneturn, the lipid content doubles during germination (Gottlieb and Ramachandran, 19601, lipid components, including ergosterol, increasing most markedly about the time of transition to vegetative growth. From Table 8 (p. 123), i t appears that synthesis of lipids during germination has been observed mostly in the higher fungi, particularly basidiomycetes, although in many cases an initial decline in storage lipids masks the synthetic processes. Separate analysis of individual lipid classes, or at least of polar and neutral lipid fractions, is more informative but has been attempted in relatively few studies, e.g. Gunasekaran et al. (1972). In some cases, the nature of the lipid consumed or synthesized during germination has been more precisely identified. In Aspergillus niger (Nishi, 1961) and A. nidulans (Shepherd, 19571, it has been claimed that the energy requirements in germination are met by polyphosphates and phospholipids. Germinating conidia of A. Javus incorporate acetate into phospholipid. Gunasekaran et al. ( 1972) found neutral lipids, particularly free fatty acids, decreased in the early phases of germination of Rhizopus arrhizus spores while contents of polar lipids increased. Langenbach and Knoche ( 197 1) detected a rapid fall in the phospholipid content of Uromycesphmeoli urediospores during the first 20 min of germination followed by phospholipid synthesis for five to ten hours, while Lin et al. (1972)demonstrated active sterol synthesis by U. phaseoli urediospores during the germination period. Teliospores of

P. J. BRENNAN AND D. M. LOSEL

130

the corn-smut fungus, Ustilago maydis, consumed triglycerides, free fatty acids, diglycerides and phospholipids during germination (Davidoff, 1964) at the same time incorporating [14C1-acetateinto diglycerides, phospholipids and sterols. Remarkably little information is available on lipid changes during formation and germination of ascospores, apart from the classical investigations of Sussman and his coworkers on Neurospora crassa (Sussman, 1966)and the studies on Sacch. cereuisiae mentioned earlier in this section. Similarly, although rust urediospores have been intensively studied, relatively little is known of the lipid metabolism of germinating basidiospores of Homobasidiomycetes. To some extent, these gaps probably reflect dormancy problems and the greater difficulty of investigating spores with slow or variable, non-synchronous germination.

C.

LIPID RESERVES AND SECONDARY METABOLITES

Triacylglycerols of fungi are the major constituents of the oil droplets suspended in the cytoplasm which are generally regarded as a store of utilizable energy and anabolic precursors for growth and reproduction. The fairly constant occurrence of such reserves in reproductive structures has been discussed in the previous section (p. 122). Depending on strain characteristics, phase of growth and environmental conditions, vegetative hyphae may also accumulate substantial amounts of lipid. Most mycelial species contain 6-9% lipid when grow? under favourable conditions, and most yeasts species contain 7-15% lipid. I n cases whZe the fat content rises above these median values, such as vegetative hyphae of some strains, particularly when growth is slowing down or in the “fat yeasts”, most of the excess lipid is triglyceride which is deposited in the form offat globules (liposomes). In yeasts of the genera Candida, Hansenula and Rhodotorula, triglyceride accounts for about 80% of total lipid (Thorpe and Ratledge, 1972). In submerged mycelium of the basidiomycete Tricholoma nudurn, 92% of the weight of all lipids is triglyceride (Leegwater et al., 1962). Triglycerides are again the principal lipids in Phycomyces blakesleeanus, Lipornyces lipoferus, Glomerella cingulata and Coprinus cornatus (Jack, 1965). Five different classes of triglyceride were demonstrated in G. cingulata. One group contained saturated and mono-unsaturated fatty acids. Another group contained saturated and both mono- and di-

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

131

unsaturated fatty acids. Two additional groups had in addition triunsaturated fatty acids. Over 50% of the triglycerides of a number of yeasts are of the 1,3-disaturated-2-mono-unsaturated type (Thorpe and Ratledge, 1972). Fatty acids occur in fungi mainly as constituents of acyl glycerides, phospholipids and various complex lipids. Significant amounts of free fatty acids may also be present. Since the distribution and metabolism of fungal fatty acids have been reviewed by Shaw (1966) and Erwin (1973) and Weete (19741, and a brief account was given by Brennan et al. ( 19751, a discussion of fatty acids will be omitted from the present review, except where particularly relevant to other topics discussed. The functional dividing line between certain secondary metabolites of fungi and the normal lipid reserves of reproductive structures or mycelium is not clear. Both seem to undergo substantial depletion on ageing or when insufficient carbon source nutrient is present. Of the lipid secondary metabolites, the sphingosine-type bases have received the greatest attention. A number of these are produced in large quantities by Hansenula aferri and excreted into the extracellular medium in the partially or fully acetylated forms (Table 5 , p. 80). Tetra-acetylphytosphingosine is the major type comprising about 90% of the mixture of acetylated long-chain bases. Stodola and Wickerham ( 1960) showed that the base in this lipid was a C,,-phytosphingosine and not a C,,-phytosphingosine of the type found in Sacch. cerevisiae (Prostenik and Stanacev, 1958; Oda and Kamiya, 1958). A minor component accompanying the tetra-acetylphytosphingosine was a triacetyl-C,,dihydrosphingosine. Barenholz et al. (197 1 ) compared 25 strains of the genus Hansenula for their levels of extracellular acylated long-chain bases and found that they varied considerably in the amount of sphingolipid produced. Low producers synthesized only 2-5 pmoles of base per litre of growth medium compared to 120-260 pmoles for high producers. Earlier Maister et al. (1962) reported as much as 175 g of the crude acetylated base from a single culture of H . cferri in a 750-litre aerated tank grown in a 4% glucose solution. For each gram of glucose consumed by the mating-type strain of H . clferrz, 5 mg of tetra-acetylphytosphingosine were formed; for each gram of yeast solids produced, 15 mg of tetra-acetylphytosphingosine were synthesized. A 750litre pilot-plant run yielded 175 g of crude tetra-acetylphytosphingosine, which was readily obtained by hexane extraction of centrifuged cells. Tetra-acetylphytosphingosine was formed only during dis-

132

P. J. BRENNAN AND D. M. LOSEL

similation of glucose; only when the glucose content of the medium was exhausted did formation of tetra-acetylphytosphingosine stop. The acetylated long-chain bases are generally regarded as secondary metabolites, i.e. they are compounds having a limited distribution, being produced in large quantities by one or a few organisms and having no known function in the cell. However, it has been observed that, in the early stages of propagation of H . ciJerri F-60-10 mating type, the cells with their buds occur singly in medum. After 12 hours, refractile globules of tetra-acetylphytosphingosine appear on surfaces of yeast cells, there is an increase in the tetra-acetylphytosphingosine content of cultures and the cells begin to cluster. After 48 h, the globules crystallize and the hydrophobic cells and crystals flocculate in large masses. Therefore, besides possibly acting as a reservoir of utilizable energy, it appears that the tetra-acetylphytosphingosine is responsible for the tendency of H . czjerri to form pellicles in liquid media. The sophorosides (Table 4, p. 76) and many of the related hydroxyacid glycosides are also regarded as secondary metabolites, excreted into the growth medium in vast quantities. For instance 0.5-1 .O g of Ac,Glc,HDA (see p. 107 1 are often found per litre of culture medium of Candida bogoriensis. Generally, the glycolipid attaches itself to the yeast cells as crystals. Suzuki et al. (1969) noted that the sophorosides are produced by hydrocarbon-utilizing yeasts ( Torulopsis spp.) and that they participate in dispersion of n-paraffins and water-soluble nutrients. Thus, the detergent properties of the sophorosides are used by the yeasts for absorption of lipids. Ustilagic acids, which consist of partially esterified P-cellobiosides glycosidically linked in the P-form to the hydroxyl groups of longchain fatty acids, found in the corn-smut fungus Ustilago maydis and the loose smut of wheat, U. nudu (Table 4, p. 761, are also secondary metabolites. So also are the partly acylated 4-O-P-~-mannopyranosyl-D-erythritols of the smut fungi (Table 4, p. 761, which are often found associated with the P-cellobiosides in extracellular oil droplets. The acetylated hexitol and pentitol esters of S-~-hydroxypalmiticand S-~-hydroxystearicacid (Table 4, p. 76) found in the extracellular fluid of several species of Rhodotorula (in large amounts, e.g. 1-2 g per litre of medium in Rhodotorula graminis) are also secondary metabolites. Some of these glycolipid secondary metabolites are clearly reserve materials which disappear quickly in ageing aerated cultures (Ruinen

133

PHYSIOLOGY OF FUNGAL LIPIDS: SELECTED TOPICS

and Deinema, 1964). Esders and Light (1972a) indicated the probable sequence of this degradation in C. bogoriensis : Ac,Glc,HDA

-+

AcGlc,HDA+ Glc,HDA +. Glc + HDA

With the onset of the stationary phase of growth, Ac,Glc,HDA gradually disappeared while AcGlc,HDA and Glc,HDA accumulated. Since, in older cultures, the glycolipids disappear completely from the medium, such cultures probably secrete glycosidic enzymes after complete cleavage of the acetyl groups, releasing glucose and hydroxy fatty acids which are then available for glycolysis and P-oxidation, respectively. D.

LIPID METABOLISM I N MORPHOGENESIS

1. L$ases

As in other fat-storing tissues, lipases are to be expected in fungal structures where acyl lipid reserves are being mobilized. There have, however, been studies in relatively few fungal systems. A comparison of esterase function in Candida lipolytica, Aspergillus niger and a yeast-like fungus by Lloyd et al. (1970) showed lipases were always present in mycelial extracts during conidiation, but were not detectable in mycelial cultures which would ultimately form conidia or in mycelium of sterile cultures. A further study by the Glasgow group (Lloyd et al., 1972) on lipase activity during development of A. niger showed a low basic level during vegetative growth of mycelium. A marked increase in lipase activity was found in the developing conidiophore tip, and persisted during formation of vesicle and phialides indicating utilization of lipids during conidium formation. Although many studies have been made of lipid metabolism during fungal spore germination, little information appears to be available on the lipases of dormant or germinating spores. Knoche and Horner (1970) investigated the activity of a lipase from Puccinia graminis urediospores. The lipases of fungi have been examined in some detail in a limited number of species. They catalyse the following reactions:

triacylglycerols

a-lipases

a, B and a',B-diacylglycerols p-mono-acyclglycerols

a-lipases

0-lipases

glycerol

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P. J. BRENNAN AND D. M. LOSEL

Alford et al. ( 1964)examined several representative fungi which were active producers of lipases. Most of the enzymes exhibited distinctive positional specificity; the majority of them preferentially cleaved the fatty acid in the 1 (a) position. A lipase from Mucor jauanicus preferentially hydrolyses ester bonds at the a and a' positions (Ogiso and Sugiura, 197 1) and shows additional preference for triglycerides containing oleic- and linoleic acid residues. The extensively examined lipase from Rhizopus arrhizus has similar specificity (Semeriva et al., 1967). The lipase of Geotrichum candidum (Alford et al., 1964) shows an unusual specificity; it cleaves Ag mono-unsaturated fatty acids regardless of their location. The lipase from A . niger showed no such specificity and cleaved triglycerides at the a, ,B, and a' positions. 2 . Phospholipases Where contents of phospholipids decline during germination, phospholipase activity must be present but, so far, such enzymes have been studied in detail only in rust urediospores and yeast. Tseng and Bateman ( 1968)compared the phosphatidase activity of various phytopathogens including Sclerotium rolfsii, Botrytis cinerea, and the bacterium Erwinza carotivora, all of which caused rapid soft rots of host tissue. All showed higher phospholipase activity than Thielauiopsis basicola, Rhizoctonia solani and E . solani, which are associated with slower, less moist rots. A phosphatidase produced by S. rolfsii in bean hypocotyls was characterized. Tseng and Bateman ( 1968) suggested that the phospholipases produced by these pathogens may increase the permeability of host tissues. Hoppe and Heitefuss (197413)showed that rust infection of bean leaves was associated with increased phosphatidase activity and increased permeability. More detailed examination of phospholipases has been carried out by Angus and Lester ( 197 2 , 197 5 1. They found Saccharomyces cereviseae, Sacch. carlsbergzensis, Kloeckera apiculata and Neurospora crassa are all cap able of very active degradation of phosphatidylinositol to glycerylphosphorylinositol, although it is debatable if the primary purpose of the de-acylation is the supply of metabolizable fatty acids. Lack of glucose decreases the formation of lysophosphatidylinositol from phosphatidylinositol, but enhances further degradation of glycerylphosphorylinositol. Formation of glycerylphosphorylinositol occurs under normal growth conditions and its level can be as high as 25% of cellular phosphatidylinositol. The site of the de-acylation is the cell mem-

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brane, and glycerylphosphorylinositol is primarily extracellular in location. Extracellular glycerylphosphorylcholine and glycerylphosphorylethanolamine accumulate more slowly than does glycerylphosphorylinositol.

-

3. Glyoxylate-Cycle Enzymes

Metabolism of lipid reserves in many fungal structures is associated with activity of the glyoxylate-cycle enzymes, isocitrate lyase and malate synthetase, by which acetate from reserve lipids or external substrates is metabolized and appears in other cell components such as sugars and amino acids. This has been demonstrated in Neurospora conidia (Zalokar, 1959a, b), in conidia of Penicillium oxalicum (Gottlieb and Caltrider, 1963) and rust uredospores (Caltrider et al., 1963), and is likely to be a general feature of dormant or germinating spores which metabolize endogenous lipids. Changes in glyoxylate-cycle activity in Phytophthora erythroseptica zoospores during the motile period, when acylglycerols and free fatty acids were being utilized, were studied by Bimpong (1975). The glyoxylate cycle appeared to be more important in providing metabolites during the motile phase than during germination of encysted zoospores. In various fungal systems, phases of morphogenesis are controlled by switching between tricarboxylic acid-cycle activity and the glyoxylate path, the key enzyme being isocritrate lyase, which is influenced by some of the reproductive and germination triggers already discussed. Transfer to a medium containing acetate may profoundly affect growth and morphogenesis by derepressing isocitrate lyase. Such effects have been studied in Blastocladiella sp., yeast and Neurospora sp. The metabolic control of morphogenesis of Blastocladiella sp. was elucidated by Cantino and Turian ( 1959). Whether encysted zoospores develop into thick-walled, resistant sporangia, with their characteristic carotenoid pigment and lipid content, rather than into ordinary colourless sporangia, is determined by the concentration of bicarbonate in the medium. This can arise by accumulation of carbon dioxide in ageing cultures, but is readily triggered by transfer to medium containing bicarbonate. Reductive carboxylation of a-oxoglutarate blocks the tricarboxylic acid cycle and induces isocitrate lyase activity. Transamination of the glyoxylate formed to yield glycine is required for production of amino acids during acetate metabolism.

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P. J. BRENNAN AND D. M. L ~ S E L

The change from vegetative budding growth of Sacch. cereuisiae to ascus formation, meiosis and development of ascospores is brought about in media lacking nitrogen and having a non-fermentable carbon source (Miller, 1963). Sporulation is normally induced by transferring yeast cells from medium with a relatively high nutrient concentration to sporulation medium containing acetate as the sole carbon source. Most yeasts are unable to utilize acetate during the logarithmic phase of growth. Sporulation depends on the development of the capacity to utilize acetate via the glyoxylate cycle which is induced by the sporulation medium. Acceleration of the rate of formation of asci by addition of glyoxylate to the acetate-containing medium was demonstrated by Bettelheim and Gay (1963), who also found that glyoxylate could replace the carbon dioxide requirement for yeast sporulation, previously noted by Adams and Miller (1954). Tingle et al. (1973), reviewing differentiation of yeasts, quote the observations of Darland ( 1969) which show that even brief exposure to acetate-containing sporulation medium for 5-10 minutes, followed by transfer to water, can trigger sporulation and that, after commitment to sporulation metabolism, the process is no longer inhibited by glucose. Isocitrate lyase activity increases prior to conidiation in Aspergillus niger (Galbraith and Smith, 1969) and in Neurosporu crassa (Turian and Combkpine, 1963) but with little malate synthetase activity. In both cases, acetate-containing medium induces activity of the glyoxylate path and increases production of conidia. Enzyme changes in differentiation of Aspergillus sp. have been reviewed by Smith and Anderson ( 1973).The relative activities of glyoxylate-cycle enzymes and tricarboxylic acid-cycle enzymes have been correlated with stages of morphogenesis in synchronous cultures of Aspergillus niger (Ng et al., 1973). From the low level of malate synthetase in these conidiating systems, Ng et al. (1973)concluded that glyoxylate, resulting from isocitrate lyase activity, is transaminated to form glycine. Again, as in yeast ascospore formation, it may be glyoxylate rather than isocitrate lyase activity which is required, since addition of glyoxylate to the medium induced heavy sporulation. Whether mycelium from Neurospora sp. produces conidia or protoperithecia is largely determined by whether metabolism is by the glyoxylate path or the tricarboxylic acid cycle (Turian, 1962). Inhibition of the tricarboxylic acid cycle by malonate or induction of the glyoxylate-cycle enzymes by acetate results in production of conidia

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without protoperithecia, formation of which involves also tyrosine metabolism and formation of melanin in hyphae sheathing the female organ, the ascogonium (Hirsch, 1954). From information available on species of Aspergillus, Neurospora, Blustocludiellu and yeast, Galbraith and Smith ( 1969) suggested that reductive carboxylation of a-oxoglutarate to isocitrate, followed by isocitrate lyase derepression, may be of universal importance in fungal morphogenesis. There would, however, appear to be a need to examine this question in a wider range of fungi. Cotter et ul. (196913) failed to find evidence that sustained derepression of isocitrate lyase synthesis was linked with normal fruiting in Schizophyllum commune or its inhibition by carbon dioxide, although the possibility of transient derepression being effective was not ruled out. I n various other systems where carbon dioxide influences morphogenesis, information is not available on whether a switching from tricarboxylic acid-cycle metabolism to glyoxylate-cycle metabolism is involved or the extent to which other aspects of carbon-dioxide fixation and acid metabolism may be important. In the cultivated mushroom, Aguricus bisporus, it has long been known that accumulation of carbon dioxide inhibits fruiting (Lambert, 1933; Tschierpe, 1959). An investigation by Long and Jacobs (1969) of growth in air streams with accurately controlled low levels 'of carbon dioxide revealed a hormonal role of carbon dioxide at concentrations from 340 to 1000 p.p.m. in inducing the change from normal growth of A . bisporus mycelium to formation of mycelial strands and fructification initials in unsterilized soil. Concentrations of carbon dioxide favourable to fruiting were associated with retardation of vegetative growth but, above this, vegetative growth increased at concentrations from 1000 to 6700 p.p.m. Below the sporophore-inducing concentrations, growth was proportional to carbon dioxide concentration from 0 to 104 p.p.m. S o far, there is no information concerning the glyoxylate-cycle path in either morphogenesis of fruiting structures or in spore germination of A . bisporus. Investigation of carboxylation reactions of A. bisporus sporophore tissue (Rast and Bachofen, 1967; Bachofen and Rast, 1968) indicated that pyruvate and phospho-en01 pyruvate functioned as carbon dioxide acceptors. Obviously there is a need for much more information concerning the involvement of glyoxylate metabolism and carbon dioxide-fixation processes in differentiation of other fungi besides the few so far investigated.

4

w m

P

TABLE 9. Lipids in biotrophic fungi and host tissues

!-

Host-parasite system

Club root Plasmodiophora brassicae in cabbage hypocotyl Wart disease ofpotato Synchytrium endobioticum o n potato Downy mildew Bremia lactucae o n lettuce White rust Albugo candida Powdery mildew Erysiphe graminis Erysiphe graminis Erysijhe cichoracearum

Sphaerotheca mrs-uvae Rusts Sugar beet rust

Location

Reference

W II]

rn 2

z D 2

Lipid globules in plasmodium

Williams et al. ( 1968)

Lipid in resting spore

Sharma and Cammack (1976)

P

Large lipid drops in oosphere, periplasm rich in small lipid droplets Single lipid drop occupying most of oospore

Tommerup et al. (1974)

5

Lipid i n fungal haustorium

Berlin and Bowen (1964)

Mycelium, older haustoria and conidia Haustorium Haustorium Lipid drops in ascus, ascospore and cleistocarp wall

McKeen et al. (1966) Bracker (1968) McKeen et al. (1966) Stavely et al. (1969) Jackson and Wheeler (1974)

Lipid drops in epidermis in vicinity of infection

Schmidt (1932)

i 0 r-

0 v)

rn r

Pucciniapoarum (aecial stage on Tussilago) Lipid globules between chloroplast membranes Mesophyll cells, older hyphae, pycnium, aecium and aeciospores Mesophyll cells, older hyphae, haustoria and Pucciniapoarum (uredial stage on Poa) urediospores Plastoglobuli in host chloroplasts Melampsora h i on flax Lipid bodies in intercellular hyphae, older haustoria Puccinia helianthi on sunflower and axenic mycelium Lipid in haustorium Uromycesphaseoli on Phaseolus appendiculatus Lipid in host cells Cronartium ribicola on Pinus tissue culture Smuts Abundant lipid in dormant spores and germ tubes Ustilago hordei Tilletia controversa Tilletia caries Mycorrhiza Oil vacuoles occupying much of resting spore Endogone Lipid in root tissue Endogone in mycorrhizal roots Lipid in root cortex and fungus Endogenous mycorrhiza of ferns Lipid in arbuscules and intracellular hyphae Glomus mossei vesicular-arbuscular mycorrhiza on Allium cepa (onion) Lichen Plastoglobuli in lichenized alga cells Ramalina muciformis with Trebouxia as phycobiont

Orcival(1968) Losel and Lewis (1974) Losel (1978) Coffey et al. ( 197 Za, b) W

I

Hardwick et al. (1970)

< v,

Robb et al. (1975)

c)

g <

% Hess andweber (1976)

-n

C

z c)

F

Mosse (1970) Mosse (1973)

Cooper (1976) Cox and Sanders (1974) Peveling(l973)

1

n0 '" v)

rn

P. J. BRENNAN AND D. M. LOSEL

140

VI. Role of Lipid in Fungus-Host Relationships

Although the greater proportion of fungi are saprotrophs, some of which may become facultative but relatively unspecialized parasites of plant or animal tissues, a large number of species exhibit biotrophic growth in association with living host cells, with which they may establish a prolonged and specialized relationship, either mutualistic or pathogenic. Such associations, which include economically important pathogens as well as mycorrhizae, lichens and certain fungusinsect symbioses, are of considerable interest in physiological studies of the movement of materials between host and parasite, but have been investigated more frequently with respect to their carbohydrate and nitrogen metabolism (Shaw, 1963; Smith et al., 1969; Scott, 1972) than their lipid metabolism. The lipids of such biotrophic fungi are likely to be of primary importance as energy reserves and as membrane components concerned in the permeability of host-parasite interfaces. A.

FUNGAL ASSOCIATIONS WITH PLANT TISSUES

1 . Microscopic Obserwations

A substantial but scattered body of observations has accumulated on lipid inclusions both in plant tissue infected with biotrophic fungi and in hyphae or spores of the fungal partner. Some of these reports are listed in Table 9 . Striking differences have been observed in the distribution of stainable lipid in the rust Pucciniapoarum and leaf tissue of its two hosts, Tussilagof a f a r a (Losel and Lewis, 1974) and Poapratensis (Losel, 197 8) during the progress of infection. Haustoria (specialized short fungal branches which penetrate host cells) are rare in the aecial host, Tussilago sp., but a dense fungal plectenchyma fills the intercellular spaces of the pustule, closely investing all cell surfaces within the mesophyll. In the early phases only, oil drops are found in host cells in the vicinity of the developing intercellular mycelium. As the aecial pustule develops, oil drops become abundant in the intercellular fungal tissue, from which they disappear later, when the stainable lipid is concentrated in aeciospores and in the cells of the fungal peridium surrounding the aecial cup. In Poa sp., on the other hand, almost all cells in the urediosorus area are penetrated by haustoria, with which are closely associated the enlarged host nucleus and one or more conspicuous oil drops (Losel, 1978). On this host, throughout growth

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on the fungus, oil droplets are abundant in host cells in and adjacent to the urediosorus and occur in the intercellular hyphae only relatively late, when urediospore production is commencing. Thus the situation in infected Poa sp. and the early stages of infection on Tussilago sp. resemble the other rust-infected tissues described in Table 9 (p. 1381, in responding to the presence of fungal hyphae by production of lipid bodies.. Heath ( 1974) demonstrated similarities between such changes in the early stages of infection of cowpea leaves by Uromycesphaseoliand those in naturally senescent tissue, and between later stages of U .phaseoli infection and development of chromoplasts in certain ripening fruit. She provided evidence that ethylene released by infected cells could induce these chloroplast changes. Light-microscope obsevations of oil droplets in roots infected by vesicular arbuscular mycorrhiza have been recorded by several workers (Table 9, p. 138).Electron micrographs produced by Cox and Sanders (1974) showed lipid droplets in arbuscules and intercellular hyphae. Mosse (1973) suggested that fat globules pass via the arbuscules from the fungus into the host cytoplasm.

2. Spore Lipids of Biotrophic Fungi Because of the problems inherent in studying metabolic activities of vegetative stages of fungi growing in their host tissue, much of the information available on the lipid composition of biotrophic fungi concerns the more accessible spore stages. Much of the material derived from host cells parasitized by fungi must be transferred to spore-lipid inclusions which, from the time of the earliest microscopic observations on fungi, have been generally accepted as a major nutrient reserve for viability and germination. Lipid may contribute as much as 20% of the dry weight of rust spores (Tulloch and Ledingham, 1962). Abundant spherical lipid bodies were observed in electron micrographs of Erysiphe cichoracearum conidia (McKeen et al., 19661, usually lying in the outer regions of the cytoplasm near the-wall but occasionally projecting into vacuoles. Behenic acid ( C z z : oaccounted ) for 42% of the spore oil in another powdery mildew, Sphaerotheca humuli (Tulloch and Ledingham, 1960) and was a prominent component of P.poarum aeciospores and of aecial pustules on Tussilago (D. M. Losel, unpublished observations).

P. J. BRENNAN AND D. M. LOSEL

142

Extensive investigations of the spore oil of rust fungi carried out at The Canadian Prairie Regional Laboratory, Saskatchewan (Tulloch and Ledingham, 1960, 1962, 1964; Tulloch, 1964) showed general similarities between different spore forms of individual species of Puccinia, but it was not clear to what extent host and rust lipid metabolism are related. Oil composition differed in rusts of the same genus on host plants belonging to different families. Much information on the lipids of rust spores and their role in germination is included in the review of rust spore-germination physiology by Staples and Wynn (1965) and in Brennan et al. (19751, together with a discussion of stimulation of urediospore germination by short-chain fatty acids (Farkas and Ledingham, 1959; Reisener et al., 1963). More recent accounts of the lipids and physiology of teliomycete spores are contained in the reviews by Hess and Weber (1976) and Reisener (1976). Hess and Weber (1976) include a comprehensive treatment of the ultrastructure and metabolism of resting and germinating spores of Tilletia spp. As well as the feeding experiments which have been carried out with rust spores in relation to the physiology of their spore germination, other metabolic activities of rust spores have been examined in order to obtain some indication of the metabolic potential of the biotrophic mycelium. Lipase preparations from Puccinia graminis urediospores were investigated by Knoche and Horner (1970). Synthesis of sterols from acetate, mevalonate and methionine has been demonstrated in TABLE 10. Sterols from rust urediospores ~

Sterol Stigmasten-3p -01 A7-Stigmasten-3P-ol

A5*’ - S ti gmastadienol Stigmasterol 28-Isofucosterol p-Sitosterol Campesterol Cholesterol 22-Dihydroergosterol Fungis terol

Fungus Uromyces phaseoli Uromycesphaseoli Melampsora lini Uromycesphaseoli Melampsora lini Puccinia graminis Melampsora lini Uromycesphaseoli Uromycesphaseoli Uromycesphaseoli Uromycesphaseoli Puccinia graminis Puccinia graminis Puccinia graminis Puccinia graminis

~

~

~~~

Reference Lin and Knoche (1974) Hoppe and Heitefuss (1975b) Jackson and Frear (1968) Hoppe and Heitefuss (1975b) Jackson and Frear (1968) Nowak et al. ( 1972) Jackson and Frear (1968) Lin et al. (1972) Lin and Knoche (1974) Linetal. (1972) Linetal. (1972) Nowak et al. ( 1972) Nowak et al. (1972) Miller et al. ( 1967 Hougen et al. (1958)

PHYSIOLOGY

OF FUNGAL LIPIDS: SELECTED TOPICS

143

Uromyces phaseoli urediosphores (Lin et al., 1972) and by cell-free preparations from urediospores (Lin and Knoche, 1974). Sterols identified from rust spores are listed in Table 10. The surface lipids of Puccinia striformis urediospores have been examined by Jackson et al. (19731, the major components being P-diketones, n-alcohols (80% being octacosanol) and hydrocarbons, especially normal CZ9,C,, and C,, alkanes. An active glyoxylate-cycle pathway has been demonstrated in urediospores (Caltrider et al., 1963) and the presence of most of the enzymes of the tricarboxylic-acid cycle (Staples and Wynn, 1965). Farkas and Ledingham (1959) postulated a shift from lipid to carbohydrate metabolism after the first hours of germination, corresponding to the stage when the germ tube has entered the host mesophyll. That the composition of spores does not necessarily correspond with that of vegetative mycelium is seen from the differences noted by Van Etten and Gottlieb (1965)between the fatty-acid composition of spores and one day-old mycelium of Penicillium atrovenetum, where mycelial growth was accompanied by rapid synthesis of oleic acid, and between the lipid analyses by Tulloch and Ledingham ( 1964) of rust mycelium growing out from infected tissues and urediospore oil. The fatty-acid composition of the mycelium was much closer to that of other higher fungi than that of rust spores. Nevertheless, however little the lipids of spores reflect the active vegetative stages, they are of interest as end products of the metabolic chain diverting material from autotroph to heterotroph, and providing the nutritional basis for future host infection processes. 3. Lipids ofBiotrophic Fungi in Axenic Culture

Although a number of rust fungi have now been grown on synthetic culture media in the absence of host cells, there appears to be no report on lipid analysis of mycelium grown in axenic cultures other than that of Tulloch and Ledingham (1964) on the fatty acids obtained from mycelium which had grown out from infected tissue. The lipids of other more readily cultured biotrophs have received some attention, particularly Ustilago maydis, the corn-smut fungus, and Claviceps purpurea, which converts the ovary of various grasses into its ergot sclerotium. Among the lipid components of U . maydis and U . nuda, the loose smut of barley and other grasses, are the ustilagic acids (Table 4, p. 76), which appear to be responsible for the antibiotic activity of Ustilago cultures (Haskins, 19501, and partially acylated

144

P. J. BRENNAN AND D. M. LOSEL

sugars and sugar alcohols, discussed in Section V (p. 132). These have not, so far, been related to host-parasite physiology. Mycelial lipids of Claviceps purpurea growing in pure culture have been compared with ergot oil extracted from naturally occurring sclerotia. Some isolated strains of this fungus exhibited plectenchymic growth, and were found to contain ricinoleic acid ( D - 1Z-hdyroxy-cis-9octadecanoic acid), which accounts for about a quarter of ergot oil, where much of it is in the form of tetra-, penta- or hexa-acid glycerides, due to esterification of the hydroxy groups with normal long-chain fatty acids (Morris and Hall, 1966). This acid was of physiological interest in being absent from isolated strains which exhibited normal mycelial growth in cultures (Mantle and Tonolo, 1969; Mantle, 1969) and in that its biosynthesis co-incided with differentiation of sclerotial-type plectenchyma tissue and formation of ergot alkaloids. In one isolate, Morris ( 1968) demonstrated that linoleic and oleic acid residues may be precursors of ricinoleic acid. There are indications (Cooke and Mitchell, 1970) that lipid-associated mannitol and trehalose may be consumed during germination of C. purpurea, and that dormancy may be due to removal of some restraints on lipid-forming reactions. The initially high lipid content of ergots falls rapidly during germination, in parallel with free mannitol and trehalose. These contents are very low in dormant sclerotia, increase considerably during cold-activation treatment and finally fall off again during germination. The basic biologically active alkaloids in the ergots of Clauiceps were shown by Stoll (1945) to be depsipeptides. Cyclic depsipeptides have since been recognized in Gibberella baccata and as the destructans formed by another biotrophic fungus, the potato pathogen Oospora destructor, which are toxic to silk worms (Kodaira, 1962). 4. Lipid Metabolism ofBiotroPhic Fungi Growing in Host Tissues

a. Plant pathogens. Investigation of the vegetative biotrophic phase of mycelial growth in host tissue by parallel analysis of infected and uninfected hQst tissue, which has proved successful in studies of carbohydrate metabolism, has been extended to lipid metabolism by a number of workers. Williams et al. ( 1968)recorded increased total lipid in cabbage hypocotyls infected with Plasmodiophora brassicae. Muckerjee and Shaw (1962) found increased levels of lipid phosphate in wheat leaves infected with Puccinia graminis. Schipper and Mirocha (19701, in-

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vestigating changes in the total fatty-acid composition of Phaseolus wulgaris at different stages of infection with Uromycesphaseoli, detected synthesis of palmitic, stearic and oleic acids between six and nine days after infection, with oleic acid reaching a level of fifty times that present in healthy tissue. The form in which host substrates are taken up by rust fungi has interested many workers. Differences in the metabolism of labelled propionate by rust spores of P. graminis and by infected wheat tissue were detected by Reinsener and Jager (1969).Propionate-[2-l4C1fed to rust-infected wheat leaf tissue was taken up by the fungus after conversion to acetyl-CoA, whereas propionate taken up directly by urediospores was oxidized, the label appearing in the carboxyl group of acetyl-CoA. Knoche (1968) investigated the lipids of P. graminis urediospores from slices of infected host tissue which had been incubated with 14C-labelledacetate, stearate and oleate, and concluded that these compounds had been incorporated directly into the cis-9,10epoxyoctadecanoic acid of the spores. Reisener (1976), in his review of the metabolism of rust spores, pointed out that Knoche’s experiments did not rule our contact of the fungus with the labelled substrates, a difficulty which was avoided in Ziegler’s ( 197 1) comparison of lipid metabolism in rust-infected leaves fed with [2-14C1-glucoseor [ l -14C1acetate, in which the hexose proved a better substrate for fatty acid synthesis. From the labelling patterns obtained by Ziegler ( 197 1) on partial degradation of spore palmitic acid, Reisener ( 1976)postulated that the acetate units for synthesis of fatty acids might be provided by a phosphoketolase system, as found in certain bacteria. By means of such an enzyme, pentoses, derived from the hexose phosphate pathway, could be split yielding triose phosphate and acetyl phosphate. Studies of the lipid metabolism of rust-infected leaves by Hoppe and Heitefuss (1974b, c; 1975a, b) in Gottingen and by Losel and Lewis (1974) in Sheffield, have analysed changes in individual classes of lipid and have attempted to distinguish changes in-host and fungus during infection. During the development of P . poarum on leaves of Tussilago f a f a r u , aecial pustules accumulate lipid to a level (600 pg/cm2), four times that present in healthy leaves and in amounts comparable to the free sugars and fructans present, which are generally regarded as the major metabolites accumulating in Tussilago tissue infected by this rust (Losel and Lewis, 1974). An apparently similar accumulation of lipid accompanied development of P. poarum on its alternate host, P. prutensis, but much of this was external to the fungus in the host

146

P. J. BRENNAN AND D. M. L ~ S E L

cells. Of the 14C assimilated in photosynthesis by Tzssilugo leaf tissue, 75% was incorporated into neutral lipids (mainly fungal), particularly triacylglycerols and free fatty acid, as well as sterols and diaclyglycerols in the aecial postule. In healthy T . fur$uru leaves, only 25% of the 14C incorporated by photosynthesis appeared in neutral lipid (Losel and Lewis, 1974). In both hosts, incorporation of 14C into lipid increased during progress of the infection to a maximum when sporulation commenced, thereafter decreasing somewhat as spores with their lipid reserves were dispersed. Investigations of membrane lipids of healthy and rust-infected leaf tissues by Hoppe and Heitefuss (197413, c, 1975a, b), using Uromyces phuseoli on Phaseolus vulgaris, and by Losel and Lewis (1974) with P. poururn on Tussilugo, produced some parallel results but different interpretations. In both systems, the amount of chloroplast lipid decreased following infection. The discrepancy between the increased levels of phosphatidylethanolamine and phosphatidylcholine per unit area of leaf tissue infected with Puccinia poururn, and the slight but steady decrease in all phospholipids per unit dry weight of Uromycesinfected bean leaves recorded by Hoppe and Heitefuss (1974131, may have been due to an increase in the dry weight of tissue with age, which occurs particularly in rust-infections. As in previous studies with their host-parasite system (Holligan et ul., 1973, 1974), and that of Von Sydow (1966) on rust-infected wheat leaves, Losel and Lewis (1974) preferred to express the amounts of substances present per unit area of leaf, rather than on a dry weight basis. Priestley (1974) recommended extracted dry weight of plant tissue as a reference basis for biochemical studies, but both dry weight and extracted dry weight present difficulties when infected tissues accumulate unusually large amounts of soluble and insoluble materials. An overall loss of membrane lipid in rust infections would be difficult to reconcile with the greater incorporation of label by infected tissue than by control leaves in bean shoots fed with 32P(Hoppe and Heitfuss, 197413) and in Tussilugo after photosynthesis in 14C02 (Losel and Lewis, 1974). In both systems, very marked increases in specific activity occurred in phosphatidylserine and phosphatidic acid following infection. Hoppe and Heitefuss ( 1974a), seeking the basis of the increased permeability to sugars and ions which they had detected in rust-infected leaves of Phmeolus, interpreted -the increased phosphatidylserine of infected tissue as an alteration in host membranes rather than new fungal

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lipid, although they found phosphatidylserine in spores of U . phaseoli, both germinated and ungerminated. The difficulty found by the Gottingen group in attributing this increased incorporation in phospholipid to phospholipid synthesis by the fungus was that “the parasite would have to contribute about 40% of the dry matter of the hodparasite complex if the phosphatidylserine increase is to be explained only with fungal lipids”. Since the presence of at least this amount of fungal material in aecial pustules on Tussilago was indicated both by chitosan estimations and direct microscopic observation, Losel and Lewis (1974) suggested that the dominant process of lipid metabolism in infected tissue was synthesis of fungal components, both storage lipids and membrane lipids, which would be likely to mask any changes in the composition of host membranes. Similarities were demonstrated between such detectable host changes in aecial pustules as losses of chloroplast glycolipids and phoshatidylglycerol, with some increases in free fatty acids and triacylglycerols, and the changes found in senescent leaves of Tussilago. Comparison of the distribution of label in healthy and infected hosts of P . poarum immediately after exposure to 1 4 C 0 2in light, and following a subsequent “chase” period in normal atmospheric conditions in the dark (D. M . Losel, unpublished observations), showed much of the activity initially incorporated into sugars was lost during the dark period, while the activity of lipids in all infected tissues rose severalfold. The 14C activity in lipids of healthy leaves and uninfected parts of diseased leaves changed relatively little in this time. The rust-infected tissue differed markedly from the uninfected tissue in the diversion of carbon from other fractions into lipids, during the “chase” period, the initially higher proportion of activity in polar lipids of diseased tissue decreasing as a greater amount of label accumulated in neutral lipids. Immediately after photosynthesis, the 14C activity of lipid fractions from both healthy and diseased tissue is mainly in diacylglycerols, chloroplast glycolipids, phosphatidylglycerol and phosphatidic acid. During the dark period, loss of the initially high activity of diacylglycerols and polar lipid adjacent to pustules is accompanied by increases in other classes of neutral lipid and membrane lipids within the fungal pustule tissue. Hoppe and Heitefuss (1974c, 1975a)investigated the fatty-acid composition of each of the polar-lipid classes present in rust-infected and healthy bean leaves and in resting and germinating urediospores. Little

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change occurred in phosphatidylglycerol and monogalactosyl diacylglycerol digalactosyl diacylglycerol or in the lipids of uninfected tissue adjacent to the rust pustules. The most striking difference between infected and healthy tissue was the higher unsaturation of phosphatidylethanolamine and phosphatidylcholine from infected half leaves, the ratio of unsaturated to saturated fatty acids reaching a maximum in phosphatidylethanolamine and, to a lesser extent, in phosphatidylcholine on the eighth day after inoculation, after which the ratio for phosphatidylethanolamine fell to just above the original level but in phosphatidylcholine, after a slight drop, continued to rise. To find whether these changes were due to the host or the fungus or both, the fatty-acid composition of uninfected tissue adjacent to rust pustules was compared with that of urediospores (Hoppe and Heitefuss, 1975a). The percentage of linolenic acid residue in urediospore lipid was much higher in phosphatidylethanolamine and slightly higher in phosphatidylcholine than in the corresponding lipids from adjacent uninfected tissue, and its level rose in both phospholipids during spore germination. Although higher unsaturation of the phospholipids of infected tissue, especially phosphatidylethanolamine, could have been responsible for leakage of sugars previously observed, Hoppe and Heitefuss (1975a) felt that the level of unsaturation of phosphatidylethanolamine from urediospores indicated that the differences were mainly due to the fungal phospholipid, although fungus-free tissue was also slightly affected. They concluded that both host and parasite are involved in changes of the lipid acyl groups which might be important for alterations in membrane permeability. A major lipid component of U . phaseoli urediospores and rustinfected bean leaf tissue was the C epoxy acid cis-epoxyoctadecenoic acid, earlier identified in urediospores of various rusts (Tulloch and Ledingham, 1962) and recently recorded as the major fatty acid of Cronartium fusforme, predominantly in the polar-lipid fraction (Carmack et d., 1976). Various workers have investigated the possibility of phospholipase activity accompanying infection being an important factor in biotrophic physiology. The observation of Lumsden and Bateman (1968) of phosphatidase activity in Phaseolus vulgaris infected with Thielauiopsis basicola was followed by Lumsden’s (1970) demonstration of phosphatidase production by Sclerotinea sclerotiorum both in culture and in bean tissue. Hoppe and Heitefuss (1974b) found increased phospho-

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lipase activity of rust-infected halves of bean leaves compared with uninfected half leaves and control leaves, the enzyme activity reaching a maximum six to eight days after infection, a time corresponding to maximum sporulation by the fungus, and cited evidence that increased phospholipase activity may damage mitochondria in infected leaf tissue. They pointed out however that, in their system, damage to mitochondria and uncoupling of respiration occurred at a much later stage of infection than the phospholipase maximum. Sterols of infected leaves have also been investigated in the search for membrane components which might be altered by the presence of a biotrophic fungus. Nowak et ul. (1972) found no differences in sterol content of healthy and rust-infected wheat leaves. Increased amounts of sterols and increased incorporation of '*C-labelled photosynthate into sterols were found in aecial pustules on Tusszlugo leaves (Losel and Lewis, 1974)and in urediospore stages on leaves of Poa (D. M. Losel, unpublished observations) following infection by Puccinia poarum. A7,24(28)S tigmastadienol was the principal sterol component of Cronartiumfusiforme aeciospores, comprising 0.3% of the spore weight (Carmack et al., 1976). Lin and Knoche (1974) found that Uromycesphaseoli appeared to stimulate sterol synthesis in rust-infected bean leaves, but that most of the increased sterol was in urediospores. The major sterol of the urediospores, A7~'4(28i-stigmastadien-3/3-ol, was accompanied by /3-sitosterol, stigmasterol, campesterol and 28-isofucosterol. In an earlier study, Lin et ul. (1972) had demonstrated synthesis of A7~24('28i-stigmastadien-3/3-01 and 7 -stigmastadien-SP-ol by germinating urediospores from [ I -I4C1-acetate, and had obtained sterol biosynthesis by cell-free preparations from urediospores. A list of sterols identified from rustinfected tissues and urediospores is given in Table 10. I t is of interest that most of the rust sterols listed in Table 10 are plant sterols, not previously recorded from fungi, and that ergosterol, which is present in all other fungi except the Oomycetes (Weete, 19741, is absent from these rust records. Hoppe and Heitefuss (1975bj detected no differences in sterol composition between healthy and rust-infected leaves of resistant bean varieties, but found that susceptible leaves infected with U . phaseoli contained in addition A7~24(L8)-~tigma~tadien-3/3-ol or 7 -stigmasten-SP-ol, which was also present in urediospores. This sterol, and smaller amounts of another which chromatographed next to it, corresponded to peaks obtained from a sample of 7-stigmastenol from wheat-stem rust urediospores. Together, these made up to 8 to 12% of total sterols

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on the sixth day after infection, rising to 27 to 28% by the ninth day. At the same time, stigmasterol and campesterol increased compared with controls. In ungerminated spores, the presumptive 7 -stigmasten01 comprised 82% of the total sterols. During germination, this decreased to 76.5%, while the component which appeared to correspond to stigmasterol increased from 17.6 to 23.5% of the total sterols and a trace of another sterol, possibly p-sitosterol, appeared. Although the amounts of stigmasterol and the characteristic major sterol of rust-infected bean leaves corresponded with the amount of fungus in the host, Hoppe and Heitefuss (197513) considered that the increase in campesterol and the unknown sterol, which appeared in infected tissue but was not detected in spores, could not be due to fungal sterols, if the mycelium and the urediospores of the parasite have the same sterol pattern. Since, however, slight changes in sterol composition during germination of bean-rust urediospores were detected in the above study, and work on two other basidiomycetes, Agaricus bisporus (Holtz and Schisler, 197 1 ; O’Sullivan and Losel, 197 1) and Coprinus sp. (Defago, 197 1 ) showed striking differences in sterol composition between mycelium and spores, this does not seem a convincing argument against regarding most of the increased sterol content of infected tissue as being due to fungal sterols. Hoppe and Heitefuss ( 1975131, however, postulated that the “host plant probably tries to compensate this permeability effect (of infection) by an increased sterol synthesis” but concluded that the sterol content of different bean varieties was not related to susceptibility or resistance. A similar conclusion has been reached by Nowak et al. (1972)for wheatstem rust. The ability of rust fungi to form sterols from simple substrates was indicated by synthesis of sterols from acetate by germinating rust urediospores (Lin et al., 1972rOther ecologically obligate pathogens may have to obtain sterols from the host plant. Since Plasmodiophora brassica resting spores, obtained from club-root infections on various hosts, were similar in sterol composition to their hosts, Knights (1970) suggested the sterols were taken up by the parasite from host tissue. In all cases, p-sitosterol was the major component (63430%) with campesterol (14-24%), stigmasterol (6-13%) and lower levels of cholesterol ( 1-3%), and variable small amounts of brassicasterol. A5-Avenasterol occurred in Plasmodiophora spores from Sinapis alba but not in healthy roots. Strandberg’s ( 1968) observation that labelled pre-

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cursors are more readily incorporated into sterols and other lipids in club-root infected cabbage hypocotyls than into healthy hypocotyls suggests that accelerated sterol metabolism of the host may result from transfer of sterols to the resting spores of the parasite. Dependence on host tissues for sterols may be an essential feature of other obligately biotrophic parasites, particularly oomycetes which have been found unable to synthesize sterols required for growth and morphogenesis. For example, sterols from potato leaves stimulate growth and sporulation in Phytophthora infestans (Langcake, 1974). I t is possible that this may be an important factor in parasitism of other oomycetes, such as the Peronosporales (powdery mildews), which have not yet been grown in axenic culture. b. Mycoparasites. Just as comparative studies on carbohydrate metabolism of different types of biotrophic associations furthered progress towards an integrated view of the movement of photosynthate from autotroph to heterotroph and subsequent sequestration within the heterotroph, so the comparative approach may yield greater insight into the lipid metabolism of such relationships and its role in parasitic physiology. The important study of Manocha (1975) on the mycoparasite Piptocephalis virgzniana may parallel and extend the picture emerging from investigations of lipid metabolism in rust fungi. When growing on the host fungus, Choanophora cucurbitarum, P. virgzniana produces branching sporophores bearing spores which contain two terminal lipid bodies. The spores on the thin, unbranched sporophores, which are formed on the limited mycelium produced in pure culture, lack these lipid bodies. Corresponding to this, the lipid content of the parasitic spores was 205 mglg dry weight, compared with 95 mglg dry weight in the axenic spores. The parasitic mycelium contained spherosome-like bodies, while axenic mycelium lacked these but contained many vacuoles. Gas-liquid chromatographic analysis indicated that the major qualitative difference was the failure of the axenic spores to synthesize y-linolenic acid, the characteristic fatty acid of the Mucorales. The proportions of other major fatty-acid components, palmitic, palmitoleic; stearic, oleic and linoleic acids, are otherwise similar in both parasitic and axenic culture. Thus, the axenic spores may have a block in polyunsaturated acid synthesis. Manocha ( 1975) mentions evidence that y-linolenic acid may be present not only in storage lipid but also in membrane lipids such as phosphatidylethanolamine and

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phosphatidylcholine, and makes the interesting suggestion that P. virginiana may have a defective membrane, which would be a disadvantage in free-living organisms but might be a useful adaptation to parasitism. The structural basis for some of these changes is indicated in an earlier study of the parasitic development of P. virginiana on another host, Mycotypha microspora, in which Armentrout and Wilson ( 1969) recorded sphaerosome changes in host and parasite. Mycotypha cells initially showed little endoplasmic reticulum, but contained numerous sphaerosomes and many small vacuoles. Following penetration by Piptocephalis, the host sphaerosomes disappeared, any remaining ones losing their membrane continuity; vacuoles began to coalesce and endoplasmic reticulum became more prominent. Sphaerosomes were Seen to move down the hypha of the parasite towards the appressorium by which it was attached to the host hypha. The entire haustorium and the sphaerosomes of P. virginiana gave a staining reaction for acid phosphatase activity, but the sphaerosome-like bodies seen in phase-contrast examination of the spores did not give a positive reaction. By 36 h after infection, host cells and the haustoria within them appeared empty. Comparable observations were made by Seymour ( 19 7 1 ) o n the chytrid Septosperma, which is parasitic on another chytrid, Rhizophydiurn macrosporurn. Oil droplets developed in the host in response to infection, then gradually became smaller and fewer as the protoplast gradually disappeared, with eventual collapse of the host wall. The protoplasm of the parasite accumulated spherical refractive globules before cleaving to form zoospores, each with a single, anteriorly placed refractive globule. During formation of resting cells later in infection, large refractive oil deposits appeared, protoplasm moved to an upper cell cut offby a wall, where the oil deposits broke up into smaller ones of uniform size, which filled the protoplasm. c. Mycorrhiral fungz. The lipid physiology of mycorrhizal associations does not appear to have been studied to any great extent apart from some observations of lipid droplets in endomycorrhizal fungi already mentioned. Increased triacylglycerol in lipid fractions of infected onion roots, compared with uninfected roots, was observed by F. E. Sanders (personal communication) in vesicular-arbuscular mycorrhiza-infected roots of onion and in roots of onion, clover and rye grass infected with vesicular-arbuscula mycorrhiza (Cooper and Losel, 1978) but with little increase in phospholipid, although the

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electron micrographs of Cox and Sanders ( 1974) showed considerable increases in the amount of host membrane present in infected cells. D. M. Lose1 and K. M. Cooper (unpublished observations) investigated the path by which host photosynthate might be transferred to fungal lipid, comparing the incorporation of 14Cby infected and uninfected roots which had been incubated in solutions of [l4C1-labelledacetate, glycerol, glucose and sucrose. Most active incorporation of label was from acetate into neutral lipids and polar lipids, mycorrhizal tissues showing greater radio-activity chan non-mycorrhizal tissue in triacylglycerols, free fatty acids, phosphatidylethanolamine and phosphatidic acid. With glycerol and sucrose as sources, radio-activity appeared in triacylglycerols, sterols and phosphatidylethanolamine but not in free fatty acids and diacylglycerols. I t would thus appear that the most direct path of labelling of lipids is from acetate, and that label from glycerol and sucrose appears first in the carbon skeletons of triglycerides and sterols. A few studies have touched on lipid physiology in sheathing or ectotrophic mycorrhiza. Krupa and Fries (1971 ) found volatile metabolites, such as isobutanol and isobutyric acid, were produced during early actively growing stages by mycorrhizal fungi, such as Boletus variegatus, and suggested these may be important during early stages of infection of roots for their proven inhibition of root pathogens such as Phytophthora cinnamomi, Fomes annosus and Rhirina undulata. More recently, Lindeberg and Lindeberg (1974) investigated the effects of short-chain fatty acids a n the growth of some mycorrhizal hymenomycetes. Five ectomycorrhizal fungi, investigated by Melhuish et al. ( 19751, showed individually distinguishable free fatry-acid patterns. Linoleic acid was most abundant in all species, with lesser amounts of oleic, stearic and palmitic acids and trace amounts of others. I t was not possible to determine whether lipids were involved in development of the mycorrhiza, but one interesting correlation was noted. Hebeloma sarcophyllum, one of the very few mycorrhizal fungi capable of forming sporophores in monoxenic culture, had much more lipid than the other mycorrhizal fungi studied, the linoleic acid content reaching 5% of the dry weight. d. Lichens. Published electron micrographs of lichens, the carbohydrate metabolism of which closely parallels other biotrophic fungi in conversion of host photosynthate to fungal polyols and trehalose, do not appear to show accumulation of lipid in the fungal tissue.

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Lichenized algae, e.g. Trebouxia in Ramalina mucformis (Peveling, 19731, differ from their free-living counterparts in the presence of lipid-containing plastoglobuli between thylakoids of the chloroplast. Undoubtedly, all of the membrane lipid of the fungus, which comprises the bulk of the lichen thallus, is derived from algal photosynthesate, but there is no evidence at present for accumulation of storage lipid by the fungus during normal thallus growth. O n the other hand, many of the characteristic lichen substances are derived from fatty acids. I t is suggested that their biosynthesis starts by condensation of a “starter” unit, usually acetyl-CoA and malonyl-CoA, with concomitant decarboxylation and formation ofaputativepoly-P-oxothio-ester (Mosbach, 1972). Experiments in which 14C-labelled acetate and succinate were fed to Cetraria islandica indicated that its y-lactone, (&I-protolichesterinicacid, is formed by condensation of palmitic acid with a C,-unit. Rocellic acid, rangiferic acid and acarenoic acid are probably derived from myristyl- or palmitolyl-CoA and oxaloacetic or n-pyruvic acid. Fox and Huneck (1969) showed that rocellic acid can be produced by the fungal symbiont of Lecanora rupicola alone in culture. Huneck (1972) has reviewed the chemistry of lichen substances. There has been much speculation concerning the physiological role of these characteristic lichen products. Mosbach (1972) suggests that, since lichens are slow growing, probably as an adaptation to nitrogen deficiency, any overflow of carbon metabolites caused through photosynthesis of the algal partner is channelled into the formation of lichen substances. He speculates that, in lichens, the fatty acid content is low but polyketides are usually found, due to a lack of NADPH required for fatty-acid formation, and point to a similar diversion in Penicillium baamense (Mosbach and Baverstoft, 197 1). Studies by Hill and Ahmadjian (1972) on incorporation of label by Cladonia cristatella from H14C0, indicated that the phycobiont alga in culture incorporated more [l4C1into ethanol-insoluble and lipid components than it did in the lichen whereas, in the lichen association, algal photosynthate was diverted from lipid and insoluble substances into the mobile polyol ribitol. The sterols of the symbionts of Xanthoriaparietina in separate culture have been studied by Lenton et al. (1973).The mycobiont contained ergosterol and lichesterol, together with lower levels of other C,, sterols. The sequence of sterols in the mycobiont suggested the same biosynthetic path as for ergosterol production in yeast. Lichesterol was thought to be produced in the dark, possibly by action of a reversible

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A 8 s A 7 isomerase on ergosterol. I t thus appears that the fungal partner can synthesize sterols which it requires from simpler metabolites of the alga. The sterols of the phycobiont Trebouxia differed from those of the mycobiont, with poriferasterol as the predominant component and lower levels of dionasterol, ergost-5-en-3P-01, brassicasterol and cholesterol. Similar sterol mixtures were found in two other Trebouxia species. B.

FUNGI ASSOCIATED WITH INSECTS

Since De Bary’s (1879) description of mycetocytes, yeast-like organisms contained in specialized organs of certain insects, an extensive but scattered literature has accumulated, describing associations between fungi and insects. Some of this was summarized by Baker (1963) and by Cooke (197 7). Although the importance of nutritional factors in the interaction of the two partners has often been indicated, relatively few detailed investigations have been made. The evidence available repeatedly implicates fungal lipids in such associations. Koch ( 1933) recorded poor growth of anobiid beetle larvae hatched from surfacesterilized eggs, unless their diet was supplemented with yeast. This observation was explained in terms of a requirement of the larvae for vitamins and sterols (Blewett and Fraenkel, 1944). Insects are dependent on a dietary source for sterols essential for development. Those which have evolved a suitable symbiotic association are able to grow successfully on a diet almost totally deficient in essential sterols and vitamins. They include scolytid beetles, which tunnel in wood and cultivate “ambrosial fungi” in their tunnels, anobiid larvae growing in flour and the attine ants, which cultivate extensive fungus gardens in tropical soils. 1. “Ambrosial” Fungi and Scolytid Beetles

During hibernation or unfavourable seasonal conditions, the ambrosial fungus survives as yeast-like cells in pockets in the integument, within which an oily secretion accumulates (FranckeGrosmann, 1956). During tunnelling, secretion increases and yeast cells are washed out onto the tunnel walls, where their germination is promoted by the oil. Francke-Grosmann ( 1956)pointed out that such an association might have arisen in ancestral types by casual colonization by fat-utilizing fungi of glands originally providing lubrication for the beetle during boring. Other groups differ in the location and type

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of the fungus-containing organs, but there is a general association of the fungi with oil secretions which are particularly characteristic of the female scolytid beetles. Since cellulose and lignin are not broken down by the insects' enzymes, the nutrition of the wood-boring scolytids is completely dependent on the ambrosial fungus growing on the walls of their tunnels. The insect normally feeds o n the spores and young hyphal tips of the fungus. The association is fairly specific and occurs most frequently in wood-boring beetles of tropical forests. North-temperate scolytids are mainly bark-boring beetles, growing in tissues which provide a richer substrate and are thus less dependent on symbionts. Mathiesen-Kaarik ( 1953) showed Trichosporium tingens, the symbiont of a bark-boring scolytid beetle, differed from typical blue-stain fungi (see p. 157) which occur in a similar habitat, in requiring inorganic nitrogenous compounds and unknown growth factors and in assimilating lipids as a carbon source. The fungus was transmitted in specialized organs by the beetles, and was dependent on the secretions of the insects for growth. I n Platypus cylindrus, the oak pinhole borer, where no special transmission organs are formed, both sexes carry fungal spores. Baker (1963) has suggested that it would be interesting to investigate the lipid metabolism of this association with Endomycopsis (Hansenula) to find whether it is lipid-producing and could stimulate growth of the principal ambrosia fungus. In the less well studied association of' Ascoidea hylecoeti, a symbiotic ambrosia fungus related to the yeasts, with Hylecoetus dermestoides, a secondary wood borer (Ships timber worm) from many hard woods and all soft-wood species in Europe, the fungus provides the insects with proteins, glycosides and lipids (Batra and Frmcke-Grosman, 196 1). Insects in general are unable to convert simple precursors into sterols, which are fundamentally involved in their differentiation. A series of studies on wood-boring scolytids (Kok et al., 1970; Chu et al., 1970) have revealed the significance of sterol production by the fungus in these symbiotic associations. Ergosterol was the only sterol detected in isolates of Fusarium solani from mycangia (fungus-bearing organs) of Xyletorus ferrugineus. Significant amounts of sterol esters were not detected. Fungus-free females, raised from surface-sterilized eggs, produced second-generation larvae, which grew but failed to pupate on cholesterol-containing medium. Either ergosterol o r 7 -dehydro-ergosterol as sole sterol source proved adequate for growth, development

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and reproduction of the fungus-free beetle through several generations. Pupation appeared to require the A’ bond which is present in both ergosterol and 7 -dehydro-ergosterol but absent from cholesterol and lanosterol. Lumisterol, which was antagonistic to progeny production or survival, differed from the useful sterols in the structural differences at C,o. Fungal ergosterol was thus shown to be a sex homone for Xyletorus, which otherwise remains infertile. Kok and Norris (1972) found the phospholipids of Fusarium solani, Cephalosporium sp. and Graphium sp. mutualistic with the ambrosia beetle, X . ferrugzneus, differed from normal fungi in lacking phosphatidylserine. Phosphatidylcholine and phosphatidylethanolamine accounted for more than 60% of ihe phospholipids of CephalosporiGm sp. and F. solani, and more than 80% of the total lipid in Graphium, where the phosphatidylethanolamine content was particularly high. In a hrther study, Kok and Norris (1972) analysed the neutral lipids of three fungi, namely Fusarium solani, Cephalosporium sp. and Graphium sp., mutualistic with Xyletorus ferrugineus. The amounts of lipid found in these ambrosial fungi were generally higher than in other Fungi Imperfecti and yeasts. Fusarium solani had nearly twice the fatty-acid and sterol content of the other two fungi. Age of mycelium up to 15 days did not significantly affect the qualitative composition and total yield of lipids. The fatty-acid compositions of the neutral lipids resembled other reports for Fungi Imperfecti, apart from the higher levels of stearic acid in the ambrosial fungi. 2. “Blue-Stain”Fungi

In the less intimate association of the “blue-stain’’ fungi and insects, lipids have also been shown to play an important part in the nutritional relationship. Some of the fungi involved and their associated beetles are listed in the reviews of Francke-Grosmann (1951), Baker ( 1963) and Cooke ( 197 7 ) . They include species of Leptographium, Ceratocystis, sometimes also associated with Fomes annosus, the wood-rotting fungus responsible for heart rots or conifers. Monilia ferruginea, the blue-stain fungus of Trypodendron lineatum, did not germinate readily on malt agar but did so on casein- or peptone-agar olive oil, giving the ambrosial form. Staining of the medium by growth of the fungus was due to an oily exudate as well as the dark hyphae. Blue-stain fungi may grow in a yeast form on the oil secetions of ambrosia beetles without colonizing their transmission organs (Francke-Grosmann, 19561,

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Ceratocystis often persisting in the tunnels after they have been abandoned by the beetles. Mathiesen-Kaarik ( 1960) found insect-dispersed blue-stain fungi of the genus Ceratocystis and the closely related Dutch elm disease fungus, Ophiostoma ulmi, partially or totally deficient for thiamin, pyridoxine and biotin, whereas air-dispersed blue-stain fungi were autotrophic. Aspartic acid, oleic acid and to a greater extent Tween 80 (polyoxyethylene (20) sorbitan mono-oleate) all had a growth-promoting effect on biotin-deficient fungi. Good growth was obtained in some cases with Tween 80 as carbon source. In media containing nutrient- rather than vitamin-levels of oleic acid o r Tween 80, biotin seemed to be unnecessary for 0.pini. Replacement of biotin by oleic acid seems to be dependent on such factors as carbon dioxide tension and pH value, varying with the species and strain of the fungus. In bacteria, Williams et al. (1947) suggested that one function of biotin is to catalyse directly or indirectly synthesis of oleic acid. These studies did not completely rule out the possibility that traces of biotin might have been present as impurities in the oleic acid, nor is the relationship ofthe fungi with the beetles completely clarified. 3. Wood-Rotting Fungz and Wood Wasps

In other fungus-insect associations, the role of lipids has not been so extensively investigated. Wood wasps of the Siricidae inoculate a wood-rotting fungus, usually a basidiomycete, while laying eggs, but the larvae feed on wood softened by the fungus, rather than directly on the fungus. In female wasps of the genera Sirex, Urocerus and Tremax, intersegmental pouches attached to the ovipositor are filled with ooidia of the basidiomycete, embedded in slime. When the ovipositer, a strongly sclerotized sting, is inserted into the wood, these fungal spores are pushed out with the egg into the wood. I n Sirex cyaneus and Urocerus gigas, deep cryptae above a layer of large glandular epithelial cells contain fatty secretions in which ooidia of the symbiotic fungus are immersed (Parkin, 1941, 1942). Shortly before the end of the larval phase, these secretions harden into waxy plates which contain ooidia. During moulting, these plates are shed and, as the emerging iemales gnaw their way out of the pupal chamber, the ooidia are transported to pouches at the base of the ovipositor. The wood-wasp fungi are effective mainly through their wood-rotting action, not by producing masses of “food” mycelium equivalent to the ambrosial types

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(Francke-Grosmann, 1967 1. The association with Xyphidiaprolongata has been demonstrated for the wood-rotting basidiomycete Daedalia unicolor on beech and Stereum spp. on soft woods, and also with the ascomycete Daldinia concentrica. In view of the demonstration by Fries and his coworkers of stimulation of growth of wood-rotting fungi by fatty acids (Glasare, 1970), it is possible that closer study might show direct stimulation of growth of these symbiotic species by secretions of the wood wasps. 4. Anobiid Beetles

Symbiosis with a yeast-like fungus enables the anobiid beetles to survive on substrates such as flour, which are deficient in essential vitamins and sterols (Jurjitza, 1962).The yeasts are present in enlarged epithelial cells of specialized organs called mycetomes, attached to the gut. Eggs passing through the gut are smeared with fungal cells and infection of the new generation occurs when the larvae, on hatching, eat the egg case with the fungal inoculum. Fungus-free larvae, hatched from surface-sterilized eggs, remain small and incompletely developed. On substrates supplemented with cholesterol, development is much better, although not quite equal to that of yeast-infected insects. The fungal symbionts of one of these beetles, Sitodrepa panicea, can be cultured on normal media, such as malt-agar, but are partially heterotrophic for asparagine or glutamic acid, possibly indicating their adaptation to intracellular existence (Kuhlwein and Jurjitza, 1967). Because of the resemblance of the symbiont isolated from S. panicea to the ascomycetous plant pathogen, Taphrina sp., Kuhlwein and Jurjitza (1967) suggested that the ancestors of the insect may have been plant eaters as are other related types. 5. Fungus Cultiuation by Leaf-Cutting Ants

Even more elaborate cultivation of fungi than that of the scolytid beetles is practised by some leaf-cutting attine ants which grow fungi as a food source in large subterranean gardens in tropical soils (Woser, 1966). These ants cut pieces of leaves and use them as a substrate for cultivation of the fungus, which originally arises from an inoculum carried in the queen ant’s head when she founds a new colony. Division of labour in the collection of leaf material, tending and weeding of the fungus gardens and of care of the eggs and young are

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ensured by a well-defined caste system and, as in the ca5e of‘ the ambrosia beetles, the nutrition of the animal, including its lipid nutrition, is entirely dependent on the fungus. C . FUNGI PATHOGENIC TO MAN AND ANIMALS

A limited number of published studies have been concerned with the lipid physiology of fungi causing mycoses of Man and animals. Aspects which have received some attention include comparison of lipid composition of pathogens and non-pathogens, possible correlation of lipid content of pathogens with virulence and with dimorphism, and the effects of various fatty acids on the growth of dermatophytes. 1. Lipid Content ofPathogenic Fungi

Lipid inclusions in host cells o r pathogen have not been the direct object of ultrastructural studies of pathogenic fungi, but can be observed in published electron micrographs from some other investigations primarily concerned with morphogenesis of pathogenic fungi and their relation to host tissue. These have been mainly on material from axenic cultures, with relatively few observations on fungal structures within infected tissue. Freeze-fractured preparations of Pityrosporium orbiculare, a lipophylic yeast-like fungus responsible for pityriasis versicolor, a skin disease of Man, showed the presence of a large lipid droplet close to the cell nucleus (Breathnach et al., 1976). Observations by Edwards et al. (1959)on the fine structure of yeast cells of Histoplasma capsulatum showed lipid bodies in very close contact with mitochondria. The multinucleate yeast-like cells of Blastomyces dermatitidis contained numerous mitochondria and irregularly-shaped lipid bodies (Edwards and Edwards, 1960). Electron micrographs of’hyphae from cultures of Phialophora werrucosa and Cladosporium carrionii, agents of chromomycosis, also showed prominent oil drops (Cooper et al., 1973).

In an early study in this field, Peck (1947) suggested that the lipid content of pathogenic fungi may be associated with their- parasitic behaviour. Peck (1947) found the most significant difference between the free lipids of pathogenic and non-pathogenic fungi to be the absence of carbohydrate-containing phospholipid in all but one of the non-pathogens. A1 Doory and Larsh (1962) investigated the lipids of ten dermatophyte species and four systemic fungi, and found that the yeast phase of each systemic species o n solid medium had a higher

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total lipid content and higher acetone-soluble fraction than did mycelium grown in liquid medium. In all of the fungi examined, shake cultures produced least lipid. Species of Cryptococcus ( Pedersen, 19631, Histoplasma and Blastomyces (A1 Doory and Larsh, 1962), causal agents of serious systemic mycoses, and the dermatophytes Microsporon and Epidermophyton (A1 Doory and Larsh, 19621, were not found to show any features of lipid composition strikingly different from non-pathogens. In contrast to this, however, Di Salvo and Denton (1963) demonstrated a direct correlation between the total lipid content of the yeast phase of Blastomyces dermatitidis and virulence towards mice. An avirulent mutant strain of Coccidioides immitis differed from the wild-type mouse-virulent strain studied by Anderes et al. (1973) in the lower lipid content of its arthrospores and in having a much higher proportion of oleic acid (79%of total fatty acids, compared with 30% in the virulent strain) and three times as much sterol, which appeared from ultraviolet spectra to be neither cholesterol nor ergosterol. The overall lipid composition was similar in virulent and avirulent strains, but the virulent arthrospores contained 18% more total lipid, 15%more free lipid and 39% more bound lipid than the mutant non-virulent spores. It was suggested that the higher lipid content may be a factor in initiation of infection. Comparison with the nutritional relationship of some of the fungi discussed in Section VI-B (p. 5 5 ) with their insect hosts might also suggest that the virulent strain is more dependent on host tissue as a source of oleic acid and sterols than is the avirulent strain. The lipid composition of dermatophytes in culture has been the subject of a number of investigations, but these have not been correlated with their pathogenic behaviour. The fatty acids of Trichophyton mentagrophytes (Audette et al., 196 11, Trichophyton rubrum (Kostiv et al., 1966) and Candida albicans (Combs et al., 1968), with linoleic, oleic, palmitic and stearic acids as the major components, did not differ appreciably from taxonomically related non-pathogenic fungi. The phospholipid fraction of T. rubrum (Das and Banerjee, 1974) contained phosphatidylcholine, phosphatidylinositol, polyphosphatidylinositol, phosphatidylserine, phosphatidylglycerol and phosphatidic acid in almost constant relative proportions throughout the life of the culture. Investigating the sterols of various dermatophytes, Blank et al. (1962) found they contained either brassicasterol or ergosterol with only occasionally small amounts of one accompanying the other. Brassica-

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sterol, the sole sterol of T. violaceum and T. discoides and the predominant sterol of T. megnini, was present in trace amounts in three other Trichophyton species and Epidermophytonjoccosum, but absent from three further species of Trichophyton and four species of Microsporum, which contained ergosterol only. First recorded in fungi, in the study by Wirth et al. (19611, from mycelium of T. rubrum and apparently absent from all other fungi, brassicasterol is a sterol of plant tissues. Knights ( 1970a) found small, variable amounts in Plasrnodiophora resting spores from different host plants, but considered that the sterols were taken up by the parasite from host tissue. 2 . Dimorphism A number of studies of dimorphism of pathogenic fungi, the characteristic alteration from filamentous habit to yeast form of growth in tissues, have investigated the involvement of lipids. In Phialophora verrucosa, isolated from a clinical case of chromomycosis, alternation of filamentous and yeast-type growth was controlled by the presence of Mn2+,and which seems to function as a cofactor in wall synthesis (Reiss and Nickerson, 197 1).Another species, P. richardsiae, showed similar morphogenetic sensitivity to iron. The chlamydospores, corresponding to the “sclerotic form” in host tissues, were characterized by large refractive globules. Manganese was similarly implicated in an alteration of morphology in Aspergillus parasiticus somewhat suggestive of hyphal-yeast morphogenesis (Garrison and Boyd, 197 1). Electron micrographs of A . parasiticus conidia during incubation on Mn2+-deficient medium showed a striking deposition o f new wall material inside the cells, increasingly compressing the protoplasmic contents, and a rapid accumulation of lipid droplets which, by 72 h, occupied most of the cytoplasm. Raising the concentration of TABLE 1 1. Cell-wall lipid content of dimorphic fungi Fungus

Lipid (percent of dry wall material) Yeast

Mucor rouxii Candida albicans Candida albicans Paracoccidioides brcrtiliensis Blastomyces dermtitidis Histoplasma capsulatum Histoplasma capsulatum

5.7 0.6-10.6 1.8 8.3-10.7 4.6 1.5-2.0 6.8

Reference

Hyphae 7.8 5.1-5.5 4.5 4.8-10.4 8.9-9.6 0.8-1.0 6.2

Bartnicki- Garci ( 1968) Chattaway et al. (1968) Bianchi (1967) Kanetsuma et al. (1969) Kanetsuma et al. (1969) DomerandHamilton(1971) Domer et al. (1967)

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manganese ions by a factor of ten resulted in a rapid change from “yeast-like” development to normal uniform-walled cells, which also contained lipid-storage bodies. Differences in the amount of lipid present in cell walls of yeast and mycelial phases of dimorphic fungi have interested a number of workers. Comparison of some of the results obtained (Table 11, extracted from the table of fungal’wall-lipid contents, in Weete, 1974) indicate great variations from species to species and between different determinations for individual species. Hyphal walls of Mucor rouxii, Candida albicans and Blastomyces determatitidis had lipid contents much higher than walls of the corresponding yeast phase, but Paracoccidioides brasiliensis and Histoplasma capsulatum tended in the opposite direction. 3. EJects $Fatty acids on Dermatophytes

Medium chain-length fatty acids, particularly undecylic (C,, ) and caprylic ( C 8 ) , are commonly used in the therapy of fungal diseases due to dermatophytes. Rothman et al. (1957) linked the spontaneous cure of scalp ringworm of children at puberty with the increase in the free fatty-acid content of the sebum. Baxter and Trotter (1969), examining the responses of certain keratophilic fungi to fatty acids from various keratin sources, found human- hair fat exerted a strong growth inhibition, whereas pigeon-feather fat had very little effect. Shorter chain-length fatty acids were shown to be important fungistatic agents but were not the only active components in the complete fat extract. Squalene was also markedly inhibitory. Microsporon adouinii, the cause of scalp ringworm of children, was particularly sensitive to fatty acids and showed 15%greater inhibition of growth on adult hair than that of children. Adult human-hair fat, which was the type most inhibitory to all of the fungi tested except Trichophyton rubrum, contained a high proportion of C,, fatty acids and differed from other fats in the proportions and amounts of shorter-chain fatty acids, especially C,, and C15. D.

DISCUSSION

That there is no clear barrier between fungi pathogenic to Man and those better known as soil saprophytes, spoilage organisms or plant pathogens, is increasingly obvious. The barrier has been rather between the groups of workers concerned with the different types of hosts, the

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plant pathologists and the medical mycologists. Emmons (19601, in his essay “The Jekyll-Hydes of mycology”, already pointed out, “Medical mycology does not deal with a bizarre group of fungi but with moulds and yeasts with which we may be in almost daily contact”. Investigations of the pathogenicity to humans and mice of Schizophyllum commune (Greer and Bolanos, 197 l), a familiar basidiomycete normally growing on wood and intensively investigated with regard to its genetics and morphogenesis, further emphasize the point. Similarly, Botryodzplodia theobromae (Sphaeropsidales), a common saprophyte and secondary parasite of plants, has been implicated in keratomycoses and isolated from keratinous nail layers (Rostrepo et al., 1976). Many other nondermatophyte moulds are associated with nail lesions. Rostrepo et al. (1976) quote this as “one more example of the intrinsic capacity of fungi to adapt themselves to multiple and diverse organic substrates”. A vast number of other examples could be cited. Species of Aspergillus can cause deep mycoses in almost every body organ (Austwick, 1965) and Aureobasidium (Pullularia) pullulans, a ubiquitous, yeast-like leaf-surface species, has been isolated from inflamed joints of patients with rheumatoid arthritis, from onychomycosis (Vieira, 19591, from cutaneous infection of porcupine (Salkin et al., 1976)and in symbiotic association with insects (Cooke, 1959). So far little is known of the physiology of pathogenic fungi in uiuo, and the subject of the involvement of lipids in host-parasite relationships of fungal pathogens of Man and animals has hardly been touched. Because of the more convenient manipulation of the experimental system, much greater progress has so far been made in studies on plant pathogens, the results of which may provide pointers of value in the investigation of pathogenicity in animal tissues. There is a need for basic information on incorporation of animal host metabolites by pathogenic fungi. Comparative studies of host-parasite physioloe based on both plant and animal material could greatly contribute to an understanding of the activity of fungi as biotrophs and pathogens and the role of lipids in such relationships.

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Compatible Solutes and Extreme Water Stress in Eukaryotic Micro-Organisms A. D. BROWN Department of Biolog y, University of Wollongong Wollongong, N.S.W. 2500, Australia

. . . . . . . I. Introduction . . . . . 11. Role of Polyhydric Alcohols . . . . . . . . . A. General . . . . . . . . B. Specific 111. Physiology of Xerotolerance . . . . . A. Xerotolerant Yeasts . . . . . . B. Xerophilic Yeasts . . . . . . C. Xerotolerant Fungi . . . . . . D. Halophilic Algae . . . . . . E. Intermediate Xerotolerance . . . . IV. RegulationofCompatibleSolute Accumulation . . . . . . . . . V. Summary . VI. Acknowledgements . . . . . . . . . . . . . . . References

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I. Introduction

Growth, as distinct from survival, of unicellular organisms requires them to be in liquid suspension at a liquid/solid or liquid/gas interface, or on the surface of a gel, because they must obtain their nutrients directly from solution. The exceptions are protozoa which engulf particulate food but, in any case, this method of feeding is simply an ecological extension of a basic situation. The consequence of taking food from solution is that the water relations of growing microorganisms are determined by the concentration of solutes in the 181

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aqueous solution in which they are growing. An “arid” environment for a growing protist is thus a concentrated solution. The problem is qualitatively different from that of multicellular plants and animals which respond to arid conditions primarily by complex physiological mechanisms of scavenging for and preventing loss of water, although some halophyte plants make limited use of‘microbial mechanisms at the cellular level. The problem is also different from that of microbial survival of desiccation as, for example, in soil. Under those circumstances, microflora are faced with the problem of the loss of water from physically discrete zones. When liquid water is present in soil, it is usually readily available thermodynamically, that is to say the solutions which occur in pockets of occluded water are dilute. When the water has evaporated, the residual microflora must equilibrate thermodynamically with whatever gas phase they are in contact, and their future depends on their ability to survive equilibration. The present article is not concerned with survival; it is concerned with microbial activity under the former type of condition, that is in a concentrated solution. There are several parameters by which the amount of thermodynamically available water in a solution can be described. Of these, biologists have commonly used osmotic pressure, water activity and water potential. Microbiologists have tended to use the first two; plant physiologists have tended to use osmotic pressure and water potential. All three parameters are rigorously interrelated (see, for example, Nobel, 1970). There are some conceptual problems associated with the use of osmotic pressure which, in the writer’s view, amount to a significant disadvantage (Brown, 1976). The parameter, water activity (aw), received a major impetus in microbiology from Scott (1957) who applied it to the study and prevention of microbiological spoilage of foods. Water activity lacks the mechanistic overtones of osmotic pressure (see Brown, 19761, and it has some practical advantages over water potential to which, via its natural logarithm, it is directly related. The advantages include the relative ease by which it can be experimentally determined and mathematically manipulated. The rigorous derivation of a, is described in a number of publications (such as Nobel, 1970) and has been summarized by Brown (1976). For present purposes, it is sufficient to state that the water activity of a solution is numerically equal to 0.01 multipled by the percentage relative humidity (R.H.) of the atmosphere with which the solution has equilibrated. For example, an atmosphere of 95% R.H. will equilibrate with

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a solution of 0.95 a,. Pure water, equilibrates with an atmosphere of 100%R.H. and has an a, value of 1.OO. A major limitation on the use of a,, which will become apparent below, is that enzyme activity in concentrated solutions is not generally a function of a, but is, instead, a function of the type and concentration of solute used to adjust a,. Nevertheless, water activity is a valuable and probably the most useful parameter currently available to describe microbial water relations in complex media. Multicellular organisms can rarely tolerate conditions which lower cellular water activity below about 0.99 a, (which corresponds to about 0.3 molal sodium chloride or about 0.5 molal sucrose). This is also true of many micro-organisms, but there are others including common bacteria, yeasts and moulds which will grow at water activities down to about 0.90 (which corresponds to about 2.8 molal sodium chloride or 4.0 molal sucrose). Some common bacteria, such as staphylococci are more tolerant (down to about 0.85 a,), and, in general, yeasts and fungi are more xerotolerant than bacteria. Over and above these organisms, however, there is a small group with extreme tolerance of low water availability. Outstanding among them are : (i) the extremely halophilic bacteria which are distinguished by an absolute requirement for sodium chloride in excess of 2-2.6 M and an ability to grow readily in saturated sodium chloride (0.75 a,). The growth characteristics of these bacteria suggest that their limit is determined by the solubility of salt rather than their physiology; (ii) the xerotolerant yeasts and moulds which are illustrated by Saccharomyces rouxii and Xeromyces bisporus, respectively. In growth media adjusted with suitable sugars, these remarkable organisms will grow at water activities as low as 0.62. In the presence of other solutes, their tolerance is much less. Unlike the halophilic bacteria, however, they do not in general have an effective upper limit. Thus they will grow in media from slightly less than 1.00 a, down to their lower limit. In other words, they tolerate rather than require high solute concentrations (there are some exceptions which are discussed on p. 212); (iii) the halophilic algae which are best represented by some species of the genus Dunaliella. Like their bacterial counterparts, they tolerate saturated sodium chloride and, under appropriate conditions, have a requirement for a minimal salt concentration as well. This minimum can be varied, however, and there are many fundamental differences between the halophilism of algae and bacteria. This review is concerned with the physiological basis of the environmental tolerances of the micro-organisms in groups (ii) and (iii)above.

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At the time the invitation to write it was received, the physiological basis of xerotolerance was fairly clear and had the task been undertaken immediately it would have been quite straightforward. With the passage of some time, the accumulation of more information and perhaps some wisdom, the explanation has become far less clear, although some of the essential truths remain. This review will therefore be in the nature of a progress report and will contain substantially more speculation than was originally contemplated. The author has recently reviewed the subject of microbial water stress (Brown, 1976) with emphasis on physicochemical aspects of the problem. The present article complements that, and places more emphasis on biological and regulatory aspects. It is also largely confined to eukaryotes. 11. Role of Polyhydric Alcohols A.

GENERAL

Micro-organisms which grow in an environment of low water activity have an interior of comparably low water activity; in fact, it is usually a little lower as revealed by the turgor of flexible cells. There is no doubt whatever on this point in spite of the uncertainties of a generation ago (see Brown, 1964, 1976). It is self-evident that, for a micro-organism to grow, its enzyme complement must be functional; in the present context this means that it must be functional at a greatly decreased water activity. Therefore, no cytoplasmic solute can be generally excessively inhibitory at the prevailing concentration and obviously the concentration of some must be very high. Such a situation can be achieved basically either by producing enzymes which are inherently resistant to inhibition or by producing an intracellular environment which, for one reason or another, is not excessively inhibitory. This argument has been expanded by Brown (1976). The extremely halophilic bacteria function by employing both mechanisms but, on present evidence, the eukaryotes depend solely on modification of their interiors. This they do by accumulating a " compatible solute". Xerotolerant eukaryotes accumulate high concentrations of polyhydric alcohols, or related compounds, in response to a water stress; when the stress is extreme, the polyol is usually glycerol. Moreover, as discussed on p. 188, polyols, especially glycerol, confer a remarkable

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degree of protection on enzymes at low a, values. So much is certain. The trouble is that non-tolerant species of the same eukaryotic genera can also accumulate glycerol in response to a water stress. Recognition of the physiological significance of polyol accumulation firstly requires evidence that corresponding enzymes in tolerant and non-tolerant organisms are essentially similar in their “water relations”. This has been done for an NADP-specific isocitrate dehydrogenase from Saccharomyces rouxii (tolerant) and Sacch. cerevisiae (non-tolerant) (see Brown, 1976) and for glycerol- and glucose 6phosphate dehydrogenases from Dunaliella teriolecta (marine) and D , uiridis (halophilic) (Borowitzka and Brown, 1974; Borowitzka et al., 1977). Although, in a strictly statistical sense, such a small sample is scarcely representative of the full complement of a cell’s enzymes, 1 have argued elsewhere (Brown, 1976) that it is indeed sufficient to discount inherent differences in protein chemistry as an explanation of different environmental tolerances. For example, anyfunctional enzyme selected at random from extremely halophilic bacteria will reveal peculiar salt requirements. Moreover, I am not aware of any report which suggests that any enzyme from a xerotolerant organism is itself inherently xerotolerant. There is also the possiblity that xerotolerant species as a group depend on different central metabolic pathways from those of nontolerant species and that enzymes which catalyse the different pathways are inherently different in their “water relations”. Xerotolerant yeasts do have some metabolic differences from their non-tolerant counterparts (see p. 198) but there is no evidence that the enzymes associated with these differences have peculiar “water relations”. The major consequence of the metabolic difference seems to lie in polyol production. There is no evidence of any such difference between halophilic.and non-halophilic species of the alga, Dunaliella. The final possibility among the improbable mechanisms is that tolerant organisms “bulldoze” their way through their environmental exigencies by producing and diverting a lot more energy into growth so that a certain degree of inhibition leaves them with enough in hand to keep going. In fact, the opposite seems to be true. Under optimal conditions, both xerotolerant yeasts and halophilic algae grow much more slowly than their non-tolerant equivalents (Anand and Brown, 1968; Borowitzka and Brown, 1974). When the foregoing possiblities are eliminated, it becomes virtually axiomatic that the intracellular environments of the two broad groups

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of organisms must be sufficiently different to enable enzymes to function more effectively in tolerant than in non- tolerant organisms under conditions of desiccation. The differences between them include polyol content but, as the following discussion will show, this is not always very great and it cannot be the only relevant difference. The others are less clear. Accumulated intracellular polyols have at least three major physiological functions, namely they act as: (i) osmoregulators, (ii) food reserves and (iii) protectors of enzyme activity at low levels of a,. Substances with the third function have been called “compatible solutes” (Brown and Simpson, 1972). Compatible solutes must always behave as osmoregulators, however, and the definition can logically be extended to include solutes which combine functions (i) and (iii). Potassium chloride (and K + ) has this role in halophilic bacteria (Aitken and Brown, 1972). Osmoregulation is a concept which is used more often by plant and animal physiologists than by microbiologists. If a cell is subjected to a water stress by exposing it to a concentrated solution (low a,) it will either (a) equilibrate thermodynamically with the solution by losing water and perhaps by admitting some of the extracellular solute(s),in which case it has died or become dormant, or (b) suffer a temporary loss of water but use energy to accumulate a solute(s1to a concentration thermodynamically commensurate with the extracellular a,. The solute can be a metabolite which is retained or an extracellular substance which is accumulated. The effect of solute accumulation is to lower intracellular a, values to the extent that water again enters the cell to achieve thermodynamic parity with the outside. Thermodynamic parity normally includes a factor for an appropriate level of turgor pressure. If the actual regulatory mechanism is for the moment ignored, the process of osmoregulation is thus a simple one. The essential criterion is that the osmoregulator is largely retained within the cell or, more precisely, its intracellular concentration is much greater than its extracellular concentration. Normally the process is associated with the effective exclusion of the major external solute (e.g. NaCl). In a dilute environment, all intracellular solutes, that is “pool” intermediary metabolites, proteins, nucleotides and salts, contribute to the osmotic or water status of the cell. With increasing desiccation, however, a single substance tends to emerge as the osmoregulator.

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Some examples of osmoregulators include a -galactosylglycerol (Kauss, 1967, 1973), cyclohexanetetrol (Craigie, 19741, aspartate, glutamate, y-aminobutyrate and proline (Tempest et al., 1970; Christian and Hall, 1972; Measures, 1975; Singh et al., 1973a, b; Stewart and Lee, 1974) and miscellaneous polyols (Lewis and Smith, 1967). Under extreme conditions, the osmoregulators are K + and KC1 in halophilic bacteria and glycerol in extremely tolerant yeasts and algae (Brown, 1976). The situation in xerotolerant moulds has not been investigated, but an educated guess suggests that they too should accumulate glycerol. It should be noted, however, that there is no a priori reason for attributing any specificity to an osmoregulator. Theoretically, any solute should do, provided of course, that it is not toxic. The significance of this condition shows up in function (ii) and especially (iii). There is ample evidence that polyols do function as food reserves in eukaryotes. Lewis and Smith (1967)have discussed this at some length for polyols other than glycerol in fungi, algae, lichens and higher plants. Corina and Munday (197 1) attempted to distinguish between the functions of mannitol and ribitol in Aspergillus clauatus. They attributed a storage role to mannitol but suggested that ribitol accumulated primarily as a result of “hydrogen-acceptor mechanisms”. There are constraints on the use of polyols as food reserves. To the extent that they act as compatible solutes, they are needed for physicochemical reasons as long as the organism is exposed to a low a,value. Consumption of accumulated polyols should thus be followed by continued growth of the organism only after dilution (i.e. raising the a , value) of the environment. It is probably no co-incidence that polyol accumulation commonly occurs together with storage of other carbohydrate reserves such as trehalose, glycogen or starch (see, for example, Lewis and Smith, 1967). Since a substantial part of the ensuing discussion deals with the interaction between enzyme proteins and a range of non-electrolytes including polyhydric alcohols, it is sufficient to state here that the polyols, in general, have a remarkable ability to preserve enzyme function at low levels of water availability. Glycerol is outstanding in this respect, and there is no doubt that, for such reasons, glycerol accumulation is a major determinant of the special water relations of xerotolerant micro-organisms.

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A. D. BROWN B.

SPECIFIC

Enzymological Role of Polyols As stated earlier, microbiologists with an interest in microbial water relations have commonly discussed the problem on the basis of some measure of the amount of water which is thermodynamically available; this is useful when considering populations in a complex environment. There were, however, some early disparities in the evidence of‘whether or not water availability was itself the primary determinant of the “water relations” of subcellular processes, especially of enzyme-catalysed reactions. For example, Conover ( 1969) studied the effect of concentrated solutions of non-electrolytes (glycerol, ethylene glycol and sucrose) on reactions associated with oxidative phosphorylation in beef-heart mitochondria, and concluded that “inhibition appeared to be related solely to the water content of the solvent mixture”. 0ther compounds, dimethyl sulphoxide and dimethylf‘ormamide,were more inhibitory. On the other hand, Anand (1969)reported in effect that solute type and concentration, not a, values, were the variables determining the V,,, of yeast isocitrate dehydrogenase in solutions of sucrose and of polyethylene glycol (mol.wt. 200). In 1972, Brown and Simpson contrasted the inhibition by glycerol of the NADP-specific isocitrate dehydrogenase of Saccharomyces rouxii with that caused by sucrose, and demonstrated unequivocally that the effect was attributable to the solute, not to a , . Glycerol at water activities down to 0.92 caused negligible inhibition of enzyme activity whereas sucrose caused about 90% inhibition under the corresponding conditions. At that time, the term “compatible solute” was introduced. In the meantime, studies of enzyme function in Halobacterium salinarium had revealed a direct interaction between salts and the corresponding isocitrate dehydrogenase of that organism (Aitken et al., 1970; Aitken and Brown, 1972). Specifically, potassium chloride, which accumulates in halophilic bacteria, inhibited to a very limited extent at physiological concentrations whereas sodium chloride, which is largely excluded, was severely inhibitory at comparable concentrations. Moreover, the kinetics implied a direct participation of salt in the reaction sequence of the enzyme (Aitken and Brown, 197 2). Direct interactions between electrolytes and proteins do not normally raise any conceptual problems; we are accustomed to thinking of salt effects in this way and salt-enzyme interactions (in dilute solu-

EUKARYOTIC WATER R ElATlO NS

I

I

I

I 2 3 Polyol concentration ( molal)

I 4

189

-

OO

I 2 3 Sugar concentration ( molal )

4

FIG. 1. A plot of reciprocal enzyme activity against concentration of selected nonelectrolytes. (a) Polyhydric alcohols. The lines, from top to bottom, represent glucitol, co-incident), erythritol, glycerol, ethylene glycol and methanol. arabitol and ribitol (0, (b) Selected sugars. The lines, from top to bottom, represent sucrose, ribose, xylose, glucose and fructose. The figure does not indicate that disaccharides generally gave the steepest slopes. For example, maltose, which is less soluble than the other sugars and is not illustrated, gave a slope (up to 2 molal) similar to that of xylose. Note the different scales on the ordinates of Figs. (a) and (b). The figure was derived from results of Simpson (1976).

tions) are normally assessed at this level. Indeed, the “water relations” of organisms in saline environments are more commonly discussed in terms of salinity than of water availability. The interaction of nonelectrolytes with enzymes has been studied far less than that of salts, however, and conventional microbiological thinking has tended to overlook this as a possible major determinant of microbial water relations. The absence of long-range powerful electrostatic forces is no doubt a factor which influences this attitude. Simpson (1976) investigated the effects on the NADP-specific isocitrate dehydrogenase from Sacch. rouxii of a series of acyclic polyhydric alcohols, partially hydroxylated di- and triols, mono- and disaccharides. All caused some degree of inhibition and in all cases, except for the straight-chain aldoses, erythrose and glyceraldehyde,

A. D. BROWN

190

inhibition was fully reversible. A comparison of the inhibition patterns caused by some of these solutes is shown in Fig. 1 . Several generalizations can be drawn from this figure and from other related results which are not illustrated. These are: ( 1) fully hydroxylated polyols, as a group, are less inhibitory than sugars although their respective ranges overlap (fructose

O r

0

FIG. 2. A plot of the slope (of reciprocal plots of enzyme activity against polyol concentration, as in Fig. 1) against chromatographic R f value in butanol-acetic acid water (6 : 1 : 2, by vol.). Points, from left to right, represent glucitol, mannitol, arabitol, ribitol, erythritol, glycerol and ethylene glycol. A plot of slope against molecular weight (or chain length) is sigmoidal (see Brown, 1976). Slope was calculated from Simpson's (1976) results (see Fig. 1) and has the dimensions (mg.min).(pmol. molal)-'. Values for &were obtained from Block et al. (1958).

being about as inhibitory as glycerol); inhibition by the polyols was always linear with respect to polyol concentration but that caused by some sugars, conspicuously ribose and sucrose, was nonlinear (concave up): (2) inhibition caused by the partially hydroxylated di- and triols (not illustrated) was biphasic, the steeper slope occurring at higher solute concentrations: (3) the inhibition caused by the fully hydroxylated polyols increased with molecular weight, but inhibition by sugars was unrelated to molecular weight: (4) of all the solutes

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examined glycerol and fructose were by far the least inhibitory of those known to occur naturally in significant concentrations. The relation between chain length (molecular weight) of the polyols and inhibition is sigmoidal (Brown, 19761, but a linear relation obtains between inhibition (slope) and chromatographic R, value of the solute (Fig. 2). The significance of Rfvalue is that it is related to an oil-water

Ethylene glycol

0 Methanol

I

I

I

I

2

3

4

u 5 6

Dielectric constant ( slope, E /C 1

FIG. 3. A plot of the slope of reciprocal enzyme activity (as in Fig. 2) against “slope” of dielectric constant, that is the rate of change of dielectric constant as a function of solute concentration in aqueous solutions. The only other relevant solute for which dielectric constants were found was sucrose, the value for which does not fall on the line. The extent of its departure is illustrated by the following numerical values of the function, “slope enzyme activity”/(“slope 6 ’ )- 1, (the denominator being corrected for the x-intercept of 1): methanol, 0.33; ethylene glycol, 0.44; glycerol, 0.36; glucose, 0.34; sucrose, 1.44. This figure was compiled from results of Simpson (1976) and dielectric constants collated by her.

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A. D. BROWN

partition co-efficient although, in a series of homologous compounds such as the polyhydric alcohols, there is also a major contribution by molecular weight. Nevertheless, the effect of the variable, R , , in straightening out a relation between chain length and inhibition suggests that oil-water partitioning and hence, presumably, hydrophobic interactions, are involved at some level in the enzyme inhibition caused by the polyols. Additional evidence on this point is given in Fig. 3 which shows a relation between inhibition (slope) and dielectric constant (slope) of aqueous solutions of the solute. Data were available for five relevant solutes, namely methanol, ethylene glycol, glycerol, glucose and sucrose. The significance of slope in comparisons of this kind is that i t expresses the rate of change of the relevant parameter (reciprocal reaction velocity o r dielectric constant) as a function of inhibitor concentration. Figure 3 (and its legend) shows that glucose forms a series with the polyols but that sucrose does not. The departure of sucrose from the numerical relation is substantial. The sugars as a group gave n o correlation between inhibition and molecular weight o r chromatographic R, value. The other factors which must therefore determine their interaction with the enzyme presumably stem from the stereochemistry of the sugars. Stereochemical implications have been discussed in some detail by Simpson (1976) who has proposed some correlations, and by Brown (1976) who was a little less optimistic. Whatever the real significance of stereochemistry, however, the evidence to this point is reasonably clear that the acyclic polyols and at least one sugar inhibit isocitrate dehydrogenase in a pattern which is systematically related to their “hydrophobicity” and to the dielectric constants of their aqueous solutions. Since the solutes function solely as inhibitors, that is to say they d o not also have a n activating role as d o salts for halophile enzymes (see Brown, 1976), it is scarcely surprising that the more inhibitory solutes appear to bind more tightly to the enzyme than d o the less inhibitory solutes. This is shown in Table 1 which lists apparent inhibitor constants for the NADP-specific isocitrate dehydrogenase of those nonelectrolytes which gave “linear” inhibition. The values were derived kinetically and are apparent constants inasmuch as they were obtained with subsaturating substrate concentrations. They show clearly, however, that the inhibitor constants o f the least inhibitory solutes are very high indeed and physically impossible to achieve in some cases (pure methanol is 31 molal, pure ethylene glycol 16 molal). Obviously, iso-

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TABLE 1. Apparent inhibitor constants ( K , ) of yeast isocitrate dehydrogenase with polyols and selected sugars Solute

Inhibitor constant Ki(mola1)

Methanol Ethylene glycol Glycerol Erythri to1 Arabi to1 Ribitol Sorbitol (glucitol)

90 42 26 12 9 9

Fructose Glucose Xylose Rhamnose Arabinose

25

8

9

4 3 2

The selection was of- those sugai-s which gave linear plots of 1/V against ill. The values were calculated from results of Simpson 11976). They are apparent constants obtained for h e d concentrations of NADP+ and o-isocitrate (the active isomer) of 1 and 2 mM. respectively.

citrate dehydrogenase would scarcely be affected at least over short time intervals, by any concentrations of these solutes which could be achieved in practice. Table 1 also reveals a virtual identity of K l ( g , \ ~ c l , ~ l l and Kl(~luctose) ; it is probably no co-incidence that the lowest values of a,, at which yeasts and moulds will grow have been reached in solutions of fructose. The value of the inhibitor constant is affected by substrate concentration. Simpson's ( 1976) more detailed kinetic studies were confined to comparisons of glycerol and sucrose, the latter giving a severe, complex type of inhibition. The true value for Kligl\cc.lol) (obtained with both substrates saturating) is about 13 molal. A similar derivation for sucrose could not be made with any accuracy because of the nonlinearity of the plot of l/Vl,,,, against sucrose concentration, but a graphical extrapolation suggested a value of 1. e l .5 molal. To this point, the evidence has been confined to the inhibitory action of non-electrolytes over a period of some minutes. I t is self evident, of' course, that compatible solutes should not only be non-inhibitory in

194

A. D. BROWN

this sense but neither should they inactivate or “denature” an enzyme over a longer period. The distinction between inactivation and inhibition can be easily demonstrated with salts and the NADP-specific isocitrate dehydrogenase of Halobacterium salinarium. Salt is required to maintain this enzyme in an active form; high concentrations of either sodium chloride or potassium chloride will do this equally well. Concentrations of sodium chloride which cause almost complete inhibition will stabilize the enzyme indefinitely. Similar concentrations of potassium chloride will cause very little inhibition (Aitken et al., 1970; Aitken and Brown, 1972). The apparent discrepancy between the extremely low affinity of methanol for isocitrate dehydrogenase and the general toxicity of methanol is likely to be resolved partly at this level. Simpson’s (1976) evidence on inactivation is limited and essentially qualitative (apart from the irreversible inhibition caused by the straight-chain aldoses). She showed by gel electrophoresis that crude preparations of yeast isocitrate dehydrogenase exist in mutliple forms. The slower moving bands are apparently polymers since they are converted to the fastest band on ammonium-sulphate fractionation. Both glycerol and sucrose at concentrations above 0.4 molal separately have a similar effect, that is, they depolymerize the slower bands. Contaxis and Reithel ( 197 1) had previously shown a depolymerizing effect of several polyols on jack-bean urease. The rate of dissociation of urease polymers in aqueous solutions (go%, v/v) o f the polyols increased in the order: glycerol, propanediol, ethylene glycol. There was little loss in enzyme activity during this process. O n prolonged exposure (twodays), conspicuous differences between glycerol and the other two solutes became apparent. Under these conditions, the enzyme reassociated “abnormally” (my terminology) in the two diols to give an enzymically inactive polymer but, in glycerol, the enzyme remained active indefinitely. Douzou (19741, in studies of enzyme activity at subzero temperatures, used ethylene glycol, propanediol and glycerol (separately) to lower the freezing point of his reaction mixtures, and reported that these polyols were much less effective “denaturing” agents than the monohydric alcohols. These and other aspects of enzyme inactivation have been discussed by Brown (1976). Although the evidence so far is by no means complete, the picture which emerges about solute compatibility, broadly speaking, is as follows. A substance which functions as a compatible solute must neither inhibit excessively nor inactivate enzymes. The short-chain

EUKARYOTIC WATER RELATIONS

195

fully hydroxylated polyhydric alcohols (allowing methanol as the first member of this series) were by far the least inhibitory of the substances tested, and had very low affinities for the enzyme protein. Affinity (and hence inhibition) of the polyols and at least one sugar for the enzyme is related to physicochemical properties which suggest that hydrophobic interactions are at least partly responsible for the binding which occurs within short time intervals. Hydrophobic interactions in no way exclude, and might even embrace, solvent perturbations such as those discussed by Brown (1976). Hydrophobic interactions were also proposed by Lanyi (1974) to explain the specific action of salts at high concentrations on bacterial enzymes. Affinity of sugars for the enzyme is dominated by stereochemical factors which have not been fully evaluated. Information about inactivation is of a more general nature. Ethylene glycol, and especially methanol, inactivate enzymes much more severely and much more extensively than does glycerol which, by and large, seems to have little or no action of this kind. The higher polyols must also be assumed to cause little or no inactivation since they occur at substantial concentrations in the cells of many organisms (see Lewis and Smith, 1967). Although sucrose gave the most pronounced inhibition of the reversible inhibitors, it is not generally associated with inactivation as shown by its common use in mitochondrial preparations. It might also be assumed that the other sugars cause little inactivation; the assumption is largely intuitive, however, since there seems to be little direct information on this point. On balance, glycerol emerges as the outstanding candidate for the role of compatible solute since the only solutes found to be less inhibitory cause inactivation. Fructose might be as effective, but it is a much more biochemically reactive substance than glycerol and, under natural conditions, it is not known even to approach the intracellular concentrations which glycerol can attain. The higher polyols have the characteristics of “second line” compatible solutes since they are more inhibitory than glycerol, but probably not much worse as inactivators. Furthermore, since they are reduced products of metabolism, they are relatively stable metabolically and capable of reaching high intracellular concentrations. There remains the question of how widely can these conclusions, drawn largely from a yeast isocitrate dehydrogenase, be applied to proteins in general, Whether the binding of polyols to enzyme proteins occurs by hydrophobic interactions as proposed or by any other mechanism, different proteins can be assumed to have different

196

A. D. BROWN

binding characteristics because of variations in amino-acid composition. For example, Heimer (1973) reported different responses to glycerol of the nitrate reductase from the XD cells of tobacco, Chlorella pyrenoidosa and Dunaliella parva. Glycerol was compared systematically with sodium chloride but not with other non-electrolytes. I t was far less inhibitory than the salt in all cases. Hart et al. ( 197 1 ) examined the effects of glycerol and various diols on the thermal stability of collagen. The alcohols were used over a range of concentrations up to 10 M , and thermal stability was assessed from “melting” temperature. Some solutes, such as propane- 1,Z-diol and butane-2,3-diol, diminished thermal stability whereas others, such as propane- 1,3-diol and ethylene glycol, enhanced it. Glycerol was conspicuous, however, in causing the greatest increase in stability, reflected in a rise of more than2O0C in the “melting” temperature of collagen in 10 M glycerol. Although these results and those already quoted make n o claim to be exhaustive, they emphasize clearly that glycerol neither inhibits excessively nor inactivates enzymes. and can even make a special contribution to the stabilization of protein molecules. Finally, Fig. 4 illustrates the effect of glycerol on the glucose 6-phosphate dehydrogenase of Dunaliella tertiolecta and D . viridis. Although no comparison was made with other non-electrolytes, the polyol was completely noninhibitory under the experimental conditions at concentrations up to 4 M.

FIG. 4. Effect of glycerol concentration on glucose 6-phosphate dehydrogenase from Dunaliella tertiolecta (0)and Dunaliella uiridis (0). The maximum specific activities were 0.183 and 0.053 pmol NADP+ reduced/min/mg protein for Dunaliella tertiolecta and Dunaliella uiridis, respectively. Reprinted from Borowitzka and Brown ( 1974).

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197

This section has emphasized the role of various non-electrolytes as general enzyme inhibitors, and concluded that glycerol is one of the least inhibitory over a very wide concentration range. The emphasis on inhibition should not obscure its physiologically protective role. If glycerol were not present at an appropriate concentration in a solution of low a,, value, then some other solute o r solutes would be. Within a cell, those solutes would be “pool” intermediary metabolites, some inorganic salts and a small contribution from polymeric molecules. Nearly all of these solutes have ionizing groups, all have been shown to be, o r can be expected to be, severely inhibitory at moderately high concentrations. The generally protective nature of glycerol is also illustrated by its use, both experimentally and naturally, as an antifreezing agent in the protection of‘blood cells, spermatozoa, insects and fish (see Schmidt-Nielsen, 1975). Higher acyclic polyols can also protect insects against freezing (Miller and Smith, 1975). 111. Physiology

of Xerotolerance

The term “osmophilic” was introduced in 1912 by Richter (see Onishi, 1963). I t has been useful but it is misleading, firstly because of the connotations which osmotic pressure can have (see Brown, 1976) and secondly because its suffix, “-philic”, implies a requirement for, rather than a tolerance of, a concentrated growth medium. Sugar- and salt tolerance can be used in specific cases, but Brown (1976) proposed the term “xerotolerant” to cover this entire group of microorganisms. The general biology of xerotolerant yeasts and, to a lesser extent, fungi has been discussed by Ingram (19571, Scott (19571, Onishi (1963) and Pitt ( 1975). Although the first three of these reviews are old, little has happened to invalidate the factual material which they contain. In particular, Onishi’s (1963) treatment of the subject is sufficiently comprehensive to enable the present article to confine its attention specifically to those areas of yeast physiology which seem, at least to the writer, to be of immediate relevance to water relations. The natural habitats of the xerotolerant yeasts include floral nectaries; the yeasts are commonly associated with bees and honey. If honey is fermented, xerotolerant yeasts are usually responsible. Their commercial significance lies primarily in the food industry since they can cause spoilage of wine must, syrups and conserves, fruit juices, dessert wines, dried fruits, molasses and malt extract. They are also

198

A.

D. BROWN

used in the preparation of various oriental fermented foods including soy sauce and miso paste. Their habitats therefore normally place them in contact with high concentrations of non-electrolytes and sometimes, in special circumstances, with moderately high concentrations of salt. Their biological significance must include their remarkable ability, shared with the xerotolerant fungi, to thrive at a lower level of water availability than the cells of any other type of orgaism. As already emphasized, this ability is usually the expression of a tolerance rather than a requirement for a low water activity, but there are some apparent exceptions which are discussed in Section II1.B (p. 212). The theoretical considerations and enzymological evidence outlined in the preceding section lead firmly to the conclusion that xerotolerance demands the accumulation of a compatible solute. The remainder of this section will therefore deal with the comparative physiology of that process. A . X E R O T O L E R A N T YEASTS

The predominant xerotolerant yeast species is Sacch. rouxii (see Onishi, 1963). The following discussion is concerned largely with investigations made in the author’s laboratory with that species (strain YA; Anand and Brown, 1968) and, for comparative purposes, with the non-tolerant Sacch. cerevisiae (strain Y4 1 ; Anand and Brown, 1968). Production of polyhydric alcohols by xerotolerant yeasts is well documented by Onishi ( 1963) and Spencer ( 1968) who have discussed in detail environmental factors which influence polyol yields. The earlier investigations of this process, however, were directed entirely to extracellular polyols with emphasis on their possible significance as fermentation products. The possibility that the alcohols might accumulate within the cell and be an influential factor in the water relations of the yeasts was not at that time considered. With the assistance of hindsight, this now seems a little surprising since both Onishi (1963) and Spencer (1968) emphasized the effect which high concentrations of salt or sugar have in stimulating polyhydric alcohol production. Moreover, in 1967, Lewis and Smith published a lengthy review on the distribution, physiology and metabolism of polyols (other than glycerol) in plants and fungi. They discussed a possible osmoregulatory function of the polyols. Brown and Simpson ( 1972) reported intracellular accumulation of polyhydric alcohols by each of a number of xerotolerant yeasts and the

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absence of detectable polyol in any non-tolerant strain. The first analyses were made on yeasts grown in a conventional “dilute” medium (a, 0.997) except for a strain of Zygosaccharomyces nectarophilus (strain YE; Anand and Brown, 1968) which has an apparent requirement for low water activity and was grown at about a , 0.97 in polyethylene glycol (mol.wt. 200). Under these conditions, arabitol was the sole or predominant polyhydric alcohol which accumulated in all of the tolerant yeasts except one. The exception was a small unidentified yeast with a slightly narrower tolerance range than the others. I t accumulated a hexitol, tentatively identified as mannitol. Zygosaccharomyces nectarophilus accumulated glycerol and a hexitol in addition to arabitol. Yeasts which were subjected to quantitative analyses were harvested in mid-exponential growth phase and, in early experiments, washed with water before drying and extraction. The amount ofarabitol which accumulated in the test strain, YA, after growth in a dilute medium and treatment in this way, was 10-14% of the dry mass, after correction for loss of polyol in the washings (about 1% wlw). The intracellular concentration of arabitol was calculated to lie within the range 0.6-0.9 molal (Brown, 1974).When yeast in this state was suspended in phosphate buffer (Na’ + K’, 0.05 M ; pH 6.98) at 3OoC, it leaked arabitol at the rate of 2.8 pmollminlg dry weight yeast. The rate of.leakage was slightly greater in water, and there was evidence of a non-specific effect of extracellular solutes at low concentrations ( 10-3-10-1 M ) in diminishing the leakage rate. The polyene antibiotic amphotericin B (methyl ester) caused arabitol to leak at rates directly proportional to amphotericin concentration in the range 2-8 pglml (Brown, 1974). When the water activity of the growth medium was lowered to a, 0.95 by addition of either glucose o r polyethylene glycol (mol.wt ZOO), there was a large increase in the intracellular content of polyol and also in the amount which leaked during washing. The yeast cannot grow, but it can produce polyol at a, 0.91 in media adjusted with polyethylene glycol. Under these circumstances, there was again a very large increase in the amount of polyol accumulated by the yeast during incubation, and a proportionately large increase in the amount which leaked during washing with cold water. There are technical problems in estimating polyols in the presence of very high concentrations of either glucose or polyethylene glycol, but the preliminary evidence suggested that intracellular concentrations of at least five molal were reached (Brown, 1974).This concentration is similar to that reached by

2 00

A. D. BROWN

K’ in halophilic bacteria (Christian and Waltho, 1962; see also Brown, 1976). Thus the polyol appeared to be functioning as an osmoregulator in the yeast. In addition, there was the evidence, already discussed, that polyols protect enzymes against inactivation and inhibition at low levels of water activity. I t therefore seemed certain that the polyol was the compatible solute for Sacch. rouxii. There was a minor anomaly in the finding that arabitol was the compatible solute for Sacch. rouxii whereas enzymological evidence indicated that glycerol should be a better enzyme protector under extreme conditions. Preliminary analyses (Brown and Simpson, 1972; Brown, 1974) had been both qualitative and quantitative for yeast grown in dilute basal medium, but only quantitative in media of decreased water activity. Later, more comprehensive analyses showed the situation to be more complex than this. In order to diminish analytical difficulties associated with high concentrations of sugar, the yeasts were grown in media adjusted with sodium chloride and confirmatory experiments were made with polyethylene glycol (mol.wt. 200). Both of these solutes have the disadvantage of causing a much narrower a,. tolerance range than is obtained with sugars. Growth of Sacch. rouxii in saline media affected neither total arabitol production nor the concentration of intracellular arabitol, but it profoundly affected the intracellular content of glycerol (Fig. 5 ) . I n basal medium (0.997 a,<),up to 20% of the total intracellular polyol was glycerol; most glycerol, however, accumulated extracellularly in the growth medium. Increasing the salt concentration up to 18% (w/v) resulted in proportionately greater retention of glycerol, but it had relatively little effect o n total glycerol production. O n the other hand, Sacch. cereuisiae, which does not produce arabitol, responded metabolically to increased salinity by synthesizing much more glycerol. Most of this glycerol (in absolute yield, not concentration) was excreted into the growth medium but, over the salinity range common to both species, Sacch cereuisiae retained as much glycerol within the cell as did Sacch. rouxii. In medium containing 10% sodium chloride, it produced 3-4 times as much glycerol as did Sacch. rouxii to achieve this result. The effect of these two types of response on the “retention factor” (ratio of internal to external concentration) is shown in Fig. 6. The factor for arabitol in Sacch. rouxii was essentially unaffected by salinity. The factor for glycerol in Sacch. rouxii increased sharply to very high values, whereas the glycerol retention factor in Sacch. cereuisiae

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201

Sodium chloride concentration lmolal)

InIracellular

lntracellulor

__-_-

Extracellular

088 Final a,

086

value

FIG. 5. Polyol production and accumulation by Saccharomyces rouxii (panel A) and Saccharomyces cerevisiae (panel B) as a function of a, value adjusted with sodium chloride. Open columns indicate production of glycerol, hatched colums of arabitol. The yeasts were grown at 3OoC in a complex medium containing glucose (0.5% w/v) and harvested in mid-exponential growth phase. Similar effects were obtained with polyethylene glycol. These are previously unpublished results of Margaret Edgley.

responded little to increasing salinity and remained in the range 200500. Gancedo et al. (1968) reported a retention factor for glycerol by Sacch. cerevisiae of up to 150, and contrasted this with the much greater permeability of Candida utilis. They concluded that the “real permeability” of C. utilis to glycerol is some lo5 times as great as that o f S a c c h . cerevisiae. The range of permeabilities or retention factors between C. utilis and Sacch. rouxii is obviously very wide indeed. The thrust of the argument so far is that a compatible solute is essential for growth at low a,,,values, and that the intracellular concentration of the compatible solute must respond in direct proportion to the

202

A. D. BROWN Sodium chloride concentration ( m o l a l )

Final ow value

FIG. 6. The “retention factor” (ratio internal concentration to external concentration) ofarabitol(0)and glycerol ( 0 )in Saccharomyes rouxii and glycerol(.) in Saccharomyces cemuisiae as a function of a, value adjusted with sodium chloride. The yeasts were grown at 3OoC in a complex medium containing glucose (0.5%w/v) and were harvested in mid-exponential growth phase. Similar effects were produced by polyethylene glycol. Previously unpublished results of Margaret Edgley.

total intracellular solute concentration. The fact is, however, that this is achieved with glycerol in both the tolerant Sacch. rouxii and the nontolerant Sacch. cerevisiae when they are grown in medium containing either sodium chloride or polyethylene glycol. The question must therefore be asked why it is, if the premise is valid, that the two species are so different in their tolerance of water stress when they accumulate the same amount of glycerol, at least over a limited range of water activity. Inevitably, the answer is complex and is still incomplete. Nevertheless it is possible to identify a primary reason and some subsidiary ones. One of the subsidiary reasons stems from the experimental conditions used in this work, and is related to the initial stress on the inoculum. Under our experimental conditions, each yeast was first transferred from the stock culture (slope) to a dilute basal “preinoculation” medium (aw0.997 1, thence to an “inoculation” medium

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at the same water activity as the culture medium, and thence to the culture medium. Thus the transfer from the “pre-inoculum” to the “inoculum” imposed a transient stress on the yeast, such as described earlier. Subsequent growth depended on the ability of the yeast to survive this stress. Saccharomyces rouxii has arabitol to help it in this transition stage, but Sacch. cerevisiae does not. Part of the limited tolerance of Sacch. cerevisiae might, therefore, be ascribed to its failure to withstand

TABLE 2. Polyol and glucose concentrations during a growth cycle in media of different salt concentrations. Unpublished results of Margaret Edgley

Yeast and medium

Saccharomyces rouxii basal

Sodium chloride (5%w/v)

Growth Concentration of Concentration of Concentration of phase‘ glucose arabitol glycerol in medium (m molal) (m molal) (mmolal) medium cell medium cell

1 2 3

19.3 11.8 0.06

1.5 1.2 0.4

450 450 5 20

1.5 1.3 0.5

350 270 7

1 2 3

24.1 6.7 0.08

0.7 1.4 1.1

460 520 890

0.9 1.7 0.7

1280 1170 36

1 2 3

21.1 15.3 0.07

1 .o 1.3 0.9

7 20 450 680

0.8 1.3 0.7

2420 2900 1960

1 2 3

22.2 12.7 0.07

0 0 0

0 0 0

2.0 1.5 1.8

53 47 47

1 2 3

5.4 1.1 0.03

0 0 0

0 0 0

2.6 7.7 4.4

1580 1020 630

1 2 3 4

3.5 1.3 0.02 0.0 1

0 0 0 0

0 0 0 0

4.6 6.8 9.9 9.9

27 10 2500 2500 1080

Sodium chloride (lO%W/V)

Saccharomyces cerevisiae basal

Sodium chloride (5%w/v) Sodium chloride ( 10%wlv)

~~~~~

‘1 indicates early-exponential growth phase, 2 mid-exponential growth phase, 3 early-stationary phase, 4 advanced-stationaryphase. Similar results were obtained in mediaadjusted with polyethylene glycol (mol.wt. 200).

2 04

A. D. BROWN

the shock of this initial stress. If that is so, it should be possible to “train” the yeast by serial transfer through media of decreasing a, values because it would then have a substantial content of glycerol to act as a compatible solute during the period of adjustment to its new environment. Training was found to extend the tolerance range ofSacch. cerevisiae but only to a very limited extent; in fact, it could be trained to grow in 12.5%but not in 15%(w/v)sodium chloride (Margaret Edgley, unpublished results). Edgley has also observed that Sacch. cerevisiae will grown well on “synthetic honey agar” (containing 48%, w/v, glucose) when streaked from a saline medium ( 10%sodium chloride) whereas it grows with difficulty, if at all, when streaked from basal medium. Another subsidiary difference is also a result of arabitol production by Sacch. rouxii. Under our experimental conditions in which the yeasts are grown in batch culture, the end of exponential growth is associated with exhaustion of glucose from the medium. When this happens in media of low a , , Sacch. rouxii consumes glycerol and presumably uses it as an energy source. Both intra- and extracellular glycerol concentrations are greatly diminished under these circumstances (Table 2). At the same time, extracellular arabitol is taken up by the yeast while intracellular arabitol content increases. There is possibly a specific solute effect, however, since arabitol uptake from media adjusted with polyethylene glycerol is uncertain (Margaret Edgley, unpublished results 1. Thus, under conditions of water stress, exhaustion of a n energy source under aerobic conditions can lead to consumption of the primary compatible solute, glycerol, and substitution of arabitol as a “second line” compatible solute. Since arabitol is not such a good enzyme protector as glycerol, metabolic activity should be diminished by arabitol’s assumption ofthe role of major compatible solute but, in the stationary phase, this would happen anyway. Arabitol can therefore serve not only as a compatible solute during the transition from a dilute to a concentrated environment but also as a reserve compatible solute capable of maintaining viability when glycerol is used as a food reserve at low ah, values following exhaustion of the extracellular energy source. Oxygen is essential for this sequence of events. Saccharomyces cerevisiae does not have this option since i t does not produce arabitol. The response of this species to exhaustion of the glucose supply in media of low a, value is dependent on solute concentration. I n medium containing 5% sodium chloride, near exhaustion of the glucose supply was accompanied by a 40% decease in

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cellular glycerol content, and uptake of’some glycerol from the growth medium. In medium containing 10% sodium chloride, however, disappearance of glucose led to halving of the cellular glycerol content without any compensating uptake of glycerol from the medium (Margaret Edgley, unpublished results). Of course, this physiological distinction between the two species has little relevance to a “natural” situation for several reasons, not least of‘ which is the practical impossibility of exhausting the energy source in a concentrated sugar solution when the source ol‘nitrogen is limiting. The major relevant difference between the two species, however, lies in the mechanisms employed to achieve the intracellular concentration of glycerol necessary for growth at low a,, values. Saccharomyes rouxii, the tolerant species, achieves this largely through a conservative mechanism, namely retention of proportionately more glycerol as extracellular solute concentration increases. Lowering water activity therefore does not cause Sacch. rouxii to divert a major proportion of its normal biosynthetic activities to producing additional compatible solute.

Cell moss ( m g dry weight

I

FIG. 7 . Glucose consumption during growth of Saccharomyces rouxii and Saccharomyces cerevisiae in a complex medium containing glucose (0.5% w/v) and sodium chloride as specified. Open symbols indicate behaviour of Saccharomyces rouxii, closed symbols of Saccharomyces cereuisiae. Circles indicate basal medium without added salt (a, 0.997); squares, basal medium containing sodium chloride (1.8 molal, final a, 0.94). Glucose consumption is shown as a function of cell density of- the growing culture. Unpublished results of Margaret Edgley.

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A. D. BROWN

O n the other hand, Sacch. cereuisiae approaches the problem by what might well be called “the American method”. The vastly increased but physiologically wasteful production of glycerol in response to water stress diverts biosynthetic capacity to this end. Since no pit is bottomless, we must conclude that, even if no other factor intervenes earlier to stop growth, the progressive diversion of energy into glycerol production must eventually reach a proportion sufficient to starve other essential biosynthetic activities. At that point, growth will cease. Furthermore, the rate of glucose consumption is itself increased by low water activity (sodium chloride o r polyethylene glycol) in growing cultures of Sacch. cereuisiae but not of Sacch. rouxii (Fig. 7 ) . Sodium ions

I

I

I

1

5

10

15

20

Sodium chloride concentration (% w/v)

FIG. 8 . Total glycerol yield (intracellular and extracellular) as a function of sodium chloride concentration in the growth medium. The yeasts were grown at 30’OC in a complex medium containing glucose (0.5% w/v) and sodium chloride as specified. Glycerol yield is expressed as mol glycero1/100 mol glucose consumed. Open symbols indicate production by Saccharomps T O U X Z Z ; closed symbols by Saccharomyces cereuisiae. Unpublished results of Margaret Edgley.

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207

have long been known to accelerate glucose consumption by yeast (Hunter and Rose, 197 1). Yields of glycerol as a function of salt concentration are shown in Fig. 8. The patterns of response to sodium chloride were very different in the two species. Glycerol yield reached a maximum in Sacch. rouxii at 2.5% sodium chloride, whereas it increased in Sacch. cerevisiae to a salt concentration of 7.5% at which point the yield was nearly twice that given by the tolerant species. Moreover, salt increases lipid production by yeast (Hunter and Rose, 197 1).We have not analysed our species for lipid but the physical state of Sacch. cerevisiae after growth at an elevated salt concentration suggests that this indeed does happen. Saccharomyces cerevisiae also responds to water stress (sodium chloride or polyethylene glycol) with at least one other biosynthetic diversion by producing more of a compound tentatively identified as trehalose. Saccharomyces rouxii produces traces of the same substance but, in that yeast, its content does not respond significantly to water stress (Margaret Edgley, unpublished results). Thus, in Sacch. cerevisiae, there are certainly two and probably at least three specialized biosynthetic pathways which are stimulated, relative to cell growth, by lowering environmental water activity. This does not happen in Sacch. rouxii which retains a balance of its biosynthetic pathways. There is possibly a range of types of response of glycerol production to water stress, even among the xerotolerant yeasts. Onishi (1963) tabulated extracellular polyol (as glycerol) production and yields for six xerotolerant strains. Production or yield increased for three species, remained unchanged for one and declined €or two species in response to 18% sodium chloride in medium in shake cultures. These results are difficult to relate to our own since, not only were the analyses confined to extracellular polyol, but they were made after 610 days’ growth. Nevertheless, one may speculate whether there is a correlation between these patterns of extracellular glycerol production and degree of xerotolerance. The interpretation advanced above suggests that glycerol conservation should be a characteristic of the most tolerant strains. The fundamental difference in glycerol conservation between Sacch. rouxii and Sacch. cerevisiae is obviously related to substantial differences in permeability or transport mechanisms. In the following discussion, the term “permeability” is used in the widest sense without attempt to distinguish between passive, active or facilitated processes. When such

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distinction is necessary, it is made with other terminology. Plasmamembrane permeability is potentially involved in the water relations of the yeasts in at least three ways. These are: (i) the direct specific effect on the amount of glycerol retained by a yeast; (ii) a possible contribution to the general solute status of the yeasts if permeability differences were general and not confined to glycerol; (iii) a possible contribution to a,-mediated metabolic regulation; this is related to (ii). The evidence for major differences in glycerol permeabilities is unequivocal even though we do not yet know details of the mechanisms involved. There is little information available about transport mechanisms for acyclic polyols in yeast. Canh et d.( 1975)reported that the C,-C, acyclic polyols permeated into Sacch. cereuisiae by a process resembling simple diffusion. These polyols were not metabolized and glycerol was not examined. Gancedo et al. (1968)stated that Sacch. cereuisiae admits glycerol by simple diffusion, and explained the better ability of Candida utilis to grow on glycerol by the much greater (by a factor of lo5) permeability of that species to the polyol. This contrasts with some bacteria which take up hexitols by group translocation (Lengeler, 1975a, b) and also possibly with Sacch. rouxii which might well have an active mechanism for transporting glycerol and arabitol. Brown’s (1974) exploratory comparisons of solute uptake by Sacch. rouxii and Sacch. cereuisiae suggested that the former species actively accumulated glycerol whereas Sacch. cereuisiae admitted it passively. In those experiments, the yeasts were incubated to “equilibrium” with glycerol at a series of concentrations up to 2.0 M. The experiments showed that, although Sacch. cereuisiae has a much greater total capacity (which is not necessairly related to permeability) than Sacch. rouxii for glycerol, it accumulated the solute by a passive process. The evidence for this and for the active accumulation of glycerol by Sacch. rouxii lay in the extrapolation of plots of internal versus external glycerol concentrations. The plot for Sacch. cerevisiae extrapolated to the origin, but that for Sacch. rouxii did not; it gave a positive intercept on the axis denoting internal concentration. Although evidence of this kind needs support by more rigorous evidence before an active-transport process can be accepted unequivocally (see, for example, Christensen, 19751, such plots can provide a simple and reliable indicator of active transport provided the measurements are not complicated by vigorous metabolism of the solute. The latter circumstance is significant when internal concentration is measured merely by estimating a radio-isotope.

EUKARYOTIC WATER RELATIONS

209

Nevertheless, there is other evidence for active transport of glycerol and possibly also of arabitol by Sacch. rouxii. Uptake and subsequent dissimilation of glycerol, and uptake of arabitol (without dissimilation), occur in both cases against concentration gradients when Sacch. rouxii exhausts its glucose supply (see p. 204).

Time ( min)

FIG. 9. Glucose consumption by Saccharomyces rouxii and Saccharomyces cerevisiae in phosphate buffer (0.067 M , pH 5.9) at 30OC. Open symbols indicate consumption by Saccharomyces rouxii, closed symbols by Saccharomyces cereviriae. In this experiment, the yeasts were grown at 3OoC with rotary agitation under air, and the washed suspensions were incubated under air. Similar patterns were obtained when the washed suspensions were incubated under pure oxygen or nitrogen and with yeasts which had been grown under pure oxygen. The patterns also persisted over at least a two-fold range of initial glucose concentration. Cell densities for this series of experiments were 1.5-2.6 mg dry wt./ml. Unpublished results of the author.

The permeability differences are wider than this, however. Saccharomyces rouxii is virtually cryptic to lactose, whereas Sacch. cereuisiae admits it freely but passively (Brown, 1974). Of more immediate relevance to their metabolic activities is the pattern of glucose uptake by the two species (Fig. 9). Saccharomyces cerevisiae consumed glucose by a mechanism which gave first-order kinetics with respect to glucose concentration, whereas Sacch. rouxii apparently gave zero-order kinetics; similar patterns were obtained anaerobically. If Sacch. r o ~ x i i ’ s

210

A. D. BROWN

kinetics were not zero order but of a higher order which went unrecognized because of the very small rate of change of slope, then the rate constants of the process were greatly different for the two yeasts. Of course, results of this kind do not necessarily mean that the differences in glucose consumption arose at the level of permeation. The alternative explanation, namely that glucose metabolism accounted for the kinetic differences, implies a wide range of concentrations of free glucose within each yeast. At least under some experimental conditions, glucose-borne 14Crapidly reaches a constant level which, in both species, is independent of extracellular glucose concentration (Brown, 1974). Some free glucose does accumulate in actively growing Sacch. rouxii. A. J. Markides (personal communication) has observed that Sacch. rouxii growing under nitrogen-limitation in continuous culture (harvested and washed twice at O°C with 0.05 M phosphate buffer, pH 7.0) accumulated glucose in the range 0.1-0.2 pmol/mg dry weight yeast. Thus, glucose transport is evidently not the rate-limiting step in glucose consumption by Sacch. rouxii, at least under those conditions. Margaret Edgley (unpublished results) has found free glucose contents in Sacch. rouxii, in the mid-exponential phase of batch culture, to lie within the range 0.02-0.04 pmollmg and to be independent of a, value (sodium chloride or polyethylene glycol). Contrary to some earlier reports (see Hunter and Rose, 197 11, she also found free glucose in Sacch. cereuisiue. In mid-exponential phase cultures in basal medium (0.997 awl, it amounted to 0.005 pmol/mg in 10% (w/v) sodium chloride. Similar results were obtained with polyethylene glycol. On balance, the limited evidence does suggest that the different patterns of glucose consumption by the two species were caused by permeability differences. Rose ( 1975) reported cell-volume measurements of three xerotolerant and three non- tolerant yeasts following their shock exposure to solutions of polyethylene glycol (mol.wt. 200) and of sucrose. The three tolerant species were Sacch. rouxii, Torulopsis apicola and T . globosa; the non-tolerant yeasts were two strains of Sacch. cereuisiae and one of Schizosaccharomyces pombe. The solutes were used at concentrations within ranges equivalent to 0.88-0.98 a,,. The three tolerant yeasts responded similarly. Cell volumes changed more or less linearly with a,. value and were always greater by a constant amount in sucrose. The yeasts were smaller by 2 5 4 0 % (in

EUKARYOTIC WATER RELATIONS

21 1

sucrose) at a, 0.88 than at a, 0.98. On the other hand, cell volumes of the three non-tolerant strains were virtually unaffected by polyethylene glycol but changed sharply in sucrose. These three yeasts were 65-75% smaller at a, 0.88 than at 0.98 in sucrose. Thus, the cells behaved as osmometers in sucrose but not, apparently, in polyethylene glycol suggesting that they equilibrated by admitting this solute extensively. A complication in this comparison is the probable metabolism of sucrose by the non-tolerant species but not the tolerant ones (see Brown, 1974). Corry (1976a, b) also studied volume changes of Sacch. rouxii and Schirosacch. pornbe, and concluded that the extent of uptake of several non-electrolytes was in the order: sucrose < sorbitol < fructose and glucose < glycerol. Interestingly, she observed that heat resistance of the yeasts was greatest in solutions of the least permeant solutes when equilibration was achieved by loss of water (even to the point of plasmolysis) rather than by solute uptake. Rose’s (1975)results again point to general systematic differences in permeability of the two types of yeast, and add to the evidence that relevant physiological differences are not confined to mechanisms of glycerol transport. In general, the non-tolerant yeasts seem to be more freely permeable, or else have a greater equilibrium capacity for nonelectrolytes, than do the tolerant species. I am not aware of comparable information about salts. The question must therefore be asked if part of the susceptibility of non-tolerant yeasts to low a , values lies not only in their difficulty in maintaining an adequate glycerol content but also in their greater admission of extracellular solutes. As already shown, almost every extracellular solute which a yeast is likely to encounter at high concentration in a potential growth medium is more inhibitory than glycerol. On present knowledge, it is difficult to judge the importance of solute uptake even approximately and impossible to discuss it quantitatively. If the assumption is made that turgor pressure is about the same in both Sacch. rouxii and Sacch. cerevisiae under similar growth conditions (as in Fig. 5, p. 201, for example), then there is no reason for expecting a rnaJor difference in accumulation of salt or polyethylene glycol. This is because glycerol, the major contributor to turgor pressure at lowered a,values, has a similar concentration in both species. There could well be a minor but significant difference in salt (or polyethylene glycol) uptake to balance the contribution made by arabitol in Sacch. rouxii. These comments are applicable only to circumstances

212

A.

D. BROWN

generally similar to those depicted in Fig. 5. (p. 201). Under other conditions, such as transition to a new environment, the yeasts could certainly be affected differently by solute penetration so that again there are powerful reasons for looking to the adaptation period for critical differences between the two types of yeast. Onishi (1963) has commented on salt accumulation by tolerant and non-tolerant yeasts. He referred to the early experiments of Conway and Moore (1954) in which growth of non-tolerant yeasts in media containing sodium but not added potassium ions led to a substitution of intracellular K+ by Na+ . He cited other experiments in which such “sodium yeast” colonies could be obtained from Sacch. cerevisiae var. ellipsoideus growing on a solidified medium containing 8% sodium chloride. Onishi also showed that, when Sacch. rouxii was grown in medium containing 18% sodium chloride, i t apparently became more permeable to K+ in the sense that this ion was easily washed from the cells with distilled water. This did not happen when the yeast was grown in the presence of glucose (50%)which gave a similar water activity to 18% sodium chloride. Furthermore, Norkrans and Kylin ( 1969) have shown that the salt-tolerant yeast, Debaryomyces hansenii, excludes Na+ more effectively than does Sacch. cerevisiae but accumulates more K’. They commented, however, that “total salt level of the cells is not sufficient to counteract the osmotic potential of the medium, so that additional osmoregulatory mechanisms must be involved in determining halotolerance”. 9 . XEROPHILIC YEASTS

As stated earlier, there are some yeasts which differ from those already described by having an apparent requirement for low water activity. Onishi reported in 1960 the isolation o f a yeast, Torulo@i halonitratophila, which was obligately halophilic, that is it had a requirement for salt. The yeast was first isolated in a medium containing 18% (w/v) sodium chloride, and “trained” by serial transfer down to 6% salt. It grew with difficulty after a long lag in 3% salt, but would not grow at a lower concentration. All of this was done at 3 O O C . At 20°C, however, the yeast did grow in a dilute medium without added salt. The yeast was described by Onishi as obligately halophilic at 3OoC and facultatively so at 20°C. The effect of non-electrolytes on this aspect of its physiology was not examined but, in the light of later evidence, it seems reasonable to assume that they would have had a similar effect.

EUKARYOTIC WATER RELATIONS

213

In 1963, Onishi discussed in more general terms the relation between temperature and water relations, and noted the general tendency of’ increasing temperature to enhance tolerance of high solute concentrations (see also Brown, 1964). He commented “but it is not too much to say that most of the osmophilic yeasts may be apparently transformed into an obligate type when cultivated at a higher temperature although they show only an osmotolerant property at a lower temperature”. This prediction has been neither verified nor disproved but i t is certainly reasonable. 0ther examples of xerophilic yeasts have been encountered. One is Zygosaccharomyces nectarophilus which, at 3OoC, apparently requires low awvalues; adjustment of a,, value can be made with polyethylene glycol (Anand and Brown, 1968). This yeast will grow in a dilute medium (a, 0.997) in the temperature range 16-23OC (Margaret Edgley, unpublished results). Obligate “xerophilia” has also occurred for different reasons. Koh (1975)isolated a mutant of Sacch. rouxii which would not grow with less than 20% (w/v) sucrose (at 30OC). I t grew fastest in the presence of 60% sucrose (+2% glucose) and, at intermediate sucrose concentrations, it was morphologically variable with a tendency to form filaments. I t was also osmotically fragile which was evidently the cause of its solute requirements; the mutation affected wall synthesis. An example of‘ what, superficially, might seem to be a related phenomenon was given by Karst and Lacroute (1974) who isolated a mutant of Sacch. cereuisiae whose growth was stimulated by any of several monohydric alcohols at a growth temperature of 22OC but not at 34OC. Neither glycerol nor ethylene glycol was effective in this respect. The mechanism was thought to be at the level of RNA synthesis which was apparently impaired by the mutation. Both of these phenomena are different from the temperaturesensitive “xerophilia” discussed on p. 212; the latter is more difficult to explain. It is certainly unlike the requirement of halophilic bacteria for salt which is specific and can be attributed to peculiarities of the protein chemistry of those organisms (Lanyi, 1974; Brown, 1976). Moreover, a temperature-sensitive salt requirement, different from the bacterial one but generally similar to that of the yeasts, can be recognized in halophilic algae (see p. 2 18). I t is generally true that electrolytes, at least partly by virtue ofthe socalled “relaxation effect”, diminish molecular agitation and thereby oppose the effects of raising temperature. I t would be a serious oversimplification, however, to extend this kind of argument to non-

214

A. D. BROWN

electrolytes where other factors are involved in solute-protein interactions. The contribution of the various “other factors” can vary substantially among non-electrolytes of’ closely similar properties as shown, for example, by the results of’ Hart el al. (197 1) with collagen stability. The relevance of that work to the present problem lies in the distinctive ability of glycerol to enhance the thermal stability of collagen. Apart from drawing attention to interesting comparisons of’ this sort, however, it is impossible, at present, to do more than speculate about the mechanism underlying a requirement for diminished a, value. The problem can be rephrased in terms of temperature relations of‘ growth as a function of a, value which, if it helps at all, does s o by emphasizing the complexity of the situation and perhaps by suggesting another way of looking at it. With two types of exception, the ternperature relations of micro-organisms have not been satisfactorily explained. The first exception is that of the temperature-sensitive mutant in which a specific reaction can be identified as the determinant of the special temperature relations of these organisms. The second, in rather more general terms, relates to the thermal stability of proteins and to the thermal tolerance of thermophilic microorganisms. In a much more general sense, microbial temperature relations can be seen as a property of an open system, the continued viability of which requires all of its components (enzymes, metabolites, salts) to remain within certain concentration limits relative to one another; these limits must be strict, even ifquite wide. Since it must be true that all of the processes which constitute such a system do not have identical temperature coefficients, a progressive shift in temperature (up or down) must cause progressive changes in the proportion of these components, relative to one another. Progressive changes of this nature, if allowed to proceed far enough, will eventually lead to at least one component’s transgressing a limit. At that point, growth will cease. Attempting to identify the limiting component/reaction/event is likely to be a futile exercise since it could well be different for every organism. In principle, the same kind of situation should occur in xerophilic yeasts either in response to a change in temperature or water activity. For example, it is quite reasonable to assume that, at 3OoC, there is at least one vital process which is sufficiently fast, relative to the rest of the system, to bring about an unacceptable degree of imbalance and that

EUKARYOTIC WATER RELATIONS

21 5

this reaction has both a temperature coefficient and an “a, coefficient” greater than those of most other processes in the system. Lowering the temperature or lowering the a, value (increasing internal glycerol concentration) slows that reaction down in relation to the rest of the system sufficiently to restore the balance and enable growth to proceed. At the present time, there is no indication o f what this process or processes might be; its identification would probably require a good deal of luck. Nevertheless, because of the relatively restricted nature of “xerophilia”, the chances are fair that the hypothetical unbalancing process might be relatively specific or at least restricted to a smaller range of possiblities than those which determine microbial temperature relations.

C.

XEROTOLERANT FUNGI

Little can be said about these organisms other than to acknowledge their existence, draw attention to articles describing their general biology (Scott, 1957; Pitt, 1975), to the ability of many fungi to produce polyols (Lewis and Smith, 1967) and to predict that extremely tolerant species, such as Xeromyces bisporus, accumulate glycerol to much the same extent as the xerotolerant yeasts.

D.

HALOPHILIC ALGAE

1. General Biology

The term “halophyte” is normally used to describe plants with an enhanced salt tolerance and is usually extended by plant physiologists to include salt-tolerant unicellular algae. Microbiologists more commonly use the term “halophil” for algae, although in neither case is a precise definition of salt relations intended or warranted. In this article, the term “halophil” will be used, partly because of the microbial emphasis but mainly because there are algae which do have an apparent requirement for salt. The term will be used loosely to apply to algae with a recognizable optimum in the general region of 1 M sodium chloride. Its use in this way is somewhat arbitrary. There are no eukaryotes known to have absolute and clearly defined salt requirements such as those which characterize the extremely halophilic bacteria.

216

A. D. BROWN

Nevertheless, there are algae which thrive in saturated saline environments and which are thus bedfellows of the halobacteria. The outstanding algal genus is Dunaliella (phylum Chlorophyta, order Volvocales). Within the same order, the genus Chlamydomonas also includes halophilic species. Both Dunaliella and Chlamydomonas spp. are flagellates but are easily distinguished morphologically by the lack of a cell wall in members of the former genus. The salt relations of Chlamydomonas spp. have received little study, but ecological evidence suggests that members of this genus are generally less tolerant than Dunaliella spp. For example, Brock (1975) has reported that species of Dunaliella are the only algae encountered in the Great Salt Lake (Utah, U.S.A.)when it approaches saturation. Species of Dunaliella (notably D . s u l k ) , alone or jointly with halophilic bacteria, can contribute to the red colouration which frequently occurs at the edges of salt lakes as they reach saturation, although apparently a red colour can sometimes also be caused by ferric oxides or hydroxides (Baas-Becking, 1931). The best known halophilic species of Dunaliella are D . parva, D. viridis and D . salina; their salt relations are not identical. The marine species, D. tertiolecta, has also been the subject of a number of physiological studies. The genus is distinguished not only by the halophilism of some of its species but also by the very wide range of salt

Concentration of sodium chloride in the medium (M)

FIG. 10. Exponential growth rates of (“untrained”; see p. 217) Dunaliella tertiolecta (A) and Dunaliella uiridis (M)as a function of salinity. Reprinted from Borowitzka and Brown (1974).

EUKARYOTIC WATER RELATIONS

21 7

concentrations tolerated collectively by species within the genus and also by individual species. For example, D . salina can grow within the range 0.3 M-saturated sodium chloride (Loeblich, 1972); this is wider than reported for any other type of organism. The same species can also withstand stresses generated by sudden severe changes in salinity (MarrP et al., 1958; Marre and Servettaz, 1959). Under suitable conditions, D. parva will withstand dilution of its suspending solution from 1.5 to 0.6 M sodium chloride without apparent leakage (Ben-Amotz and Avron, 1973a). A simple characteristic of these algae which distinguishes their salt relations sharply from those of the extremely halophilic bacteria lies in the position of their salt optima (for growth rate). Halophilic bacteria require a minimum of about 2.8 M sodium chloride and have an optimum in the region of 4 M (depending on species and temperature). Dunaliella spp., on the other hand, have an optimum very close to the bottom of their salt concentration range (Fig. 10). Thus, over most of their tolerance range they are, in fact, displaying a tolerance of, rather than a requirement for, salt. As Fig. 10 shows, this is true for the marine species. D . tertiolecta as well as the halophile, D. viridis. Furthermore, the tolerance ranges of Dunaliella spp. can be changed by “training”, a capacity which further distinguishes them from their bacterial counterparts and implies that their apparent salt relations are determined partly by their previous experience. Thus, Craigie and McLachlan (1964) extended the upper limit of D . tertiolecta from 2.0 to 2.6 M by serial subculturing through intermediate salt concentrations. Latorella and Vadas (1973) extended the range to 3.6 M salt. Borowitzka et al. ( 197 7 ) showed that D . tertiolecta could also be trained to grow in the presence of 3.5 M salt, the highest concentration tested, and trained D . viridis down to 0.3 M salt. In the experiments of Borowitzka and Brown (1974)and subsequently Borowitzka et al. (1977),the standard inocula of D . tertiolecta and of D. uiridis were grown at 26OC in 0.17 M and 3.4 M sodium chloride, respectively. Thus, there was a considerable stress when the inocula were introduced into some growth media; this stress was greatly diminished by training. I t is, therefore, evident that the so-called salt relations of Dunaliella spp. in many experimental situations are determined by the ability of the algae to survive the transition stage discussed in Section 1I.A. (p. 186)of this review. When D. viridis is transferred from a standard inoculum into a medium more dilute than 3.4 M sodium chloride, a major part 01‘ the

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stress is, of course, osmotic. It is perhaps not surprising, therefore, that this species will accept a single-step transfer from 3.4 M salt to media containing 0.05 M sodium chloride together with sucrose in the concentration range 1.0-2.0 M (Borowitzka et al., 197 7 ) . Growth rate in the sucrose-containing media was very slow, however, and virtually independent of sucrose concentration. Growth-rate measurements revealed an apparent anomaly in the behaviour of D . tertiolecta and D . uiridis, at least in relation to their respective designations as marine and halophilic. As Fig. 10 (p. 216) shows, exponential growth rates of the marine species are much greater than those of the halophil when each is grown in its “normal” (i.e. untrained) salt concentration range. This in itself is to be expected. When they are trained, however, growth rates of D . tertiolecta remain much higher than those of D . uiridis. For example, in 2.0 M salt, the exponential growth rate of D . tertiolecta is about twice that of D . uiridis; in 3.5 M salt, the ratio is about 1.5 (Borowitzka et al., 1977). When D . uiridis was trained to grow at lower salt concentrations, the ratios were 2-3 : 1, again in favour of D . tertiolecta. Thus, what might have been a logical expectation, that the halophil would always show up better at high salt concentrations, was not realized. The low growth rates of halophilic algal species are reminiscent of the xerotolerant yeasts (see Anand and Brown, 1968). It is true that growth rate of (trained) D. tertiolecta declined more rapidly with increasing salt concentration than did that of D . viridis, as shown by the ratios already quoted. Furthermore, a comparison of the growth rates of trained and untrained algae revealed an optimum for D. uiridis of about 1.O M , whereas that of D. tertiolecta remained unchanged at the low level shown in Fig. 10 (p. 216). Temperature modified the salt relations of D . tertiolecta along similar lines to those already discussed for xerophilic yeasts. Increasing incubation temperature from 26OC to 3OoC permitted D . tertiolecta to grow in 2.2 M sodium chloride after a single transfer from a standard inoculum. At 14OC, the highest salt concentation tolerated on a single transfer was 1.4 M (D. S. Kessly, unpublished observations). Thus, when results obtained with trained and untrained algae are combined, differences in the salt relations of the two species can be recognized but, even so, the halophilism of D. uiridis is not a matter of simple definition. I t evidently has a higher optimum than D . tertiolecta, implying a significant salt requirement, but the physiological basis of that requirement has yet to be elucidated (see p. 219). The fact remains

EUKARYOTIC WATER RELATIONS

21 9

that, even without such a salt requirement, D . tertiolecta demonstrably can grow at salt concentrations almost as great and, by extrapolation, just as great as D . viridis. Furthermore, when it does so, it grows faster than D . uiridis. The ecological implications of this are obscure. An important supplementary comment is that Borowitzka et al. ( 197 7 ) used continuous illumination for growing the algae. Latorella and Vadas (1973) have reported that D . tertiolecta requires continuous illumination for growth at salt concentrations above 2.5 M . There is in this at least a superficial similarity with the extra energy needs imposed on Sacch. cerevisiae by water stress. 2. Physiology of Algal Salt Relations At least two enzymes, glucose 6-phosphate dehydrogenase and glycerol dehydrogenase, from the halophilic and marine species respond identically to salt and to glycerol (Borowitzka and Brown, 1974; Fig. 4, p. 196). Thus, although this is scarcely a comprehensive selection of enzymes i t is, for all practical purposes, sufficient to eliminate generalized differences in protein chemistry as the basis of the different salt relations of the two species. The same argument was invoked for the xerotolerant yeasts (see also Brown, 1976). Nor does the difference lie in their gross internal composition, at least as far as compatible solutes are concerned. Because of technical difficulties in estimating salt concentrations within Dunaliella spp., which are mechanically delicate organisms, there has been substantial disagreement about whether or not they admitted salt to a concentration similar to that of the growth medium. Most of the argument has been based on indirect methods rather than on direct analyses. For example, Trezzi et al. (1965) measured volume changes in response to environmental salinity. They concluded that the plasma membrane is freely permeable to salt, and that the organism should therefore contain salt at something like the extracellular concentration. Marri: and Servettaz ( 1959) had already reached a somewhat similar conclusion as did Ginzburg in 1969. Johnson et al. in 1968 used a different indirect approach by assaying the salt sensitivity of various enzyme preparations from D . uiridis, and concluded that the alga did not, in fact, accumulate salt. Ben-Amotz and Avron (1972) made similar observations and also noted the pronounced salt sensitivity of isolated chloroplasts. They interpreted their results to signify compartments of low salt concentration within the algal cell. Borowitzka and Brown

220

A. D. BROWN

( 1974) confirmed

the salt sensitivity of two enzyme preparations from D. tertiolecta and D . uiridis and, in so doing, demonstrated that the response to salt was identical for the equivalent enzymes from both species. The most revealing and direct information about intracellular composition of Dunaliella spp., however, came from a series of demonstrations that various species accumulated glycerol which behaved as an osmoregulator in response to external salinity. Craigie and

Too

Concentrotion of sodium chloride in the medium (MI

FIG. 11. Glycerol (open symbols) and glucose (closed symbols) contents of Dunaliella tertiolecta (triangles) and I). uiridis (squares) as a function of salinity of the growth medium. Reprinted from Borowitzka and Brown (1974).

McLachlan ( 1964) reported glycerol production by D . tertiolecta; Wegman (197 1) recognized its osmoregulatory role, as did Ben-Amotz and Avron (1973b) who showed that it accumulated to a concentration of about 2 M in D.parua when that species was adapted to 1.5 M sodium chloride. Borowitzka and Brown (1974) demonstrated that glycerol accumulated in direct response to environmental salinity in both D . tertiolecta and D . uiridis (Fig. 11). The regulation of glycerol accumulation in all species of Dunaliella so far studied is achieved metabolically, not by the control of leakage. Ben-Amotz ( 1975) could not detect glycerol leakage from D . parua at salt concentrations above 0.6 M. D. S. Kessly (unpublished observations) detected traces of leak-

EUKARYOTIC WATER RELATIONS

22 1

age from D. uiridis below 1.5 M sodium chloride, and from D . tertiolecta below 0.4 M salt. The amount leaked in every case was so slight as to be negligible either as an explanation of the differences between the species or of the mechanism responsible for regulating glycerol accumulation. Calculation of internal concentrations of a solute, especially in a mechanically delicate organism such as Dunaliella, is beset by a number of technical problems not least of which is obtaining a reliable estimate of intracellular water content. Statements of intracellular glycerol concentration should therefore be treated as approximations. The values which have been obtained so far suggest that internal glycerol content is insufficient to balance external salinity. Although this conclusion is probably qualitatively valid, an attempt to identify accurately the magnitude of the discrepancy would be unwarranted on present data. In general terms, however, the high glycerol contents achieved by Dunaliella spp. preclude the possibility that salt would be admitted to anything like the extracellular concentration. If that did happen, it would seriously disturb the thermodynamic balance between the cell and its environment and cause the osmotic disruption of the cell. The small discrepancy which undoubtedly does exist between internal glycerol concentration and external salt concentration should be accounted for by “pool” metabolites and a small but significant uptake of K+ (see Borowitzka and Brown, 1974). Previous evidence for a general enzyme-protective role of glycerol together with its demonstrable osmoregulatory function in Dunaliellu identify the polyol as the compatible solute for members of that algal genus. The conclusion is further supported by the demonstration that, when D. tertiolecta and D. viridis are trained up and down respectively, they accumulate glycerol to a concentration appropriate to their new environment (Borowitzka et al., 1977). In other words, when D . tertiolecta was trained to grow at salt concentrations within the range “normal” for D . uiridis, it accumulated the same amount of glycerol as did D . viridis at those concentrations, and vice versa. Furthermore, the function of salt as a determinant of glycerol production is not specific. In the experiments of Borowitzka et al. ( 197 71, in which D. viridis grew on a single transfer from a standard inoculum to a medium containing salt (0.03 M ) together with sucrose, glycerol content responded to sucrose concentration. In fact, when expressed on the basis of water activity, the glycerol content of sucrose-grown algae was higher than that of algae grown in saline media. Thus, although

22 2

A. D. BROWN

growth rate was a function specifically of salt concentration, internal glycerol content was, broadly speaking, a function of a,, value. This is an additional point of similarity with the yeasts. Up to this point, the interiors of both the yeasts and the algae have been treated as if they were “bags of enzymes” or perhaps buckets of glycerol of uniform overall composition which, of course, they are not. I am not aware of any evidence which bears directly on the compatible solute status of organelles in cells adapted to low a,, values. The argument that microbial cells cannot maintain a dilute interior in a concentrated environment (see Brown, 19641, however, must apply equally to organelles. Thus, the water potential or water activity of nuclei, mitochondria and chloroplasts cannot be greater than that of the surrounding cytosol. What is not known is whether the compatible solute(s) of the organelles is the same as that of the cytosol. Again, theory and limited evidence suggest that it is. Notwithstanding the special ability of xerotolerant yeasts and halophilic algae to retain glycerol, that polyol is classically quite permeant through many types of lipoprotein membranes. There is evidence that the chloroplast envelope of D . salina is no exception to this generalization (Ben-Amotz and Avron, 1972; Pfeifhofer and Belton, 1975). Moreover, total organelle volume is significant; if the organelles contained a compatible solute other than glycerol it is unlikely that the solute would have escaped detection. This is especially true for the single giant chloroplast of Dunaliella spp. but the argument does raise questions about the distribution of arabitol in Sacch. rouxii. The surface area of the D . parua chloroplast is about twice that of the plasma membrane (Rabinowitch et al., 1975) which is a factor favouring gross glycerol flux across the chloroplast envelope relative to its flux across the plasma membrane. The site of glycerol synthesis is discussed on p. 226. I t is likely therefore that glycerol is distributed more or less evenly throughout the cytosol and organelles of Dunaliella and of non-tolerant yeasts such as Sacch. cereuisiae. Some doubt must exist, however, about the relative distribution of the two polyols, glycerol and arabitol, in Sacch. rouxii. In summary, the general situation with halophilic and salt- tolerant algae is as follows. The genus with outstanding salt tolerance is Dunaliella. Within that genus or, indeed, within some individual species of the genus, there is a range of tolerances from almost no salt up to saturation. Differences in salt relations can be recognized between halophilic species, such as D . viridis, and marine species such as D. tertiolecta, but these differences do not show why, or even if, D . uiridis is

EU KARYOTIC WATER RELATlONS

223

more tolerant than D. tertiolecta. The evidence from growth rates of trained algae in pure culture suggests that, not only can D . tertiolecta grow at salt concentrations as high as does D . viridis, but it can do so more vigorously. Differences between the salt relations of the two species are actually more significant at low salt concentrations. Dunaliella tertiolecta has a significantly lower salt optimum than D . viridis, and has a considerable advantage over it, in terms of growth rate, at low o r relatively low salinities such as occur under marine conditions. The physiological distinction between them might therefore be not so much that D. uiridis is halophilic but that D . tertiolecta is marine. As with the xerotolerant yeasts, the salt tolerance of these algae is achieved at least partly through accumulation of glycerol as a compatible solute, but this in itself does not explain the different salt optima of the two types. At present, there is no clear explanation of the relatively high optimum of D. viridis but theory again suggests a “systems” type of mechanism or a significant involvement of salt in membrane function. E. INTERMEDIATE XEROTOLERANCE

There are numerous micro-organisms with intermediate levels of xerotolerance. In general, yeasts are more tolerant than bacteria and it is not unreasonable to assume that many yeasts achieve this moderate degree of tolerance by enhanced glycerol production such as described earlier for Sacch. cerevzszue. Plants, including algae, respond to slight and intermediate levels of water stress in many cases by using as osmoregulators other solutes such as higher polyols, a-galwtosylglycerol and the imino acid, proline. Information about their enzyme protective capacity is limited but, what there is, together with some inferences, suggests that they do have such a role and can therefore be regarded as intermediate types of compatible solute. The problem is discussed more fully by Brown ( 197 6). IV. Regulation of Compatible Solute Accumulation

Until now, the two major microbial groups with which we are concerned, namely the xerotolerant yeasts and the halophilic algae, for the most part have been treated separately. Nevertheless, the two groups have much in common in that they each respond to the same type of water stress by accumulating glycerol which has essentially the same

224

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physiological significance for both of them. A fundamental question with extensive biological ramifications remains, namely by what mechanism does the concentration of an extracellular solute regulate accumulation of a metabolite? The question becomes more intriguing with the recognition that the chemical nature of the extracellular solute is not specific, and that glycerol is produced by different pathways in yeasts and algae (see p. 226). For that reason, the following discussion of regulation will deal with both yeasts and algae together. Although there is a significant amount of fringe information, there is, as yet, no clear answer to the basic question and much of the following attempt at explanation is speculative. Notwithstanding this reservation, however, a non-metabolic regulatory mechanism is clearly recognizable in Sacch. rouxii (strain YA) which responded to increased concentrations of either salt or polyethylene glycol partly by retaining more of the glycerol which it was already producing. This aspect of the regulatory mechanism is relatively simple and presumably occurs at the plasma membrane. Whether the extracellular solute causes this change by modifjling a transport mechanism or merely by .“tightening up” (decreasing the permeability of) the membrane is not known, but there is some evidence relevant to the question. The surface structures of yeasts can change in response to solute concentration in the suspending solution. For example, incubation in solutions of D-mannitol (0.7 M ) , Lrhamnose (0.7 M ) or KC1 (0.6 M ) causes changes in the surface structures of Sacch. cerevisiae, Schizosacch. pombe and Endomycopsis jibuligera (Kopecka et al., 1973). The changes apparently involve wall synthesis and include deepening of wall ridges which intrude into the cytoplasm. Similar effects are caused by glycerol ( 10-20%) and by high concentrations of urea. High concentrations of sucrose can also induce changes in the surface structures of growing Sacch. rouxii. “Periplasmic bodies” (vesicular structures located between the wall and the plasma membrane) which are frequent in the yeast when grown in a dilute medium were fewer when grown in the presence of 10% sucrose and not detected in yeast grown in 20% sucrose (Arnold et al., 1974). Sodium chloride ( 10%w/v) causes extensive morphological changes in Sacch. cerevisiae and similar, but less pronounced, changes in Sacch. rouxii. The changes include formation of filaments similar to those described by Koh (1975) for a xerophilic mutant of Sacch. rouxii (Margaret Edgley, unpublished results). Polyethylene glycol does not have this effect. Whether either of these structural changes is associ-

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225

ated with changed permeability or transport kinetics is not known. Low concentrations (50 mM) of any of several types of external solute diminished polyol leakage from Sacch. rouxii (Brown, 1974), but this does little to resolve the basic question about glycerol retention since, under Brown’s (1974) conditions, the major polyol which leaked was arabitol. When either species is harvested by centrifuging, most of the glycerol is lost if the yeasts are washed in water at O O C ; virtually none is lost if they are washed in sodium chloride isotonic with the growth medium (Marget Edgley, unpublished results). This effect is unlikely to involve a specific transport mechanism. None of this is to say, of course, that a transport mechanism does not exist; indeed a case has already been made for such a mechanism in Sacch. rouxii. The point is merely that a simple physicochemical process involving a non- specific decrease in membrane permeability is probably sufficient to explain the increased retention of glycerol in response to decreased a, value. The less efficient retention of glycerol by Sacch. cerevisiae could then be explained by a more generally permeable and less effective exclusion of the extracellular solute. It is also possible that the more permeable membrane of Sacch. cerevisiae encourages metabolic regulation of glycerol accumulation by admitting an extracellular solute with regulatory properties. Such a solute is likely to be an ion (see p. 232). The absence of a correlation between arabitol retention by Sacch. rouxii and a, value suggests that arabitol distribution is determined by a transport mechanism which functions independently of a,,,value. Although a substantial part of the regulation of glycerol content in Sacch. rouxii involves changed permeation, as distinct from metabolism, there is a significant metabolic contribution as well. In Sacch. cerevisiae, the regulatory process is mainly, perhaps entirely, metabolic. In Dunaliella spp. it is apparently solely metabolic. At this point, it is as well to identify two levels at which glycerol concentration can be regulated. The first is the response to a water stress in which decreased a, value leads to accumulation of more glycerol, or increased a,value leads to depletion of glycerol. The second is the means by which a concentration is maintained when the organism is adapted to its environment. We are concerned with the first, and it must follow that the transition phase is critical in the regulatory process. The second process is no different, in principle, from the homeostatic maintenance of “pool” concentrations of any metabolite. As Krebs (1975) has commented, the time is hardly ripe for this type of

A. D. BROWN

226

analysis; more information on K, values and other parameters is needed. Moreover, comprehensive interpretation of the homeostatic process would probably require the technique of systems analysis. In Dunaliella spp., glycerol is an early product of photosynthetic fixation of carbon dioxide (Craigie and McLachlan, 1964; Wegman, 197 1 ; Ben-Amotz, 1975); it can also be formed in the dark, again in direct response to increased salinity, by degradation of starch (Borowitzka et al., 197 7). Thus, both photosynthetic and glycolytic pathways to glycerol are regulated in the same manner by external salt or sucrose which suggests that the regulatory site is on a reaction sequence common to both pathways, that is, close to glycerol itself. Normally, the major product of photosynthesis to cross the chloroplast envelope into the cytosol is dihydroxyacetone phosphate (Walker, 1974). Since starch also occurs in the chloroplast, we can assume that, in the dark, degradation of starch goes as far as dihydroxyacetone phosphate in the chloroplast. Whether dihydroxyacetone phosphate is then converted in the chloroplast to glycerol which passes into the cytosol, or whether dihydroxyacetone phosphate migrates to be converted to glycerol in the cytosol, is not yet known. What is known, however, is that Dunaliella spp. contain an active NADP-specific glycerol dehydrogenase which catalyses the reaction (Ben-Amotz and Avron, 1973a; Borowitzka and Brown, 1974): dihydroxyacetone

+ NADPH + Hf

glycerol + NADP'

The final stages in production of glycerol, both photosynthetically and glycolytically, are therefore presumably : Pi

dihydroxyacetone phosphate

NADPH

NADP'

A dihydroxyacetone W

g1ycerol

Although Sacch. rouxii contains a number of polyol dehydrogenases, it does not have the ability to oxidize glycerol, at least via a nicotin-

amide nucleotide coenzyme (Simpson, 1976). The final steps i n glycolytic glycerol production in both Sacch. cereuisiae and Sacch. rouxii are : Pi NADH NAD'

2

dihydroxyacetone p h o s p h a t e ~ ~ - g l y c e r o p h o s p h a ~ e

(seealso Gancedo et al., 1968).

glycerol

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227

As a first approximation, a major regulatory mechanism might be expected to operate at the reversible dehydrogenation step in each organism, namely dihydroxyacetone + glycerol in the algae, and dihydroxyacetone phosphate + a-glycerophosphate in the yeast. The possible regulation of these steps is discussed on p. 230. First, however, another question should be asked: does the regulation occur at the level of enzyme catalysis or at the level of inductionlrepression of enzyme synthesis? Little can be deduced from the time scale of adaptation since both types of organism require several hours to adapt to a new environment of different water activity. In Dunaliella spp., however, glycerol production has been demonstrated specifically to respond to increased salinity in the absence of protein synthesis (Borowitzka et al., 1977). I am not aware of a similar experiment with yeast. This is about the extent to which direct comparisons can be made between Dunaliella spp. and the yeasts although there are additional comments, of varying degrees of relevance to the central problem, which can be made about one or other type ofmicroorganism. Aeration classically inhibits ethanolic fermentation in yeast, and is reported to favour polyol production by xerotolerant yeasts ( 0nishi, 1963; Spencer, 1968).This is not true of glycerol production (as would be expected), and the effect of aeration on arabitoi production is complicated by another important parameter, namely growth rate. Table 3 shows polyol production (intracellular and extracellular) of Sacch. rouxii growing in medium containing 20% (wlv) glucose in continuous culture under conditions of nitrogen limitation. TABLE 3. Effect of growth rate a n d aeration on polyol production by Saccharomyce~ rouxii Dilution rate

Polyol

(h-ll

0.05 0.124.13

Arabitol Glycerol Arabitol Glycerol

Aerated culturen Rate of polyol Total polyol produced production (pmol/mg$ (pmollmglhi 3. I 11.3 1. I 36.4

0.16 0.57 0.14 4.7

Non-aerated culture Total polyol Rate of polyol produced production (prnol/mg) (pmollniglh) 2.8 35.7 0.48 36.7

0.14 1.80 0.06 4.4

"Aerated cultures maintained at 70-8096 saturation with oxygen at atmospheric pressure. *Total polyol is the sum of intracellular and extracellular polyol yield/mass (dry)ofyeast. The cultures were grown at 30°C in a defined rnediuni containing 20% (w/v) glucose. Unpublished results of A. J . Markides.

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A. D. BROWN

At the lower dilution rate, aeration did not affect arabitol production, but it did affect glycerol production by a factor of about three. At the higher dilution rate, aeration affected arabitol production by a factor of about 2.5 but did not affect glycerol production. Expressed another way, in aerated cultures, increasing the dilution rate from 0.05 to 0.12 h-’ had little or no effect on the rate of arabitol production but increased the rate of glycerol production approximately eight-fold. In non-aerated cultures, the same increase in growth rate halved the rate of arabitol production but increased the rate of glycerol production 2.5 times. Information of this kind is difficult to interpret at present. I t does serve to emphasize, however, that growth rate is itself a parameter of major significance in regulating polyol biosynthesis, and that rates of production, not simply final yields, are likely to be determinants of an organism’s physiological response to water stress. I t is a simple statement of the obvious that the relation between polyol biosynthesis and growth rate reflects changes in the balance of various biosynthetic pathways which, in turn, might well be expected to reflect steady-state “pool” concentrations of ATP/ADP and NAD+/NADH. Variations in growth rate should also correlate with levels of enzyme activity. The evidence that the specific response of glycerol production in Dunaliella spp. to salinity does not require protein synthesis does not exclude an influence of protein synthesis on the overall process either in algae or yeasts. At this level, another difference between Sacch. cereuisiae and Sacch. rouxii is apparent. The respiratory capacity of Sacch. cerevisiae is classically subject to catabolite repression by glucose, but this does not happen with Sacch. rouxii. Brown ( 1975) compared the respiratory capabilities of the two test species after growth in a dilute medium (a, 0.9971, the medium adjusted with polyethylene glycol (mo1.wt. 200 33.5%w/v; a,,, 0.95) and with glucose (24 and 36% w/v). Additional measurements were made with Sacch. rouxii after growth in the presence of 48% glucose. Properties compared were rate of oxygen uptake by whole cells ( Q o 9 ) , NADH oxidase activity, cytochrome content and mitochondria (number and morphology). The two solutes acted differently. Growth in the presence of polyethylene glycol diminished 9.0, in Sacch. cereuisiae but not Sacch. rouxii, approximately halved NADH oxidase activity in Sacch. cereuisiae but not Sacch. rouxii and increased the content of “cytochrome b + c” in both species. Glucose greatly decreased the Qo, and NADH oxidase activities in Sacch. cereuisiae, but had a complex effect which left the

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respiratory capacity (Qo,) of Sacch. rouxii changed only slightly. Glucose caused a slight diminution in NADH oxidase activity, but enhanced the cytochrome content of the tolerant species. Brown commented that the effect of glucose was presumably expressed at two levels, at least. One was the direct physicochemical impact of a high concentration of solute exemplified in these experiments by polyethylene glycol; at this level, the dif'f'erences between the two species were minor. The other level was the much more specific one of catabolite repression at which the differences between the species were profound. In summary, this work implies that the shift to enhanced glycerol production by Sacch. cereuisiae in response simply to diminished a, value is not primarily the result of an impaired ability to respire. On the other hand, growth in a high concentration of'glucose would (classically) shift Sacch. cereuzsiae away from respiration to fermentation which would, of course, favour glycerol production. Saccharomyces rouxii (strain YA), however, is unlikely to shift its metabolic balance to fermentation because of diminished respiration at high concentrations of solute (inert or metabolizable sugar). The stability of the respiratory system under these conditions is also consistent with the ability to produce arabitol at high concentrations of' solute and the need for aerobic conditions for this activity. Brown ( 1975) drew attention to the possibility that differences between the two species in their response to catabolite repression might have been effected by polyol in Sacch. rouxii. Although no experiment has been done specifically to test this hypothesis, recent evidence suggests that such a mechanism is unlikely, at least as a primary reason, for Sacch. rouxii's immunity to repression. As we now know, it is glycerol which is the major compatible solute under conditions of all but minor stress and glycerol accumulates also in Sacch. cereuisiae. Nevertheless, there is a real possibility that, in addition to its role as a compatible solute, glycerol has a more specific function in other areas of the physiology or biochemistry of yeasts; glycerol itself should not be overlooked as a regulatory agent. Wallis and Whittaker (1974) noted that incubation of starved Sacch. cereuisiae in glycerol (2% w/v) produced a high proportion of petite mutants. The authors were unable to explain the mutagenic action but speculated that, in the context of a deficient nutrient supply, glycerol caused a breakdown in the control system necessary for maintaining mitochondria1 DNA levels. Saccharomyces rouxii does not normally give rise to petite mutants (Kreger-van Rij, 1969).

230

A. D. BROWN

Glycerol has also been shown to activate DNA transcription. The experiments were done with a cell-free bacterial system, but the glycerol concentrations used (up to about 2 MI were high enought to be relevant to the yeast situation. Nakanishi et al. (1974) used a cell-free transcription system containing A gal DNA and DNA-dependent RNA polymerase from Escherichia coli. They found that total RNA synthesis was stimulated in direct proportion to glycerol concentration up to about 2 M . In addition, glycerol has a number of specific effects, namely: (i) transcription of “ A early r strand” RNA was greatly stimulated by glycerol whereas transcription of “d early 1 strand” was not; (ii)when the synthesis of the early 1 strand RNA was impaired by a defective promotor mutation, activity was restored to normal by glycerol; (iii) glycerol was able to replace cyclic AMP and cyclic AMPreceptor in transcription of the gal site and, furthermore, it restored gal RNA synthesis to normal when it was impaired by defective promotor mutations. Nakanishi et al. (1974) also cited evidence to suggest that glycerol acted by changing the conformation of the DNA template rather than by altering the kinetics or specificity of RNA polymerase. Part of the reason for the contention lay in the response of the glycerol effect to temperature. The authors pointed out that glycerol and salt have opposite effects on DNA stability; glycerol lowers and salt raises the DNA melting temperature (T, value). Other organic solutes, such as sucrose, ethylene glycol, dimethylsulphoxide and 1,3-propanediol, stimulated both gal and total RNA synthesis; these solutes also lowered DNA T, values. Additional evidence that solute concentration can change the quantitative details of gene expression was provided, again with bacteria, by Piovant and Lazdunski (1975).They showed, with various strains of Escherichia coli, that higher concentrations of cyclic AMP were needed for induction of synthesis of tryptophanase than pgalactosidase. Addition of sucrose (0.3 MI’ to the growth medium, however, decreased intracellular cyclic AMP concentration to the point where synthesis of tryptophanase could no longer be induced but /3galactosidase could. Notwithstanding these extensive and complex effects by a, value or solute concentration on control of enzyme synthesis, the fact remains that, in Dunaliella spp. and probably also in Sacch. cereuisiae, there is a regulatory mechanism which operates at the level of reactiods) catalysed by preformed enzymes. There is a reversible hydrogenation immediately responsible for glycerol formation in the algae, and one step removed from it in the yeasts. Both of these reactions use nicotin-

EUKARYOTIC WATER RELATIONS

23 1

amide nucleotide coenzymes. The ratio of oxidized to reduced coenzyme must influence the steady-state or equilibrium (see below) concentration of glycerol. It is a reasonable hypothesis, therefore, to suggest that extracellular solute concentration (or a, value) regulates glycerol accumulation by modifying the ratio of oxidized : reduced coenzyme. The type of contribution likely to be made by this ratio will depend on how far from equilibrium is the glycerol-forming reaction. Reactions requiring either NAD+/H or NADP+/H might seem unlikely candidates for a near-equilibrium situation because of their intimate involvement in cellular energetics. Nevertheless, Krebs has argued that equilibrium conditions can indeed be approximated, if necessary by coupling with another reactionh) which brings the multiple reaction complex into a state close to equilibrium (Eggleston and Krebs, 1974; Krebs, 1973, 1975).If the reaction is close to equilibrium, then changes in the ratio of oxidized to reduced coenzyme will control the reaction merely by displacing the equilibrium. If, on the other hand, the reaction is far from equilibrium, the coenzyme ratio will not work in such a simple manner, and one might then expect an effector role of either the reduced or oxidized form of the coenzyme or, alternatively, involvement of a coupled reaction such as already described. We do not have quantitative information on the equilibrium status of glycerol, but qualitative considerations suggest that, at least in Dunaliella spp., the reaction might indeed be close to equilibrium in those algae which are fully adapted to their physicochemical environment. Two factors which lead to this opinion are: (i) glycerol concentration is very high relative to all other metabolites but it is, nevertheless, of the same order as the K, value (glycerol) of the glycerol dehydrogenase (Borowitzka and Brown, 1974). Polyol dehydrogenases frequently have high Michaelis constants for polyols. This is certainly true of a number of yeast dehydrogenases of higher acyclic polyols (Barnett, 1968) and is scarcely surprizing in view of the likelihood that these polyols can be compatible solutes and reach intracellular concentrations of the order of 1 M ; (ii) glycerol is metabolically at a “dead-end”, and is removed apparently by a simple reversal of the glycerol-forming reaction. Moreover NADP+/NADPH demonstrably does not have an effector role in the kinetics of the algal glycerol dehydrogenase (Borowitzka and Brown, 1974; Borowitzka et al., 1977). The situation in yeast is somewhat different inasmuch as glycerol removal is not a simple reversal of the biosynthetic reaction but requires glycerol kinase and ATP. Nevertheless, if the assumption is

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made that, in each type of organism, we are dealing with a reaction close to equilibrium, there are still some complications arising from participation of different coenzymes in the two cases. The regulatory mechanism must also explain not only why a reduced end product (glycerol) accumulates but also why, in yeast, carbon is diverted from ethanol to glycerol if, in fact, that does happen. In actively metabolizing yeast, as well as in animal tissues, cytoplasmic ratios of NADVNADH are very high, of the order of lo3 (Krebs, 1973). The ratio is thus very sensitive to small changes in the concentration of NADH. Moreover, a slight change in hydrogen ion concentration should also have a marked effect on the ratio and hence on the equilibria : dihydroxyacetone phosphate +glycerophospha5 and acetaldehyde e ethanol Of the reactants mentioned, H + above all is known rapidly to cross membranes in association with energy transduction; furthermore, H+ fluxes are usually intimately associated with fluxes of other solutes, including other cations (Hamilton, 1975). Energy metabolism in yeast can mediate a proton efflux which is coupled to an uptake of K+ (Suomalainen and Oura, 197 1) and can be coupled to transport of non-electrolytes as well. Recent examples of this have been given by Riemersma and Alsbach (1974) and by Misra and Hofer (1975). The H+ efflux is fastest when a yeast suspension lacking an extracellular energy source is provided with a metabolite such as glucose, when there is a “shock” exchange of various solutes. Arabitol is one solute which is rapidly extruded by Sacch. rouxii under such conditions (Brown, 19 74) providing, incidentally, added evidence for an arabitol transport system. Recent preliminary and unpublished experiments of the writer have shown substantial quantitative differences in shock H + efflux between Sacch rouxii and Sacch. cereuisiae and differences between the yeasts in types of response of the H+ efflux to a, value. I t seems highly probable that, if definitive information about the regulatory process is to be found, it will be most easily recognized in the transition of the yeast from one water activity to another. There are good reasons for investigating the phenomenon of energy-mediated ion fluxes during this transition. Another factor of potential importance is the ATP/ADP ratio. Krebs (1973) has pointed out that this ratio is directly proportional to the NAD+/NADH ratio in the cytosol of animal cells but inversely pro-

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portional to it in mitochondria. Presumably, a similar relation exists in yeast. Given that glycerol (and ethanol) are formed in the cytosol, it should follow that any event which decreased the ATP/ADP ratio should enhance glycerol (and ethanol) production. Water stress is likely to be such an event. None of these comments, however, distinguishes between glycerol and ethanol production. Basically a shift of metabolism from ethanol to glycerol might be expected to be implemented at triose phosphate isomerase, aldehyde dehydrogenase or alcohol dehydrogenase. The interpretation of the so-called “third form of fermentation”, in which alkaline conditions direct the fermentation away from ethanol to glycerol, was formerly based on a shift of the triose phosphate equilibrium in favour of dihydroxyacetone phosphate. More recently it has been explained in terms of the activation of an aldehyde dehydrogenase with a high pH optimum (pH 8.7; Sols et al., 197 1).The enzyme converts acetaldehyde to acetic acid, removes thereby the normal hydrogen acceptor in the ethanolic fermentation and generates more NADH in the process. The only other glycolytic hydrogen acceptor is dihydroxyacetone phosphate, reduction of which leads in due course to glycerol. Again, a regulatory mechanism at this point could be reconciled with K+/H+ exchange across the plasma membrane, especially since yeast normally contains at least two aldehyde dehydrogenases, one of which is activated by Kf (Black, 1955). Paradoxically, Gancedo et al. ( 1968) have reported that crude cell-free preparations of the a-glycerophosphate dehydrogenase of Sacch. cerevisiae are inhibited by physiological concentrations of K+ (0.1 M ) and Mg2+(0.01 MI. Yeast alcohol dehydrogenase is susceptible to modification by glycerol. Myers and Jakoby (1975) have reported that glycerol, at concentrations up to 30% (w/v?), decreased the (apparent) Michaelis constants for ethanol and NAD+, but increased the K, (NADH) value. The effect occurred only above pH 8.0. Values obtained at pH 8.8 were (mM, without and with 30% glycerol): K, (ethanol), 11 and 2.8; K,,, (NAD’), 0.22 and 0.063; K,, (NADH),0.53 and 0.83. The K,n (acetaldehyde) was not reported. The results were attributed to a conformational change in the enzyme, although there can be several forms of yeast alcohol dehydrogenase (Lutsdorf and Megnet, 1968);Myers and Jacoby ( 1975) used an unfractionated commercial preparation. Nevertheless this does seem to be another example of glycerol’s acting as an effector, this time at the level of a preformed enzyme. The effect

2 34

A. D. BROWN

of' glycerol's lowering the Michaelis constants for both ethanol and NAD+, while increasing that for NADH, would be to inhibit acetaldehyde reduction which, in turn, would favour glycerol production at the expense of'ethanol. Alcohol dehydrogenase activity also correlates with petite mutation in yeast. Normally there are two cytosol alcohol dehydrogenases. One is constitutive, has a high K, (ethanol) value and is primarily responsible for reducing acetaldehyde to ethanol. The other has a low K, (ethanol) value and can oxidize accumulated ethanol to acetaldehyde. Both use NAD . Petite mutants produce only the first enzyme; the second has been found only ingrande mutants (see Wills, 1976).The reported failure of xerotolerant yeasts to produce petite mutants (Kreger-van Rij, 1969)should be mentioned again at this point. Complications arise when some of the preceding arguments about nicotinamide nucleotide ratios are applied to Dunaliella spp. The first complication lies in the different coenzyme used by the algae, and a second is our ignorance of the site of glycerol production (chloroplast or cytosol). I am not aware of analytical data on NADP+/NADPH ratios in marine or halophilic algae, but some information is available for the leaves of vascular land plants and for Chlorella spp. Heber and Santarius ( 1965) determined nicotinamide nucleotide concentrations in Spinacea oleracea, Beta uulgaris and Elodea densa. All gave similar results for NADP+/NADPH ratios. The chloroplast ratio was about three in the dark and about 0.7 in the light. The cytoplasmic ratio was in the range of about 0-0.4, and was essentially unaffected by light. Green and Israelstam (1970) made similar determinations on extracts of whole Chlorella spp. (no distinction was made between chloroplast and cytoplasmic coenzymes). They found that, in the dark, the ratio of NAD+/NADH was 11-27 and that of NADP'/NADPH was about unity. In white light the NAD+/NADH ratio increased rapidly to about 190 and subsequently dropped to about five; in blue light it rose to 35 and thereafter fell progressively to about 13. Ratios of NADP+/NADPH increased to two in white light to fall to about 0.7. In blue light, this ratio initially dropped to about 0.1 and thereafter rose to 0.7. These values are very different from NAD+/NADH ratios in heterotrophs, and the comments made about the sensitivity of the ratio to changes in the proportion of reduced coenzyme are not applicable. They would seem to be equally inapplicable to Dunaliella spp. The glycerol dehydrogenase from Dunaliella spp., however, does

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have regulatory properties and is thus likely to be implicated directly in control of glycerol production. I n the “forward” direction (i.e. in the direction of glycerol dehydrogenation), the substrate glycerol has a positive co-operative effect on the reaction (Borowitzka and Brown, 1974). In the “reverse” direction, dihydroxyacetone has a negative cooperative effect (Borowitzka et al., 1977). These authors have proposed the following scheme to describe the reaction mechanism of glycerol dehydrogenase G.E 11 E 11 D.E

+

G.E.N

+

11 E.N

+

11 D.E.N

+

G.E.N.G 11 E.N.G 11 D.E.N.G

+ G.E.N’.D e

G.E.N’

11 E.N’.D 11 D.E.N’.D

11

+

+

G.E

It E.N’ G E 11 11

+

D.E.N’

+

D.E

Scheme 1 D indicates dihydroxyacetone, E enzyme, G glycerol, N NADP’, and N’ NADPH. Effector molecules are shown before the enzyme and

substrates after. Again glycerol, the dominant intracellular solute under conditions of desiccation, can apparently contribute to the regulation of its own formation in a manner which is more complex than a mass action effect. Scheme 1 implies positive feedback which would have the advantage of augmenting or accelerating the response to increased salinity. I t also implies, of course, that there must be a negative control as well. This remains to be identified. This discussion, however speculative, should not end without paying some attention to arabitol production in Sacch. rouxii. Arabitol is synthesized via the pentose phosphate pathway (see Ingram and Wood, 1965) which uses NADP+. At least in our test strain of Sacch. rouxii, arabitol production is not affected by a, value nor is it affected as much as glycerol by other cultural factors such as given in Table 3 (p. 227). The pentose phosphate pathway also operates in Sacch. cerevzsiue but the failure of this species to accumulate detectable quantities of any higher polyol implies that the pathway is of less quantitative significance than it is in the xerotolerant yeasts. There is as yet no explanation for this quantitative difference, but a possibility which can be tested experimentally is that the xerotolerant species are deficient or defective in phosphofructokinase activity. There is some suggestion

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that this is so with the xerotolerant fungus Moniliella tomentosa (Hanssens et al., 1974). If that is the case, then the C, leg of the glycolytic pathway in the tolerant species would have to be entered at the point of triose phosphate from the pentose phosphate cycle. This could explain both their ability to accumulate a higher polyol and their low growth rates (see Anand and Brown, 19681, since a significant proportion of their metabolic carbon is diverted away from glycolysis to a pathway which, in terms of ATP production, is relatively non-profitable. I t is also possible that the quantitative importance of the pentose phosphate pathway is significantly enhanced by the resistance of xerotolerant yeasts to catabolite repression of the respiratory system (see p. 228). The cycle in yeast seems to be regulated primarily by glucose 6 phosphate dehydrogenase (Osmond and Rees, 1969) which, at least in rat liver, is inhibited by NADPH at physiological concentrations (Eggleston and Krebs, 1974). In rat liver, oxidized glutathione relieves the inhibition, but it can be expected that any physiological conditions which raise redox potentials will generally favour the pentose phosphate pathway in yeast. Certainly the well documented evidence of enhancement of polyol production by aeration in xerotolerant yeasts supports this opinion as do more specific reports such as that of Woodhead and Walker (1975) who noted a proportionately greater contribution of the pentose phosphate cycle in Penicillium expansum when it was vigorously aerated. In the relevant yeasts, glycerol production is regulated by a, value and requires NADH which, in turn, is intimately associated with ATP metabolism. Arabitol production requires NADPH which is not intimately associated with ATP metabolism and is not regulated by a, value. In Dunaliella spp. glycerol production, which is regulated by a, value, requires NADPH which is intimately associated with ATP metabolism. There seems, therefore, to be a prima facie case for looking to ATP/ADP ratios as a central factor involved in regulating glycerol production in response to a water stress. Finally, it should be remembered that glycerol is not the only substance regulated by water activity or salinity. Other solutes, such as those already listed (a-galactosylglycerol, cyclohexanetetrol, glutamate and proline) are also subject to this type of environmental regulation. I t is not always apparent, however, whether the regulatory mechanism is simply a changed retention, as described for Sacch. rouxii, or involves a metabolic control. When metabolic pathways are found to be

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regulated in this way, it would be instructive to identify those steps which have features in common with glycerol biosynthesis in species of Saccharomyces and Dunaliella. V. Summary

The accumulation of a compatible solute is essential for growth of micro-organisms in environments of low water availability. A compatible solute functions as an osmoregulator in response to water stress and as a protector of enzymes at substantially lowered water activity (h). Several types of solute have this function under slight to moderate stress. Under severe conditions, however, eukaryotes apparently resort to polyhydric alcohols and, under extreme conditions, to glycerol. The ability of polyols, especially glycerol, to protect enzymes under these conditions stems firstly from their failure, even at very high concentrations, to inactivate enzymes or to inhibit them to an appreciable extent. Evidence derived predominantly from enzyme kinetics and, to a lesser extent from physical chemistry, suggests that the acyclic polyols and some sugars bind to an enzyme at least partly through hydrophobic interactions. The binding of other sugars is strongly influenced by stereochemical factors which have not been fully identified. The highly protective polyols, such as glycerol, are characterized by a very high inhibitor constant (low affinity) for a test enzyme, yeast isocitrate dehydrogenase, whereas inhibitory sugars such as sucrose or ribose have much lower values for their inhibitor constants (higher affinities). Compatible solutes rarely activate enzymes; they function primarily as very poor inhibitors but in no sense does this conflict with their protective role. If a compatible solute were not present, other more inhibitory substances would be at a higher concentration, commensurate with the prevailing a, value. Xerotolerant yeasts generally accumulate at least one acyclic polyol (commonly arabitol) when grown in a “normal” dilute medium; nontolerant yeasts do not. When grown at low water activity, the test species, Sacch. rouxii, responds by accumulating glycerol while its arabitol content remains constant. Arabitol can serve as a compatible solute, however, during the transition from a high to low water activity and as a “second rank” compatible solute at low a, values if an energy supply is exhausted, and Sacch. rouxii is forced to metabolize its glycerol. Saccharomyces cerevisiae, which is not xerotolerant, also

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responds to water stress by accumulating glycerol. Over the range of solute concentrations (sodium chloride or polyethylene glycol)which it can tolerate, Sacch. cereuisiae accumulates as much glycerol as does Sacch. rouxii under the same growth conditions. There is, however, a major difference in the mechanisms by which glycerol accumulation is regulated in the two yeasts. Saccharomyces rouxii synthesizes relatively small amounts of additional glycerol in response to diminishing a,value; its major response is to retain within the cell a progressively greater proportion of the glycerol which it synthesizes. On the other hand, Sacch. cereuisiae leaks to the growth medium a very high proportion of the glycerol which it produces. I t responds to diminishing a, value by synthesizing much more glycerol and retaining an approximately constant proportion within the cell. A major cause of the different tolerance ranges of the two yeasts is probably that Sacch. cereuisiae eventually diverts an unacceptably high proportion of its biosynthetic capacity to glycerol production. Another important difference between the yeasts is that Sacch. cereuisiae seems to be generally more permeable than Sacch. rouxii to a variety of solutes, a factor which could itself be an important factor in initiating a major metabolic response to water stress in Sacch. cereuisiae. Algae of the genus Dunaliella also respond to salinity (or sucrose concentration) by accumulating glycerol. The marine species, D . tertiolecta, can be “trained” to grow at high salt concentrations (at least 3.5 M sodium chloride) and the halophilic species, D . uiridis, can be trained to grow at low salt concentrations (0.3 M or less). Dunaliella tertiolecta grows faster than D. uiridis over the entire salt concentration range (i.e. untrained and trained) but is distinguished by having a lower salt optimum. The differences between the salt relations of the two species are greater at low salt concentrations. Algal growth rate responds to salt concentration but glycerol accumulation responds to a, value since, for this purpose, salt can be replaced by sucrose. Neither species leaks more than trace amounts of glycerol, and their response to a, value involves synthesis or breakdown of glycerol. Three types of compatible solute regulation can be recognized in the test organisms. Saccharomyces rouxii responds to a, value physically and, to a limited extent, metabolically; Sacch. cereuisiae responds metabolically by a reaction sequence involving NAD’ whereas Dunaliella spp. respond metabolically by a reaction sequence involving NADP +. Protein synthesis is not required for the immediate response of Dunaliella spp. to increased salinity, but there is evidence that total

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adaptation of the yeasts to water stress is determined by control of respiration and other factors as well as by regulation of preformed enzymes. There is some evidence that glycerol itself has a regulatory function in both yeasts and algae. The article speculates on various possible factors involved in regulation of glycerol production, but it should be remembered that much of the experimental evidence has been obtained under restricted and somewhat artificial experimental conditions, namely growth of the yeasts in salt or polyethylene glycol and growth of the algae under continuous illumination. It is likely that additional factors are involved when the yeasts are grown in highly concentrated sugar solutions because of metabolism of the sugar and because the yeasts will tolerate much lower a, values than they do in salt or polyethylene glycol. Finally, the evidence in toto reveals that enzyme activity is determined not by a,,,value or water potential but rather by the type and concentration of the solute(s1 used to adjust a, value. On the other hand, the concentration of compatible solute is determined by a, value, not by solute type, at least in species ofSaccharomyces and Dunaliella but, in spite of this, a, value is not the primary determinant of growth rate or xerotolerance. Here again, the type of solute is important, presumably because of different degrees of penetration of the cell and, in some cases, specific effects on the plasma membrane. VI. Acknowledgements

I am greatly indebted to my colleagues, Margaret Edgely and Andrew Markides, for the use of some of their results before publication. I also wish to thank Margaret Edgely for criticism of this article and for bringing to my attention a number of valuable references. REFERENCES

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The Yeast Nucleus BRUCE L. A . CARTER Department of Genetics. Trinity College. University of Dublin. Dublin. 2. Ireland

and Rosenstiel Basic Medical Sciences Research Center. Erandeis University. Waltham. Massachusetts 02 154 U S A. I . Introduction . . . I1. Nuclear Morphology .

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IV . Yeast Chromosomes . A . Introduction . . . . . . . . B . Histones . . . . . . . . . C. Sizeofyeast Chromosomes . . . . . D . Localization of Genes on Chromosomes . . V . Initiation of Nuclear DNA Synthesis . . . . . . . . . VI . Nuclear DNA Replication . VII . Nuclear Control Over Mitochondrial-DNA Replication VIII . Nuclear DNA Enzymes . . . . ' . . A . DNA Polymerases . . . . . . . B . DNA-Dependent RNA Polymerases . . . C . Poly(A) Polymerases . . . . . . IX . ExpressionofNuclear Genes . . . . . . . X. Expression of Yeast Genes in Escherichia coli . XI . Integration of Growth and Nuclear/Cell Division XI1 . SomeTechnicalConsiderations . . . . . A . Isolation ofyeast Nuclei . . . . . B . Isolation of a Nucleolar Fraction from Yeast Nuclei C. Isolation of Chromatin from Yeast Nuclei . . D . InhibitionofNuclearFunctions . . . . E. DNA Estimation . . . . . . . F. Nuclear Staining . . . . . . . XI11 . Conclusions . . . . . . . . . XIV . Acknowledgements . . . . . . . References . . . . . . . . . 243

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6. L. A. CARTER

1. Introduction

In this review, I have tried to illuminate recent progress on the structure and activities of the yeast nucleus. I have limited my attention to budding yeasts and even within this group of organisms I have concentrated on Saccharomyces cerevisiae. This organism is currently much in favour because it is a eukaryote that can be grown and analysed genetically with the same ease as bacteria. Many investigators emphasize that, although yeasts can be manipulated like prokaryotes, they are essentially similar to higher eukaryotes in cellular structure and behaviour. For instance, it is often pointed out that yeasts have a nucleus bounded by a nuclear membrane, mitochondria, vacuoles and Golgi bodies. In addition, macromolecular synthesis, chromosome replication and segregation in yeast are similar to these processes in higher organisms. There are, however, features of yeast structure and behaviour that are rather different from those of higher eukaryotes but, in our anxiety to label yeast as a typical eukaryote, these are rarely mentioned. In this review, many of the similarities between yeast and higher eukaryotes will be apparent but I will mention now some of the atypical features of yeast which will be discussed at length later. Gray et al. ( 1973) found that mitosis in yeast was comparable in its general aspect to that in higher eukaryotes. The details, however, are somewhat different; spindle formation is quite different from that observed in higher eukaryotes and the nucleus does not break down during mitosis in yeast. In addition, in yeast, unlike other eukaryotes, mitosis and cell separation are temporally distinct. Yeast is unusual in that mitosis occurs in both haploid and diploid cells. Cell division by budding and nuclear migration into the isthmus of mother cell and bud prior to nuclear division are not a common feature of eukaryotic cells. Yeast chromosomes are unusual for a eukaryote in that they are less than a third the size of those of bacteria and, although yeasts do have some histones somewhat similar to higher eukaryotes, histones corresponding to H1 and H3 histones have not yet been detected in Saccharomyces cerevisiae. The absence of H 1 histones may explain why yeast chromosomes do not appear to condense at mitosis, Repetitive DNA is a feature of the chromosomes of higher eukaryotes but, in yeast, there is little repetitive DNA. Another feature that distinguishes yeast chromosomes from higher eukaryotes is the arrangement of the 5s ribosomal cistrons which in higher eukaryotes are unlinked to the

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18s and 26s ribosomal cistrons. In yeast they appear to be grouped together on a DNA molecule but are arranged in separate transcriptional units and may well be on different strands of a DNA molecule. The RNA polymerases of yeast are similar to the corresponding polymerases in mammalian cells, but higher eukaryotes possess a DNA polymerase with a sedimentation coefficient of 3.4s which appears to be absent from yeast and other lower eukaryotes. Instead yeast has two DNA polymerases having a sedimentation coefficient of 8s. This short list is sufficient to serve as a warning that yeast is not a mammalian cell, and that what is true for yeast may or may not be true for mammalian cells. This in no way detracts from the usefulness of experiments with yeasts. An initial conceptual framework established in an easily manipulated system (yeast) provides direction for experiments on higher-cell systems. 11. Nuclear Morphology

Our present knowledge of the morphology of the yeast nucleus is largely the result of careful observations of both living and fixed cells by Robinow who has recently reviewed (Robinow, 1975) the cytological methods used to investigate nuclear structure and behaviour. Phase-contrast examination of yeast in aqueous media reveals a large central inclusion which the untrained observer concludes is the nucleus. However, studies in nutrient media containing 18-20% gelatin (Robinow and Marak, 1966) indicate that this structure is a large central vacuole; the nucleus is somewhat smaller and usually occupies a position between the vacuole and the bud. Within the nucleus an optically dense crescent-shaped area, the nucleolus, can be observed in isolated nuclei and whole cell preparations. At the opposite pole of the nucleus, a small dense granule can be seen at the outer edge of the nucleus. This lateral granule is seen at the margin of other ascomycete nuclei (Robinow and Caten, 1969) and, during mitosis, it is transformed into an intranuclear spindle apparatus. The spindle itself is sometimes seen as a thin line traversing dividing nuclei in the light microscope and as bundles of microtubules extending from electron opaque bodies, the centriolar plaques, at each pole of the nuclear envelope in the electron microscope (Robinow and Marak, 1966; Moens and Rapport, 197 1). While some microtubules form a straight continuous fibre connecting the two spindle plaques, others extend only a short distance into the nucleus and yet others extend from the

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6. L. A. CARTER

FIG. 1. Electron micrographs of spindle plaques (sp) and associated structures in a haploid strain of Saccharomyces cerevisiae fixed during vegetative growth. (a)The spindle plaque of a n unbudded cell ( ~ 4 5 , 0 0 0 )(b) . The double plaque at early budding. Extranuclear microtubules (mt) project toward the vesicle-filled early bud (b). Three serial

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spindle plaque out into the cytoplasm (Robinow and Marak, 1966; Moens and Rapport, 1971; Byers and Goetsch, 1973). The nucleus is-surrounded by a pair of double unit membranes, the nuclear membrane, as in higher eukaryotes but, unlike higher eukaryotes, it is not broken down during mitosis (Matile et al., 1969). Circular pores 80-90 nm in diameter are spread over the whole nuclear envelope. They generally occur in a density of 10-15 units per square micron such that a nuclear envelope is perforated by ab%t 200 pores. Robinow and Marak (1966)noted that nuclear pores are usually filled with a cluster of closely packed granules less dense and smaller than the ribosomes which fill the cytoplasm. This material is not seen elsewhere in the nucleus. 111. Nuclear Division

Nuclear morphology during mitosis has been studied by Robinow and Marak (1966), Moens and Rapport (1971) and Byers and Goetsch (1973). Unbudded cells early in the cell cycle have a single centriolar plaque (Fig. 1). The single plaque consists of a dense disc with an adjacently staining inflected membrane of the nuclear membrane, the half-bridge, on one side (Byers and Goetsch, 1975).Before budding, this gives way to a satellite-bearing single plaque (Byers and Goetsch, 1975). The satellite is at the opposite end of the half-bridge and is a sphere of dense amorphous material similar to, but not identical with, plaque material. I t differs from a true plaque in being on the cytoplasmic side of the nuclear membrane rather than embedded in it as in a true plaque. In addition, it is of smaller dimensions than a plaque and has no spindle microtubules attached to its intranuclear side. At the time of bud initiation, plaque duplication occurs such that two plaques lie side-by-side in the nuclear envelope connected by a bridge (Fig. 1). Budding occurs in the region of the plaques, and both intra- and extranuclear microtubules emanate from these plaques. The plaques separate as the bud increases in size and the intranuclear spindle extends between the two plaques (Fig. 1). Moens and Rapport ( 197 1) found that the plaques separated to a distance of 0.9 ,um rather sections away, in (d), these microtubules enter the bud (both ~ 3 0 , 0 0 0 )(c) . The short complete spindle of a budded cell shortly after separation of the spindle plaques ( ~ 4 5 , 0 0 0 ) (el . The long complete spindle of' a cell with a large bud ( ~ 3 6 , 0 0 0 ) . Reproduced with permission from Byers and Goetsch (1973).

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quickly and the spindle remains stable at 0.9 pm as the bud continues to enlarge. In cells growing at fast rates, the bud reaches the size of the mother before the nucleus migrates into the neck between mother cell and bud and the spindle elongates to 6-8 pm (Byers and Goetsch, 1973) forcing the poles of the nucleus into the far ends of the dividing cell. The nucleus subsequently pinches apart within the neck just before the cytoplasmic masses separate, close to cell separation. While, at fast growth rates, nuclear migration occurs when mother cell and bud are almost equal size, at slow growth rates nuclear migration occurs when the bud is smaller than the mother cell (B. L. A. Carter, unpublished results). In fact, at slow growth rates, the bud never reaches the size of the mother even at cell division. Whether or not the yeast intranuclear spindle plays an active role in nuclear division has not been established. Byers and Goetsch (1973) examined nuclear pore morphology in temperature-sensi tive cell-cycle mutants (Gulotti and Hartwell, 197 1 ; Hartwell et al., 1974)blocked (at the restrictive temperature) in nuclear division, and found they all possessed a complete spindle with a spindle plaque at either end. In mutants blocked in nuclear division, it was noted that, as in wild-type strains, the spindle is shorter than the greatest dimension of the nucleus. In every strain blocked in late nuclear division, the much more elongate nucleus contains a spindle reaching to its furthest extremes. These data do not support the idea of a mechanical role for the spindle in medial nuclear elongation, but do not rule out a role in later elongation. Byers and Goetsch (1973) integrated their observations on the morphology of nuclear behaviour during the cell cycle with some of the temperature-sensitive cell cycle (cdc) mutants which block development at various stages in the cycle. They found that cells arrested at the cdc 28 block did not duplicate their plaque and that, while the plaque could duplicate in cdc 4 at the restrictive temperature, a complete spindle was not formed. Thus, expression of the cdc 28 gene is necessary for plaque duplicaton, and expression of the cdc 4 is necessary for separation of the spindle plaques and formation of a complete spindle. While bud emergence and DNA synthesis are co-incident, they are independent processes; mutants blocked in the initiation of D N A synthesis can initiate bud emergence, and mutants blocked in bud emergence synthesize DNA (Hartwell, 1974; Hartwell et al., 1974). Byers and Goetsch ( 1973)have shown that plaque duplication is necessary for both bud emergence and DNA synthesis, and they have

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speculated that plaque duplication or some closely associated event serves to integrate the normal cell cycle by co-incidentally triggering bud emergence and DNA replication. There is, as yet, no evidence for this interesting hypothesis and while, at fast growth rates, bud initiation and DNA synthesis co-incide they may not do so in all conditions, for instance in slowing growing cells. Byers and Goetsch (1973) noted that both haploid and diploid strains of yeast duplicate a single plaque in G1 phase. How do cells “lose” one plaque when haploids fuse to form a diploid? Byers and Goetsch ( 1973) have shown that nuclear fusion which accompanies cell fusion is not a random process, but that nuclei fuse directly at the site of the plaques, which themselves fuse. A corollary of this must occur when meiosis results in production of haploid cells from diploids. Moens and Rapport (197 1 ) showed that plaques duplicate twice in a manner consistent with the two meiotic divisions. Culotti and Hartwell ( 19 7 1 ) isolated organisms with temperaturesensitive mutations in seven genes of Sacch. cerevisiue which were thought to confer a defect in nuclear division. However, it is now considered that three of these are involved with DNA replication which leaves cdc 9 and cdc 13, two mutants that terminate development at a medial stage of nuclear division, and cdc 14 and cdc 15 which arrest at late nuclear division. For all of these mutants, the execution point (the time in the cycle that the thermolabile gene product completes its function) is much earlier than nuclear division. Subsequent to arrest at the termination point (either medial or late nuclear division), these mutant strains do not undergo cell separation or bud initiation. Evidently nuclear division is a prerequisite for cell separation and bud initiation. The gene products encoded by these mutants are not known. IV. Yeast Chromosomes A.

INTRODUCTION

Genetic analysis indicates that there are at least 1 7 chromosomes in the nucleus of Sacch. cerevisiae (Fig. 2) although these are not normally visible in mitotically dividing cells (Mortimer and Hawthorne, 1973). Recently, however, Wintersberger et al. (1975) reported in sphaeroplasts the presence of chromatin material in dispersed bodies which they consider to be chromosomes. The number of such bodies is quite variable, but the most frequent number is 18 which corresponds rather

B. L. A. CARTER

250 Chiornosone number

I I[

Iu

m P

PI

FIG. 2. Genetic map of Saccharomyces cereuisiue. Linkages established by tetrad analysis or random-spore analysis are represented by solid lines. Dashed and dotted lines indicate linkages established by mitotic analysis and trisomic analysis, respectively. The sequence of genes within parenthesis has not been determined relative to ourside markers. Reproduced with permission from Mortimer and Hawthorne (1973).

THE YEAST NUCLEUS

25 1

closely with the number of chromosomes as judged by genetic mapping. It would be interesting to examine the behaviour of the chromatin bodies at different stages of the mitotic cell cycle. Tamaki ( 1965) published photographs of presumed chromosomes at meiosis, but Matile et al. (1969)suggested that, since meiotic cells are crammed with a variety of granules, it is possible that some of these granules were mistaken for chromosomes. Wintersberger et al. (1975)noted that in some of their preparations the bodies containing chromatin material appear to be joined to one another, which might arise if all of the DNA in the nucleus consisted of one long pice of circular DNA in which condensed regions (chromosomes) alternating with uncondensed regions as suggested by DuPraw (1970).More upon this later. Chemical estimates of the total DNA content of a haploid cell indicate that there are 0.84-1.2 x 10" daltons of DNA per cell (Ogur et al., 1953; Ciferri et al., 1969; Schweitzer and Halvorson, 1969). The size of the yeast genome has also been determined by renaturation studies of DNA. Bicknell and Douglas (1970) estimated a minimum genome size of 9.2 x lo9 daltons. Renaturation studies.on DNA from nuclei of Sacch. carlsbergensis suggested a genome size of 9.4 x lo9 daltons (Christiansen et al., 1971). P. A. Whitney and B. D. Hall (personal communication) found that nuclear DNA prepared from cells broken with glass beads is 80%single copy and 20% repeated, containing 2.8% ribosomal DNA. Renaturation experiments further showed that the kinetic complexity of yeast single-copyDNA is 4.2-4.6 x lo9 daltons assuming a value of 2 x lo9 daltons for the genome of Bacillus subtilis, and the complexity of the total nuclear DNA is 5.2-5.8 x lo9 daltons. The reasons for the discepancy between this estimate and that of Bicknell and Douglas (1970)are not clear. B.

HISTONES

The presence of histones in yeast similar to those of calf thymus, but without the H1 fraction, was first reported by Tonino and Rozijn (1966). Similar results were obtained by Wintersberger et al. (1973)for histones of nuclei from Sacch. cerevisiue. These results were, however, based on migration in electrophoretic gels and were not substantiated by amino-acid analyses. Recently, Franco et al. (1974) used both methods to show that Sacch. cerevisiue does not contain H1 or H3 although fractions resembling H2al and H2a2 were obtained (Table 1). Yeast chromosomes, unlike those of higher eukaryotes, do not

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TABLE 1. Amino-acid composition of arginine-rich histones from Saccharomyces cereuisiae and calf thymus. Amino-acid residue

Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine BIAa Lysine/Arginine

Saccharomyces cereuisiae H2A2 4.5 4.4 4.8 10.2 4.6 7.4 12.8 trace 4.7 trace 4.9 11.1 2.2 2.9 12.0 1.6 11.7 1.72 1.03

Calf thymus H2A2

6.6 3.9 3.4 9.8 4.1 10.8 12.9 0.0 6.3 0.0 3.9 12.4 2.2 0.9 10.2 3.1 9.4 1.38 1.08

Saccharomyces cereuisiae H2A1 7.5 5.1 5.6 11.0 2.6 9.8 6.2 0.0 6.3 0.6 6.1 10.4 2.2 2.5 10.2 1.7 11.2 1.25 0.9 1

Calf thymus H2A 1

4.9 6.9 2.0 5.9 1.o 16.7 6.9 0.0 8.8 1 .o 5.9 7.8 3.9 2.0 10.8 2.0 13.7 2.45 0.79

"B/A indicates the ratio of basic to acidic amino acids. Reproduced with permission from F r a n c o d d . (1974).

appear to condense at mitosis and it may be significant that histone H 1 which has been implicated in chromosome condensation in other eukaryotes is absent from yeast. It is possible, however, that yeast-does possess H 1 histone but that it is broken down during extraction by the action of proteases, since H1 histones in other organisms are very sensitive to proteolytic attack. Chromatin from a number of mammalian sources has been digested in situ with endogenous and exogenous nucleases, and DNA fragments of discrete sizes have been obtained (Hewish and Burgoyne, 1972; Noll, 1974; Shaw et al., 1974). I t has been suggested that these discrete size classes probably are a result of protection of sections of DNA by bound chromosomal proteins rendering these sections inaccessible to nucleases. Kornberg ( 1974b) suggested that chromatin

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exhibits a periodic pattern of organization produced by chromosomal proteins. Lohr and Van Holde (1975) treated isolated nucleic from Sacch. cereuisiae with micrococcal nuclease for varying lengths of time, isolated the DNA fragments obtained and, after ribonuclease digestion, separated the fi-agments by electrophoresis and stained the resultant gels with ethidium bromide. They found that the exogenous nuclease produced distinctive avd rather sharp size classes of DNA whose distribution was time dependent. The possibility that the enzyme was recognizing and cleaving specific base sequences of yeast DNA to produce discrete sizes was ruled out because micrococcal nuclease activity on isolated naked yeast DNA produced a continuous smear of small DNA sizes. Digestions of nuclei produced several bands on gels, but one class was pivotal and probably represented the basic “chromatin unit”. Short digestion produced fragments of this size class together with larger fragments while longer exposure to nuclease produced this class as well as smaller fragments. If these discrete fragments are the result of sections of the DNA being protected by repeated globular particles in protein, the structure of the repeating unit must be different in yeast from that proposed by Kornberg (197413) for higher eukaryotes. He suggested that the unit in higher eukaryotes covers about 20 nm of DNA and is based on a repeating unit of two of each of HZal, H3, H2a2 and H2B histones. The absence of H3 histones from yeast means that the structure of the repeating unit in yeast must be rather different. Nucleohistone fibres from Sacch. cereuisiue sphaeroplasts have been examined in the electron microscope by Gray et al. (1973). They observed that water-spread chromatin from yeast had an average diameter of 16.5-18.5 nm which is in the range of fibre chromatin of other eukaryotes including those with a full histone complement. Individual fibres have a rather “knobby” appearance which may represent the repeating unit. Moll and Wintersberger ( 1976) pulse labelled synchronously growing cells with tritiated lysine at various times during the cell cycle, and isolated histones from purified nuclei by acid extraction. They found that incorporation of lysine into histones occurs predominantly during the S phase. Experiments in which hydroxyurea was used to inhibit DNA synthesis indicated that histone synthesis was dependent on DNA synthesis.

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8 . L. A. CARTER

C.

SIZE OF YEAST C H R O M O S O M E S

The size of individual chromosomes has been investigated by sedimentation analysis of nuclear DNA released from sphaeroplasts of Sacch. cerevisiae. Because large DNA molecules are extremely susceptible to shear forces, sphaeroplasts were lysed after being layered on sucrose gradients. Blamire et al. (1972) found that nuclear DNA molecules exhibited a narrow range of molecular weights between 4 x lo8 and 6 x lo8 daltons. Petes and Fangman (1972) found that chromosomal DNA molecules range in size from as small as 5 x lo7 to as large as 1.4 x lo9 daltons, with a number average molecular weight of 6.2 x lo8. If, for the moment, we take 9.2 x lo9 as the total nuclear DNA content and divide by 17 (the chromosome number), we get 5.4 x lo8 for the average size of a yeast chromosome. This is close enough to the data derived from sedimentation analysis to suggest that each chromosome contains a DNA duplex and is unineme. A recent genetic map (Fig. 2 , p. 250; Mortimer and Hawthorne, 1973) shows that the distribution of 142 mutants on the 17 chromosomes is significantly different from a normal distribution. The maximum number of mapped genes on the 1 7 chromosomes is 18 on chromosome VII and the minimum is three on chromosome I. If the number of genetic markers per chromosome is proportional to the physical size of each chromosome, then chromosome VI I contains 0.12(17/142) of the total chromosomal DNA of 1.1 x lo9 daltons. This is close to the largest DNA molecules observed by Petes and Fangman (1972). Subsequently, Petes et al. (1973) examined the contour lengths of yeast chromosomes by electron microscopy, and found molecules in the range 1.2 x lo8 and 8.4 x lo8 daltons. They surmize that one reason why molecules a,s large as lo9 daltons were not observed was that the largest molecules were sheared during preparation for electron microscopy. The average contour length of chromosomal DNA was 165 pm, corresponding to a molecular weight value of 3.8 x lo8 daltons. Again, this is close to the value of 5.4 x lo8 daltons which is expected if there is one DNA duplex per yeast chromosome and supports the view that at least some and probably all yeast chromosomes are unineme. Lauer and Klotz (1975) determined the molecular weight of yeast nuclear DNA by visco-elastic retardation time experiments. They conclude that, assuming the DNA is linear and unbranched, the largest DNA molecules released from sphaeroplasts have a molecular weight

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of 2 x lo9 daltons and, assuming circular unbranched DNA molecules, a value 4.3 x lo9 daltons is obtained. Lauer and Klotz (1975)suggested that Blamire et al. (1972)and Petesand Fangman( 1972)underestimated their DNA molecular weights because they used too high rotor speeds and that, under different centrifugation conditions, the largest moleculesTeen by Petes and Fangman (1972) (1.4 x lo9) would have revealed a molecular weight of 2 x lo9 daltons. A value of 2 x lo9 daltons represents 22% of the figure of Bicknell and Douglas (1970) for total nuclear DNA (9.2 x lo9 daltons) and is almost twice the length of chromosome VII (based on the assumption that the amount of DNA in a chromosome is proportional to its genetic length). This assumption may be false; there may be one (or more) very large chromosomes in yeast. If, however, Whitney and Hall’s figure of 5.4 x lo9 daltons is correct for the molecular weight value of the Sacch. cerevisiae genome (see p. 2511, and Lauer and Klotz (1975)are measuring circular DNA of 4.3 x lo9 daltons then, as they suggest, the largest piece of DNA in the yeast nucleus may, at least during part of the cell cycle, consist of all of the genome. This is surprising considering the genetic evidence (Mortimer and Hawthorne, 1973)and biochemical evidence (Blamire et ul., 1972; Petes and Fangman, 1972; Finkelstein et al., 1972; Cryer et al., 1973) that there are several chromosomes in Sacch. cerevisiae. Lauer and Klotz (1975) point out that, if yeast nuclear DNA consists of chromosomes connected by non-gene-containing DNA lengths longer than 50 map units, then genes on two chromosomes connected by a linker would assort randomly because of frequent crossing over between them. They also suggest that the linkers may, for some unknown reason, be sensitive to shear, and breakage of the linkers would yield chromosomes which would be equivalent to those observed by sedimentational analysis. I t is interesting that Wintersberger et a/. (1975) noted that their chromatin bodies were sometimes joined to each other by thin threads. The Lauer and Klotz (1975) data establish that either at least one yeast chromosome is larger than hitherto supposed or that yeast “chromosomes” are linked, at least during part of the cycle. The size of individual chromosomes has been investigated in Marmur’s laboratory (Blamire et al., 1972; Cryer et al., 1973). Sedimentation analysis of total DNA does not allow the DNA of individual chromosomes to be resolved because the size variations of yeast chromosomes are not great enough (Fig. 3). This problem was overcome rather neatly by sedimentation analysis of DNA from aneuploid

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B. L. A. CARTER

Fraction number

FIG. 3. Sedimentation of total DNA from Saccharamyces cereuzszae. Culture ( 5 ml) was labelled with 5 pCi [SHladenine/mland grown for 15 h at 30OC. Sphaeroplasts were layered onto detergent mixture over 50 ml of a sucrose gradient (5-2596, w/v) and centrifuged in a Spinco SW25-2 rotor for 16 h at 11,000 rev/rnin at 18OC. Fractions were collected, alkali digested and assayed for $H in DNA. The internal standard was T2 phage DNA, and the values refer to the molecular weights in daltons calculated from the Friefelder equation. Repr6daced with permission from Blamire et al. (1972).

strains of yeast. Two types of aneuploids have been used, namely monosomes which are diploid strains in which one or more chromosomes are haploid, and disomes which are haploid strains diploid for only one or a few chromosomes. The rationale of the experiments is that an aneuploid, for example a disome for one chromosome ( n + 1) in which the DNA is labelled with one isotope (for instance 3H),can be cosedimented with a haploid (n) strain labelled with a different isotope (for instance 14C) and the region of the gradient containing the chromosome under investigation is revealed by the enhanced 3H to 14C ratio. A disome for chromosome XI was investigated in this way and, on the basis of the enhanced isotope ratio in the gradient, a molecular weight of 7.5 x lo8 daltons was assigned to this chromosome (Cryer et al., 1973). Similar experiments using a disome for chromosome VIII indicated that the molecular weight of this chromosome was 4 x lo8 daltons (Blamire et al., 1972) and a monosome for chromosome I has been shown to have a molecular weight of approximately 4.5 x lo8 daltons (Cryer et al., 1973). D. L O C A L I Z A T I O N O F G E N E S O N C H R O M O S O M E S

S. K. Welsh and B. D. Hall (personal communication) have estimated that the minimum amount of transcriptionally active yeast

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nuclear DNA is 4000 genes, of average length 1000 nucleotides. It is clear from the genetic map shown in Fig. 2 (p. 250) that our knowledge of the location (and indeed the function) of these genes is rudimentary. There are, however, a few examples of clusters of genes of related or co-ordinated function. Genes for the pyrimidine and purine biosynthetic pathways are spread throughout the genome as are those for many amino-acid pathways, but three genes on the galactose pathway are closely linked on chromosome I1 and there is clustering of histidine biosynthetic genes on chromosome I11 although neither cluster shows evidence of an operator segment like bacterial operons. The techniques of molecular biology have been used to determine the number and localization of genes coding for rRNA and tRNA. Schweitzer et al. (1969) found that 0.06-0.08% of total yeast nuclear DNA hybridized with tRNA. This is equivalent to approximately 350 tRNA cistrons per haploid genome implying a large amount of redundancy for these genes. This is consistent with the demonstration of at least eight different tyrosyl-tRNA genes (Gilmore et al., 1968). Marker I

I

I I I

I I

t'

7 a I

Marker

7 a

I

I I

I

Density(g /crn3)

FIG. 4. Appearance of y DNA as a function of-molecular weight of nuclear DNA. High molecular-weight of DNA was subjected to controlled shearing to yield halves, quarters and eighths. The molecular weight of each fraction was determined by analytical band sedimentation, and the DNA was centrifuged in analytical CsCl gradients. The molecular weights of the fractions were as follows: (a) 123 x lo6 daltons, (b) 61 x lo6 daltons, (c) 32 x lo6 daltons, (d) 15 x lo6 daltons. Reproduced with permission from Cramer et d.(1972).

6. L. A. CARTER

258

There are around 140 copies of the rRNA genes in a haploid yeast (Finkelstein et al., 1972; Schweitzer et al., 1969; Qlyen, 1973). Two components of nuclear DNA can be distinguished by their different densities in CsCl equilibrium gradients (Fig. 4). The major component 1.707 1.706

E'" 1.705-

Y 0, Y

0-0-z

1.704-

O\oo

4

.\,

1703L 't

1.702t

$

1.701-

0

1.700 1.699

0

\o'pp-ol I

1

1

1

1

I

I

,

,o

10 20 30 40 50 60 70 00 90 100 110 120 130 Molecular weight(x10-6daltons)

FIG. 5 . Density of ribosomal-DNA as a function of molecular weight ot' nuclearDNA. The density of the heavy satellite containing the 1-ibosomal cistrons was calculated in a series ofexperiments like that described in Fig. 4 (p. 257), and plotted as a function of the molecular weight of the various preparations. Reproduced with permission from Cramer et d . ( 1972).

( n DNA) has a buoyant density of 1.699 g/cmYand a minor satellite ( y

DNA) has a density of 1.705 g/cm3 (Cramer et al., 1972). The y DNA represents about 12% of total nDNA, and hybridization with the yeast rRNA indicates that it contains most if not all ofthe rRNA genes. The y band was not resolved unless nDNA in sheared to 70 x lo6 daltons, and shearing below 25 x 106daltons led to no further enhancement of the heavy satellite (Figs. 4 and 5 ) . If the molecular weight of the rRNA precursor is 2.5 x lo6, these shear data suggest that the ribosomal cistrons are grouped in clusters of 10-32 (Cramer et al., 1972). Localization of the ribosomal cistrons has been investigated using disomic ( n + 1) and monosomic (2n + 1) yeast strains. Hybridization experiments using rRNA and DNA have shown that the DNA of' strains monosomic for chromosome I has decreased hybridization plateaux with rRNA (Goldberg et al., 1972; Finkelstein et al., 1972; Kaback et al., 1973). These experiments indicated that approximately 70% of the ribosomal genes are on chromosome I ; location 01' the remaining ribosomal genes is unknown. Finkelstein et al. (1972) separated DNA, in which shear was kept to a minimum, on sucrose gradients and found

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that rRNA hybridized in two regions of the gradient corresponding to two size classes of'chromosome DNA, one with a molecular weight of' chromosome I (4-4.5 x lo8 daltons) and the other with a molecular weight of5-6 x lo8 daltons. It is, therefore, possible but has not been proven that the ribosomal cistrons are accommodated on only two chromosomes. Rubin and Sulston ( 1973) have shown by RNA-DNA hybridization, using 5 . 8 s and 5s RNA, that these rRNA species are represented 140150 times in the yeast genome. Thus, 5S, 5.8S, 18s and 28s RNA are represented in equal numbers in the genome. There is conflicting evidence about the arrangement of these within the genome. Rubin and Sulston (1973) denatured DNA and annealed in a large excess of 18s and 28s RNA. Hybrids (RNA-DNA) were separated from renatured DNA by virtue oftheir higher buoyant density, and it was noted that 5s RNA hybridized to DNA -from the RNA-DNA hybrids rather than from the denatured DNA. Similar results were obtained even when the DNA was sheared to a molecular weight only slightly larger than the 35s ribosomal RNA precursor. Rubin and Sulston (1973) indicated that the 5S, 5.8S, 18s and 28s cistrons are linked together. Rather different conclusions were drawn by Aarstad and Oyen (1975) who isol-

Fraction number

FIG. 6 . Banding of' the regions complementary to the ribosomal-RNA species upon centrifugation of isolated y-band DNA in alkaline CsCl density gradients. Parts of each h-action were fixed to membrane filters and annealed with 13*P].5S RNA (3 p g l m l ; L O ) , o r with [32P] 18s + 26s rRNA ( 5 pg/ml; L O ) . Reproduced with permission from Aarstad and 0 y e n (1975).

260

B.

L. A. CARTER

ated v band DNA and centrifuged this DNA in alkaline CsCl gradients where they obtained overlapping bands of single stranded DNA and found (Fig. 6) that 5s RNA hybridized to the heavy strand while the 18s RNA and 26s RNA hybridized exclusively to the light strand. They concluded that genes for the larger rRNA species and for the 5s species may be located on the same DNA molecules but arranged in separate transcriptional units. They postulate that they could be o n opposite strands, and hence be transcribed in an antiparallel manner, or they could represent clusters on the same strand but having a very different G + C content. If the 5s cistrons are on the opposite strand to the 18s and 26s cistrons, Rubin and Sulston’i (1973) results are difficult to explain. Two other observances that may be pertinent are that Udem and Warner (1972) found that 5s RNA could be synthesized independently of precursor of 18s and 26s rRNA, and Fraser and Creanor ( 1974) found (in Schizosaccharomyces pombe) that 8 - hydroxyquinoline in low doses inhibits 18s and 28s RNA formation although 5s RNA continues to be synthesized. I t would be of interest to see if 8-hydroxyquinoline has the same effect in Sacch. cereuisiae. That some 5s RNA cistrons, like some 18s and 26s RNA cistrons, are located on chromosome I has been shown by Cryer et al. (1973). They showed that DNA of a strain monosomic for chromosome I had a smaller hybridzation plateau with 5s RNA. V. Initiation of Nuclear DNA Synthesis

In fast-growing cultures (generation times of 2-3 hours), DNA replication is initiated in the first quarter of the cell cycle co-incident with bud emergence, and takes about a quarter of‘a cycle for complete replication. This has been shown in synchronous cultures (Ogur et al., 1952; Williamson, 1964; Williamson and Scopes, 1960) and in asynchronous cultures in which cells are positioned with respect to cell-cycle age using cell size to monitor progression through the cycle (Williamson, 1965; Sebastian et al., 197 1). In Escherichia coli there is, at a variety o f growth rates, a constant time between initiation of DNA synthesis and cell division (Cooper and Helmsletter, 1968). I have examined whether such a relationship holds in a eukaryotic organism like yeast. That such a relationship might exist was suggested by the observation by von Meyerburg (1968) and Beck and von Meyerburg ( 1968) that, in Sacch. cereuisiae grown at a variety of rates in a glucose-limited chromostat, the length of time

261

THE YEAST NUCLEUS

from bud emergence to cell division was almost unaffected by growth rate; .at slow growth rates, the unbudded phase expanded in time. If bud emergence and initiation of DNA synthesis are always co-incident, it follows that the time from initiation of DNA synthesis to cell division will be constant at a wide variety of growth rates. Several authors (reviewed in Mitchison, 1971) have shown that, if one knows the percentage of cells in an asynchronous culture that are

24°C

38OC

I

I

1

2

I

I

I

I

3

4

5

6

Time ihours)

FIG. 7. Effect of shifting a culture of Saccharomyces cereviszae cdc7 (temperature sensitive for initiation of DNA synthesis) from steady-state growth (p = 0.09) in a chemostat at 24OC to fresh medium at 38OC. A culture of’ cdc7 was grown in a glucose-limited chernostat (p = 0.09), and a sample was removed and diluted in lresh medium. H a l f o l the culture was incubated at 24OC and half was incubated at 38°C (the restrictive temperature). The increase in cell number was monitored at both temperatures. After a short lag, the number of cells in the culture at the permissive temperature increased exponentially; however, the culture at the restrictive temperature only increased from 2.3 x lo5 cells/ml to 2.7 x lo5 cells/ml before a plateau of.cell numbers was reached. The percentage increase in cell numbers after the shift to the restrictivr temperature ( 17%) is equal to the percentage of the population that had initiated DNA synthesis at the temperature shift. These data were used as in Table 2 (p. 263) to calculate the stage of the cycle at which initiation of S phase occurs.

6 . L. A. CARTER

262

.. .. .. ...

D

D

D

I

I

> I

Constant

>I

D I

Constant

D

Fasl

> !?wth e

D

9 :

D Constant

>

ow Fr owth te

FIG. 8. A model of-the cell-division cycle in Saccharomyes cereuzszae. The cell cycle from cell division (D) to cell division is shown at three growth rates. Immediately preceding

division is the constant phase ( 0) which is constant in time regardless of growth rate. is growth-rate dependent. The remainder of the cell cycle, the expandable phase ( -1, As growth rate slows, the expandable phase takes longer (in time) to complete.

at a particular stage of a cycle, one can determine what fraction of a cell cycle that stage occupies. Howell and Naliboff (1973) have shown using, temperature-sensitive mutants that are blocked at a particular stage in the cycle, that it is possible to determine in asynchronous cultures, at what fraction of a cell cycle this block occurs. If an asynchronous culture growing at the permissive temperature is shifted to the restrictive temperature, then only cells that are beyond the cellcycle block can go on to divide. Thus, the percentage of cells in the population beyond the block can be determined and used to calculate at what fraction of the cycle the block occurs (Howell and Naliboff, 1973; Hartwell et al., 1973). I have used this method to determine the time during the cycle of the block in cdc 7, a temperature-sensitive mutant defective in initiation of DNA synthesis. This mutant was grown at a variety of rates in a glucose-limited chemostat at 24OC, and then removed from the chemostat, diluted in fresh medium, incubated at the restrictive temperature 38OC and the number of cells which divided (and thus the number of cells past the block) were noted (Fig. 7 ) and used to calculate when the block occurred in the cell cycle.

THE YEAST NUCLEUS

263

The results (Table 2) indicate that, as growth rate is slowed, the block is later in the cycle. Indeed, except at very slow growth rates, there is a constant time from initiation of DNA synthesis to cell division. These results suggest that the cell cycle can be divided into two phases, namely an expandable phase prior to initiation of DNA synthesis which expands in time at slow growth rates, and a constant phase from initiation of DNA synthesis to cell division which is independent of-growth TABLE 2. Time of initiation of DNA synthesis at different growth rates for Saccharomyces cereuisiae cdc 7

Specific growth rate (h-')

Generation time ( h )

0.059 0.080 0.090 0.099 0.104 0.167 0.219 0.240

11.70 8.66 7.70 6.67 6.60 4.55 3.16 2.89

Stage of cycle at which Time before cell division of the initiation of initiation of DNA synthesis begins ( X , ) DNA synthesis (h) 0.73 0.77 0.77 0.66 0.70 0.43 0.38 0.22

3.1 1.99 1.77 2.38 1.98 2.37 1.95 2.25

Cells of Saccharmyes cereuisiae cdc 7 were grown at various growth rates in a glucose-limited chemostat a t 24°C. Cells were then removed from the chemostat, inoculated into fresh medium at 38OC, and the percentage of cells that divided at the restrictive temperature were measured as in Fig. 7 (p. 26 1 ), T he strain is defective in initiation of DNA synthesis at the restrictive temperature, and the percentage of cells which go o n to divide at the restrictive temperature is a reflection of the percentage of cells that are past this stage of the cycle at the time of the temperature shift. From these data, Howell and Naliboff (19731 showed that one can calculate the stage of the cell cycle a t which the block occurs using the equation X I = I -In (N/No)/ln 2 where X I = the stage of the cycle of the block, N o = cell number at the time of the temperature shift, and N = final cell number attained.

rate (Fig. 8). Perhaps the way of looking at the cycle as a progression of events from division to division is wrong; rather I should consider that a cell prepares, during the expandable phase, to get ready for the cell cycle which is the constant phase and which, once started, progresses to division in a constant time regardless of growth rate. At slow growth rates, the time from initiation of DNA synthesis to cell division expands somewhat (Table 2). The reasons for this are not clear, but one possibility is that the constant phase only extends from initiation of S phase to nuclear division and that at, slow growth rates, a gap is introduced between nuclear division and cell separation.

264

8. L. A. CARTER

I t would obviously be of interest to carry out this kind of anlaysis o n Sacch. cereuisiae cdc 28, a mutant which Hartwell has called “start”. This mutant blocks earlier in the cell cycle than cdc 7, and it may be that cdc 7 is not the start of the constant phase but merely a very early event in this phase. What triggers the constant phase of‘ the cell cycle? It has been suggested in bacteria that cells trigger DNA synthesis when they reach a certain size. I n Sacch. cereuisiae, the size (volume) at which cells initiate DNA synthesis is not constant at various growth rates. Size (volume) at the initiation of DNA synthesis increases slightly with growth rate (B. L. A. Carter and M. N. Jagadish, unpublished results). If, however, the start of the constant phase is earlier than initiation of S phase, then it is possible that there is a size control at this earlier point because, in the time between “start” and the initiation of S phase, growth will occur and, since in a constant time more growth will occur in faster growing cells, these will be larger at the initiation of S phase than slow growing cells. Biochemical and genetical methods have shown that, in Sacch. cerevisiae, protein synthesis is required for initiation of DNA synthesis but that, once initiated, DNA replication can be completed in the absence of protein synthesis (Hereford and Hartwell, 1973; Williamson, 1973; Slater, 1974). Hereford and Hartwell (1973) cultured a temperature-sensitive mutant, defective in initiation of DNA synthesis, at the restrictive temperature (36OC) for a generation to align cells at the cell-cycle block. The culture was then incubated at the permissive temperature (23OC) and at intervals two samples were taken. One was incubated at 36OC and the other remained at 23OC, but in the presence of the protein synthesis inhibitor cycloheximide. I t was observed that samples shifted to 36OC made the same amount of DNA as samples at 23O to which cycloheximide had been added, and it was concluded that cycloheximide has the same effect as a temperaturesensitive block in initiation of DNA replication. Williamson ( 1973) added cyclohexamide to cells at different stages of the cell cycle, and noted that 10 min before initiation of DNA synthesis cells could initiate and complete S phase in the presence of cyclohexamide. Thus, the proteins required for a complete round of replication are made 10 min prior to initiation of DNA synthesis which means that the programme of multiple replication initiation cannot be due to sequential synthesis of initiator proteins during S phase as it appears to be in other eukaryotes (Muldoon et al., 197 1 ; Hori and Lark, 1973).

THE YEAST NUCLEUS

265

Our knowledge concerning initiation of S phase is rudimentary, but it is to be hoped that analysis of some of the cell-cycle mutants isolated

by Hartwell and his colleagues will shed light on this area. As Pringle (1975) has .pointed out, analysis of cell-cycle mutants is likely to be useful in a variety of ways. These mutants will prove useful in identifying previously unknown functions. Indeed, analysis of such mutants has already revealed a sequence of at least three steps that precede, are are vital for, initiation of DNA synthesis (Hereford and Hartwell, 1974). These authors analysed three mutants of Sacch. cerevisiae (cdc 4, cdc 7 and cdc 28) defective in initiation of DNA synthesis, and showed that the order of the steps mediated by these unknown gene products is: cdc 28-cdc 4-cdc 7. The existence of this sequence would have been difficult to detect without genetic analysis. Hereford and Hartwell (1974) also discovered at what stage in this sequence the proteins required for DNA initiation are synthesized. The three mutants (cdc 4, cdc 7 and cdc 28) were grown at the restrictive temperature until they aligned at their respective blocks. They were then shifted to the permissive temperature and grown in the presence of cycloheximide. Only cdc 7 completed a round of replication, implying that the protein-synthesis requirement for yeast DNA replication is completed before the cdc 7-mediated step. Although analysis of these mutants has identified previously unknown functions and shown that they form a dependent sequence in which late gene products (e.g. the cdc 7 gene product) cannot express themselves until after early gene products (e.g. cdc 28 and cdc 41, we do not know what products these genes code for nor do we know the molecular basis of the dependence of gene expression in the cell cycle. I t would, of course, be a major step forward if these gene products in cell-cycle mutants could be identified, but the difficulty of finding a needle in a haystack should not be underestimated. Hartwell ( 1974) has speculated on the molecular basis of the dependence of gene expression in the cell cycle, and suggested two broad possibilities. The product of one gene (e.g. cdc 4 ) might be necessary for synthesis of another (e.g. cdc 7 ) in which case the product of cdc 4 is a positive regulator of the synthesis of that formed by cdc 7 . Alternatively, he suggests that the product of one gene (e.g. cdc 4) is necessary for the function of another gene (e.g. cdc 7 ) perhaps because they are sequential steps in a pathway and the product of one gene is a substrate for the next.

266

B. L. A. CARTER

VI. Nuclear DNA Replication In prokaryotes, the chromosome is usually replicated from a single initiation fork (Klein and Bonhoeffer, 1972)but, in higher eukaryotes, chromosome replication proceeds via multiple initiation sites (Huberman and Riggs, 1968; Callan, 1972). Replication of the yeast chromosome is similar to that of higher eukaryotes (Petes et al., 1973; Newlon et al., 1974) although individual chromosomes of Sacch. cereuisiae are less than a third the size of the Escherichia coli chromosome. Analyses of DNA molecules isolated from Sacch. cereuisiae (Newlon et al., 1974) revealed that 66% of such molecules isolated form S-phase cells contained Y-shaped forks or “eye forms”. These were assumed to be replication intermediates because DNA molecules from cells in G 1

FIG. 9. Tracing of an electron micrograph of‘ a replicating DNA molecule from Saccharomyes cerevcszae. The molecule shown is 11 7 prn long. Replication forks are indicated by arrows. The bar represents 5 pm. Reproduced with permission from Newlonetal. (1974).

phase of the cell cycle were predominantly linear structures. A high proportion of molecules contained multiple “eye-forms”, and these are presumed to arise by initiation of replication at several sites on a chromosome (Fig. 9). Newlon et al. (1974)noted that “eye-forms” were of different sizes suggesting that, if the rate of replication at all sites is the same, different replicons in a chromosome initiate at different times. The distance between centres of “eyeforms” ranged from 3 to 86 pm with clusters at 15-20 and 30-35 pm, which is similar to the distance between initiation sites in animal somatic cells (Huberman and Riggs, 1968; Callan, 1972). Independent evidence for the existence of multiple initiation sites has been obtained by DNA-fibre autoradiography (Petes and Williamson, 1975). These authors suggested an average distance between initiation sites of about 30 pm, and they investigated the rate and direction of DNA synthesis from each initiation site by labelling cells with two consecutive isotopic pulses of different specific activities.

m

3 J w 0 3 Z

Lu

I I-

268

6 . L. A. CARTER

Figure 10 shows examples of a DNA fibre after a shift from low to high specific-activity [3H]uracil.These and other experiments revealed that replication was bidirectional, and the rate of fork migration at 24OC was 0.7 pmlmin which is similar to that of higher eukaryotes. If the initiation sites in yeast are 20 to 30 pm apart, then the average yeast chromosome ( 165 pm) will have between five and eight replicons (Petes and Williamson, 1975).The data of Petes and Williamson (1975) suggest that the entire nuclear genome could be replicated in 15-23 min which is well within the 30-40 min time that it takes at 24OC. Replication of the 140 ribosomal cistrons in Sacch. cereuisiae has been examined in cultures synchronized by the feeding-starving technique. Samples of DNA were removed at eight times during the S period and hybridized with ribosomal-RNA prepared from an exponential culture (Gimmler and Schweitzer, 1972). A constant percentage of DNA hybridized to ribosomal-RNA in all samples, demonstrating that the proportion of rDNA to total DNA remains constant throughout S phase. This result indicates that ribosomal cistrons replicate throughout S phase at the same rate as total nuclear DNA, and this in turn suggests that chromosome I which contains 70% of these cistrons r e p i r e s the whole of S phase to replicate. Three groups of workers (Burke and Fangman, 1975; Dawes and Carter, 1974; Kee and Haber, 1975) have used genetic techniques to study DNA replication at the gene level. All three studies are based on the idea that, if N-methyl-N’-nitro-N-nitrosoguanidine causes enhanced mutation at the replication form in Succh. cereuisiae as it does in bacteria (Cerda-Olmedo et al., 1968; Guerola et al., 197 11, and in the eukaryote Chlamydomonas rheinhardii (Lee and Jones, 1973) it should be possible to order temporally the replication of genes on a chromosome. In addition, it should be possible to determine whether some chromosomes replicate early during S phase while others are replicated later, or whether particular chromosomes take the whole of the S phase to replicate. Dawes and Carter (1974) treated cells at various stages of the cell cycle with nitrosoguanidine, and observed that mutations for erythromycin and oligomycin resistance, known to have nuclear (and cytoplasmic) characteristics, occurred at high frequencies during a small period of the cycle corresponding to the period of nuclear DNA synthesis. There were three peaks of enhanced mutation during this period, and genetic evidence suggested that mutants from at least two and possibly all three peaks were in different nuclear genes. The fact that these peaks were resolved indicates that the

THE YEAST NUCLEUS

269

multiple initiations of DNA synthesis are highly ordered in time and are not random from cycle to cycle. If gene replication were a random process, discrete peaks would not be seen. To study the effect of nitrosoguanidine on a single chromosome, Kee and Haber (1975) looked at the frequencies of reversion to prototrophy of six auxotrophic markers located along one arm of chromosome VII of Sacch. cerevisiae (Fig. 11) at various stages of the cell cycle. They observed that for five of these markers the frequency of reversion is greatest during the period of DNA replication. They observed that the two markers most distant from each other (ade 5 and leu 1) both

-

20 c M

FIG. 11. (a) A genetic map ot'chrornosome VII of Saccharomyces cerevzsiae. ib) Relative reversion frequencies o t tive auxotrophic markers located o n one arm ot chromosome VII. Asynchronous cultures were treated with N-methyl-N'-nitro-N-nitrosoguanidine (200 pg/mli for 5 min and then separated according to size and stage ofthe cell cycle by zonal rotor sedimentation. Cells were collected, and the relative reversion frequencies of certain fractions calculated for the following markers: ade 5 (A),leu I (01, lys 5 (m), trp 5 ( 0 )and tyr 3 ( x ) . For each type of revertant, two dropout plates were counted. I n the fractions containing the largest number of revertants, there were on average 20 colonies per plate. Reproduced with permission from Kee and Haber 19751.

6 . L. A. CARTER

270

have their highest reversion frequency early during DNA replication. The order of peak reversion frequency appears to be : ade 5 -+ leu 1 + lys 5 -+ tyr 3 trp 5 (ade 5 and leu 1 were close together in time as were tyr 3 and trp 5). Kee and Haber (1975) suggested that the complete replication of chromosome VII requires most of the S period because, while the reversion frequencies of ade 5 and leu 1 occur early in S phase, those of trp 5 and try 3 occur very late in S phase. Because the maximum reversion of ade 5 and leu 1 both occur early during S phase, although these markers are at oppodite ends of one arm of chromosome VII, it was concluded that there must be at least two widely separated points of initiation of DNA replication along one arm of this chromosome. This conclusion is consistent with the observations of Newlon et al. (1974), mentioned earlier, that DNA strands with three or more replication structures could be seen in DNA from Sacch. cereuisiae. Burke and Fangman (1975) also demonstrated, by nitrosoguanidine mutagenesis, that particular genetic loci replicate in a particular temporal order during the S period; however, they observed that, while maximum reversion to leu 1 was obtained during early S phase, ade 5 was reverted later. The reasons for this rather disquieting discrepancy are not clear although the difference might be a result of the different methods used for mutagenesis or to obtain synchrony prior to mutagenesis. Hartwell (197 1, 1973) isolated two temperature-sensitive mutants that cease DNA synthesis when they are shifted to the restrictive temperature. These mutants rode for functions required for ongoing DNA replication. Since hydroxyurea addition has the same effect on wild-type cells, and as this inhibitor acts by preventing synthesis of deoxynucleotide precursors, one must beware of jumping to the conclusion that these mutants are defective in polymerization of deoxyribonucleotides. However, at least in the case of cdc 8 , it is unlikely that this mutant is defective in precursor synthesis because it is is temperature-sensitive for D N A synthesis in a permeabilized cell system that incorporates deoxynucleoside triphosphates into DNA (Hereford and Hartwell, 1973). The other mutant (cdc 21) has not been tested in this system. Recently, L. H. Hartwell (personal communication) has reinvestigated two mutants (cdc 2 and cdc 6) which were thought to play an essential role in nuclear division. Both of these mutants synthesize DNA, but the DNA made at the restrictive temperature is probably faulty or incomplete. Hartwell interprets the cdc 6 function as a step --f

THE YEAST NUCLEUS

27 1

necessary to initiate DNA correctly. Likewise, the proper activity of the cdc 2 gene product is necessary to replicate DNA properly. VII. Nuclear Control Over Mitochondrial-DNA Replication

A haploid yeast cell contains about 50 mitochondrial DNA molecules per cell (Williamson, 1970). I t has not yet been established whether the components of mitochondrial-DNA replication are, encoded in the nuclear or mitochondrial genome, but it is known that mitochondrial-DNA replication is partially independent of nuclearDNA replication. Mitochondrial-DNA replication continues for at least two generations when nuclear-DNA synthesis is prevented by addition of a factor (Petes and Fangman, 1973). Furthermore, mitochondrial-DNA synthesis is only partially affected after nuclear-DNA replication has ceased in cells treated with cycloheximide ( Grossman et al., 1969). Four groups of workers (Cottrell et al., 1973; Cryer et al., 1973, Wintersberger et al., 1974; Newlon and Fangman, 1975) have shown near-normal mitochondrial-DNA synthesis at the restrictive temperature in temperature-sensitive mutants defective in initiation of nuclear-DNA replication, which suggests that the amount of mitochondrial-DNA is regulated in response to some component other than the amount of nuclear DNA. Mitochondrial- and nuclear-DNA replication are not, however, completely independent events because the two gene products defective in cdc 8 and cdc 21, two mutants temperature-sensitive for elongation of nuclear DNA chains, are also essential for mitochondrial-DNA replication (Newlon and Fangman, 1975). VIII. Nuclear DNA Enzymes A.

DNA POLYMERASES

Higher eukaryotes possess, in addition to mitochondrial-DNA polymerase, at least two other DNA polymerases (Kornberg, 1974a). One of these enzymes is present in the cytoplasm and has a sedimentation coefficient of 6-8s and a molecular weight-of 200,000 to 230,000 daltons. Another appears to be nuclear, and has a sedimentation coefficient of 3.4s and a molecular weight of 400,000 daltons. Wintersberger and Wintersberger (1970) found two nonmitochondrial DNA polymerases in cell-free extracts of Sacch. cere-

272

B. L. A. CARTER

uisiue which sediment at about 8s. Helfman (1973) used DEAEcellulose column chromatography to separate two DNA polymerases from yeast; DNA polymerase A eluted at a lower salt concentration than DNA polymerase B. Activity of DNAse was associated with polymerase B but, since this enzyme was not purified to homogeneity, the two activities may be separable. Subsequently, Wintersberger (1974) investigated DNA polymerases present in isolated nuclei of Sacch. cerevisiae. She used similar methods to those used by Chang and Bollum (1972) to isolate and characterize the 3.4s DNA polymerase from nuclei of mammalian cells. No evidence for DNA polymerase with a sedimentation co-efficient of 3.4s was obtained, but a DNA polymerase with a sedimentation co-efficient of about 8s was detected. This suggests that both DNA polymerases in Sacch. cereuisiae have sedimentation co-efficients of about 8s. Holmes and Johnston (1975)

I

I

I

I

2

3

4

5

I

Time ( h 1

FIG. 12. Specific activity of DNA polymerase at different stages of the cell cycle in synchronized Saccharomyes cereuzszae. The second and third cycles after initiation of synchronized growth are shown. Reproduced with permission from Golombek et al. (1974).

THE YEAST NUCLEUS

273

noted that the small ( 3 . 4 s )DNA polymerase is absent from many lower eukaryotes, although it is possible that it is present in small and as yet undetectable amounts. There are two reports of activities of DNA polymerases in cells synchronized by the feeding-starving method of Williamson and Scopes (1960). Eckstein et al. (1967) found that the total DNA polymerase activity in cell-free extracts oscillated such that polymerase activity was maximal at the onset of DNA-synthesis and decreased to a minimum value during replication. Somewhat surprisingly, the oscillations were not dependent on DNA replication; treating cells with Xrays delayed DNA replication but did not alter the timing of the oscillations in DNA polymerase activity, although the amount of enzyme activity in the maxima was decreased. Golombek et al. (1974) have also reported oscillations in total DNA polymerase activity in synchronized Sacch. cereuisiae. Their data, however, indicate two oscillations per cell cycle, a major oscillation at the start of DNA replication and a minor oscillation after completion of a round of DNA synthesis (Fig. 12). Oscillations continue in the presence of the protein synthesis inhibitor cycloheximide (200 ,ug/ml) suggesting that neither the increase nor decrease of DNA polymerase activity depends on protein synthesis. Golombek et al. (1974) also considered the possiblity that fluctuating activity resulted from cyclic association and disassociation of enzyme molecules, but it was found that DNA polymerase size did not alter during the cell cycle although these measurements may not be sensitive enough. I t is not known which of the two DNA polymerases found in Sacch. cerevisiae is responsible for the oscillations although, in animal cells, it has been observed that the activities of cytoplasmic DNA polymerases fluctuate during growth whereas the nuclear DNA polymerases remain fairly constant in activity (Loeb et al., 1970; Chang et al., 1973). B.

DNA-DEPENDENT RNA POLYMERASES

Multiple forms of RNA polymerases are found in many different species of animals, plants and fungi (Chambon, 1975). Initial studies on RNA polymerases in yeast were confused with respect to the number of RNA polymerases present in yeast nuclei. Some investigators (e.g. Dezklee et al., 1972; Sebastian et al., 1973a) found only RNA polymerase I and I1 whereas others (e.g. Ponta et al., 1971; Adman et al., 19 7 2 ) observed these two together with another. It is now

0 . L. A. CARTER

274

A

B

C

-z

-

'0

10

20

30 40 Fraction number

50

60

70

FIG. 13. Chromatographic separation of yeast RNA polymerases on DEAE-sephadex A-25. Cellular extracts of yeast were prepared. Protein ( 4 . 5 mg) was applied to a column ( 2 x 1 7 cm) of DEAE-sephadex and chromatographed according to the method of Roeder and Rutter (1970).Fractions (2.5 ml) were collected, the absorbance of the eluate recorded automatically with a n LKB UV-cord (---), and 25 pl of each fraction used for determination of RNA polymerase activity (-0). indicates the molarity of ammonium sulphate. Reproduced with permission from Ponta et al. (197 I).

established that this third RNA polymerase is not an artifact due to a trivial alteration of RNA polymerase I during chromatography, but is RNA polymerase 111. These three polymerases elute in the order I, I1 and I11 on DEAE-sephadex chromatography with increasing salt concentration (Fig. 13). Methods to purify RNA polymerases from yeast have been reviewed by Ponta et al. (1975).More recently, Valenzuela etal. (1976a)published a purification procedure for RNA polymerase I which permits isolation of 48 mg of pure RNA polymerase I fi-om 3.0 kg yeast in five days. Their procedure involves batch absorption on phosphocellulose and DEAE-cellulose,followed by chromatography on DEAE-sephadex and sucrose-gradient centrifugation in 25% glycerol. DNA-Cellulose chromatography is used to remove contaminating RNA polymerase 111. Valenzuela et al. (197613)purified RNA polymerase I11 by batchwise treatment with phosphocellulose and DEAE-cellulose, and ionfiltration chromatography on DEAE-sephadex (these steps are common to their purification of polymerase I). DNA-Cellulose chromato-

THE YEAST NUCLEUS

275

TABLE 3. Molecular weights of subunits of RNA polymerases I, I1 and 111 from

Saccharomyces cerevisiae RNA

polymerase I 185 137

48 44 41'

RNA 170 145

41

36 28 24 20 14.5 12

RNA

polymerase 11 polymerase 111

33 28 24 18 14.5 12

160 128 82 53

41 40.5 37 34 28 24 20 14.5 11

Numbers indicate subunit molecularweight x Reproduced with permission from Valenzuela (1976b).

el al.

graphy was used to separate enzymes I and 111; enzyme I is eluted at 0.45 M KCl whereas enzyme I11 is eluted in a highly purified form at 0.7 M KC1. Concentration and further purification are achieved by DEAE- sephadex chromatography. All three RNA polymerases are multimeric protein complexes containing two high molecular-weight subunits and a collection of smaller polypeptide chains (Table 3). The molecular weights of the large subunits of the three polymerases differ (Valenzuela et al., 1976b) indicating that the RNA polymerases of Sacch. cerevisiae have a distinct molecular structure. The molecular weights of some of the smaller subunits (41,000, 28,000, 24,000 and 14,500 daltons) appear to be identical, and suggest that the three enzymes may have common subunits. I t is possible, however, that while these subunits are identical in size they may have unique sequences. Bell et al. (1976) suggest, on the basis of two-dimensional polyacrylamide electrophoresis and phosphorylation of the small polypeptides of RNA polymerases, that at least the 24,000-dalton polypeptide is common to all three poly-

276

0 . L.

A. CARTER

merases and that the 20,000-dalton polypeptide is common to polymerase I and 111. Immunological studies of RNA polymerases I and I1 (Buhler et al., 1976) have confirmed that the large subunits of these enzymes are unrelated, while fingerprint analysis of some of the smaller subunits has revealed that the subunits of 29,000, 24,000 and 16,000 daltons are identical whereas those of 14,000 daltons are clearly different. This is in keeping with immunological studies indicating that RNA polymerases I and I1 have some common antigenic determinants (Hildebrand et al., 1973). Thonart et al. ( 1976) examined four temperature-sensitive mutants of Sacch. cerevisiae in which RNA synthesis is specifically and immediately blocked after a shift to the restrictive temperature (37O C ) . They observed that all mutations were recessive, segregated in a Mendelian fashion and belonged to three complementation groups. RNA Polymerases I, I1 and I11 were separated on DEAE-sephadex after mutants were grown at 37OC for 3.5 h and all three enzymes showed low activity. I n vitro thermodenaturation studies on the mutants showed that the half lives of the three RNA polymerases in the mutants are about ten times less than in the wild type. The existence of three genes, each of which is indispensable for the activity of all three polymerases i n vivo as well as i n vitro, suggests that the polymerases have at least three subunits in common. Schultz and Hall ( 197 6) compared the properties of the three nuclearRNA polymerases of Sacch. cerevisiae and concluded, on the basis of subunit composition, salt dependence on enzyme activity, template specificity and chromatography behaviour on DEAE-substituted cellulose and sephadex columns, that there is homology between yeast and mammalian RNA polymerases. Both mammalian and yeast RNA polymerase I elute early from anion exchangers and have a salt optimum of less than 0.07 M (NH,)$O, (Chambon, 1975; Adman et al., 1972; Valenzuela et al., 1976b; Ponta et al., 1972; Schultz and Hall, 1976). Yeast RNA polymerase 11, like mammalian RNA polymerase 11, is eluted from both DEAE-sephadex and DEAE-cellulose at approximately 0.22 M and is much more active on denatured than on native DNA (Chambon, 1975; Adman et al., 1972; Dezelke and Sentenac, 1973; Ponta et al., 1972; Sebastian et al., 1973).Yeast RNA polymerase 111 is like the corresponding enzyme in mammalian cells in that it has a double salt optimum for activity on native DNA (Adman et al., 1972; Shultz and Hall, 1976; Valenzuela et al., 197613).

277

THE YEAST NUCLEUS

I'

-

\

0

I

10-2

IO-I

I

100

18\-10'

'01

I

103

I 104

a-Amanitin (,ug/ml)

FIG. 14. Sensitivity of yeast RNA polymerases I, I1 and 111 purified o n DEAEsephadex to a-amanitin. Assay conditions were 0.05 M tris-HCI ( p H 7.9), 5% (v/v) glycerol, 1.6 mM MnCI,, 1 mM dithiothreitol, 0.05 mM ['HI UTP (specific activity was 300 Ci/mol) and 0.5 mM each of ATP, CTP and GTP. Heat-denatured salmon-sperm DNA was present at 100 pg/ml in assays with RNA polymerases I and 11. Poly[d(A-T)] was present a t 50 mg/ml in assays with polymerase 111. Final ammonium sulphate concentrations i n the assay were 30 mM, 60 mM and 100 mM ammonium sulphate for RNA polymerases I , I1 and 111, respectively. Appropriate concentrations of a-aminitin were added to the substrate solution prior to initiation of the reactions by addition of enzyme. Reaction mixtures were incubated 5 min at 3OOC. Maximal activities were 268, 123 and 153 pmol of UMP incorporated per ml for RNA polymerases I, I1 and 111. respectively. A indicates polymerase 1 activity, 0 polymerase I1 activity, and polymerase 111 activity. Reproduced with permission from Schultz and Hall (1976).

While yeast polymerases I , I1 and I11 are similar to the corresponding polymerases in mammalian cells as judged by subunit structure, catalytic activities and chromatographic properties, they contrast with mammalian polymerases in their sensitivities to a-amanitin (Fig. 14). Yeast RNA polymerase I1 is similar to that of other eukaryotes in that it is inhibited by relatively low concentrations of a-amanitin. However, in contrast to results with higher eukaryotes, RNA polymerase I11 is resistant (1-2 mglml) to a-amanitin (Schultzand Hall, 1976; Valenzuela et al., 1976b). RNA Polymerase I from higher eukaryotes is resistant to a-amanitin but yeast RNA polymerase I is inhibited 50% by between 300 pglml (Valenzuela et al., 1976b) and 600 pglml (Shultz and Hall, 19761, concentrations of a-amanitin at which class-I vertebrate RNA polymerases are fully resistent. The relative a-amanitin sensitivities of yeast RNA polymerases I and I11 (Fig. 14) are the reverse of those found for mammalian polymerases.

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RNA Polymerase I of mammalian cells is thought to be responsible for ribosomal-RNA synthesis (Roeder and Rutter, 1970). In yeast, there is some indirect evidence which suggests that RNA polymerase I is involved in ribosomal-RNA synthesis. Sebastian et al. (1973) found that, in yeasts grown at different rates, the level of RNA polymerase I is correlated with total RNA content (85%of which is ribosomal-RNA). Cramer et al. (1974) attempted to obtain more direct evidence. They showed that [32Pl-labelledribosomal-RNA hybridized to the light strand of y-band DNA. They used y, a , and total DNA as templates for purified RNA polymerase I and 11, but no quantitative differences existed between the activities of the two enzymes on any of the templates. When the product of in uitro transcription of y-band DNA was hybridized to DNA, it was found that, while RNA polymerase I1 transcribed both heavy and light strands of y-band equally, RNA polymerase I transcribed from the light strand with a greater frequency than the heavy strand. If RNA polymerase I is responsible in viuo for transcription of the light strand of y-band DNA, its lack of complete fidelity may be a result of alterations in enzyme or template during isolation. More recently, Valenzuela et al. (1976b) quote unpublished work by M. Holland, G. Hager and W. J. Rutter in which they claim that yeast polymerase I as isolated has inherent specificity for ribosomal genes from naked yeast DNA. In mammalian cells, it has been shown that RNA polymerase 111 transcribes 4s and 5s RNA genes (Price and Penman, 197 2 ; Weinmann and Roeder, 1974). Fraser and Creanor (1974) have suggested that an analogous situation exists with regard to yeast RNA polymerase 111. They showed that, in Schizosacch. pombe, low concentrations of the chelating agent 8-hydroxyquinoline decreased ribosomal- RNA and polydisperse RNA (mRNA) synthesis, but not 5s RNA and 4s RNA synthesis. They argue that, while all three enzymes will operate with Mn2+,only polymerase 111 is fully active with Mg" alone (Ponta et al., 1972; Brogt and Planta, 1972). As 8-hydroxyquinoline binds Mn2+ very much more strongly than it binds Mg2+,they anticipate that at low concentrations of 8-hydroxyquinoline the Mn2+concentration might be drastically lowered with little change in Mg2+concentration thereby decreasing the activity of polymerases I and I1 but having less effect on polymerase I11 activity. Although it is thought that, in yeast, polymerase I transcribes ribosomal genes, polymerase I1 produces heterogeneous RNA and polymerase I11 produces 4s and 5s RNA, this has not yet been firmly

THE YEAST N U C L E U S

279

established. I t is likely that this problem will be resolved fairly soon by examining the RNA products made by yeast nuclei subject to varying degrees of a-amanitin inhibition to differentiate between the three RNA polymerases. The activity of DNA-dependent RNA polymerases from yeast grown at various rates has been examined by Sebastian et al. (1973b) and Carter and Dawes (1975). It was observed that the specific activity of

1

I

1

0.1

0.2

0.3

I

0.4

1

0.5

Growth rate ( h - ' )

FIG. 15. Specific activity of RNA polymerase I ( 0 )and I1 (0) in Saccharomyes cervzszae 1L46 grown at different rates in a glucose-limited chemostat. The RNA polymerases from cells grown at the growth rate indicated were separated by DEAE-sephadex chromatography and the activity of each of' the enzymes measured. Specific activity is given in units per mg protein. One unit is the incorporation of one pmol U M P into trichloroacetic acid-insoluble material in 15 min. Reproduced with permission from Sebastian et al. (1973).

R N A polymerase I1 remained fairly constant at a variety of growth rates, but that of RNA polymerase I increased with growth rate (Fig. 15). Thus, although RNA polymerases I and I1 have common subunits, activities of the two enzymes are independently regulated. Further evidence for this came from their differential activities during the cell cycle (Sebastian et al., 1974; Carter and Dawes, 1975). Bell et al. (1976)have conceived the possibility that some changes in the activities of RNA polymerases may not be due to alterations in the rate of synthesis or degradation but may be a result of modulation of the activities of pre-existing RNA polymerases. They have investigated phosphorylation of RNA polymerases from yeast both in uivo and in

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vitro. They observed that in vivo labelling with [32Plphosphateresulted in phosphorylation of five of eleven polymerase-I peptides. Purified RNA polymerase I was also phosphorylated in vitro resulting in phosphorylation of the five polypeptides phosphorylated in uiuo together with the 48,000 dalton polyeptide. Purified RNA polymerase I1 and I11 were also phosphorylated in vitro. Bell et al. (1976) suggest that phosphorylation of RNA polymerase could affect transcription by altering the activity of the enzyme either by controlling the extent of interaction between enzyme polypeptides or between the enzyme and other molecules or chromatin. Huet et al. (1975) proposed that the structural complexity of RNA polymerases suggests that each enzyme may be composed of a fundamental enzyme together with regulatory components (presumably some of the smaller subunits). They surmized that, if some of the small polypeptides of the polymerases were regulatory, they might be loosely and reversibly associated to the basic enzyme. Huet et al. (1975) isolated two forms of RNA polymerase I from yeast; the modified form lacked the 48,000- and the 37,000-dalton polypeptide chains. Removal of the two components alters the template specificity of RNA polymerase I , particularly its ability to transcribe native calf thymus DNA. Further studies by Huet et al. (1976)have shown that ribonuclease I1 activity is inseparable from RNA polymerase I by a variety of fractionation procedures. The nuclease activity specifically degrades RNA-DNA hybrids. Although the phyiiological role of the nuclease is uncertain, Huet et al. (1976)'speculated that, since yeast RNA polymerases seem to prefer unpaired DNA structures to initiate (Frederick et al., 1969; Dezelee et al., 1974), transient RNA-DNA hybrids could also be formed at the initiation sites and removed by RNAse 11. c.

P O L Y ( A ) POLYMERASES

Haff and Keller (1973, 1975) obtained two major forms of poly(A) polymerase from Sacch. cereuisiae and a third minor species which they suggest might be mitochondria1 in location. They found that enzyme I utilized the mixed polynucleotide poly(A, G, C, U ) or ribosomal-RNA most efficiently, and they speculated that it may be responsible for initiation of the poly(A) tract at the 3'- end of the primary transcript messenger RNA. Enzyme 11, which eluted at a higher ammonium sulphate concentration from a DEAE-cellulose column, required oligo(A) or an RNA primer containing a 3'-oligo(A) tract, and may be

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Fraction number

FIG. 16. Chromatography of poly(A)polymerases I and 11 from Saccharomyces cereuisiae in logarithmic-phase and stationary-phase cultures o n columns of DEAE-cellulose. Crude extracts from both types of cell were separately chromatographed o n DEAEcellulose. A linear 500 ml gradient of 0.0 1-1 .O M ammonium sulphate was used to elute the column. Fractions ( 5 ml) were collected and assayed for poly(A)polymerase activity. The peak eluting at 0.25 M ammonium sulphate is form I while the peak eluting at 0.75 M is form 11. Open circles represent activity in logarithmic-phase cells and closed circles that in stationary-phase cells. Unpublished results of I. C. Saunders and S . J. Sogin.

responsible for elongation of poly(A) tracts initiated by enzyme I. C. Saunders and S. J. Sogin (unpublished observations) obtained only two forms of poly(A) polymerase even in cells grown in a glycerol-containing medium which might be expected to enhance the activity of a mitochondrial enzyme. These authors noted that, after phosphocellulose chromatography, in vitro length addition of poly(A) polymerase I was decreased 10-fold but that this length-addition activity could be 50% restored by addition of flow-through material from the phosphocellulose column. This material itself had negligible poly(A) polymerase activity. Saunders and Sogin speculate that poly(A) polymerase I, which had a nuclear location, is responsible for addition of a poly(A) tract to the primary transcript in the nucleus, and poly(A)polymerase I1 found primarily in the cytoplasm may have a maintenance function continually repolymerizing poly(A) onto existing mRNAs to insure a more or less constant poly(A) length. These workers also investigated the relative amounts of forms I and 11in cells from the logarithmic phase and stationary phase of growth. They found that there was no change

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8. L. A. CARTER

in the activity of the nuclear enzyme (form I ) but that the cytoplasmic enzyme (form 11)was lost as the culture enters the stationary phase (Fig. 16). If, as they suggest, form I1 enzyme provides a maintenance function, then the total poly(A) per microgram of RNA found in stationary-phase cells should decrease. Such measurements indicated that stationary-phase cells had half the amount of poly(A) per microgram total RNA compared with logarithmically growing cells. The results could be a reflection of a constant number of mRNA sequences with shorter poly(A) tails, or a smaller number of sequences with fulllength poly(A) tails or a combination of both. Using reverse-phase chromatography, it was observed that the average poly(A) size was slightly smaller in stationary-phase cells but, to account for the twofold difference, it is likely that the number of poly(A)-containing RNAs is also decreased. IX. Expression of Nuclear Genes

S. K. Welch and B. D. Hall (personal communication) investigated the fraction of yeast nuclear-DNA expressed as RNA transcripts, in both vegetatively dividing cells and sporulating cells. They isolated labelled non-repeated yeast DNA, and hybridized this with a vast excess of total cell or polysomal RNA extracted from vegetative cells or sporulating cells. The fraction of DNA forming hybrid was assayed by hydroxyapatite chromatography. Approximately 25% of the DNA was present in RNA-DNA hybrids which were shown to have the high thermal stability characteristic of well matched duplexes. These results indicate (assuming assymetric transcription) that at least 50% of the non-repeated DNA of both vegetative and sporulating cells is transcribed. Since similar results were obtained with RNA from polysomes as with total cell RNA, it is likely that these transcripts are translated. If a gene has an average length of 1000 nucleotides, these results, together with the estimates of P. A. Whitney and P. D. Halls (unpublished observations) of the apparent genome size of yeast, indicate that the minimum amount of transcriptionally active yeast nuclear DNA is 4000 genes. L. M. Hereford (personal communication), using similar experimental procedures, found that, when single-copy DNA was hybridized to vast excesses of total cell RNA or poly(A)-containing RNA, 20% of the DNA formed DNA-RNA hybrids. From DNA renaturation kinetics. a value of 7 x lo9 daltons was obtained for the

THE YEAST NUCLEUS

283

kinetic complexity of yeast-nuclear DNA and, assuming that 1500 nucleotides is the average length of a transcript, it was concluded that 4000 genes are transcriptionally active in yeast. Hereford has also exploited the ability of reverse transcriptase to synthesize very radioactive DNA copies of poly(A)-containing RNA of Sacch. cereuisiae. Analysis of the kinetics of hybridization of cDNA copies of messenger RNA to excess of the template RNA allows a determination not only of the number of different sequences present as messenger-RNA, but also of the relative abundance of these sequences within the messenger population. This procedure, first used by Bishop et al. (1974), revealed three abundance classes in HeLa-cell mRNA. The kinetics of annealing of cDNA to its template yeast-mRNA demonstrate discrete frequency classes with most of the different mRNA species (3400)present with about one copy per cell, 600 mRNA species present at about 10 copies per cell and 20 mRNA species present at about 100 copies per cell. These conclusions are only valid if mRNAs are copied to produce cDNA in proportion to their frequency in a given population. I t is of interest that Hereford, using two different experimental approaches, calculated that 4000 genes are transcriptionally active in yeast. What is the significance of these abundance classes? What do these messenger RNAs code for? L. M. Hereford has repeated these experiments at the permissive (23OC)and restrictive temperature (37OC)for a mutant rna 2, defective in ribosomal protein synthesis (Gorenstein and Warner, 1976). When cDNA is made from mRNA of cells grown for one hour at the restrictive temperature, 200 of the sequences present at about 10 copies per cell are absent, but the sequences present at 100 copies per cell and one copy per cell are not noticeably affected. That this is not an artifact resulting from growth at two temperatures is shown by the fact that there is no difference in wild-type cells at the two temperatures. The absence of 200 mRNA species is perhaps more than one might expect, given that there appear to be only about 60 ribosomal proteins in yeast (Gorenstein and Warner, 1976), but Hereford suggests that there may be other mRNA species involved with ribosome synthesis and assembly that are co-ordinately regulated in this mutant. I t is not known what the species represented 100 times per cell code for but, given the fermentative ability of yeast, it is tempting to suggest that they code for glycolytic enzymes. Enzymes are, however, catalysts and thus may not be present in vast amounts, and it may be that these mRNAs code for structural components of the yeast cell.

284

B. L. A. CARTER

One of the earliest observations of a metabolically unstable RNA with a base composition similar to that of DNA was made in yeast (Ycas and Vincent, 1960; Kitazume et al., 1962).However, recent research o n heterogeneous nuclear RNA and messenger RNA in yeast has lagged behind many other eukaryotes. Saccharomyces cereuisiae, like other eukaryotes, has a poly(A1 sequence at the ?,‘-end of some messenger RNAs, but the poly(A) sequences have an average length of about 50 residues (Edmonds and Kopp, 1970; McLaughlin et al., 1973)somewhat smaller than the chain length ( 150200) found in mammalian cells. The exact proportions of heterogeneous nuclear-RNA and messenger-RNA which do or do not have poly(A) sequences is not known with certainty. One of the problems of making such measurements is that RNAs bearing short tracts of poly(A) do not bind to oligo-dT-cellulose and may be mistaken for poly(A)-minus RNA. Groner and Phillips (1975) studied poly(A)containing RNA obtained from nuclei and cytoplasm of yeast, and noted that both RNA species have similar base compositions. Electrophoretic analysis indicated that poly(A)-containingRNA in the nucleus i s heterogeneous in size with an average molecular weight of 1.3 x lo6 daltons, or about twice the average molecular weight of poly(A)containing RNA in the cytoplasm. Similar conclusions were reached by Sripati and Warner (1974).Neither group found any evidence for giant heterogenous RNA but, as Groner and Phillips (1975) point out, this does not prove its absence from yeast. Detection would be very difficult if the compound was present in very small steady-state levels and was processed rapidly. Nuclear, heterogeneous and messenger RNAs from yeast have modified 5’-termini similar to but not identical with those present in higher eukaryotes (Sripati et al., 1976).The 5’-nucleotide is linked by a 5’-5’-triphosphate bridge to the unusual nucleoside rl-methylguanosine. An appreciable proportion ( 75%) of poly(A)-containing RNA contains methylated blocked 5’-terminal structures of composition M’G (5‘)pppAp, and 25% have the composition M7G(5’)pppGp. No other methylated nucleosides were found within poly(A)containing RNA. It is thought that terminal methylation is essential for ribosome binding of mRNA (Both et al., 1975; Muthukrishan et al., 1975). Petersen and McLaughlin (1973) have shown that most if not all yeast messenger-RNA is monocistronic, or rather, they have shown that the average size of a nascent polypeptide chain on a polysome is

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directly related to the size of the polysome. This demonstrates that Sacch. cerevisiae messenger-RNA is monocistronic in the sense that each messenger-RNA molecule codes for one protein molecule which is released intact from the ribosome upon completion of translation. X. Expression of Yeast Genes in Escherichia Coli

Struhl et al. (1976) have shown that genes from a eukaroyte, namely Sacch. cerevisiae, can be transferred to Escherichia coli and can express themselves in this organism. They used a selection technique employing the lysogenic state of phage /1 to obtain a specific hybrid which contains DNA from Sacch. cerevisiae. The selection is based on transferring yeast DNA into E. coli having a non-reverting mutation (hisB-) that renders the cells incapable of making active imidazole glycerol phosphate (IPG) dehydratase, an enzyme needed for synthesis of histidine. After transferring yeast DNA into E . cob, those bacteria that can grow on minimal media are selected on the assumption that the bacteria are synthesizing a functional gene product (IPG dehydratase) from genetic information encoded in yeast DNA. Struhl et al. (1976) used a A vector which contained all of the necessary genes for phage growth, but was too small to be packaged. Yeast DNA and viral DNA were then mixed, and treated with the restriction endonuclease EcoRI. The ends of the fragments generated by this treatment join together randomly with consequent production of some yeast-viral hybrid DNA which was sealed by the use of a ligase. Hybrid DNA molecules were selected by transference to E . coli since only viral cells with additional DNA are large enough to be packaged. These viruses were then grown in another E. coli strain which methylates the foreign DNA. The viable hybrids are thereby modified, and can infect any K12 strain of E. coli which is-sensitive to infection by bacteriophage A. Hybrids were then integrated into the chromosome of an E. coli hisB- mutant using an integration helper virus. His' colonies were selected on plates of minimal media. The fact that some cells can now grow in the absence of histidine suggests that these cells have acquired yeast genes which complement the E. coli mutant lacking imidazole glycerol phosphate dehydratase activity. Struhl et al. (1976) carried out a number of tests to demonstrate that it is the yeast DNA that is functioning and not a contaminant. They found that the E . coli his- strain did not revert to his', thereby ruling out the most trivial explanation of their results. I t was found that

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B. L. A. CARTER

removal of hybrid DNA rendered the bacteria his- but, when hybrid was re-inserted, the cell; regained the ability to make their own histidine. In addition, if the inserted phage is removed and used to reinfect a fresh colony, the frequency of his+ cells increases by a factor of lo4 over that when random virus hybrids were used. The hybrid DNA could complement another hisB- strain, but did not complement any histidine auxotroph belonging to other complementation groups (HisA, C, D, E, F, G, I). An important observation was that RNA made from the inserted hybrid DNA hybridized with certain bands of yeast DNA fragments separated on polyacrylamide gels after EcoRI digestion, indicating that the hybrid insertion contains at least some yeast DNA sequences. Struhl et al. (1976) suggest that transcription is probably initiated in the yeast fragment because expression occurs in a /1 lysogen, in which /1 promotors are either strongly repressed or have been deleted in the hybrid, and because inversion of yeast DNA (with respect to A and the bacterial chromosome) has no effect on expression which would require two as yet unknown d promotors if the promotor did not reside in the yeast DNA fragment. There is, however, no evidence as to whether transcription begins from the correct yeast promotor or merely from a sequence fortuitously recognized by E . coli RNA polymerase. These results indicate that yeast genes can be transcribed and, perhaps more surprisingly, translated in a prokaryote, and suggest that genes from higher organisms can be transferred to prokaryotes and can function and express themselves in these organisms. XI. Integration of Growth and Nuclear/Cell Division Dividing cells, under constant conditions, display a fairly constant cell size. There must be co-ordination between the processes of growth and nuclear/cell division to maintain this constancy. Hartwell ( 1974) suggested two ways in which growth and nucleadcell division might be co-ordinated. Either continued growth may be dependent on completion of one or more events in the pathway to nuclear/cell division, or progression beyond certain cell-division cycle events may require growth beyond some specific size. A combination of both models is also possible. The former hypothesis can be ruled out because, when specific cell-cycle events are arrested by shifting temperature-sensitive cell-cycle mutants to the restrictive temperature, growth continues such that cell size increases to several times that of normal cells (Culotti

287

THE YEAST NUCLEUS

and Hartwell, 1971). In addition, if DNA synthesis is inhibited with hydroxyurea, cell growth continues for several cell cycles (Slater, 1973). Pringle (personal communication in Hartwell, 1974) noted that, when stationary-phase cells are inoculated into fresh medium, there is an inverse relationship between the size of a cell and the length of time before bud initiation. Hartwell (1974) suggested that the simplest explanation of these data is that cells need to attain a certain size before bud emergence is possible. This observation is consistent with the idea that growth to a certain size is necessary for the completion of certain cell-cycle event(s). I have observed, however, that the size at which a cell produces a bud increases with increasing growth rate. This is not inconsistent with the interpretation placed on Pringle’s data; it simply means that the cell-cycle event which required attainment of a certain size is some time before bud emergence and, if bud emergence occurs a constant time after this cell-cycle event, then obviously fast growing cells will be larger at bud initiation than slow growing cells. This is, in fact, observed (B. L. A. Carter, unpublished results). This model also explains the well known fact that unbudded cells in a population appear to get larger each time they produce a bud. If the trigger for the constant phase of the cycle (see Section V, p. 260) occurs prior to bud formation and a cell produces a bud after a certain time, cell growth during this time interval ensures that a cell will be larger at the time of bud formation. After cell division, the mother cell starts the next cell-cycle programme, but a period of time and growth occurs before the next bud initiation so that the unbudded cell gets even bigger; and so on in successive cell cycles. If yeast cells require to be a certain size before a limiting nuclear/cell-cycle event can be initiated,

I

D GI

I

I

S

I

G2

I

ND

G1

I D

FIG. 1 7 . The cell cycle of Saccharomyces cereuisiae growing with a 3-h generation time from cell division (D) to cell division (D). After division, a single unbudded cell is in the G1 phase of the cycle. Initiation of DNA synthesis (S phase) is co-incident with bud emergence. The S phase occupies approximately 0.25 o f a cycle and is followed by the G2 phase. Nuclear division occurs at about 0.8 of the cycle and, because of-temporal separation of nuclear and cell division, a short G1 phase is introduced at the end o f t h e division cycle.

0 . L. A. CARTER

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and the cell then proceeds to division in a constant time regardless of growth rate (see Section V, p. 2601, then the size of cells at division will increase with increasing growth rate for the same reasons put forward above (viz. in a constant time, more growth will occur at fast growth rates). McMurrough and Rose (1967) together with Mor and Fiechter (1968) have shown that the size of yeast cells increases at fast growth rates. Saccharomyces cerevisiae is an unusual eukaryote in that nuclear

...

D

D

I

L

...

D

[

Constant

Constant

>

D

D Constant

D constant

- -

D

D

>

D

Constant

>

,t wth

>

D

Constant

>

>

D Constant

>

D IW

growth rote

FIG. 18. A simplified model of the cell-division cycle in Saccharomyces cereuzjiae to show the behaviour of cells at different growth rates. The cell cycle is divided into a constant )which varies in phase ( 0), independent of growth rate, and a n expandable phase ( time depending o n growth rate. Cells enter the constant phase when they reach the appropriate size/mass. The size control can trigger the constant phase any time after nuclear division or possibly cytokinesis. At fast growth rates, the mother cell and bud are the appropriate size at nuclear division (or cytokinesis) and the constant phase is triggered. At slower growth rates, the bud is not the appropriate size at nuclear division (or cytokinesis). Indeed, after division, the bud requires a period of growth (the expandable phase) before it reaches the appropriate size/mass to trigger the constant phase. The size of the bud at cell division decreases as the growth rate is slowed, so that increasing amounts of time are required after division to reach the appropriate size to initiate the constant phase. Thus, the length of time a cell spends in the expandable phase of the cell cycle increases at slower growth rates.

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289

division and cell separation are temporally distinct. Thus, a cell (Fig. 17) is in the G1 period just prior to cell separation; the nuclear cycle and the cell-division cycle do not co-incide. I t is possible that, after nuclear division or cytokinesis, a cell, if it is the appropriate size, may begin the programme of events culminating the next generation as cell division. If, however, the cell is not the appropriate size/mass, it must grow until it achieves it sometime after cell division, and then the cell procedes in a constant time to nuclear/cell division. Thus, we have the situation in Fig. 18. If the cell cycle is divided into an expandable phase prior to a size control triggering a constant phase then if, after nuclear division the cell is the appropriate size, the expandable phase will be absent. This probably occurs at very fast growth rates. The length of the constant phase defines the minimum cell-division time of Sacch. cerevisiae. If cells are grown at mass-doubling times faster than the time of the constant phase, then cells will grow filamentously. Filamentous growth has been observed in the second state of two-stage chemostats where cells can be grown at extremely fast growth rates (,u = 09 h-'; Vrana, 1973). Because of the peculiar mode of cell division in yeast, viz. budding, a mother cell (the cell that produces the bud) is roughly the same size after division as at the time of bud initiation. The daughter cell (bud), however, may or may not be the same size as the mother cell. If, as I have mentioned earlier, once a cell has attained a critical size it proceeds to cell division at a constant rate then, while at fast growth rates the bud may (and does) reach the size of the mother, at slow growth rates it cannot grow to the same extent in the constant time and, therefore, will not reach the size of the mother by the time of cell division. Thus, at slow growth rates, cell division will produce one mother cell and a smaller daughter cell. As the mother cell is still at least the same size that it was at the time of the proposed size control, it should be ready to begin the next cycle. The daughter cell, on the other hand, will not have attained the critical size and will take a period of time to grow. sufficiently to reach the critical size. The result of this is that at slow growth rates there will be two populations having different generation times (Fig. 18). The slower the growth rate, the smaller the daughter cell (bud) will be at division and the longer its subsequent cell-cycle time to the next cell division. B. L. A. Carter and M. H. Jagadish (unpublished results) found that, at generation times of 9.1 hours (mass doubling), daughter cells (buds)required 35.4 hours to complete the next cell division.

h, (D

0

/Q\

/".\

0

Q

Q

0

0 0/Q\ 0

d d

J '10 0

ci

FIG. 19. Representation of cells growing at fast growth rates (a) and slow growth rates (b).At fast growth rates (a),the bud reaches the size of the mother cell at cell division. Both cells are the appropriate size to initiate the constant phase of the cycle, and consequently proceed to the next cell division in an equivalent time. At slow growth rates (b), the bud does not reach the size of the mother cell at cell division. Thus, division results in the production of a mother cell and a smaller daughter cell. The mother cell is the appropriate size to initiate the constant phase of the cycle. The daughter cells, however, require a period of time during which they grow to reach the appropriate size to initiate the constant phase of the cycle. Thus, at slow -,.-..+I. ,-..I+..-:" nfmnther r e l l ~anr4 r l a i i w h t e r r e l l ~ which hive different veneration times.

-----

. 1

m r ?

THE YEAST NUCLEUS

291

At fast growth rates, as shown in Fig. 19, where one cell at division forms two equal size cells, the mass-doubling time is equivalent to the cell-number doubling time. At slow growth rates, it is evident from Fig. 19 that mother cells do not double in mass during the cell-division cycle. On the other hand, the daughter cells (buds) more than double their mass during a cell division cycle. Therefore, at slow growth rates, a yeast population is composed of mother cells which divide in a time faster than the mass-doubling time of the culture, and daughter cells which divide in a time considerably slower than the culture massdoubling time. To demonstrate the heterogeneity of cells at slow growth rates, B. L. A. Carter and M. N. Jagadish (unpublished results) grew cells in an ethanol-containing medium which supports a mass-doubling time of

'0

2

4

6

8

10 12 14 Time ( hours 1

16

18

20

22

FIG. 20. Synchronous culture growing with a mass doubling time of 7.4 hours. Saccharomyces cerevisiae (cdc7)was grown at 24OC in an ethanol-containing medium for 48 hours with one dilution. The culture was then harvested, separated according to size by zonal centrifugation (Carter et al., 197 1; Sebastian et al., 197 I ) , and the smallest cells used to inoculate fresh ethanol-containing media. Cell number measurements were employed to follow the development of the synchronous culture. Not all cells divide in the first generation (the remainder are presumably inviable). The second plateau is reached after half the cells which produced a bud in the first generation divide again. These cells divide yet again before the other half of the viable culture divides a second time.

B. L. A. CARTER

292

7.4 hours, and then separated cells according to size by zonal centrifugation (Carter et al., 1971; Sebastian et al., 1971). Small cells (daughters)were removed and used to inoculate fresh ethanol-containing media. These cells were allowed to develop into a synchronous culture. If these cells represent the small subpopulation, the prediction is that they would develop synchronously and, at division, the population should double in number releasing equal quantities of large mother cells and small daughter cells. I t is clear from Fig. 20 that cell numbers do not double exactly at the first division. This is probably because, in isolating the smallest cells, we enriched for inviable small cells. After division, the large mother cell will be an adequate size to start the programme for the next cell division, but the small daughter cells will need to grow before they attain the critical size. Therefore, one would expect half of the population to divide again, while the daughter cells develop to the critical size. This is, in fact, observed (the second plateau in Fig. 20). Indeed, the newly formed unbudded mother cells divide yet again before the original daughters divide. Since the original population contained primarily daughter cells, it is not surprising that they take longer than the average mass doubling time (7.4 h) of asynchronous cultures to divide. These results are what would be predicted if an asynchronous culture growing at a slow generation time is composed of mother cells having a generation time shorter than the mean of the asynchronous culture, and daughter cells having a generation time longer than the mean of the asynchronous culture .

XII. Some Technical Considerations A.

I S O L A T I O N OF YEAST N U C L E I

Methods to isolate yeast nuclei have recently been reviewed by Duffus (1975). Nuclei may be isolated from sphaeroplasts (Rozijn and Tonino, 1964; May, 1971; Wintersberger et al., 1971) or from whole cells (Bhargava and Halvorson, 197 1 ; Sajdel-Sulkowska et al., 1974; Wintersberger et al., 1973). Comparisons of nuclei obtained by the sphaeroplast method of May (1971) with the whole-cell method of Bhargava and Halvorson (1971) led Wintersberger et al. (1973) to conclude that the nuclei were very similar with regard to DNA, RNA and protein content, but yelds from the whole-cell method were lower and less reliable. The method of Bhargava and Halvorson (197 11, derived

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from that of Duffus ( 19691, depends on freezing cells in an Eaton press or French press and, from my own experience, the freezing step is critical. Published reports recommend freezing in a dry ice-ethanol mixture for about seven minutes. In practice, best results are obtained when freezing is permitted until a drop of water placed on the press just freezes without running down the outside of the press. If the water runs down the press before freezing, the press is not cold enough. Because of the time taken to prepare sphaeroplasts, it is impossible to use nuclei from sphaeroplasts to investigate different physiological states, for instance, the properties of nuclei from cells at different stages of the cell cycle. For such studies, the methods of Bhargava and Halvorson (1971) and Sajdel-Sulkowska et al. (1974) must be used, and it would be very useful if more reliable methods could be found to isolate nuclei from whole cells. B.

I S O L A T I O N O F A N U C L E O L A R FRACTION FROM YEAST NUCLEI

The nucleolus appears as a dense crescent within the yeast nucleus. Sillevis-Smitt et al. (1973) have isolated from the nuclei of Sacch. curlsbergensis a fraction rich in dense crescent material after treatment with DNAse. Their published electron micrographs reveal that the dense crescents are still attached to, and often partially encircled by, a nuclear membrane which has many ribosomes attached to the outer nuclear membrane. C.

I S O L A T I O N O F C H R O M A T I N FROM YEAST NUCLEI

Using methods similar to those used to isolate chromatin from nuclei of animal cells, Butteworth et al. (197 1 ) and Wintersberger et al. (1973) have prepared a chromatin fraction from nuclei of Sacch. cerevisiae. D.

INHIBITION O F NUCLEAR FUNCTIONS

Schindler and Davies ( 1975) have recently reviewed inhibitors of macromolecular synthesis in whole cells of yeast, and I will restrict my remarks to those inhibitors which appear to be the most useful. Hydroxyurea inhibits ribonucleotide reductase activity and prevents formation of deoxynucleotide substrates. It has no direct effect on replication of DNA. Hydroxyurea at 0.075 M has been used successfully to inhibit DNA synthesis (Slater, 1973, 1974) and, at this con-

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centration, rates of RNA and protein synthesis are only slightly altered. The rapidity of inhibition of replication indicates that the pools of deoxyribonucleotides must be small in size. In my experience, with cells grown at slow rates, higher concentrations of hydroxyurea (up to 0.2 M ) are necessary for rapid cessation of DNA replication. Trasiquone (Trenimon)at 10 pglml inhibits yeast DNA synthesis selectively, and is the other drug of choice of inhibiting DNA synthesis (Jaenike et al., 1970; Hartwell et al., 1974). There are very few specific inhibitors of RNA synthesis and, of these, perhaps lomofungin and 8-hydroxyquinoline, both chelating agents, are the best. 8-Hydroxyquinoline is likely to be the more popular because of its easy availability and its greater reliability (Fraser and Creanor, 1974). 8-Hydroxyquinoline, at a concentration of about 50 pglml, is a rapid and selective inhibitor of RNA synthesis in fission yeast. Although DNA synthesis was significantly inhibited by this drug, protein synthesis, cell growth and isotope uptake were not (Fraser and Creanor, 1974). At low concentrations, 8-hydroxyquinoline inhibits synthesis of ribosomal-RNA and messenger-RNA, but has no effect on synthesis of transfer-RNA or 5.5 RNA. It is thought that this differential effect occurs because of differing cation requirements of the three yeast RNA polymerases (Fraser and Creanor, 1974). Anderson and Roth (1974) isolated an adenosine-requiring strain of Sacch. cerevisiae which is sensitive to the action of cordycepin to which wild- type cells are impermeable. Cordycepin blocks addition of poly(A) tracts to heterogenous RNA (Penman et al., 1970) and use of this mutant should facilitate studies on yeast RNA metabolism. E.

D N A ESTIMATION

Saccharomyces cerevisiae lacks thymidine kinase (Grivell and Jackson, 1968) and takes up thymine and deoxythymidine rather poorly (Grenson, 1969;Jannsen et al., 1968; Lochmann, 1965).Therefore, for a long time, it was not possible to label yeast DNA specifically with either of these two compounds. Instead, the laborious and somewhat inaccurate diphenylamine method (Burton, 1956) has been used. A recent detailed account of this method is given by Stewart (1975). Recently, however, it has become possible to label DNA specifically with isotopes. Jannsen et al. (1968) were the first to demonstrate that exogenously supplied deoxythymidine 5’-monophosphate (dTMP) could be incorporated in some strains specifically into DNA although

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the efficiency of labelling was poor. To improve this efficiency, successful attempts were made to isolate mutants able to grow with exogenous dTMP after inhibition of endogenous dTMP biosynthesis with folic acid antagonists (Brendel et al., 1975; Fath et al., 1974; Jannsen et al., 19 73 ; Laskowski and Lehmann-Brauns, 19 73 ; Wickner, 1974, 1975).

These mutants should be very useful for the routine labelling of DNA. In addition, because some of these mutants can incorporate deoxybromouridine 5'-monophosphate (dBrUMP) into DNA and as a result become sensitive to 313 nm light (Jannsen et al., 19731, they should be very helpful for isolating mutants temperature-sensitive for DNA synthesis. F.

NUCLEAR STAINING

A variety of techniques have been used to stain yeast nuclei. These include staining with acridine orange (Royan and Subramanian, 19601,

FIG. 2 1. Nuclear staining in vegetatively dividing yeast cells. Logarithmic-phase cells in minimal medium were exposed to 0.1 M hydroxyurea to accumulate cells in niidnuclear division. A sample was stained, embedded in agar, and photographed using a Leitz vertical illumination fluorescence microscope with blue excitation filter KP400, a 510 nm dichroitic mirror and suppression filter K460 (position 2). Reproduced with permission from Slater (1976).

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FIG. 22. Nuclear staining in sporulating yeast cells. Samples removed after 5 and 7 hours in sporulation medium were suspended in 70% ethanol and refrigerated overnight. The samples were then stained, mixed to exhibit various phases ofmeiosis, and photographed as in Fig. 21. Reproduced with permission from Slater (1976).

Giemsa (Robinow and Marak, 1966; Robinow, 1975)and acetic orcein (Day and Jones, 1972). These techniques suffer from one or more of the following handicaps : multiple manipulations are required, a critical incubation step which can lead to irreproducibility is necessary, the procedure is lengthy. Recently, Slater ( 1976)reported a rapid and simple method to stain nuclei of Sacch. cerevisiae which suffers from none of these defects. He found that it was possible to stain nuclei with mithramycin without hydrolysing RNA or removing the stain before observation. The method he used to stain vegetatively growing cells involved incubating a sample ofcells with an equal volume of50% (v/v) aqueous ethanol containing mithramycin (0.4 mg/ml) and 30 mM MgCI,. After 15 min counterstain (acridine orange at a final concentration of 1 pglml may be added if desired), a drop is examined as a wet mount (Figs. 2 1 and 22). XIII. Conclusions

“The structure and reproduction of the nucleus of the yeast of baking and brewing, Saccharomyces cerevisiae was the subject of bitter

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polemics at the turn of the century and a mycology text of our time informs us that the controversy over the structure of' the nucleus . . . continues undiminished. Some progress there has been" (Robinow and Marak, 1966). In this review I have tried to indicate just what progress there has been on the structure and functions of the yeast nucleus in the last decade. XIV. Acknowledgements

I thank B. D. Hall, L. H. Hartwell, C. Saunders, S . J . Sogin, M. Jagadish, S. K. Welch, P. A. Whitney and L. M. Hereford f'or unpublished information, J. Haber and S. Sogin for discussions during the preparation of this manuscript and H. 0. Halvorson for extending to me the hospitality of his laboratory. Unpublished work from my laboratory was supported by the Irish Medical Research Council, and by Arthur Guinness, Son and Co. Ltd. REFERENCES

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Biology, Physiology and B iochemistry of Hyp ho microbia W. HARDER AND MARGARET M. ATIWOOD Departments of Microbiolog y, The Universities of Groningen, The Netherlands, and Sheffield, England I . Introduction

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11. Biologyand PhysiologyofHyphomicrobia

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A. The genus Hyphomicrobium . . . . . . B. Enrichment and Isolation . . . . . . . . C. Nutrition . . . . . . . . . . . . D. Life Cycles and Pleomorphism . . . . . . . . E. Effect of Environment on Morphology . . . . . . F. Ecology . . . . . . . . . . . . 111. Biochemistry of Hyphornicrobia . . . . . . . . A. Biochemistry of Growth on Reduced One-Carbon Compounds . B. Biochemistry of Growth on Two-Carbon Compounds. . . . C. Possible Role of Cytochromes . . . . . . . . D. Biochemical Basis for Restricted Methylotrophy in Hyphomicrobia IV. Conclusion . . . . . . . . . . . . . V. Acknowledgements . . . . . . . . . . . References . . . . . . . . . . . . .

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303 305 305 311 314 319 326 329 332 332 349 350 352 355 356 356

I. Introduction

Hyphomicrobia are stalked bacteria which reproduce by a budding process. These organisms have been recognized for a long period of time and were observed originally in enrichment cultures for nitrifying bacteria. Rullman ( 1897) considered the stalked organism in such an enrichment to be a nitrifying bacterium and named it Nitrosobacterium formae novae. Stutzer and Hartleb (1898) observed an organism with a similar morphology and proposed the name Hyphomicrobium vulgare. These authors described “thin filaments with terminal knobs” on this organism and considered the filaments to be mycelium and the knobs 303

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to be chlamydospores. They did not associate these features with any mode of reproduction and, although doubtful of any role in nitrification, did not rule out this possibility. This situation remained until Prouty ( 1929) reported the organism to be a non-nitrifying contaminant of the nitrifying cultures. Despite this early recognition that the organism was a contaminant, it proved difficult to isolate the Hyphomicrobium sp. in pure culture. As a result little research was undertaken with the organism. In 1934 Kingma Boltjes succeeded in isolating Hyphomicrobium uulgare from an enrichment culture of nitrifying bacteria, and he identified the filament of this organism as a stalk and described the mode of reproduction as a budding mechanism. Henrici and Johnson (1935) also identified the filament of Hyphomicrobium uulgure as a stalk, and grouped the organism with other stalked bacteria. However, Stanier and van Niel (1941) considered that, since the mode of reproduction was a budding mechanism, these organisms could not be included in the Schizomycetes and placed them in a provisional appendix along with other organisms whose taxonomic position remained to be defined. Thus, until recently, despite the early recognition of these stalked bacteria, apart from a further appreciation of their unusual morphology and reproduction (Zavarzin, 1960; Leifson, 1964) and biology (Hirsch and Conti, 1964a, b; 19651, little was known about the physiology, biochemistry (Mevius, 1953; Naveke, 1957)and taxonomy of the group. This was due basically to, on the one hand, the lack o f a rapid method for their isolation from natural habitats into pure culture and, on the other hand, the low growth rates and poor growth yields encountered with the few pure isolates available. In addition, many of these isolates characteristically grew in clumps or rosettes, and often adhered to the walls of the culture vessel (Hirsch and Conti, 196413, 1965). With the recent development of enrichment procedures based upon denitrification in the presence of methanol (Sperl and Hoare, 197 1 ; Attwood and Harder, 19721, a rapid procedure with a high degree of selectivity became available. Furthermore, the isolates of Hyphomicrobium spp. thus obtained generally appeared to have higher growth rates and gave higher growth yields than the? pi-evious isolates. Moreover, the cultures grew homogeneously with little clumping or rosette formation and no adherence to the walls of the culture vessels.

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Thus many isolates of Hyphomicrobium spp. which were easy to handle in the laboratory became available in pure culture. As a result, in the last few years, work has been undertaken in an attempt to understand aspects of the biology, physiology and biochemistry of these organisms. Therefore, it seemed appropriate to review this work so that information available might be evaluated and important areas of further research identified. Hyphomicrobium spp. as a group of organisms have formed part of previous reviews on freshwater bacteria (Henrici and Johnson, 1935), bacterial diversity with respect to unusual morphology (Starr and Skerman, 1965) and as examples of stalked or budding bacteria (Zavarzin, 1961; Schmidt, 1971; Hirsch, 1974a). 11. Biology and Physiology of Hyphomicrobia A.

THE GENUS

HYPHOMICROBIUM

At the present time it is difficult to give a concise definition of the genus Hyphomicrobium. This is due to the fact that, although many different isolates have been purified (Zavarzin, 1960; Hirsch and Conti, 1964a; Sperl and Hoare, 197 I ; Attwood and Harder, 1972; Shishkina and Tro tsenko, 1974; Takada, I9 7 5 ) , their identification has been based largely on morphological criteria and little or no reference to their physiology has been made. In the few instances where the latter information is available, and in particular when the nutritional requirements of the organisms have been published, there are certain areas of disagreement between different groups of workers. These areas will be discussed in the appropriate sections. In the most recently published bacterial classification scheme (Bergey’s Manual of Determinative Bacteriology 8th Edition, 19 741, bacteria which reproduce by asymmetrical binary fission have been subdivided into two major groups with respect to their ability to produce prosthecae. Those organisms which produce prosthecae, the so-called prosthecate bacteria, are further subdivided on the basis of the mechanism of cellular reproduction. Prosthecate bacteria which reproduce by a budding process are placed in one of three genera, namely Hyphomicrobium,Hyphomonas or Pedomicrobium.This last genus is separated from the other two on the basis of a morphological criterion. Members of this genus produce prosthecae from several sites

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over the cell surface whereas Hyphomicrobium spp. or Hyphomonas spp. produce prosthecae from only one or both poles of the cell. The criteria which have been used to distinguish between the genera Hyphornicrobiurn and Hyphomonas are less distinct. Organisms in these genera cannot be differentiated on morphological grounds, but are separated on the basis of DNA-DNA base sequence homology studies (Moore and Hirsch, 1972). Furthermore, Hyphomonas spp. appear to be unable to utilize one-carbon compounds for growth (Pongratz, 1957 ), whereas these compounds are the preferred, if not the only, carbon sources for the growth of Hyphomicrobium spp. (Hirsch and Conti, 196413; Sperl and Hoare, 197 1 ; Attwood and Harder, 1974; cf. Hirsch, 1974a). Using the above criteria to characterize the different genera, the currently accepted description of the genus Hyphomicrobium was formulated (Hirsch, 197413). However, in the light of the reported literature now available, this description is inaccurate. For example, many Hyphornicrobiurn spp., particularly the more recent isolates, have been reported to grow in liquid culture with a uniform turbidity (Attwood and Harder, 1972; cf. Hirsch, 1974a)whereas the description of the genus states that liquid media are never turbid. Similarly evidence has accumulated which suggests that it is not usual, if at all possible, for Hyphornicrobium spp. to show good growth with carbon sources containing from three to six carbon atoms as the sole source of carbon and energy (Zavarzin, 1960; Hirsch and Conti, 1965; Sperl and Hoare, 1971; Attwood and Harder, 1974; see Section 11-C, p. 317). Indeed, Kingma Boltjes ( 1936) showed that, although Hyphornicrobium vulgare Stutzer and Hartleb appeared to be able to utilize the carbon compounds cane sugar and asparagine, if stringent precautions were taken to ensure that all trace organic compounds were removed from both the inorganic growth medium and the atmosphere, then growth was restricted to one-carbon compounds and acetate. Despite this evidence, in the description of the genus it is stated that good growth occurs with carbon sources such as 0.2% (w/v) lactate, succinate and mannitol. Finally strains of Hyphomicrobium are available which, when grown under anaerobic conditions in the presence of nitrate, accumulate nitrite (Sperl and Hoare, 1971; J. B. M. Meiberg, unpublished data). Again, this is contrary to the published description of the genus (Hirsch, 197413). Therefore, for the purpose of this review, the genus Hyphomicrobium will be defined as that group of organisms which consists of cells (0.5 to 1.O x 1.3 pm) which are either rod-shaped with pointed ends, oval- or bean-shaped, and produce mono- or bipolar filamentous outgrowths

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FIG. 1. (a)Phase-contrast photomicrograph of a Hyphomicrobium sp. (b)Motile swarm cell. (c) Non-flagellated “swarm” cell entering the maturation phase. (d) Filament formation. (e) “Daughter” cell synthesis. (f) Asymmetric-division yielding a mature “mother” cell and an immature “daughter”. The arrow shows the point of division. Organisms were grown in the mineral salts medium of Attwood and Harder ( 1972).

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(prosthecae or hyphae) of varying length (Fig. la). The length of'these filaments depends upon the environmental conditions but the diameter remains approximately constant (see Section 11-E, p. 326). The filaments may show true branching (Zavarzin, 1960; Hirsch and Conti, 1964a). Multiplication occurs by the formation of a bud or daughter cell at the tip of the filament. During maturation, the bud develops one to three polar or subpolar flagellum(ae)(Kingma Boltjes, 1936; Zavarzin, 1960; Leifson, 1964) and, after a period of active movement, the bud breaks away from the filament of the original cell (mother cell) and becomes an actively motile swarmer. After some time, the swarmer loses its flagellum(ae) and forms a filament to become a mother cell and repeat the life cycle (see Fig. 1 and Section 11-D, p. 320). The mother cells may aggregate with the filaments and buds to the outside to form the so-called rosette colonies (see Fig. 6, p. 328).This tendency to show rosette formation appears to be strain-dependent, but may also be influenced by environmental conditions (Attwood and Harder, 1972). All Hyphomicrobium spp. are chemo-organotrophs with a limited range of carbon and energy sources for growth. They can all utilize a range of one-carbon compounds and, in addition, a number of strains can grow on some two-carbon compounds and four-carbon compounds which are metabolized via acetyl-CoA (see Section 111-B, p. 349). They can be considered therefore as obligate methylotrophs or restricted facultative methylotrophs depending upon their ability to utilize two- and four-carbon compounds (Colby and Zatman, 19 75a). Hyphomicrobium spp. form polyhydroxybutyrate as storage material. They are aerobic, although many strains are able to grow under anaerobic conditions with nitrate as the terminal electron acceptor and show optimum growth rates when grown in a well buffered medium at neutral or slightly alkaline pH values in the dark at temperatures between 25-37OC. The type species is Hyphomicrobium uulgare Stutzer and Hartleb, 1898. Other species have been named, but recently Hirsch (1974b) has challenged the validity of three of these isolates to be included as Hyphomicrobium sp. Hyphomicrobium neptunium Leifson ( 1964) was isolated from stored sea water and, as a result of morphological similarities with Hyphomicrobium uulgare, was considered to be a Hyphomicrobium sp. Studies of DNA-DNA base sequence homologies with known Hyphomicrobium isolates including Hyphomicrobium neptun-

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ium (Moore and Hirsch, 1972)resulted in the suggestion that the latter isolate was more closely related to another budding bacterium Hyphomonas polymorpha (Pongratz, 1957 ) than to Hyphomicrobium spp. Transfer of this species into the genus Hyphomonas has been recommended (Hirsch, 197413). Hyphomicrobium indicum, Johnson and Weisrock (19691, was isolated from mud at 400 m depth in the Indian Ocean. The cells were rod or coccus shaped ( 1.0 to 2.5 ,urn) and were joined by slender filaments. This morphology was taken to be indicative of' a budding mechanism and, on this evidence, the authors named this organism a Hyphomicrobium sp. However, since there is now evidence that this organism does not form buds and has a quite different metabolism and a G + C base ratio outside the range of 59.2 to 66.8% reported for Hyphomicrobium spp. (Mandel et al., 19721, transfer of this organism to another genus has been recommended (Hirsch, 197413). Hyphomicrobium T3 7 was isolated from deposits of manganese compounds found in hydro-electric pipes in Tasmania (Tyler and Marshall, 1967a, b ; Bauld et al., 1971a, b). This organism was reported to show pleomorphism. When growing on the mineral medium 33 7 (Hirsch and Conti, 1964a) with methanol as the carbon source, and supplemented with 0.02% MnSO,. 4H,O, the morphology of the population was typically that of a Hyphomicrobium sp. The mother cells had filaments showing little branching and the pear-shaped buds had a polar flagellum. However, when growing with methanol as the carbon source on Pringsheim mineral medium (Pringsheim, 1949) with 0.002% MnSo4.4H,0 and 0.005% yeast extract added, or on the mineral medium 337 with methylamine as the carbon source, the cells assumed bizarre shapes with several short but richly branched filaments which projected from all Gver the cell surface. This morphology had been described previously as characteristic of the genus Pedomicrobium (Aristovskaya, 1961). The swanners retained the pear shape. O n this evidence, Bauld et al. (1971b) questioned the validity of the genus Pedomicrobium, and suggested that this reflected one growth form of Hyphomicrobium. O n the other hand, Hirsch ( 1968) had claimed that the genus Pedomicrobium contained those organisms which produce filaments from one or several points over the cell surface, whereas Hyphomicrobium spp. produce filaments from the poles only, The isolate Hyphomicrobium T37 was deposited in the National Collection of Industrial Bacteria at the Torry Research Station, Aberdeen (U.K.) and has been designated Hyphomicrobium 10099 in their catalogue. In the hands of the present authors, this organism

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exhibits the classical morphology of Hyphomicrobium spp. with an accepted amount of branching of the filaments and mono= and bipolar filament formation when growing in mineral media with methanol or methylamine as the carbon source and in the presence or absence of manganese ions (M. M. Attwood and W. Harder, unpublished results). However, since under certain conditions a Pedomicrobium-like morphology has been observed with this organism, it clearly cannot be accommodated in the genus Hyphomicrobium as already defined. This problem could be resolved if the definition of the genus was qualified so that morphological criteria only apply under optimal growth conditions. This leaves the option of accommodating isolates which show a somewhat broader pleomorphic variability than that accepted in the current definities of the genus, when growing under unfavourable environmental conditions (Hirsch and Conti, 1964a; Shah and Bhat, 1968; Johnson and Weisrock, 1969; Whittenbury and DOW, 1977). Since, at the present time, it may not be appropriate to qualify the definition of the genus because of this one observation with Hyphomicrobium 10099 (T37),we suggest that this organism is retained within the genus at least for the time being. Further developments in this area may indeed require the suggested modification of the definition. Isolation of two new strains of Hyphomicrobium has been reported recently. The isolation of a new Hyphomicrobium sp. which showed a strong preference for one-carbon compounds as sources of carbon and energy was reported by Shishkina and Trotsenko ( 1974).These authors suggested that the organism should be classified as a methylotroph and, on the basis of a comparison of the new isolate with Hyphomicrobium uulgare N Q they identified the organism as a new strain of Hyphomicrobium uulgare. However, a formal description of this organism has not been published. Similarly Takada ( 1975) described the isolation from rice fields of a strain of Hyphomicrobium which had different characteristics as compared to those described for other Hyphomicrobium isolates, and the author claims that this isolate should be regarded as a new species. He proposes the name Hyphomicrobium coagulans. However, a final decision on this isolate must await further detailed documentation. Thus, despite the recorded isolation of a number of new species, Hyphomicrobium uulgare remains the only valid species formally described. For more detailed accounts of the relationships between Hyphomicrobium sp. and other stalked bacteria, see Zavarzin (19611, Starrand Skerman (1965)and Hirsch (1974a).

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

B.

ENRICHMENT AND

31 1

ISOLATION

The original attempts to isolate Hyphomicrobium spp. from enrichment cultures of nitrifying bacteria proved rather difficult (Kingma Boltjes, 1936; Mevius, 1953).In view of these difficulties, attempts have been made to develop enrichment procedures for hyphomicrobia. Various workers found that enrichment procedures based upon aerobic incubation in the presence of an added carbon and energy source were generally unsuccessful. This has been attributed to the fact that any Hyphomicrobiurn spp. initially present in the inoculum were rapidly overgrown by other micro-organisms. Zavarzin ( 1960)reported a successful enrichment procedure using a liquid medium without an added carbon source, but incubating the inoculum aerobically in the presence of methanol vapour. Under these conditions, after 3 to 5 days a thin pellicle of growth was observed on the vessel walls and the surface of the medium. Subsequent isolation of pure cultures was easily obtained using standard methods; however, the growth rates of the isolates were rather low. Using the characteristic ability of Hyphomicrobium spp. to grow in the absence of any added carbon- and energy source, Hirsch and Conti ( 1964a)developed an enrichment procedure which has since been used successfully to isolate Hyphomicrobium spp. from a wide range of natural habitats. These workers used a mineral medium without an added carbon source to discourage growth of other organisms. The inoculum was added, and aerobic incubation led to the formation of thin surface pellicles which contained stalked organisms from which Hyphomicrobium spp. could be isolated. I t was assumed that the concentration of organic carbon added with the inoculum was negligible so that the organisms were growing predominantly in an oligocarbophilic manner at the expense of volatile carbon compeunds present in the laboratory atmosphere. This procedure successfully overcame the problem of other organisms outgrowing Hyphomicrobium spp. present in the inoculum, but necessitated incubation periods of 4 to 18 weeks before growth became apparent. Using this method, Hirsch and Conti (1964a, b) isolated 1 1 new strains and compared them with cultures of a strain isolated in Germany by Mevius (1953). Hyphomicrobium spp. have been isolated from a wide variety of'soil and water samples using modifications of this procedure. These modifications involved addition of low concentrations of' carbon compounds as a vapour or slides to encourage attachment 01' stalked

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W. HARDER AND MARGARET M. ATTWOOD

bacteria (Kriss, 1963; Hirsch and Rheinheimer, 1968; Marshall et al., 197 1 ) and the use of dilute mineral media o r even distilled water in the presence of carbon monoxide (Hirsch, 1974a). The finding that aerobic incubations with one o r more added carbon sources never lead to a successful enrichnierit of'Hyphomicrobium spp. except when the carbon source (methanol)is present as a vapour suggests that, if low but constant concentrations of' suitable nutrients could be produced and maintained in a certain eiiviroimieiit over a period of time, enrichment of these organisms may occur. Such an approach is riow possible using an apparatus designed by Caldwell and Hirsch (1973).These workers described a two-dimensional steady-state diffusion system in which micro-organisms can be grown in nutrient concentration gradients. In this system, the organisms present in the inoculum are exposed to various concentrations and combinations of' two test nutrients. Unfortunately no data are available to show isolation of hyphomicrobia from a natural environment; however, using this procedure, the authors have reported successful separation of an artificially mixed population of a Hyphomicrobium sp. Rhodomicrobium uannielii and a Thiopedia sp. when gradients of' methylamine and sodium sulphide were employed. Thus, the potential use of' this method to enrich for specific fastidious bacteria like hyphomicrobia has been demonstrated. This method could be used to enrich for and simultaneously isolate several members of' a mixed population whose growth requirements differ only slightly. Indeed, this method need not be limited to isolation procedures but, with sollie further development, could be a useful tool for ecological studies. Recently, an enrichment procedure was developed based upon the preference of' hyphoniicrobia to grow o n one-carbon conipounds, in particular methanol. The method involves incubation of' a mineral salts medium with methanol as the carbon- and energy source under anaerobic conditions in the presence of nitrate as the terniiiial electron acceptor, and has resulted consistently in rapid and highly selective enrichment of Hyphomicrobium spp. from a wide range of natural habitats (Sperl and Hoare, 197 1 ; Attwood and Harder, 1972). When the mineral salts medium is prepared in natural or artificial sea water rather than demineralized water, inocula from marine erivironmeiits can be used successfully. With this procedure, the initial incubation period can be decreased from a matter of weeks (Hirsch and Conti, 1964a) to 3-9 days. The time required to ensure a high ratio 01' Hyphomicrobium cells to other organisms in the initial growth from the

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inoculuni was found to be dependent upon the concentration of' organic material in the inoculum and the nitrate concentration in the medium. If the concentration of organic compounds in the inoculurn was high then the degree of selectivity with the method was decreased (Attwood and Harder, 1972).This probably results from the ability of other organisms to utilize organic compounds in the inoculum at the expense of the nitrate in the medium, and consequently these organisms can outgrow the Hyphomicrobium spp. present. If' the initial concentration ofnitrate in the medium is low, this may lead to nitrate depletion with the consequent failure to enrich for Hyphomicrobium 5pp. However, if the initial nitrate concentration is increased, the method can be used to isolate hyphomicrobia from habitats rich in organic content. Successful enrichment has been obtained from a variety of'soil types, mud and aquatic environments including an industrially polluted canal water rich in both organic and sulphur compounds. With this latter inoculum, despite an initially rich microbial population with respect to the number and microbial type, after 1 7 to 20 days microbial growth in the enriched cultures was predominantly due to Hyphomicrobium spp. Methylamine (Attwood and Harder, 19721, dimethylamine and trimethylamine (Meiberg and Harder, 1976) can be substituted for methanol as the carbon- and energy source without any loss in the selectivity of the method. However, the use of formate, although able to support growth of Hyphomicrobium spp., resulted in a loss of specificity. Microbial growth then consisted predominantly of Pseudomonas spp. After 4 to 5 transfers of the enriched culture into the same medium and using anaerobic incubations, pure cultures can be obtained by streaking onto solid media consisting of the mineral salts medium with methanol or methylamine as the carbon- and energy source, but without added nitrate and solidified with agar. The streak plates are incubated aerobically, and pure single colonies of the isolates are obtained. Failure to transfer the initial enrichment cultures through the series of anaerobic incubation procedures recommended may result in apparent production of isolated colonies on the agar plates, but each colony may consist of a mixture of Hyphomicrobium cells and a small actively motile contaminant. This contaminant cannot easily be removed by repeated plating procedures. Microscopic observations alone generally are not able to differentiate these small organisms from the true Hyphomicrobium swarmer cells. A criterion routinely used to establish the purity of cultures of Hy~homicrobium spp. was the

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inability to grow within 7 days on a range of nutrient agar media (Mevius, 1953; Attwood and Harder, 1972).A word of caution should be included with respect to this method, however, because it will only detect contaminants which are able to grow on the media used. Once pure cultures have been obtained they can be maintained o n methanol or methylamine agar at 4OC in the dark for months and can be preserved by lyophilization or cold storage in liquid nitrogen.

C. NUTRITION

The ability of members of the genus Hyphomicrobium to show optimal growth in mineral salts media containing calcium ions with added trace elements, nitrate or ammonia as the nitrogen source, and carbon compounds containing no carbon-carbon bond as the carbon- and energy source has been known for many years (Hirsch and Conti, 1964b; Hirsch and Rheinheimer, 1968; Sperl and Hoare, 1971; Attwood and Harder, 1972; Caldwell and Hirsch, 1973). The substrates most widely used for growing these budding bacteria are methanol, methylamine and formate (Kingma Boltjes, 1936; Mevius, 1953; Korte and Engel, 1955; Naveke, 1957; Hirsch and Conti, 1964b; Attwood and Harder, 1972; Shishkina and Trotsenko, 1974). A detailed investigation into growth of the pellicle-forming isolates Hyphomicrobium uulgare NQ and Mev in batch culture was undertaken by Hirsch and Conti ( 1964b).The isolates were tested for growth in the mineral salts medium 337 supplemented with various carbon compounds. The cultures were incubated at 3OoC in the dark for a number of weeks, and growth was then recorded. After 5 weeks’ incubation, no growth was found with the following compounds: glyoxylate, oxalate, glycine, threonine, glycollate, pyruvate, tartronate, 3phosphoglycerate, methylsuccinate, methylmalonate, P-hydroxypyruvate, citramalate, fumarate or alanine. 0 nly methanol and methylamine supported good growth of both isolates after 2 weeks; in addition, Hyphomicrobium vulgare N Q showed similar growth over this period with formate. These workers reported a third group of compounds whereby measurable growth was obtained after prolonged incubation periods. However, the extent of these prolonged incubations was not recorded. Compounds in this group include acetate, lactate, acetamide, forrnamide, hydantoate, serine, glutamate, malate, succinate and aspartate. Similar results showing that the preferred carbon sources for the growth of Hyphomicrobium spp. were one-carbon

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compounds but that other compounds such as glycerol, lactate or succinate could support growth of these organisms have been reported by other workers (Sperl and Hoare, 1971). However, care should be taken when interpreting poor growth yields, particularly when associated with long incubation periods, since misleading results can occur with organisms which are able to grow oligocarbophilically at the expense of volatile carbon compounds in the atmosphere (Kingma Boltjes, 1936). Furthermore, since more than one morphological form is found in the life cycle of hyphomicrobia (Zavarzin, 1960; Leifson, 1964; C. S. Dow and A. Lawrence, unpublished observations) it is extremely difficult to differentiate between the activity motile swarmers of hyphomicrobia and small motile contaminants using microscopic observations alone. With these problems in mind, M. M. Attwood and W. Harder (unpublished observations) screened a number of isolates for their ability to use a wide range of compounds as sole source of carbon and energy. Organisms were grown to the midexponential phase in media containing methanol or methylamine and used as the inoculum (3.0%, v/v) for a mineral salts medium containing ammonium ions as the nitrogen source and supplemented with different carbon compounds over the range of 10 to 150 mM. The cultures were incubated with shaking in the dark at 3OoC and observed daily for an increase in turbidity. Growth was detected after 2 days’ incubation in media containing the carbon sources most readily used by the isolates. All cultures were incubated for a total of 2 weeks. After this period the cells were harvested and growth recorded (Table 1 ) ; in all cultures were growth occurred this was found to be due exclusively to the development of hyphomicrobia. In these experiments, no attempt was made to measure accurately either growth rate or growth yield of the organisms and therefore the data given only reflect the ability of the different organisms to grow on the compound. All isolates screened showed very similar growth patterns. Good growth was observed on carbon compounds with no carbon-carbon bond (methanol, methylamine, dimethylamine and trimethylamine). Formate was not included in the screen because of the problem of drift of pH value with growth. However, in pH-controlled continuous cultures, formate did support growth of Hyphomicrobium spp. (unpublished observations). Some isolates could grow using the two-carbon substrates ethanol and acetate and the four-carbon substrate 3-hydroxybutyrate. This last compound is metabolized as a two-carbon substrate (Attwood and Harder, 1974). Growth was not detected with any of the

TABLE 1 . Growth ofHyphornicrobium isolates 011 vaIiou5 carbun conipouiidh Cayboli corllpounda

Coneenti-ation

Isolate

(mM)

N one Methanol Methylamine Dimethylamine Trimethylamine Ethanol Sodium acetate 3 - H ydroxybu tyra te

-

I56 74 24.5 38.8 55.6 15 19.2

X

WA/B

0.04 0.26 0.28 0.22 0.20 0.19 0.24 0.28

0.07 0.34 0.63 0.19 0.13 0.15 0.12 0.S1

Ti

(T37) 10099

0.01 0.03 0.790.29 1.20 0.23 0.41 0.03 0.31 0.04 0.73 0.05 0.48 0.03 0.10 0.04

3T+M

Har3

B522

H526

L530

0.03 0.34 0.55 0.01 0.05 0.08 0.04 0.04

0.03 0.36 0.16 0.02 0.03 0.09 0.05 0.02

0.01 0.17 0.22' 0.00 0.00 0.00 0.03 0.00

0.02 0.10 0.26

0.00 O.lb

0.01

0.08 0.01 0.01 0.01

0.23 0.00 0.07 0.00 0.01 0.00

MC748 COiN2

0.01 0.24 0.30 0.02 0.10 0.01

0.01 0.01

0.01 0.24' 0.38 0.01

0.03 0.01 0.01 0.01

NQIIO.i2

Mcv533 ZVbL'O GI.

0.0 I (1.40

0.01 0.1(i

0.20 0.01 0.04

0.33 0.01 0.01

(1.18 0.02 0.02

N.D.' 0.00 0.02

0.0 I

0.5.5 0.20 0.01

0.06 0.42 0.211d 0.Oi

a Each compound was tested over a rauge 01 concentrations. T h e value 1-ecoi-ded is the giowth (mg PI otein/5 nil 01 ccll suspeii~iou)aftel. 2 weeks incubation at 30OC with shaking at the optiniuni conecutration ol tlir carbon compound added. GI-owth was not obsel-ved with any isolatr when the tollowing carbon coinpounds were added : urea, glyoxylate, glycero1,l lactate, pyruvatc, malate, succ inate, citi-ate, ribose, Iructosc, glucose, glycine, oL-serine, p-alanine, butyrate, formamide or asparagine. The value indicates thc optinial growth at a methanol cunceiitration of62.4 m M . The value indicates the optimal growth at a methylamine concentration of 29.6 mM. dThe value indicates the optimal growth at an acetate concentration of 8 mM. ' N.D. indicates that the value was not dctci-mined.

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other carbon compounds (C,-C6) screened. These compounds included some of those named in the third group reported by Hirsch and Conti (196413). From this survey, M. M. Attwood and W. Harder (unpublished observations) concluded that the growth substrates for these organisms were one-carbon compounds, and that Hyphomicrobium isolates were either obligate methylotrophs or restricted facultative methylotrophs (Colby and Zatman, 1975a), depending upon their ability to grow on acetate, ethanol o r 3-hydroxybutyrate. The authors were careful in the selection of isolates used in this survey. The isolates included those enriched by denitrification in the presence of methanol ( X and WA/B), by denitrification in the presence of trimethylamine (T,) and using the oligocarbophilic procedure (Har. 3 ) and B522, H526, L530, MC748 and C0582 which were kindly supplied by Professor P. Hirsch, University of Kiel, West Germany. One isolate (ZV 620) had been enriched by Zavarzin in the presence of gaseous methanol and two (NQI 1052 and Mev533 Gr) were pellicle-forming isolates. Furthermore, amongst these were isolates which had been enriched from different soils, muds, marsh swamps and river water. I n this way, any significant difference which could be associated with groups of isolates based upon either the procedure used for their isolation o r their natural habitat as suggested by Hirsch (1974a) would be detected. However, no significant differences were observed. This is somewhat unexpected in view of the widespread distribution of these organisms in Nature. Thus, our conclusion is at variance with those of Hirsch and Conti (1964b)and Sperl and Hoare ( 197 1). For, although these authors reported a distinct preference for growth on one-carbon compounds, especially methanol and methylamine, they considered several other substrates including lactate, glycerate, serine, succinate and glutamate able to support growth. Similarly, Shishkina and Trotsenko (1974) observed slight growth of Hyphomicrobium vulgare strain 3 after two weeks in media containing valeric, butyric, malic, lactic or pyruvic acid. Slow growth of the organism was also observed in a medium lacking a carbon source and, because a wide variety of other carbon sources did not support growth, these authors considered the above five carboxylic acids as (rather poor) growth substrates. In view of the differences in the results reported by these groups of workers, two isolates, Hyphomicrobium X and G,obtained by denitrification in the presence of methanol (Attwood and Harder, 1972) and Hyphomicrobium B522 isolated by the oligocarbophilic procedure and supplied by Professor Hirsch, were investigated in more detail.

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Enzymic analysis of cell-free extracts from Hyphomicrobium X and G failed to detect any one of several pyruvate-metabolizing enzymes and in particular pyruvate dehydrogenase. This last enzyme was also absent from Hyphomicrobium B522 (Attwood and Harder, 1974; Harder et al., 1975; see Section 111-D, p. 354). This limited ability to utilize pyruvate means that these organisms are unable to grow on any compound which requires pyruvate to be oxidized to acetyl-CoA for entry into the TCA cycle in order to generate energy. Therefore, it would be difficult to understand how such isolates could grow on compounds containing more than two-carbon units unless the compounds are initially catabolized to two-carbon or one-carbon compounds. Little information has been published on the growth rates of' these organisms. Doubling times of 6.9 and 9.8 h have been reported for Hyphomicrobium X when grown in batch culture on methanol or ethanol, respectively, at 30OC (Harder et al., 1973). A survey of the growth rates of six isolates of Hyphomicrobium grown in batch culture at 3OoC on methanol, methylamine and ethanol gave a range of doubling times from 6 to 23 h on methanol, 10 to 23 h on methylamine and 13 to 15 h on ethanol (Attwood and Harder, 1977).A double time for the first generation of Hyphomicrobium B522 was reported to be approximately 14.5 h when grown on agar slides or agitated in liquid media containing methanol or methylamine (Hirsch and Jones, 1968; Moore and Hirsch, 1973a). When a species of Hyphomicrobium was grown in an oxygen- and pH-controlled fermenter with methanol as the carbon source, a generation time of 4.7 h was recorded (Wilkinson and'Hamer, 1972). The range of nitrogen sources which support growth of all Hyphomicrobium isolates appears to be very similar, although it has been reported that isolates enriched from brackish water prefer nitrogen in the form of nitrate, whilst others show optimum growth yields when nitrogen is added as ammonium nitrogen (Hirsch and Conti, 1964b; Moore and Hirsch, 1972; Shishkina and Trotsenko, 1974). A recent survey of eight isolates (M. M. Attwood, unpublished observations) has shown that all of the strains tested have a very similar growth pattern with respect to the nitrogen source added. In the presence of methanol as the carbon and energy source, none of the isolates grew when glycine, aspartic acid, glutamic acid or nitrite was used as the nitrogen source. Some isolates could utilize nitrate, urea, formamide or L-serine and all isolates grew in the presence of ammonium nitrogen, asparagine or glutamine. When methylamine was added as both the

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carbon- and nitrogen source, growth was recorded with all of the isolates tested. Within the range of organic compounds screened as the nitrogen source, glutamine rather than urea reported by Hirsch and Conti (196413)was best for growth. Thus Hyphomicrobium spp. grew little, if at all, in rich organic media such as nutrient broth; their range of single growth substrates does not merely show a preference for one-carbon compounds as suggested by Hirsch ( 1974a) but appears to be limited to one-carbon compounds and some two-carbon compounds. The best conditions for growth have been found to be aerobic incubation in the dark at 3OoC in neutral buffered media containing calcium ions, trace elements, ammonium ions as the nitrogen source, and methanol or methylamine as the carbon source. Inhibition of growth by certain amino acids has been noted. Hirsch (1974a) reported inhibition of growth with most Hyphomicrobium isolates when growing on one-carbon compounds in the presence of’ D L - threonine, DL-phenylalanine, L-tryptophan or L-glutamine. The inhibitory effect of L-glutamine has not been observed by either Shishkina and Trotsenko (1974) or the present authors, but growth inhibition by glycine and serine has been observed with all isolates grown on methanol (M. M. Attwood, unpublished observations). Certain isolates, particularly those which form pellicles, are reported to accumulate on the glass walls of culture vessels (Large et al., 1961; Hirsch and Conti, 1964b). ‘Normal laboratory light’ will prevent most, if not all, of this attachment. However, incubation in the presence of high light intensity results in a decreased growth yield (Hirsch and Conti, 196413; M. M. Attwood, unpublished observations). Indeed, growth inhibition by light was suggested by Kingma Boltjes in 1936 on the basis of findings made with agar slide cultures. Preliminary action spectra have indicated that the effect may be associated with two areas in the visible spectrum, one around 420 nm and the second at approximately 568 nm (Hirsch, 1974a). However, as yet, no mechanism has been postulated to explain this light inhibitory effect. D.

LIFE CYCLES A N D

PLEOMORPHISM

I t has been pointed out that Hyphomicrobium spp. were originally recognized on the basis of their distinct morphology (Section I, p. 304) and later seen to reproduce by an unusual method, namely through a process of bud formation (Kingma Boltjes, 1936). Consequently,

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Hyphomicrobium spp. have been included in the group of budding bacteria of which different forms are now recognized (Hirsch, 197413; Section 11-A, p. 305). The process of reproduction by budding has been assumed to be fundamentally different from binary fission seen in other bacteria. Whittenbury and Dow (1977) pointed out that this assumption stems from a comparison of the cell cycles of “rod-shaped’’ bacteriasuch as Escherichia coli and budding bacteria. In the latter organisms, cell separation usually results in two asymmetric cells (one with a filament and the other without) whereas, in E. coli, cell division usually results in two siblings. These authors suggest that the division process which is binary in both cases is fundamentally similar. The apparent difference lies in the fact that, in budding bacteria, wall growth is always polar whatever the growth environment whereas in E. coli and other “rodshaped” bacteria, growth rate affects the mode of wall growth. When these bacteria are growing at high rates, growth occurs from multiple points along the envelope, whilst at low growth rates (in minimal medium with a generation time of 60 min or longer) growth occurs polarly (Donachie and Begg, 1970). In this latter case, the only difference between true budding bacteria and E . coli is that, in the former, division is asymmetric (see Fig. 2 ) . The obligate nature of this asymmetric division in Hyphomicrobium spp. results in f o k a t i o n of two different cell types, namely a mature mother cell and an immature daughter cell. These two cell types will differ in their consequent growth cycles. Studies of the growth cycle of Hyphomicrobium spp. have been made by a number of authors (Kingma Boltjes, 1936; Zavarzin, 1960; Leifson, 1964; Moore and Hirsch, 1973a; C. S. Dow and A. Lawrence, unpublished observations). During the reproductive cycle of hyphomicrobia (see Fig. 31, a swelling at the tip of the filament (or hypha) begins the process of bud formation. The bud continues to increase in size and, at some point before its completion, develops one to three polar or subpolar flagellum(ae1.A septum is then formed in the hypha just below the bud and vigorous movements of the flagellum(ae1 eventually cause the bud to become detached from the h ha of its mother cell. The liberated bud, still immature, is now yp referred to as a swarmer. During its maturation, the swarmer becomes non-motile and grows in size to reach the final mature size of a mother cell. The so formed “daughter” cell then produces a hypha from the pole of the cell opposite to where it was formerly attached to the mother cell and o n this new hypha a bud may form (see Fig. 1, p. 307).

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..

321

..

(b)

(a)

(C)

FIG. 2. Diagrammatic representation of (a)growth-fonn multiple points in Escherichiu coli (division time less than 4.0 m i d ; (b) polar growth in Escherichiu coli (division time greater than 60 min); (c) obligate polar growth and asymmetric division in a Hyphomicrobiumsp. After Whittenbury and Dow (197 7).

A detailed study of this rather elaborate life cycle was made by Moore and Hirsch (1973a). An approximately synchronous culture of Hyphomicrobium B522 was obtained by isolating swanner cells from random cultures using procedures based on centrifugation and filtration. In the heterogeneous swanner population thus obtained,

0 Daughter cycle

Mother cycle

FIG. 3. Diagrammatic representation of the life cycle of hyphomicrobia showing mother and daughter cycles. After Leifson (1964).

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hyphal outgrowths were initiated between 0 and 10 h indicating that the cells in the initial swarmer population were at different stages of maturation and that this maturation process may last for up to 10 h. The period of hyphal development required 3.5 to 4.5 h, whilst bud formation and separation required an additional 5.5 h. Therefore, under the experimental conditions used, the maximum mother generation time would be 20 h and the minimum 9 h. Moore and Hirsch (1973a) pointed out that their study examined only the events leading to the production of the first daughter cell, and that evidence had been obtained in an earlier study which showed that the second and later generations of progeny were produced more rapidly. An average mother generation time, calculated from these data, was 14.5 h; this value is the same as that found on agar slide cultures (Hirsch and Jones, 1968). Of this period, 28 to 39% is used for hyphal growth, 33 to 38% for bud formation and detachment, and the remaining 28 to 34% for swarmer maturation. Evidence obtained from staining with primuline showed that, after formation of the first swarmer cell, the hypha of the mother cell recommences growth but the growth area is now at the tip of the hypha rather than at the initial growth point at its base (Hirsch, 1974a). The new increment of hyphal length is approximately one half to one third of the previous one; each consecutive daughter-cell formation is followed by less hyphal growth until a finite hyphal length is reached which is constant for that given set of growth conditions. In slide cultures, a new bud is formed every 5 to 6 h, and up to eight daughter cells have been observed during the active period of mother-cell reproduction (Kingma Boltjes, 1936; Mevius, 1953).When the growth conditions are still favourable, a new budding hypha is formed from the cell’s other pole. Still older cells may have hyphal side branches which are capable of bud formation. I t is important to note, however, that, as with Rhodomicrobium spp. (Whittenbury and Dow, 19771, in Hyphomicrobium spp. at any one time bud formation occurs only at one hyphal tip (Hirsch, 1974a). It has been suggested that the shorter intervals of hyphal growth during the formation of second and subsequent daughter cells may be the cause of the shorter cell doubling times recorded during subsequent “daughter generations” (Weiner and Blackman, 1973; Hirsch, 1974a). Replication of D N A during the life cycle of Hyphomicrobium spp. has been studied by Zavarzin (1960) and Moore and Hirsch (1973b).The latter authors have studied the nuclear apparatus of Hyphomicrobium B522 both in cells of various ages and in synchronized cultures. The

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results obtained indicated that the young swarmer cell contained a single nucleoid which consisted of a DNA molecule of molecular weight 3.1 x lo9 daltons with multiple membrane-attachment sites. Replication of DNA occurred during swarmer maturation, hyphal development and the initial stages of bud formation. However, separation of the two daughter nucleoids did not take place until after initiation of new bud development. The experimental results indicated that, within a short time interval, transfer of one of the nucleoids from the mother cell into the bud occurred. Such a rapid transfer of DNA is a rather unusual process, particularly when it is realized that it may involve transfer of DNA over a considerable distance. The mechanism used for transportation of the DNA is uncertain, but the authors suggested thal protoplasmic streaming might provide a reasonable explanation. I t is of interest to note that, in this study, it was found that the new genome arrived in the developing bud together with stable mother-cell RNA. The bud immediately commenced DNA and RNA synthesis, and then the formation o f a septum was observed. The above results are at variance with those reported for a rosette-forming Hyphomicrobium sp. (Zavarzin, 1960) in which DNA was found in the buds from the first moment they became visible. When considering the two conflicting sets of data, Hirsch (1974a) concluded that more work with synchronous populations of hyphomicrobia was necessary before a more conclusive picture-can be put forward. Recently, Whittenbury and Dow (1977) have proposed a model for Rhodomicrobium Rm5 which is compatible with Zavarzin’s observation. These authors suggest that DNA replication is initiated in the developing bud immediately prior to stalk formation, and that the DNA is attached to the cytoplasmic membrane. During stalk formation, the DNA membrane-attachment point, which is located in close proximity to the tip of the developing stalk, moves inside the stalk thus pulling the DNA along. The model further suggests that the membrane-attachment point appears in the developing bud as a n initial event, and thus is consistent with the fact that DNA is present in the bud from the first moment it becomes visible. Since Rhodomicrobium Rm5, under certain growth conditions, exhibits a simplified life cycle analogous to that of’hyphomicrobia (A. France and C. S . DOW,unpublished observations), this model may also apply to the latter group oforganisms. Various authors have noted a n effect of environmental conditions on both the morphology of Hyphomicrobium spp. and their life cycle (see Section 11-E, p. 326). Kingma Boltjes (1936)in his investigations on the

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'.

/

In the presence of I

formation

anol

In the presence of methylamine

'b\ 1

\

&

Microcolony formation

Complex matrix formation

FIG. 4. Polymorphic variation in hyphomicrobia in response to different carbon sources (methanol or methylamine) or ionic environment (high concentrations of Fez+, Mn2+,NO,). After A. Lawrence and C. S . Dow (unpublished observations).

life cycle of Hyphomicrobium vulgare observed in partly dry slide culture preparations, where the daughter cells were unable to swim away from the hypha, that bud formation without prior formation of a hypha frequently occurred. Also under these conditions new cells were occasionally formed from the stalk half-way between the mother and daughter cell, again without prior formation of a branch on the existing hypha. These processes led to the formation of cell aggregates of a variable and complex nature. Following the observation of Y shaped cells in older cultures of methanol-grown Hyphomicrobium C (M. M . Attwood 'and W. Harder, unpublished observations), the polymorphic variation in this and similar strains of Hyphomicrobium as a response to environmental conditions was investigated (Whittenbury and Dow, 1977). When methylamine was substituted for methanol as

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FIG. 5. Electron micrographs of: (a)“mother” and developing “daughter” cell; (b)Y cell formation in 0.2% (w/v) methylamine hydrochloride; (c) a released Y cell; (d) dichotomous branching characteristic of Y cells; (e) “normal” filament and “daughter” cell formation from a Y cell; (0 branch formation in medium containing 2% (w/v) potassium nitrate; (g) chain formation in medium containing 2% (w/v) potassium nitrate. Organisms were grown in the mineral salts medium of Attwood and Harder (1972), and shadowed with gold-palladium.

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the source of carbon and energy for growth, Y-shaped daughter cells were produced (see Fig. 4). These cells developed into complex structures which may form a normally shaped daughter cell whilst the mother cell continued the branching process. On resupplying methanol to the cultures, the Y -shaped cells produced filaments of “normal” dimensions which subsequently gave rise to normal motile swarmer cells. In addition, high concentrations of Fez+ or Mn2+ ions in the medium or excess of K N O , (2%) caused production of microcolonies (Fig. 5 ) similar to those observed with Rhodomicrobium (A. Lawrence and C. S. DOW,unpublished observations). Recently an interesting observation has been made with Rhodomicrobium Rm5 (A. France and C. S. Dow, unpublished observations); under conditions of high carbon dioxide concentrations, this organism does not produce microcolonies but shows a life cycle similar .to Hyphomicrobium spp. These observations indicate that polar growth and bud formation in species of Hyphomicrobiumand Rhodomicrobium are basically similar processes. E.

EFFECT O F ENVIRONMENT O N THE MORPHOLOGY

It has been indicated already that branching of the hyphal filaments in hyphomicrobia may occur under certain environmental conditions (see Section 11-D, p. 324). Many workers have observed that, with these organisms, cell dimensions, the length of hyphal filaments, the degree of hyphal branching, incidence of bud formation and rosette formation are dependent on environmental conditions (cf. Hirsch, 1974a). Rapid growth under favourable conditions resulted in formation of large cells with short hyphae and only a few hyphal branches. Growth at very low rates in mineral media without added

TABLE 2. Relationships between mother-cell size, mean h p h a l length and growth rate in Hyphomicrobium X grown in a methanol-limited continuous culture Growth rate (h-’) 0.02 0.05 0.11

Mean dimensions of mother cell (pm) 0.7 x 0.4 0.9 x 0.6 1.0 x 0 . G

Mean hyphal length (pm) 3.9

2.6 1.2

T h e organisms were grown in the mineral salts medium of Attwood and Hartlci- ( 1072)

pH 7.2 and 30%.

at

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carbon or nitrogen sources or in media with added methylamine but lacking phosphate gave small cells with hyphae over 100 pm long (Zavarzin, 1960; Hirsch and Conti, 1964a). Furthermore, cells from changing environments showed bipolar hyphal outgrowths, whereas those grown under more stable conditions had one hypha only (Hirsch, 1974a). These observations suggest a possible relationship between growth rate of Hyphomicrobium spp, and the average length of the hyphal filaments of the mother cells. A further indication of the effect of growth rate on hyphal length was obtained during growth of’ Hyphomicrobium X in a methanol-limited chemostat where growth rate is controlled by the concentration of the limiting factor (W. Harder, unpublished observations). The organism was grown at three different rates and, when a steady state was obtained, photomicrographs were taken of the cells and using these the hyphal length measured. Although it was found that the length of the hyphae varied by approximately a factor of two at any growth rate, the mean hyphal length calculated from 200 measurements at every growth rate was inversely proportional to growth rate (Table 2 ) . This suggests that the length of the hypha is related to the concentration of nutrients in the medium. The data also indicate that the average size of the mother cell increased with increasing growth rate. This phenomenon has been observed frequently with various bacteria (Herbert, 196 1). Aggregation of mother cells to form rosette colonies (Fig. 6) has been observed with a number of strains (Conti and Hirsch, 1965; Kingma Boltjes, 1936; Zavarzin, 1960). Some Hyphomicrobium isolates when growing under optimal conditions characteristically form these colonies (Zavarzin, ISSO), whilst other isolates only produce such growth forms when growing under suboptimal growth conditions with respect to, for example, pH value or temperature (Attwood and Harder, 1972). This may suggest both a genotypic and phenotypic variability amongst the Hyphomicrobium isolates which may be related to the ability of the young swanner cells to produce monopolar excretions of sticky “hold-fast” material (Conti and Hirsch, 1965). The function of the hyphal filaments as an obligate cell organelle of‘ Hyphomicrobium spp. is completely obscure. Since it is involved in the reproduction process, attempts have been made to relate the requirement for a hypha with certain aspects of the cellular events which accompany bud formation (Hirsch, 1974a). For instance, it has been suggested that the function of the hyphal filament could be to provide a minimal distance between the mother nucleoid and that of’ the

W. HARDER AND MARGARET

M. A r r W O O D

FIG. 6. Rosette formation by HyjDhornicrobiurn ZV 6 2 2 growing in liquid culture with methylamine as the carbon and energy source. The photographs a-d show different stages of complexity of the rosettes. The magnification of the phase-contrast micrograph is x3500. Reproduced by permission of M. Veenhuis.

growing but not yet separated bud (Moore and Hirsch, 19731s). However, the significance of the obligate nature of the hyphae in the reproduction process is far from clear. Furthermore, it could be

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suggested that the hyphae play a role similar to that in other prosthecate bacteria such as species of Ancalomicrobium and Prosthecomicrobium. In these organisms, the prosthecae are not involved in the reproductive process but are considered to represent a mechanism to increase the area of the cell surface (Pate and Ordal, 1965; Staley, 1968). Recently, prosthecae have been isolated from Asticcacaulis biprosthecurn and active transport of amino acids and glucose in these organelles has been demonstrated (Porter and Pate, 1975; Larson and Pate, 1976). These findings support the suggestion that prosthecae, in this and similar organisms, do indeed represent specialized organelles which provide increased surface area for transport of nutrients from dilute environments. In addition, this suggestion provides an explanation for the finding that, similar to hyphomicrobia, the length of the prosthecae of A. biprosthecurn increases when the concentration of nutrients in the environment decreases (Larson and Pate, 1976). In view of the relationship between growth rates and hyphal length as described earlier, it seems reasonable to postulate a similar function for the prosthecae of hyphomicrobia. However, this remains to be confirmed. F.

ECOLOGY

Little is known about the ecology of hyphomicrobia. Not only is there an almost complete lack of quantitative data on the distribution of these organisms in various ecosystems, but also their natural habitat and possible role in the various nutrient cycles in Nature are completely unknown. Several observations made in the laboratory on enrichment and isolation of these organisms, and their unusual morphology, physiology and biochemistry, may give an insight into their role in Nature and, at present, this is the only method available to evaluate the ecological position of these organisms. Current information on the distribution of these organisms indicates that they are ubiquitous. They occur in various types of soil and soil suspensions, lakes, brooks, springs, acid mine-drainage water and water-distribution pipes, and also have been observed frequently in aquarium water and laboratory water baths (Hirsch, 1974a). In addition to their occurrence in these nutritionally rather poor environments, they have also been found in eutrophic nutrient conditions such as sewage sludge o r polluted streams (Kingma Boltjes, 1936 ; Attwood and Harder, 197 2). Furthermore, there are several

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accounts of the isolation of hyphomicrobia from the marine environment (Attwood and Harder, 197 2 ; Hirsch, 1974a). Hyphomicrobia are frequently found attached to submerged slides, other micro-organisms and marine and freshwater algae, where they are associated with slime layers (Geitler, 1965). In view of the rather restricted range of energy sources which hyphomicrobia are able to use for growth (Section II-C, p. 315), this wide distribution is at first sight rather unexpected. However, when it is realized that one-carbon compounds like methanol, methylated amines and formic acid are widely distributed in Nature (Quayle, 197 21, the ubiquitous occurrence of hyphomicrobia, which have become specialized to grow on these compounds, may be appreciated. Also, the frequent observation of oligocarbophilic growth in mineral media may be explained on the basis of diffusion of one-carbon compounds from the laboratory atmosphere into the liquid media in which these organisms thrive (Kingma Boltjes, 1936). The observation that Hyphomicrobium spp. occur in enrichment cultures for methane oxidizers (R. Whittenbury, unpublished observations; Hazeu and Steenis, 1970; Adamse et al., 1972; Wilkinson and Hamer, 197 2 ;Jannasch, 197 5) is not surprising and may even indicate that, in Nature, these organisms are closely associated. The possible form of such an association has been studied by Wilkinson et al. ( 1974). These authors investigated the behaviour of a stable mixed microbial population grown on methane in continuous culture, and found that the population consisted of a methane oxidizer, a Hyphomicrobium sp. and two other chemo-organotrophs, namely an Acinetobacter sp. and a Flavobacterium sp. The latter two organisms were found to grow on complex organic compounds, whilst the Hyphomicrobium sp. was shown to be growing on the small amount of methanol excreted by the methane oxidizer. If methanol was added to this mixed culture, then growth of the methane oxidizer was inhibited whereas the percentage of the population accounted for by the Hyphomicrobium sp. increased. This suggested that the role of the Hyphomicrobium sp. in the mixed culture was to remove the inhibitory methanol in order that the methane oxidizer could grow. Consistent with this idea was the fact that the methane oxidizer alone could not be grown in liquid culture for any length of time. For continued growth of the organism in such cultures, the Hyphomicrobium sp. was required. Thus it can be considered that this may be the reason why hyphomicrobia are seen to occur consistently in enrichment cultures for methane oxidizers.

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However, a large number of other bacteria are known to utilize methanol (Quayle, 1972) and, at least in theory, may be expected to form such stable mixtures with methane-oxidizing bacteria. Yet such interactions have not been reported. This may be due to the fact that rod-shaped methanol utilizers are not easily recognized as contaminants in liquid cultures of rod-shaped methane oxidizers. O n the other hand, it may be that hyphomicrobia have a selective advantage over rod-shaped methanol utilizers particularly at very low concentrations of methanol (Wilkinson and Hamer, 1972) and therefore are selected in the enrichment cultures. If this is true, then one would expect hyphomicrobia to have an affinity for methanol greater than that of other methanol utilizers. The data available in the literature suggest that this might be correct. The K, value for methanol of Hyphomicrobium sp. in the mixed culture of Wilkinson et al. (1974)was found to be 8 x loe6 M (Wilkinson and Harrison, 1973) whereas, the K, value for Pseudomonas extorquens was found to be 2 x 10” M (Harrison, 1973). However, there is some doubt as to the validity of this explanation. Continuous selection experiments as suggested by Veldkamp and Jannasch (1972), using very low dilution rates with methanol as the limiting carbon source, invariably resulted in the selection of a Spirillum sp. (W. Harder and M. M. Attwood, unpublished observations). This indicates that Hyphomicrobium spp. do not show consistently a greater affinity for methanol than other organisms. Therefore, the situation on this point is not clear, and we must await further experimentation before it will be possible to construct a coherent picture. Many Hyphomicrobium isolates have been reported to be able to grow anaerobically in the presence of nitrate. Indeed, a specific enrichment method has been developed based on the capacity to grow on methanol and nitrate (Sperl and Hoare, 197 1 ; Attwood and Harder, 1972)and the fact that this method was repeatedly found to be highly specific for Hyphomicrobium spp. tends to suggest that no other onecarbon utilizer is able to grow under these conditions. This ability of‘ hyphomicrobia to denitrify with methanol as the electron donor may have a bearing on their role in Nature, since methanol may be produced under anaerobic conditions from compounds containing methyl esters or ethers such as pectin (Quayle, 1972). In the presence of nitrate, the methanol thus produced could be utilized by hyphomicrobia. Not only may hyphomicrobia occur in Nature when nitrate is present, but these organisms may also be expected in comparable

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environments. The increase used of nitrate fertilizers and consequent increase in the nitrate concentrations in freshwater poses a pollution problem, and several engineering groups have studied methods for the economical removal of nitrates from waste-water streams (St. Amant and McCarty, 1969; Dijkstra, 1974). Methanol was shown-to be amongst the most economical carbon sources effective in supporting denitrification. Indeed, hyphomicrobia have been found in large numbers in the denitrification tank of a pilot system operating at The Dutch State Mines, Geleen, The Netherlands (W. Harder, unpublished observations). Finally, a recent electron microscopic survey of the bacterial populations of several oligotrophic freshwater environments in England has shown that Hyphomicrobium spp. may constitute up to 25% of such populations (C. S . DOW, unpublished observations). This percentage was related to the nutrient status of the system, and showed an inverse proportionality in that the percentage of the population related to Hyphomicrobium spp. fell as the nutrient concentration in the environment increased. It would be of interest to see whether growth of hyphomicrobia in such systems is mainly due to the diffusion of one-carbon compounds from the atmosphere or at the expense of the methanol excreted by methane oxidizers which are also present in such environments. 111. Biochemistry of Hyphomicrobia A.

BIOCHEMISTRY

O F GROWTH

O N REDUCED

ONE-CARBON

COMPOUNDS

The main requirements for any micro-organism growing on a reduced one-carbon compound are the ability to generate energy and reducing power from the substrate and to synthesize cell constituents. The latter function invblves a unique biosynthetic problem since carbon-carbon bonds have to be formed in order that the one-carbon substrate can be converted into a three-carbon skeleton from which some central intermediary metabolite such as pyruvate or phosphoenolpyruvate can be derived. Once such a compound has been synthesized, there is no reason to suspect that the pathways leading to the main classes of cell constituents will be different from those of other organisms.

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LH,

CHI

----------+

CH,OH

HCHO ?

HCOOH

N-methyl compounds (e.g. trimethylamine, dimethylamine, methylamine) FIG. 7. Central pathway for oxidation of reduced one-carbon compounds by hyphomicrobia. El indicates methanol dehydrogenase, E, formaldehyde dehydrogenase, and E, formate dehydrogenase. X, Y and Z represent the electron acceptors for the enzymes involved.

1. Oxidation of One-Carbon Compounds

Energy for growth arises by oxidation of the substrate to carbon dioxide and water. Little is known about the mechanism of these reactions but it is considered that there is a central pathway which consists of three successive oxidations and which leads to the dissimilation of reduced one-carbon compounds via formaldehyde and formate to carbon dioxide (Fig. 7). Each oxidation is accompanied by the transfer of two electrons from the substrate to a prosthetic group or coenzyme. The nature of these electron acceptors is not known for all of the enzymes. For further information on energy production in methylotrophs, see the following reviews: Quayle, 1972; Anthony, 1975a; and Van Dijken and H>rder, 1975. Three different enzymes have been cited for bacterial oxidation of methanol. Initially it was suggested that the reaction was catalysed by an NAD+-linked methanol dehydrogenase (Kaneda and Roxbrough, 1959). After the failure of other workers to detect this enzyme, Harrington and Kallio (1960) suggested that catalase functioning as a peroxidase could oxidize methanol; although methanol can be oxidized by hydrogen peroxide in the presence of catalase (Keilin and Hartree, 19451, this appears to have little relevance to methanol oxidation in vivo. The most detailed studies so far on the enzymology of methanol oxidation were made by Anthony and Zatman ( 1964a, b, 1965, 1967a, b). In all methanol- or methane-utilizing bacteria studied so far, except for an actinomycete (Kato et al., 1975), oxidation of methanol is catalysed by an enzyme which requires ammonium ions or methylamine for activation and which causes dehydrogenation of methanol in the presence of phenazine methosulphate at an optimum

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rate around pH 9.0 (Anthony and Zatman, 1963; Ladner and Zatman, 1969; Patel and Hoare, 1971; Patel et al., 1972; Mehta, 1973; Goldberg, 1976). This enzyme has been shown to operate in Hyphomicrobium spp. A four-step purification procedure resulted in a homogeneous preparation, as judged by Sephadex G-200 molecular filtration, of methanol dehydrogenase from Hyphomicrobium WC (Sperl et al., 1974).The enzyme was shown to require in uitro ammonium ions and phenazine methosulphate for activity and to possess a pH optimum, a molecular weight and absorption spectrum characteristic of methanol dehydrogenases purified from other organisms. I t was unable to catalyse the transfer of electrons to NAD' and was shown to contain the same fluorescent prosthetic group as observed for the methanol dehydrogenase from Pseudomonas AM1. I t had the same broad specificity with normal primary alcohols and could catalyse the oxidation of formaldehyde. N o activity was detected with secondary and tertiary alcohols. Similarly, when the pathway of oxidation of methanol was investigated using cell-free extracts of Hyphomicrobium X and G , an ammonium- activated phenazine me thosulp ha te-linked methanol dehydrogenase was detected (Harder and Attwood, 1975). The enzyme had a dual specificity for normal primary alcohols and formaldehyde. The pH optimum for activity was 9.5. Antigenic relationships between the methanol dehydrogenase from o ne-carbon- utilizing organisms have been used to group certain methane- and methanol-oxidizing bacteria (Patel et al., 1973; Patel and Felix, 1976). The methanol dehydrogenase from cell extracts of Hyphomicrobium B522 showed complete serological identity with the enzymes from two budding methane-oxidizing bacteria of the Methylosinus group. Recently, evidence has been obtained which suggests that the enzyme from Pseudomonas AM1 can transfer electrons from methanol to an electron- transport chain through a carbon monoxide-binding cytochrome c (Anthony, 197513).A similar carbon monoxide-binding cytochrome has been detected in Hyphomicrobium spp. including HyphomicrobiumX (Tonge et al., 1974; Widdowson and Anthony, 1975). At the present time, the situation associated with the oxidation of formaldehyde is under review; at least four different enzymes have been shown to catalyse an oxidation of formaldehyde to formate in one carbon-utilizing organisms. This oxidation step is common to organisms growing on many compounds since formaldehyde is not

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

335

only the oxidation product of methanol (and hence methane) but is an intermediary metabolite in bacterial oxidation of methyl groups from methylated amines and trimethylamine sulphur compounds. I t seems probable that, in many organisms, formaldehyde is oxidized by methanol dehydrogenase as previously discussed (Pate1 and Hoare, 197 1 ; Quayle, 1972; Mehta, 1973). However, both an NAD+-linked glutathione-dependent (Harrington and Kallio, 1960; Cox and Quayle, 1975) and an NAD+-linked glutathione-independent (Kung and Wagner, 1970; Colby and Zatman, 1973)formaldehyde dehydrogenase have been reported. Other organisms appear to contain a general aldehyde dehydrogenase which oxidizes formaldehyde in vitro via an ammonium ion-independent dichlorophenolindophenollinked reaction (Johnson and Quayle, 1964). Recently Mehta (1975) reported a similar enzyme in methylamine-grown Pseudomonas RJ 1. This enzyme was less stable and did not exhibit the same broad specificity for aldehydes. Studies with cell-free extracts and purified enzymes have shown that methanol-grown Hyphomicrobium X and G (Harder and Attwood, 1975) and Hyphomicrobium WC (Sperl et al., 1974) oxidize formaldehyde via a reaction catalysed by methanol dehydrogenase. However cell-free extracts of methylamine-grown Hyphomicrobium X appear to contain, in addition to methanol dehydrogenase, another formaldehyde-oxidizing enzyme which is very similar to the general aldehyde dehydrogenase reported by Johnson and Quayle ( 1964). The enzyme is ammonium ion-independent, has a pH value for maximum activity around pH 7.5 and shows a general specificity for aldehydes (M. M. Attwood, unpublished observations). In the majority of one-carbon utilizing organisms studied so far, this central oxidation pathway is completed by an NAD+-dependent formate dehydrogenase (Kaneda and Roxbrough, 1959 ; Johnson and Quayle, 1964; Kung and Wagner, 1970; Mehta, 1973).Johnson and Quayle ( 1964) purified formate dehydrogenase three-fold from methanol-grown Pseudomonas AM 1, and showed the enzyme to be unstable, specific for formate and inhibited by low concentrations of cyanide, ferrous and cupric ions. Studies with cell-free extracts of methanol-grown Hyphom’crobium X and G led to the detection of a similar NAD+-dependent formate dehydrogenase. Furthermore, the enzyme showed the characteristic inhibition pattern associated with other formate dehydrogenases. Enzyme activity was partly inhibited by 2.0 mM iodoacetate, 0.1 mM copper sulphate, 0.1 mM hypophosphite

336

W. HARDER AND MARGARET M. A W O O D

and 50 mM EDTA (Harder and Attwood, 1975). A similar nicotinamide nucleotide- linked formate dehydrogenase has been reported in formate-grown Hyphomicrobium B522 (Hirsch, 1974). Several bacteria are known to gfoG aerobically with trimethylamine, dimethylamine or methylamine as the sole source of carbon and energy, and the pathways for growth on these compounds have been elucidated (Fig. 8). All of the pathways proposed result in the production of formaldehyde and therefore enter the central pathway (Fig. 7 , p. 3 3 3 ) at the level of formaldehyde. Two different pathways have been proposed for aerobic degradation of trimethylamine to dimethylamine (Large, 197 1; Large et al., 1972; Colby and Zatman, 1973). The first involves a trimethylamine mono-oxygenase to convert trimethylamine to trimethylamine N-oxide which is then demethylated to dimethylamine; the second route involves direct conversion of trimethylamine into dimethylamine by action of trimethylamine dehydrogenase. In both reactions, the dimethylamine produced is oxidized to methylamine by a dimethylamine mono-oxygenase. This enzyme is also present when the bacteria are growing aerobically on dimethylamine as the carbon and inergy source (Fig. 8). Hyphomicrobium spp. are able to grow both aerobically and anaerobically on trimethylamine and dimethylamine in the presence of nitrate. During anaerobic growth of Hyphomicrobium X on trimethylamine in batch culture, accumulation and subsequent utilization of dimethylamine and methylamine were observed, whilst aerobic growth o n trimethylamine led only to dimethylamine accumulation and

t,

\

Trimethylaminr

Fornialdchyde

Trirnethylaniine N-oxide

b2.b

Formaldehyde

Dimethy lamine Formaldehyde Methylamine

FIG. 8. Aerobic degradation of' trirnethylamine to methylamine. After Colby and Zatinan ( 1973). E, indicates trimethylamine mono-oxygenase, E, trirnethylamine dehydrogenase, E, trimethylamine N-oxide demethylase, and E, dimethylamine mono-oxygenase.

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

Trimethylamine

Dimethylamine

Formaldehyde

5

337

Methylamine

Formaldehyde

FIG. 9. Aerobic and anaerobic degradation of trimethylamine and dimethylamine in Hyphomicrobium X (Meiberg and Harder, 1976). E, indicates trimethylamine dehydro-

genase and E, dimethylamine dehydrogenase.

utilization. These results suggested that these compounds were intermediates in the degradation of trimethylamine by Hyphomicrobium X (Meiberg and Harder, 1976). Enzymic analysis of crude cell-free extracts showed that both aerobically- and anaerobically-grown Hyphomicrobium X contained an active trimethylamine dehydrogenase and that this was the only enzyme which was capable of catalysing oxidation of the trimethylamine present. Oxidation of dimethylamine was also mediated by a dehydrogenase which required phenazine methosulphate and dichlorophenolindophenol for activity. Under no conditions could an active dimethylamine mono-oxygenase be detected. Since it could be postulated that oxidation of dimethylamine was due to a dual specificity of the trimethylamine dehydrogenase, the dimethylamine dehydrogenase activity was partially purified ( 13-fold) by ammonium sulphate fractionation between 50 and 70% saturation and DEAEcellulose chromatography. The results showed that, in Hyphomicrobium X, dimethylamine is oxidized by a novel enzyme, namely a dimethylamine dehydrogenase (Fig. 9 ; Meiberg and Harder, 1976).

Methylamine

I

E,

Formaldehyde

y- Glutamylmethylamide \ l l

Ammonia /--Glutamatev

NH,

N-Methylglutarnate FIG. 10. Direct and indirect pathways for oxidation of methylamine. E, indicates methylamine dehydrogenase, E, N-methylglutamate synthase and E, N methylglutamate dehydrogenase.

338

W. HARDER AND MARGARET M. ATWOOD

Evidence from other organisms shows that methylamine can be oxidized either directly to formaldehyde via a methylamine-oxidizing enzyme (Fig. 10, I), as in methylamine-grown Pseudomonas AM 1 (Eady and Large, 1968), or indirectly through the formation of N-methylglutamate (Fig. 10, 11) as demonstrated with methylamine-grown Pseudomonas MA (Bellion and Hersh, 1972). The pathway of methylamine oxidation has been studied in Hyphomicrobium vulgare ZV and vulgare 3 by a group of Russian workers. These report the N methylation of glutamate to form initially y-glutamylmethylamide and then N-methylglutamate (Trotsenko et al., 1974; Logincrva et al., 1976; Fig. 10, 111). Preliminary evidence using cell-free extracts of Hyphomicrobium X demonstrated the lack of a direct methylamineoxidizing systein and suggested the presence of the indirect pathway as depicted in Fig. 10, 111 (J. B. M. Meiberg, unpublished observations). 2. Assimilation of One- Carbon Compounds Assimilation of one-carbon compounds, particularly methanol, has leceived much more attention (see reviews by Ribbons et al., 1970; Qwyle, 197 2 ; Anthony, 197 5a). Three pathways are known whereby net synthesis of a three-carbon skeleton from one-carbon compounds is accomplished. The first of these to be formulated was the ribulose diphosphate cycle of carbon dioxide fixation (Bassham et al., 1954). However this pathway is characteristically associated with autotrophic organisms, and the ability of micro-organisms to assimilate reduced one-carbon compounds via carbon dioxide does not appear to be widespread (Quayle and Keech, 1959; Stokes and Hoare, 1969; Quayle and Pfennig, 1975; Cox and Quayle, 1975). The second pathway, the ribulose monophosphate cycle of formaldehyde fixation, has recently been elucidated completely. This pathway is similar to the ribulose diphosphate pathway but results in fixation of formaldehyde rather than carbon dioxide (Kemp, 1964; Strsm et al., 1974). This pathway was originally considered to function only in organisms limited to growth on one-carbon compounds, but it has been shown to be the major carbon assimilation pathway in some organisms which are not restricted to one-carbon compounds for growth (Chalfan and Mateles, 1972; Coxand Zatman, 1974; Colby and Zatman, 1975b). The third pathway which was originally proposed by Large et al. (1961)is the serine pathway. The complete pathway associated with the id' (where id stands for isocitrate lyase) variant was initially documen-

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

339

ted by Bellion and Hersh (1972).The one-carbon units in this pathway are incorporated at the oxidation levels of both formaldehyde and carbon dioxide. Unlike the previous two pathways, there is no involvement of pentose phosphates; instead one-carbon units condense with two-carbon and three-carbon compounds (Hersh, 1975; Bellion and Woodson, 1975; Wagner and Levitch, 1975; and Fig. 1 1). There are four important features of the cycle. Firstly, two reduced one-carbon units at the oxidation level of formaldehyde condense with

2 Serine

Acetyl- CoA

2 Glycine

GI yoxylate

Citrate

2 GlyoxyIate

Isocitrate

/

FIG. 1 1. Pathway of one-carbon assimilation in methylamine-grown Pseudomonas MA. After Bellion and Hersh (1972).

two glycine molecules to form two molecules of serine. The serine is then converted through a series of reactions to phosphoenolpyruvate and, at this point, the third one-carbon unit required for synthesis of a three-carbon skeleton from one-carbon units is incorporated in the form of carbon dioxide to form oxaloacetate and finally malate. The third important feature is the activation of malate to maleyl-CoA which is then cleaved to glyoxylate and acetyl-CoA. The glyoxylate is converted to glycine and re-enters the cycle. The last feature is the regeneration of glyoxylate through oxidation of acetyl-CoA to isocitrate which is cleaved by isocitrate lyase to glyoxylate and succinate, the overall product for assimilation into cell constituents. I t is now clear that there is a group of organisms which utilize a pathway, identical in part with the id' serine pathway, but which lack

340

W. HARDER AND MARGARET M. A l l W O O D

isocitrate lyase activity. Some of these organisms (Pseudomonas AM 1) also lack malate thiokinase activity. This is known as the icl- variant of the pathway. The manner in which the methyl group of acetate is oxidized to glyoxylate is unknown (Quayle, 1975). Early isotopic studies with cells of Hyphomicrobium vulgare demonstrated that serine was an initial stable product of carbon assimilation (Naveke, 1957; Large et al., 1961; Doman et al., 1965). Incorporation of carbon from [14Cl-methanol by cultures of Hyphomicrobium vulgare (Mevius isolate) grown on methanol showed that the first stable products were serine, which accounted for 50% of the total count fixed, a four-carbon dicarboxylic acid such as malate or aspartate, and glycollate (Large et al., 196 1). Formation of glycollate has been assumed to reflect the presence of glyoxylate since glyoxylate itself would not be detected under the conditions of the chromatography employed (Heptinstall and Quayle, 1970).These results were taken to indicate that the serine pathway was being utilized. Recently, a comparative study of the specific activities of enzymes in cell-free extracts of methanol- and ethanol-grown

I 2,2-Phosphoglycerate

. ,

I

2 Serine

.1 Phosp hoenolpyruvate

residues Matate

2 Glycine

r x:;I;:

J . Acety-CoA

*

Oxaloacetate

1

Succinate

Citrate

.1

lsocitrate

FIG. 12. Pathway of one-carbon assimilation in methanol-grown Hyphornicrobium X as proposed by Harder et al. (1973).

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

341

Hyphomicrobium X and G indicated that, during growth on methanol, one-carbon units were incorporated in a pathway similar to that proposed by Bellion and Hersh (Harder et al., 1973).These results were in accordance with the report that Hyphornicrobiurn WC grown on methylamine contained an active malate lyase (CoA acetylating; Hersh and Bellion, 1972). The pathway proposed by Harder et al. (1973) differed from the previous proposal with respect to the overall product of the cycle, since attempts to detect the appropriate enzyme activities in cell-free extracts of Hyphornicrobium X and G for conversion of succinate into phosphoenolpyruvate failed. Therefore a slight variant on the pathway was proposed (Fig. 12) whereby succinate, instead of being withdrawn as the overall product of the cycle, was used to regenerate the carbon skeleton of oxaloacetic acid, the acceptor molecule for acetyl-CoA. This then permits withdrawal of a threecarbon intermediate, phosphoglycerate, as the net product of the cycle. The presence of hydroxypyruvate reductase in cell-free extracts has confirmed the use of the serine pathway in all Hyphomicrobiurn isolates tested, namely Hyphomicrobium B522 and G855 (Bellion and Spain, 1976) and Hyphornicrobiurn isolates C, 10097, ZV620, 3T + M and B522 (Attwood and Harder, 1977). A comparison of the rates of conversion of glycerate and 2 phosphoglycerate to phosphoenolpyruvate with the rate of conversion of 3-phosphoglycerate to 2-phosphoglycerate in cell-free extracts of methanol-grown Hyphomicrobium X and G resulted in the suggestion that 2-phosphoglycerate rather than 3-phosphoglycerate was the product of the glycerate kinase reaction in the serine pathway (Harder et al., 1973). Later work, using a glycerate kinase preparation purified from methanol-grown Hyphomicrobium X, confirmed P-phosphoglycerate as the product of the glycerate kinase reaction (Hill and Attwood, 1974). 2-Phosphoglycerate has been shown to be the product of the reaction in other one-carbon utilizers using the serine pathway (Hill and Attwood, 1974; Newaz and Hersh, 1975). After the original report that Hyphomicrobium X and G incorporate one-carbon units from methanol using the icl' serine pathway, Quayle (1975) calculated that the specific activity of isocitrate lyase should be 115 nmol min-' mg protein-' in order to support a doubling time of 5 h during growth on methanol o r methylamine using the icl' serine pathway. An examination of the specific activities recorded for Hyphomicrobium X and G grown on methanol (Harder et al., 1973) revealed that they were too low to support the growth rate of the

342

W. HARDER AND MARGARET M. A W O O D

culture. A re-investigation of the specific activities of isocitrate lyase in cell-free extracts of methanol-, methylamine- and ethanol-grown isolates of Hyphomicrobium was carried out. Three difftrent methods were used to measure the isocitrate lyase activity, and it was found that the initial enzyme activity recorded using the standard but nonspecific assay (Dixon and Kornberg, 1959) with methanol-grown Hyphomicrobium X and G was due to formation of‘ 2-oxoglutarate via isocitrate dehydrogenase rather than glyoxylate via isocitrate lyase. All of the isolates tested, including Hyphomicrobium B522, grown on each of the substrates which supported growth (i.e. methanol, methylamine and ethanol), failed to give measurable amounts of “true” isocitrate lyase activity (Attwood and Harder, 1977) and, therefore, hyphomicrobia are considered to assimilate one-carbon units from methanol and methylamine via the icl- variant of the serine pathway. These results agree with those of Y. A. Trotsenko using Hyphomicrobium vulgare sp. (personal communication) but are not in agreement with those reported by Bellion and Spain (1976).The latter authors, on the basis of results obtained with the non-specific assay, suggested that the level of activity of isocitrate lyase in extracts of Hyphomicrobium WC and B522 was high enough to indicate use of the icl’ serine pathway. Since data are not recorded on the growth rate of these organisms under the conditions used, and since the specific activities recorded for isocitrate lyase activity were low, it is difficult to comment on the enzyme activities with respect to the minimum value calculated by Quayle. The suggestion that 2-phosphoglycerate is withdrawn from the cycle for biosynthesis of cell material was substantiated when crude cell-free extracts from methanol-grown Hyphomicrobium X were shown to contain all of the enzymes necessary to convert 2-phosphoglycerate to fructose 6-phosphate by the reversal of glycolysis (Hill, 1976). Furthermore, it had been noted (Harder et al., 1973) that addition of 0.6 mM ATP caused an apparent 2.5-fold stimulation in the activity of phosphoglycerate mutase in cell-free extracts of methanol-grown Hyphomicrobium X and G.This stimulation was antagonized by AMP. In view of the probable position of this enzyme as the first enzyme in conversion of phosphoglycerate into cell material, it was suggested that the nucleotide effect could reflect adenylate-charge control on activity of the enzyme. If so, the mutase might play a regulatory role whereby flow of carbon can be channelled into carbohydrate synthesis when the energy status of the cell is high and wice versa.

m

g<

0

TABLE 3 . P r o p e r t i e s o t ' p h o s p h o g l y c e r a t e iiiutase purilicd tIum various

S O U I ' C C ~Ft-onl .

Hill (1976)

71

I

< PropelTy

Colactoi- drpetident pH Value optimum K, Value for 2-phosphoglycerate K, Value for 3-phosphoglycerate K, Value for 2,3-diphosphoglycerate

Wheat gem1

NU

8.9 1 to 5 x lo-' 3.3 x l o r 4

-

Rice germ

No 8.9 2 . 5 x lo-' 1.2 -

HyphomicrubiumX Y cs 1.3 9.3 x Ill-' 6.0 10-3 7.0 x lo-'

Pfeudomonus AM I

Yes 7.0 to 7 . G 3.2 x IO-' 3 . 4 IO-? 1 0 x IO-"

Escherithiu culi

Baker's ycas1

Rabbit skeletal

Cliickeii

IIluacIc

n1usclc

g D

bicabt

Yes

Y cs

Ycs

Yes

-

7.0

7 .(I

1.3

-

1 x 10-4

-

1to(ix10-' 1.13 x 10-4

2 x 10-s 5x10-3 1.25 x 10-1

fjx10-4 1.4 x lo-''

6

Molecular weight (daltons) Iaoclectric-point Polymer-ic-structure

30,000

LO

10

-

13.3

10.4

5.0

-

-

8.0 x lo-" 6.3

<

z

~

01-

Equilibriuiii cotistarit: 2-phosphoglycerate to 3-ptiosplioglycerate

?

0 0

I rn

I

v)

--I

m

< -

30,000

-

32,000 4.5 Possibly n1o11o111cr

32,000 4.2 Possibly 111o110111e1'

5ti,000 -

I 12,000 5.0 to 5.5 l'etraiiici

(~.+,ooo 5.3 Dinici-

65,(i90 -

-

I

4

I

0

5

n . I

n

!? D w

P W

344

W. HARDER AND MARGARET M. A W O O D

However, phosphoglycerate mutase has been purified and characterized from methanol-grown Hyphomicrobium X and Pseudomonas AM 1 (Hill and Attwood, 1976a) and detailed comparative studies failed to reveal any significant differences between the kinetic and molecular properties of these purified enzymes and other purified phosphoglycerate mutase preparations (Table 3). In particular, addition of nucleotides had no effect on the activities of the purified preparations. Further work using crude cell-free extracts, a partially purified phosphoglycerate mutase preparation and a purified phosphoglycerate mutase preparation from Hyphomicrobium X, demonstrated that stimulation of phosphoglycerate mutase activity in crude cell-free extracts measured in the presence of ATP is not a true property ofthe enzyme but the result of interference in the assay system by other enzymes, namely, 3-phosphoglycerate kinase arid glyceraldehyde 3-phosphate dehydrogenase. Once these enzymes were removed from the niutase preparation, no stimulation in activity occurred when ATP was added to the assay system (Hill and Attwood, 197613; Section 111-A-3,p. 345). The pathway of carbon assimilation during growth on formate has received little attention. Radio-isotope experiments using cells of forniate-grown Hyphomicrobium vulgare showed that the carbon fr-om [l4C1-carbon dioxide was incorporated into the carbon skeletons of amino acids from protein hydrolysates (Mevius, 1953; Naveke, 1957). Furthermore, in view of the rapid incorporation of [l4C1-carbon dioxide into aspartate, glutamate, glycine and serine, Mevius (1953) suggested that carboxylation reactions were important in the carbonassiniilation pathway. Further research is necessary before a more detailed account of the carbon-assimilation pathway can be reported but the information available is in accordance with operation of' the for-mate-assimilation pathway reported for Pseudomonas AM 1 (Large et al., 1961; Large and Quayle, 1963; Quayle, 1972). 3. Possible Regulation Sitesf o r the Serine Pathway When Hyphomicrobium spp. are grown on one-carbon compounds, formaldehyde is an intermediate in both the energy-generating arid the carbon-assimilating pathways (Fig. 13). Therefore, some control niust be exerted in order to channel, in relation to the requirements of' the cell, one-carbon units into one or other of these pathways. Initially, as a result of data obtained with crude cell-free extracts of Hyphomicrobium

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

I

Cellmaterial

345

J

t t. 1,3-Diphosphoglycerate

ADP ATP

3-Phosphoglycerate

Malate

\ Acetyl-CoA Methanol

-

Formaldehyde

r

/”

Methylamine

-

/’

Glyoxylate ccx c--------

Formate 7-q COP NAD+ NADH+H+

I

I Energygeneration I FIG. 13. Position of formaldehyde with respect to the energy-generating and carbonassimilating pathways in one-carbon-grown Hyphomicrobium spp. THF indicates tetrahydrofolate, ----+ indicates that the reactions are unknown.

X and G, it was suggested that this might occur by influence of adenylate charge on the enzyme phosphoglycerate mutase (Harder et al., 1973; Section 111-A-2, p. 342). Subsequent work showed that this effect was an artefact of the assay procedure used (see Section 11-A-2, p. 344).

346

W. HARDER AND MARGARET M. A W O O D

Moreover, when cell-free extracts of Hyphomicrobium X were analysed for the specific activities of other enzymes which had been implicated in control mechanisms in other organisms grown on one-carbon compounds, regulatory functions could not be detected. Phosphoenolpyruvate carboxylase activity in methylamine-grown Pseudomonas MA has been shown to be stimulated by NADH (Newaz and Hersh, 1975). As a result of this stimulation, together with the failure to detect any NAD+-dependent formate dehydrogenase activity, these workers postulated that, in this organism, the energy-generating pathway and the carbon-assimilation pathway utilized in part a common sequence of reactions, and that phosphoenolpyruvate carboxylase had a regulatory function. When the NADH level of the cell was high, phosphoenolpyruvate carboxylase activity was stimulated and the one-carbon units were channelled into carbon assimilation. In the absence of NADH, p hosphoenolpyruvate was converted into pyruvate and then, via the action of pyruvate dehydrogenase, the one-carbon units entered the TCA cycle as acetate and were channelled into energy generation. This pathway could not function in Hyphomicrobium spp. since they do not have a pyruvate dehydrogenase complex (see Sections 11-C, p. 318 and 111-D, p. 354). However, since phosphoenolpyruvate carboxylase is k n o m to be stimulated and inhibited by many compounds, the effect of such compounds on the enzyme in cell-free extracts of Hyphomicrobium X was investigated. I t was found that the enzyme belonged to class I11 (Utter and Kolenbrander, 1972)in that none of the compounds tested affected the rate of action of the enzyme (Hill, 1976). I t has been reported that, in Isolate XX, two isoenzymes of serine transhydroxymethylase are formed (O’Connor and Hanson, 1975). One isoenzyme is used for biosynthesis of glycine and one-carbon units when the organism is grown on succinate as the carbon and energy source, whilst synthesis of the other is induced during growth on the one-carbon compounds methane and methanol, and is involved in carbon assimilation using the serine pathway. Activity of the latter isoenzyme is stimulated by glyoxylate. Since this enzyme is associated with an initial step in the assimilation pathway, activation by glyoxylate could be considered to be a control mechanism. When the activity of serine transhydroxymethylase was measured in cell-free extracts of methanol- or ethanol-grown Hyphomicrobium X, there was no indication of the presence of two isoenzymes. The activity of’ the enzyme from either methanol- or ethanol-grown cells did not show

BIOLOGY, PHYSIOLOGY AND BIOCHEMISTRY OF HYPHOMICROBIA

347

any stimulation when glyoxylate was added to the assay system (Hill, 1976). Therefore, it seems unlikely that the carbon-assimilation pathway in one-carbon grown Hyphomicrobium X is regulated through the activity of either phosphoenolpyruvate carboxylase or serine transhydroxymethylase. This failure to detect regulation of any of these enzymes, together with the very limited range of compounds used by these organisms as a source of energy and the failure to observe increases in cell yield greater than 5% when exogenous carbon sources are supplied and taken up by the cells under methanol limitation in continuous culture (Section III-D, p. 3551, leads us to suggest that the carbon-assimilation pathway from methanol may be subjected to control by energy availability. I t is known that tetrahydrofolate will condense rapidly and nonenzymically with free formaldehyde to form the one-carbon-tetrahydrofolate complex used in the initial incorporation of one-carbon units into glycine to form serine (Quayle, 1972). Therefore, it can be postulated that only when all of the tetrahydrofolate in the cell is in the form of the one-carbon complex will formaldehyde be available for conversion into formate and used for energy generation (Fig. 13, p. 345). If the cell contains a high level ofATP, then the glycerate kinase in the one-carbon assimilation pathway can operate at an optimal rate. Furthermore, through action of 3 -phosphoglycerate kinase, carbon units in the form of P-phosphoglycerate can be drawn from the serine pathway via 3-phosphoglycerate and gluconeogenesis to cell constituents. When the ATP level in the cell falls, then activity of the two kinases will not be optimal and finally these enzymes will not be active. Under these conditions, the one-carbon assimilation pathway can be considered to be blocked at glycerate kinase and remains so until all of the tetrafolate of the cell is again in the form of the one-carbon tetrahydrofolate complex. Then energy generation can occur by the conversion of free formaldehyde to formate and oxidation of formate to carbon dioxide and water by an NAD+-dependent formate dehydrogenase (Section III-A- 1, p. 335). Once the ATP level in the cell rises above a minimum value, then the two kinase enzymes become active and carbon assimilation occurs. This scheme postulates that growth of Hyphomicrobium spp. on methanol is essentially energylimited. Further work is necessary in order to validate this regulatory mechanism. For instance, the above hypothesis assumes that formal-

348

W. HARDER AND MARGARET M. A T W O O D

(a)

0 Ethanol

1 F K H . H . NAD' Acetaldehyde

LKH +

NADH+H+ Acetate

Hi

Acetoacetate Succinyl- CoA Succinate Acetoacetyl CoA

-&+

o-3-hydroxybutyryl-CoA

1-CA .

FIG. 14. Pathways tor conversion of (a) ethanol, (bj acetate and (cj 3-hydroxybutyrate to acetyl-CoA in Hyphomicrobium X.

dehyde dehydrogenase(s1 of hyphomicrobia can only oxidize free formaldehyde and is (are) not able to oxidize methylene- tetrahydrofolate. I t is not known whether this assumption is correct. In spite. of these reservations, the above hypothesis seems to be attractive since all available information is in accordance with it. For

BIOLOGY, PHYSIOLOGY A N D BIOCHEMISTRY OF HYPHOMICROBIA

349

instance, using the proposed mechanism it is possible to understand the failure to increase cell yield above 5% by introduction of nonenergy-producing exogenous carbon sources in methanol-limited cultures since the growth of these organisms would be regulated by the availability of energy rather than carbon skeletons. B.

BIOCHEMISTRY OF G R O W T H

O N TWO-CARBON

COMPOUNDS

A comparative study of the specific activities of enzymes in cell-free extracts of Hyphomicrobium X and G grown on methanol, ethanol, acetate or 3-hydroxybutyrate has been made (Attwood and Harder, 1974). The results demonstrated that, .during growth on ethanol, synthesis of an NAD+-dependent ethanol dehydrogenase was induced. This was unexpected since the methanol dehydrogenase used to oxidize methanol and formaldehyde has been shown to oxidize ethanol in uitro at a rate 69 to 85% of that of methanol oxidation, and washed suspensions of ethanol-grown Hyphomicrobium spp. to oxidize methanol at the same rate as methanol-grown Hyphomicrobium spp. (Harder and Attwood, 1975). This induced NAD+-dependent ethanol dehydrogenase converts ethanol to acetaldehyde which is then either metabolized directly to acetyl-CoA or to acetate, and then activated by acetothiokinase to acetyl-CoA (Fig. 14a). This pathway is different from that found in ethanol-grown Pseudomonas AM 1 where the ethanol is converted to acetaldehyde by methanol dehydrogenase which interacts with the cytochrome chain at the level of cytochrome c (Taylor and Anthony, 1976).The acetaldehyde so formed was then oxidized to the level of free acetate by an NAD(P)+-independent aldehyde dehydrogenase rather than an NAD +-CoA-dependent acetaldehyde dehydrogenase. Activation of acetate by acetothiokinase also occurs in acetate-grown cells. Cell-free extracts of acetate-grown Hyphomicrobium spp. did not contain detectable amounts of acetokinase or phosphotransacetylase activity (Fig. 14b). Lastly, when cell-free extracts of ethanol- and 3-hydroxybutyrate grown cells were analysed for activity of enzymes associated with 3hydroxybutyrate oxidation, it was observed that 3-hydroxybutyrate was converted to acetyl-CoA by enzyme reactions already reported in other micro-organisms (Senior and Dawes, 1973; Oeding and Schlegel, 1973).3-Hydroxybutyrate is oxidized to acetoacetate which is

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then activated to acetoacetyl-CoA. Cleavage of acetoacetyl-CoA by pketothiolase results in production of acetyl-CoA (Fig. 14c). O n all pathways, energy and reducing power were shown to be generated from acetyl-CoA by oxidation in the TCA cycle. Since isocitrate lyase is not synthesized by Hyphomicrobium X or G , the route whereby cell constituents are made from C, units is not presently known. Thus the carbon assimilation pathway for Hyphomicrobium spp. growing on ethanol, acetate or 3-hydroxybutyrate remains uncharted. This is comparable to the situation with Pseudomonas AM 1 growing on ethanol or 3-hydroxybutyrate (Dunstan et al., 1972a, b).

C.

POSSIBLE ROLE OF CYTOCHROMES

The presence of several cytochrome types in methanol- and methylamine-grown Hyphomicrobium spp. has been known for several years. Using intact cells of methanol- and methylamine-grown H . vulgare, Hirsch and his coworkers ( 1963) obtained dif'ference spectra between dithionite-reduced and air-oxidized cells which showed the presence of the following cytochromes (absorption spectra of peaks are given in nanometers): type b (562, 530, 4291, type c (551, 522, 418) and a type a (608, -, 446). Carbon monoxide-reduced against reduced difference spectra revealed two carbon monoxide-binding haem proteins: an ala, (589, -, 430 and troughs at 606 and 447) and an o type (567, 532, 420 and a trough at 5 5 5 ) .The relative concentrations of the o type cytochrome were influenced by the degree of aeration of the culture. Later Hirsch (1965) reported the Hyphomicrobium spp. could tolerate carbon monoxide. When measuring the rate of oxygen uptake by Hyphomicrobium NQ52 1 using Warburg manometric techniques, uptake was stimulated in an atmosphere of CO : 0, (90: 10%). This stimulation occurred in the presence or absence of added carbon sources. The presence of cytochrome types a, b and c has been reported in other hyphomicrobia (Netrusov et al., 197 1). Recently, two groups of workers in England have begun to investigate the electron-transport chains of methylotrophs. Both groups report the presence of cytochromes a, b and c in methanolgrown Hyphomicrobium spp. Difference-spectra analysis of supernatant and particulate preparations from a Hyphomicrobium sp. gave the following distribution of cytochrome types (cytochromes are cited as

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pmoles mg protein-'). In the particulate fraction, cytochromes b, c and a were detected although the concentration of type a was not large enough to permit accurate measurements. The concentration o f cytochrome b was 285 and cytochrome c 173 of which 73 were bound to carbon monoxide. The supernatant fraction contained cytochrome c only; the concentration, however, was very high, namely 1263 of which 763 were bound to carbon monoxide. Indeed 61% of the carbon monoxide-binding cytochrome c was present in the supernatant (Tonge et al., 1974). A similar picture has been obtained by Widdowson and Anthony (1975).These authors claim that methanolgrown Hyphomicrobium X contains cytochromes of alas, b and c type and that all types have the ability to bind carbon monoxide. The distribution and concentration of the different types between the particulate and supernatant fractions were similar to those quoted by the previous workers except that, in addition, Widdowson and Anthony (1975) were able to measure the concentration of the a-type cytochrome in the particulate fraction. They quoted a concentration of 420 for alas and 140 for as type alone. Cytochrome c from the soluble fraction showed the ability to bind carbon monoxide in a manner identical with that observed with Pseudomonas AM 1 (Anthony, 1975b). This has been demonstrated with cytochrome c from Hyphomicrobium X grown on methanol, both aerobically or anaerobically. Furthermore, the carbon-monoxide difference spectra were essentially the same when cell extracts or whole cells were used. Short periods of exposure to carbon monoxide of washed membrane fractions of Hyphomicrobium X demonstrated that cytochromes a and b were able to react with carbon monoxide and may, therefore, function as an oxidase. Using these results together with similar results from other methylotrophic bacteria, mutant studies, and the fact that the carbon monoxidebinding cytochrome c can be induced- by growth on one-carbon compounds, both groups of workers have proposed tentative schemes for electron transport and energy transduction during growth on onecarbon and two-carbon compounds. Although further research is necessary before any definite scheme may be proposed, the carbon monoxide-binding cytochrome c has been implicated in oxidation of methanol (see Section III-A-1, p. 333). Methanol is oxidized to the level of formaldehyde by methanol dehydrogenase which interacts with the electron-transport chain at the level of cytochrome c (Tonge et al., 1974; Anthony, 1975b; Widdowson and Anthony, 1975).

-

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D.

W. HARDER AND MARGARET M. A r r W O O D

BIOCHEMICAL BASIS F O R RESTRICTED

METHYLOTROPHY

IN HYPHOMICROBIA

Since many hyphomicrobia have been shown to utilize only a limited range of carbon compounds for growth (Section 11-C, p. 3151, a detailed investigation was made by Harder et al. (1975)in order to find an explanation for the rather fastidious behaviour of these organisms. These authors argued that the observed inability to grow on a variety of carbon and energy sources could be due to: (i) a limited ability to transport these compounds across the cell membrane; or (ii) the lack of one or more key enzymes necessary for the metabolism of these compounds. I t has been suggested that hyphomicrobia isolated as denitrifiers, i.e. with a one-carbon compound and nitrate, in particular show a limited metabolic potential (Hirsch, 1974a). Therefore, the experiments were carried out with three organisms, namely Hyphomicrobium X, G and B522. The former two had been isolated using the anaerobic enrichment technique (Attwood and Harder, 19721, whereas strain B522 was isolated from aerobic enrichments by Hirsch and Conti (1964a). Using ['*CIlabelled substrates, the rate of transport of various carbon compounds was measured in methanolgrown cells in the presence of methanol as an energy source (Table 4). The rate of uptake of succinate, pyruvate and malate was linear with time for at least 4-5 h, and paralleled the disappearance of radioactivity from the suspending medium. N o uptake of the six-carbon compounds glucose or citrate was observed. In the absence of the growth substrate, uptake of succinate, pyruvate and malate also

TABLE 4. Rate of uptake of some organic compounds by methanol-grown Hyphomicrobium X

Rddioactive- labelled supplement

Rate of uptake (nmol h-l mg protein-I)

Methanol Acetate Citrate Glucose Succiriate

6.7 2.0 0 0 1.1 0.4 0.3

Pyruvate

Malatc

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occurred, although it was linear only for about 2 h and presumably relied upon endogenous reserves to provide energy for the transport process. Similar results were obtained with Hyphomicrobium G and B522. When ethanol was substituted for methanol as the growth substrate, essentially the same results were obtained with both Hyphomicrobium X and G ;Hyphomicrobium B522 cannot grow on ethanol (see Section 11-C, p. 317 and Table 1, p. 316) as sole carbon and energy source. From these results, it was concluded that failure of hyphomicrobia to grow on carbon compounds containing three or more carbon atoms cannot be explained completely in terms of an inability to transport such compounds into the cell, and therefore may be due to a metabolic block. This was investigated further by studying the rate of oxidation of these compounds. Washed cell suspensions of Hyphomicrobium X, grown in media containing methanol in the presence or absence of recrystallized pyruvate and malate or succinate, were incubated in the electrode compartment of an oxygen electrode system. Any stimulation in the uptake of oxygen above the endogenous rate upon addition of an organic compound was recorded. I t was found that methanol was oxidized whereas pyruvate, succinate and malate were not. Similar results were obtained with Hyphomicrobium G and B522. When Hyphomicrobium X and G were grown with ethanol or

2-Phosphoglycerate

Pyruva te- derived amino acids

It

T

Phosphoenolpyruvate

7 Pyruvate

Oxaloace ta te

Malate

FIG. 15. Metabolism ofpyruvate in Hyphomicrobiurn X.

3 54

W. HARDER AND MARGARET M. A W O O D

acetate as the carbon and energy source, essentially the same results were obtained in that ethanol and acetate were oxidized, whereas pyruvate, succinate and malate were not. From previous work on the carbon-assimilation pathway during growth of Hyphomicrobium spp. on methanol, it appeared that the metabolic potential of pyruvate was limited (see Fig. 15). These organisms lack an active pyruvate dehydrogenase complex as well as phosphoenolpyruvate synthase and pyruvate carboxylase (Harder et al., 1873; Attwood and Harder, 1974). If the pyruvate dehydrogenase complex is missing, then the organism would be limited in its ability to derive energy for growth on many threG-carbon to six-carbon compounds because acetyl-CoA required for the TCA cycle cannot be generated from these compounds (see Section 11-C, p. 318). This lack of an active pyruvate dehydrogenase complex as well as the E l component of the system was confirmed using Hyphomicrobium X and G grown under a variety of conditions. Thus it was decided to conduct studies on the distribution of pyruvate carbon into cellular amino acids in order to verify these findings. Organisms were grown for 2.5 generations in methanol-containing media in the presence of [L4Clpyruvate,["Clacetate or [14Clsuccinate and analysed for the distribution of radioactivity in the cellular amino acids. Very few amino acids became labelled after growth in the presence of [14Clpyruvate,and it was found that 93% of the total radioactivity recovered from the chromatograms was present in amino acids for which pyruvate is the direct precursor. In contrast, cells grown in the presence of ["Clacetate showed radioactivity in all classes of amino acids. Amino acids derived directly from pyruvate accounted for only 39% of the total radioactivity recovered, and the largest amount of radioactivity was found in glutamate. Similarly, cells grown in the presence of [14Clsuccinateshowed radioactivity in all classes of amino acids and the radioactivity was associated predominantly with glutamate and aspartate. These results are consistent with the enzymic data, and indicate that, in contrast to acetate and succinate, Hyphomicrobium X can make only restricted use of pyruvate. Similar results were obtained with Hyphomicrobium G and B522. Thus, when three-carbon or four-carbon compounds are present as the sole source of carbon and energy, they are unable to support growth of these organisms, and this appears to be due to the organism's inability to generate acetylCoA. The lack of an active pyruvate dehydrogenase complex has been shown in many hyphomicrobia including the following isolates : X, G, C, WA/B, T I , 10099, Har3, B522, ZV620, L530, H526 and C0582 (M.

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M. Attwood, unpublished observations). Of these isolates only X, G, C, WAD and T, were obtained from anaerobic enrichments. However, the lack of a pyruvate dehydrogenase complex is not common to all methanol-utilizing organisms since Pseudomonas AM 1 has been shown to contain such activity when growing on methanol. This is reflected in the ability of the organism to grow on succinate or pyruvate as sole source of carbon and energy (Salem et al., 1973). I t is not clear why the inability to channel pyruvate into major pathways of intermediary metabolism has persisted in hyphomicrobia during the course of evolution. Since these organisms are ubiquitous, it must be assumed that in their natural habitat loss of this potential is of no serious consequence. The situation is reminiscent of the lack of 2-0x0glutarate dehydrogenase in obligate chemolithotrophs (Smith et al., 1967; Kuenen and Veldkamp, 1973). A further point of similarity between hyphomicrobia and the obligate chemolithotrophs is that both groups of organisms show obligate dependence on special energy sources. In the presence of these energy sources, they may be able to assimilate various other compounds and use them as a source of carbon (Kuenen and Veldkamp, 1973). However, in this respect, hyphomicrobia are less versatile than the thiobacilli. This was made evident by the observation that supplements of exogenous carbon compounds when supplied under conditions of methanol limitation in continuous culture caused only a 0-5% increase in the yield of Hyphomicrobium spp. (J. B. M. Meiberg, unpublished observations), whereas an increase in the yield of thiobacilli of more than 20% has been recorded under these conditions (Kuenen and Veldkamp, 1973). A possible explanation for this limited ability of Hyphomicrobium spp. to assimilate exogenous carbon compounds during growth on methanol is given elsewhere (Section 111-A-3,p. 347). IV. Conclusion After an initial period when information was obtained on aspects concerning the biology of Hyphomicrobium spp., in particular their unusual morphology and reproduction, research in the last few years has resulted in progress being made into the physiology and biochemistry of the growth of these organisms on their preferred carbon sources. I t is very clear from this review that much more work needs to be done on almost every aspect of the microbiology and biochemistry of these organisms before a clear picture can be drawn of these unusual

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bacteria. In particular, a very rewarding field awaits the taxonomist and ecologist since little is known about either the taxonomy or the role of these organisms in Nature. However, now that Hyphomicrobium isolates are easily accessible, and therefore available as model systems for microbiological studies, one can expect a rapid progress in “Hyphomicrobiology” within the next few years. V. Acknowledgements

The authors are indebted to Professors H. Veldkamp, T. Y. Kingma Boltjes and J. R. Quayle for advice during the preparation of this review, to Professor R. Whittenbury, Dr. C. S. DOW, Dr. I. J. Higgins and Mr. J. B. M. Meiberg for providing data in advance of publication and Dr. C. S. Dow for supplying the electron micrographs. REFERENCES

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AUTHOR INDEX Numbers in italics refer to the pages on which references are listed at the end of each article

A

Arpin, M., 120, 165 Arthur, H., 66, 67, 73, 165 Ashworth, J. M., 165 Aspinall, D., 187, 241 Attwood, M . M., 304, 305, 306,307,308, 312, 313, 314, 315, 317, 318, 325, 326, 327, 329, 330, 331, 335, 336, 340, 341, 342, 344, 345, 349, 352, 354,356,357 Audette, R. S., 161, 165 Austwick, P. K. C., 164, 165 Avron, M., 217, 219, 220, 222, 226, 239, 240 Ayres, J . C., 26, 42

Aastad, K., 259, 297 Abdolrahimzadeh, H., 7 1, 173 Adams, A. M., 136, 164 Adamse, A. D., 330,356 Adhya, S., 230, 241 Adman, R., 273,276,297 Agarwall, S. S . , 273, 300 Ahmadjian, V., 154, 170 Aitken, D. M., 186, 188, 194,239 Aitken, W. B., 123, 129, 164 Albergoni, F., 2 17, 24 I Alderton, G., 18, 3 1, 35, 38, 40, 43 A1 Doory, Y., 161, 164 Alford, J . A., 134, I65 Allen, J . V., 51, 165 Allen, P. J., 116, 139, 165, 167 Alsbach, E. J. J., 232,241 Altman, R. L., 34, 40 Baas-Becking, L. G. M., 216,239 Anand, J. C., 185, 188, 198, 199, 213, Babczinski, P., 82, 165, 176 2 18, 236, 239 Bachofen, R., 137, 165, 174 Anderes, E. A,, 161, 165 Baddiley, J., 71, 97, 98, 165, 173, 174 Anderson, J . G., 118, 133, 136, 172, 175 Bailey, G. F., 34, 40 Anderson, J. M., 294,297 Baillie, E., 38, 40 Ando, Y., 31, 40 Angus, W. W., 86, 89, 91, 92, 94, 95, 96, Baker,J. M., 155, 156, 157, 165 Balassa, G., 3, 24, 42, 44 112, 134, 165, 172 Ballario, C., 33, 40 Annear, D. I . , 35, 40 Ballou, C. E., 68, 83, 84, 98, 165, 174, Ansell, G. B., 78, 165 179 Anthony, C., 333, 334, 338, 349, 350,351, Bamberger, M., 90, 171 356,357,359 Banbury, G. H., 49,58, I74 Anwar, R. A., 24, 40 Banbury, Y. G . H., 118,177 Araki, Y., 21, 22, 40 Banerjee, A. B., 161, 167 Arango, M., 165, 175 Bannerjee, A. K., 284,297 Aristovskaya, T. V., 309,356 Bangham, A. D., 70, 165 Armentrout, V. N., 56, 152, 165 Baniecki, J . E., 121, 165 Arnaud, M. V., 292, 293,301 Baraud, J., 78, 165 Arnold, W. N., 224, 239 Aronson,A. I . , 5, 8, 10, 11, 12, 16, 17, 27, Barenholtz,Y., 102, 111, 131, 165, 168 Barksdale, A. W., 115, 165, 172 33, 37, 40, 41, 42 36 1

362

AUTHOR INDEX

Barlow, A. J. E., 149, I67 Barnett, J. A., 231, 239 Barondes, S. N., 99, I68 Barr, R. M., 79, 82, 165 Bartnicki-Garcia, S., 49, 50, 121, 149, I65 Bassham, J. A . , 338,356 Basu, S., 88, 91, 112, 165, 171 Bateman, D. F., 134, 148, I77 Batra, L. R., 156, 165 Bauer, H. F., 76, 172 Bauld, J., 309,356 Baverstoft, I., 154, I73 Baxter, M., 161, 164, 165 Baxter, R. M., 66, 68, 165, 171 Beaman,T. C., 3, 4, 5 , 6, 7, 12, 40, 4 3 Bean, G.A., 122, 153, 173, I74 Bechet, J., 276, 301 Beck, C., 260, 297 Beck, D. P., 58, 165 Becker, S. A. W. E., 5 1, 176 Beckett,A., 51, 123, 125, 169, 170 Beckman, M. M., 23, 4 1 Beesley, T., 162, I79 Beevers, H., 57, 165 Begg, K. J., 320,357 Behrens, N . H., 79, 165 Bell, G., 274, 275,301 Bell, G. I., 279, 280, 297 Bellini, E., 219, 241 Bellion, E., 338, 339, 341, 342,356,357 Belton, J. C., 222, 241 Ben-Amotz, A . , 217, 219, 220, 222, 226, 239,240 Benson, A. A., 338,356 Benson, S. W., 34, 4 0 Ben-ZePv, H., 2 5 , 40 Benzonana, G., 134, 175 Bergere, J-L., 27, 4 0 Berkeley, R. C. W., 23, 40 Berlin, J . D., 138, 165 Bernlohr, R. W., 15, 42 Berry, D. R., 114, 120, 176 Bertaud, W. S., 122, I65 Berrhoud, S., 96, 168 Bertsch, L. L., 5, 26, 29, 32, 40, 43 Bettelheim, K. A., 136, I66 Bhargava, M. M., 257, 258, 273, 292, 293, 297,298,299,301 Bhat, J . V., 310,359 Bhattacharjee, S. S., 7 7 , 166 Bianchi, D. E., 50, 149, I66

Bicknell, J. N., 251, 255, 297 Billing, E., 32, 43 Bimpong, C. E . , 5 1 , 123, 124, 135, I66 Binder, M., 249, 251, 255,302 Bishop, H., 25, 40 Bishop, H. L., 25, 4 0 Bishop, J. O., 283, 297 Black, S., 233,240 Black, S. H., 4, 7, 30, 32, 40, 41 Blackman, M. A., 322,359 Blamire, J., 254, 255, 256, 258, 260, 271, 297,298 Blank, F., 161, I66 Blewett, M., 155, 166 Bloch, K., 101, 166 Block, R. J., 190, 240 Bloss, H. E., 121, 166 Blumberg, P. M., 24, 40 Boggess, S. F., 187, 241 Bohonos, N., 80, I66 Bolanos, B., 165, I69 Bollum, F. J., 272, 273,298 Bonhoeffer, F., 266, 299 Bonincontro, A., 33, 40 Bonsen, P. P. M., 6, 26, 28, 29,32, 40, 43 Bont, J. A. M., de, 330,356 Booij, H. L., 66, 73, I67 Boothroyd, B., 77, 78, 166 Boothroyde, B., 77, I69 Borer, R., 96, I68 Borkenhagen, L. F., 105, I66 Borowitzka, L. J . , 185, 196, 216, 217, 218, 219, 220, 221, 226, 227, 231, 235,240 Bosch, Van der H., 102, I66 Bose, H. R., 334,358 Bosmann, H. H., 98, 172 Both, G. W., 284,297,300 Bott, K. F., 2 5 , 41 Boulton, A. A., 69, I66 Boulton, C. A . , 336,358 Bowen, C. C., 50, 138, 165, 168 Bowman, R. D., 66, 166 Boyd, K. S . , 162, 169, 224,239 BrBcker, C. E., 49, 50, 52, 54, 57, 138, 166, I69 Bradbury, J . H., 15, 40 Bradley, D. E., 7 , 41 Bramley, P. M., 119, 120, 166 Breathnach, A. S., 160, I66 Brendel, M., 295,297, 298

AUTHOR INDEX

Brennan, P. J., 48, 68, 72, 74, 75, 76, 7 7 , 80, 84, 85, 86, 87, 88, 90, 94,95, 100, 102, 112, 116, 131, 142, 166, 169, 171 Bretscher, M. S., 59, 166 Bretthauer, R. K., 82, 166 Brewer, S. J., 23, 40 Brichti, J., 224, 240 Brock, T. D., 216,240 Brody, S., 70, 71, 72, 113, 170 Brogt, T. M., 278, 297 Brooks,J.B., 117, 174 Broquist, H. P., 158, 178 Brown, A. D., 182, 184, 185, 186, 187, 188, 191, 192, 194, 195, 196, 197, 198, 199, 200, 208, 209, 210, 211, 213, 216, 217, 218, 219, 220, 221, 222, 223, 225, 226, 227, 228, 229, 231, 232, 235, 236, 239, 240 Brown, C.M., 72, 171, 187, 241 Brown, M., 273,298 Bruff, B. S., 219,240 Brunstetter, B. C., 30, 41 Buhler, F. M., 280, 299 Buhler, J. M., 276, 280, 297, 299 Bull, A. T., 50, 166 Bulla, L. A , , 26, 41 Bu'Lock, J. D., 114, 166 Burger,M. M.,98, 107, 166, 177 Burgoyne, L. A . , 252,299 Burke, W., 268, 270,297 Burney, A., 276,301 Burr, H. K., 3 5 , 4 2 Burton, K., 294, 297 Burton, R. M., 107, 166 Butterworth, A . H. W., 79, 176 Butterworth, D. H. W., 293,297 Byk, C., 104, 177 Byers, B., 247, 248, 249, 254, 266, 297, 298,300 Byrne, P. F. S . , 68, 74, 75, 77, 88, 95, 166

C Caglioti, L., 11 7, 166 Cainelh, G., 117, 166 Caldwell, D. E., 312, 314,356 Callan, H. G., 266,298 Callister, H., 26, 41 Caltrider, P. G., 135, 143, 166, 169

363

Calvin, M., 338,356 Cameron, R., 285, 286,301 Cametti, C., 33, 40 Cammack, C. L., 124, 138, 176 Campbell, C. K., 51, 123, 166 Campbell, L. L., 7, 43 Campbell, R., 122, 166 Campbell, W. P . , 127, 166 Canh, D. S., 208,240 Cann, D. C., 3, 4 2 Cantino, E. C., 51, 58, 7 7 , 79, 124, 135, 166, 173, 176 Carbonell, L. M., 149, 171 Carlile, M. J., 118, 166 Carmack, C. L., 148, 149, 167 Carstensen, F. L., 33, 41 Carter, B. L. A., 260, 268, 279, 291, 292, 293,301 Carter, H. E., 92, 95, 167 Cislavski, J., 39, 41 Cassier, M., 17, 37, 41 Caten, E. E., 245,300 Cerbon, J . , 73, 167 Cerdi-Olmedo, E., 268,298,299 Chalfan, Y . , 338,356 Challinor, S. W., 96, 174 Chambers, T. C., 124, 167 Chambon, P., 25, 41, 275,276,298 Chang, L.M. S., 272, 273,298 Chapman, D., 60, 167 Charalampous, F., 96, 168 Chattway, F. W., 149, 167 Chen, C., 146, 170 Cheng, H. M., 17,41 Cherrier, C., 27, 40 Chesterton, C. J., 293, 297 Chevrier, G., 10, 11, 12, 42 Chichester, C. O., 119, 171, 179 Chin, T., 20, 21, 41 Ching, T. M., 122, 177 Christensen, H. N., 208, 240 Christian, J . H . B., 187, 200,240 Christiansen, C., 251, 298 Christiansen, G., 251, 298 Chu, H. M., 156, 167, 171 Ciferri, O., 251, 298 Cleveland, E. F., 21, 41 Cobon, G. S., 105, 167 Cochrane, J . C., 123, 167 Cochrane, V. W., 123, 167 Coffey, M. D., 139, 167

3 64

AUTHOR INDEX

Colby, J . , 308, 3 17, 335, 336, 338,356 Cole, R. M., 36, 4 I Coleman, R . , 72, 168 Collins, C. B., 123, 167 Cornbepink, G., 136, I 7 7 Coinbs,T. J., lG1, 167 Compe, J., 82, I 7 2 Conovcr, T. E., 188,240 Contaxis, C. C., 194,240 Conti, S. F., 3, 43, 304, 305, 306, 308, 309, 310, 311, 312, 314, 317, 318, 319, 327,350, 3 5 2 , 3 5 6 , 3 5 7 , 3 5 8 Conway, E. J., 212,240 Cooke, R. C., 144, 155, 158, 165, 167 Cooke, W. B., 164, 167 Cooper, B. H., 160, I67 Cooper, D. R., 196, 214,240 Cooper, K. M., 66, 139, 167 Cooper, S., 260, 298 Corder, J. L., 252, 301 Corina, D. L., 187, 240 C o r y , J. E. L., 21 1 , 2 4 0 Cosovic, C., 80, 88, 91, 95, 167, 174 Cotter, D. A,, 123, 137, 167 Cottrell, S. F., 27 1, 298 Cox, G., 139, 141, 152, 167 COX,R. B., 335,338, 356,357 Cox, R. F., 298,297 Crabbe, M. J. C., 3 3 6 , 3 5 8 Craig, B. M . , 67, 127, 130, 142, 170, 171 Craigie, J . S., 187, 217, 220, 226, 240 Cramer, J . H., 257, 258, 278, 298 Creanor, J.. 260, 278, 294, 298 Grocken, B. J., 70, 72, 104, 113, 167 Cronan, J. E., Jr, 106, 170 Crosby, W. H., 30, 41 Crowfoot, P. D., 105, 167 Cryer, D., 254, 255, 256,297 Cryer, D. R., 255, 256, 260, 271,298 Culotti, J., 248, 249, 262, 286, 294, 298, 299 Culotti, M., 262, 299 Curran, H. R., 41 Curry, M . V., 35, 42

D Dahl, H . L., 75, I 7 7 Daniel, J. W., 118, 167 Danielli, J . F., 59, 167 Dark, F. A,, 22, 23, 44

Darland, G. K., 136, 167 Dart, R. K., 49, I67 Das, S. K., 161, 167 Davenport, A,, 357 David, C. N., 58, 167 Davidoff', F., 123, 130, 167 Davies, B. H., 119, 120, 166 Davies, J., 293,301 Davis, R. W., 285, 286,301 Davson, H., 59, 167 Dawes,E.A.,349, 359 Dawes, I. W., 268, 279,298 Day, A . W., 296,298 Dean, D. H., 33, 4 I Dearborn, D. G., 99, 167, 171 De Bary, A , , 155, 167 De Bell, R. M., 68, 167 Dee, J., 165 Defago, G., 150, 167 Deierkauf, F. A,, 66, 73, 167, 175, 176 Deinema, M . H., 74, 76, 132, 177 Deitz, V. R., 33, 43 Delafuente, G., 233, 241 Delafield, F. P., 10, 11, 12, 14, 17, 41, 44 Delwiche, C. V., 142, 173 Denton, J . F., 161, 169 Deponder, R. 10, 11, 1 2 , 4 2 Deshusses,J., 96, 97, 104, 106, 167, 168 Desnuelle, P., 134, 175 , Deutscher, M. P., 25, 41, 42 Dezelee, S., 273, 276, 280, 298 D ' H o n d t , E., 236,240 Dijkstri, F., 332, 357 DiSalvo,A. F., 161, 168 Dixon, G. F., 342, 3 5 7 Dixon, J. F., 75, 177 Doi, R., 25, 40, 41 Doi, R. H., 25, 26, 42 Doman, N . G., 340,357 Domer, J . E., 149, 168 Donachie, W. D., 320, 357 Donike, M., 294,299 Donnellan,J. E., 33, 41 Douglas, H. C., 251, 255, 297 Douthit, H . A., 33, 41 Douzou, P., 194, 240 Dow, C., 310, 320, 321, 322, 323, 324, 359 Drabble, W. T., 350,357 Dring, G. J., 33, 35, 36, 37, 38, 39, 41 Duckworth, M., 98, 174

AUTHOR INDEX

Duffus, J. H., 292,293, 298 Dulak, N. C., 75, 177 Dunstan, P. M., 350,357 Du Praw, E. J., 251,298 Durrum, E. L., 190,240 Dutton, H. J., 81, 176

E Eady, R. R., 338,'357 Eckstein, H., 273, 298 Edelman, I., 131, 165 Edgell, M. H., 25, 41 Edmonds, M., 284,298,300 Edwards, G. A., 160, 168 Edwards, M. 160, 168 Eggleston, L. V., 231, 236,240 Eisenstadt, E., 31, 41, 43 Elmer, P. R., 50, 168 Emmons, C. W., 164, 168 Engel, H., 314,358 Ensign, J . C . , 23, 42 Erwin,J.A.,48, 49, 100, 101, 131, 168 Esders, T. W., 76,*78, 107, 108, 109, 133, I68 Esterbrook, K., 5 8 , 167 Evans, H. H., 264,300 Evans, T. E., 264,300 Eveleigh, D. E., 50, 175 Ewall, J. L., 273,300

F Falaschi, A,, 25, 41 Fan, H., 294,300 Fan, P., 23, 41 Fangman, W., 254, 266,300 Fangman, W. L., 254, 255, 266, 268, 270, 271,279,300 Farkas, G. L., 142, 143, 168 Fath, W. W., 295,297,298 Felix, A., 334,358 Ferenci, T., 338,359 Fergus, C. L., 65, 67, 173 Fiasson, J. L., 120, 168 Fiechter, A., 288, 300 Fiegert, E., 80, 89, 90, 178 Fielding, L., 105, 166 Findlayson, J. S., 15, 43 Finean, J. B., 72, 168 Fink, A. M., 27 1,30 2

365

Finkelstein, D. B., 254, 255, 256, 258, 260,271,297,298 Finlayson, A. J., 142, 175 Finley,A. A,, 161, 165 Fisher, D. J., 122, 168 Fisher, K.A., 61, 168 Fisher, P., 249, 25 1, 255,302 Fitz-James, P. C . , 2, 5 , 8, 10, 11, 12, 16, 1 7 , 18, 26, 27, 28, 33, 36, 37, 38, 40, 41, 43 Fleming, H. P., 31, 43 Fleming, R., 118, I 8 0 Fluharty,A. L., 77, 78, 168 Flynn, M. P., 77, 84, 166 Foerster, H. F., 31, 41 Forrest, H. S., 334, 335,359 Foster, J. W., 12, 13, 14, 30, 31, 41, 42, 43,44 Fox, C . H., 154, 168 Fraenkel, G., 155, 166 Francke-Grosmann, H., 155, 157, 158, 165, 168 Franco, L., 251, 252,298 Frank, J., 66, 168 Franklin, J. G., 7, 41 Fraser, R. S. S., 260, 278, 294,298 Frazier, W. A., 99, 168 Frear, D. S., 123, 142, 170 Fred, E.,357 Frederick, E. W., 280,298 Freese, E., 36, 41 Freese, E. B., 36, 41 Frey-Wyssling, A., 57, 168 Friend, J., 118, 166 Fries, N., 116, 117, 153, 168, 171 Fromageot, P., 273, 276, 280, 297, 298, 299 Froschl, N., 80, 90, 168 Fujita, T., 33, 42 Fujita, Y., 25, 44 Furuichi, Y., 284,300

G Gadot, N., 1 1 1, I65 Gahan, P. B., 5 6 , 168 Galbraith, J. C., 114, 136, 137, 168, 176 Galli, M. G., 219, 241 Gancedo, C . , 201, 208, 226, 233,240,241 Gancedo, J. M., 201, 208, 226, 233,240 Ganfield, M.-C. W., 97, 174

AUTHOR INDEX

366

Garcia, R. E., 108, 173 Cardner, J. V., 35, 42 Garrett, M. K., 1 1 7 , 168 Garrison, R. G., 162, 168, 2 2 4 , 2 3 9 Gatt,S., 102, 1 1 1 , 131, 165, 168 Gauthier, J . J., 6 , 15, 18, 21, 24, 42, 44 Gay, J. L., 50, 136, 166, 168 Gaylor, J. L., 142, 173 Geitler, L., 3 3 0 , 3 5 7 Gerhardt, P., 3 , 4, 5, 6 , 7 , 12, 28, 30, 32, 40, 41, 43

Gerisch, G . , 99, 178 Getz, G. S . , 66, 72, 73, 102, 106, 168, 170, 1 7 1 , 172, 173,271, 298

Ghosh, A , , 96, 168 Ghuysen, J. M., 20, 4 1 Gibson, D. T., 3 3 4 , 3 3 5 , 3 5 9 Gillespie, J . B., 23, 40 Gilmore, R. A . , 2 5 7 , 2 9 8 Gilpin, R. W., 25, 41 Gdvarg, C., 21, 24, 27, 34, 4 1 , 42, 43, 44 Gimmler, C . M., 268, 298 Ginzburg, B. Z., 222, 241 Ginzburg, M., 219, 2 4 0 Ghsare, P., 159, 168 Glaser, L., 20, 21, 23, 41, 42, 107, 166 Click, M.-C., 61, 178 Goad, L. J., 155, 172 Goetsch, L., 247, 248, 249, 2 9 7 , 2 9 8 Gold, M. H., 82, 169 Goldberg, I., 334, 357 Goldberg, S., 2 5 8 , 2 9 8 Goldring, E. S., 27 1, 299 Goldthwaite, C. D., 255, 256, 260, 27 1 , 298 Golombek, J., 272, 273, 299 Golub, E. S . , 17, 42 Gooday, G . W., 115, 169 Gooday, M. A . , 116, 169 Goodwin, R. W., 115, 177 Goodwin, T. W., 100, 155, 169, 172 Gordon, M . A , , 165, 175 Gorcnstein, C., 283, 2YY Gorin, P. A . J . , 76, 77, 166, 169, 177 Got, R., 82, 84, 172, Gortesman, M., 230, 241 Gottheb, D., 123, 129, 135, 143, 166, 169 Gottlieb, D. J . , 143, 178 Could, G. W., 13, 14, 1 7 , 23, 33, 35, 36, 3 7 , 3 8 , 39, ,41, ‘12 Graf; G. L. A , , 66, 67, 169

Gray, R. H., 2 4 4 , 2 5 3 , 2 9 9 Gray, W. D., 118, 169 Green, W. G. E., 2 3 4 , 2 4 0 Greenawalt, J . W., 58, 165 Greene, R. A , , 30, 41 Greenwood, A. D., 50, 51, 5 8 , 127, 139, 168, 169, 173

Greer, D. L., 164, 169 Grenson, M., 294,299 Greuter, B., 51, 57, 128, 169 Grieshaber, E., 57, 168 Griffin, P. F. S., 48, 72, 74, 75, 76, 7 7 , 84, 85, 95, 100, 102, 116, 131, 142, 166, 171

Griffiths, D. A , , 127, 166 Grinstead, K. H . , 22, 43 Grivell, A. R., 294, 299 Groner, B., 284,299 Groner, Y., 284,?01 Gross, M., 160, I 6 6 Grossmann, L. I., 27 1 , 2 9 9 Grove, S., 161, 167 Grove, S. N., 49, 50, 51, 5 2 , 5 7 , 5 8 , 169 Grover, N. B., 222,241 Guameri, J. J., 161, 167 Guening, C., 66, 67, 169 Guerola, N., 268,298, 299 Guinand, M., 24, 42 Gunasekaran,M.,51, 122, 123, 129, 169 Gunstone, F. D., 102, 169

H Haber, J. E., 268, 269, 2 7 0 , 2 9 9 Hackett, J . A , , 80, 86, 90, 95, 112, 169 Hacking, A . J., 3 5 5 , 3 5 9 Hacskaylo, E., 153, 173 Haff, L. A., 280, 299 Hager, G. L., 274, 275, 276, 277, 278, 301

Hahn, H . J . , 82, 168 Hakomori, S., 99, 169 Haley, J . E., 69, 16Y Hall, B. D., 273, 276, 277, 279, 2 9 7 , 3 0 1 Hall, J . M., 187, 210 Hall, M. M . , 21, / ? Hall, S. W., 144, I 7 3 Halvorson. H. O., 35, 39, 42, 45, 125, 136, 170, 178, 251, 257, 258, 260, 273, 276, 278, 279, 291, 292, 293, 297, 298,299, 301

AUTHOR INDEX

Hamer, G., 3 18, 330, 33 1,359 Hamilton, J. G . , 149, 168 Hamilton, W. A,, 232, 240 Hamilton-Miller, J. M. J., 61, 169 Hanahan, D. J., 68, 169 Hanawalt, P. C., 268,298 Hanh, V. T., 24, 42, 43 Hanssler, G., 56, 165 Hanson, E. W., 138, 176 Hanson, R. S., 35, 42,346,358 Hanssens, K . , 236,240 Harder, W., 304, 305, 306, 307, 308, 312, 313, 314, 315, 317, 318, 325, 326, 327, 329, 330, 331, 333, 335, 336, 337, 340, 341, 342, 345, 349, 352, 354,356,357,358,359 Hardwick, N. V., 58, 139, 169 Harkin, J. C., 149, 168 Harrington, A. A., 333, 335,357 Harris, A. Z., 338,356 Harrison, D. E. F., 331, 334, 351, 357, 359 Harrison, J. E., 338,358 Hart, G. J., 196, 214,240 Hartleb, R., 303, 308,359 Hartmann, E., 90, 169 Hartree, E. F., 333, 357 Hartwell, L. H., 248, 249, 262, 264, 265, 270, 286, 287, 294,298,299 Harvey, A. E., 139, 175 Harwood, J. L., 100, 169 Hasegawa, K., 69, 177 Hashimoto, T., 3, 30, 40, 43 Haskell, B. E., 68, 169 Haskins, R. H., 50, 77, 78, 143, 166, 169, I75 Hattori, J., 25, 40 Hawker, L., 123, 169 Hawker,L. E.,51, 116, 169, 170 Hawthorne, D. C., 249, 250, 254, 255, 300 Hawthorne, J. N., 66, 78, 92, 95, 104; 106, 165, 167, 178 Hayashi, H., 21, 22, 40 Hazen, E. L., 160, 168 Hazen, W., 330,357 Headley, C. L., 3, 42 Heath, I. B., 50, 168 Heath, M., 141, 169 Heber, U. W., 234,240 Heimer, Y. M., 196,240

367

Heintz, C. E.,51, 123, 129, 169 Heitefuss, R., 67, 134, 142, 145, 146, 147, 148, 150, 170 Helfman, W. B., 272, 299 Helmstetter, C. E., 260, 298 Hemmes, D. E., 50, 51, 124, 166, 169 Hemmings, F. W., 79, 82, 165, 176 Hendrix, J. W., 66, 67, 120, 121, 169 Henrici, A. T., 304, 305,357 Henry,S.A., 125, 136, 169, I77 Hepden, P. M., 116, 170 Heptinstall, J., 340,357 Herbert,D.,327,357 Herbold, D. R., 23, 42 Hereford, L. M., 264, 265, 266, 270, 299, 300 Hereward, F. V., 65, 170 Hersh, L. B., 338, 339, 341, 346, 356, 357,358 Hess, W. M., 51, 114, 116, 120, 121, 122, 123, 128, 129, 139, 142, 165, 169, 170, 171, I78 Hewish, D. R., 252,299 Hexum, T. D., 75, 177 Heywood, J., 66, 168 Hickman, C. J., 51, 123, 124, 166 Higgins, I . J., 334, 351,359 Hignite, C., 80, 81, 90, 170, 1 7 1 Hildebrand,A., 122, 143, 171,276,297 Hilger, F., 276,301 Hill,A., 76, 77, 1 7 7 Hill, B., 341, 342, 343, 344, 346, 347,357 Hill, E. P., 51, 124, 154, 170 Hilz, H., 273, 298 Hirashi, S., 25, 44 Hirsh, H. M., 137, 170 Hirsch, J., 27 1,302 Hirsch, P., 304, 305, 306, 308, 309, 310, 311, 312, 314, 317, 318, 319, 320, 322, 326, 327, 328, 329, 330, 336, 350,352,356,357,358 Hirschberg, R., 255, 256, 260, 271, 298 Hoare, D. S., 304, 305, 306, 312, 314, 315, 317, 331, 334, 335, 338, 358, 359 Hoch, H . C., 50, 170 Hodgkiss, W., 3, 42 Hoeks, J., 330,356 Hoeniger, J. F. M., 3, 4 2 Hofer, M., 232, 241 Hohl, H. P., 50, 51, 173

368

AUTHOR INDEX

HBhl, H . R., 51, 123, 124, 126, 168, 169, 170 Holbert, P. E., 7 , 42 Hollenstein, G. O . , 51, 174 Holligan, P. M., 146, 170 Hollway, P. J., 122, 168 Holmes,A. M., 272, 299 Holmes, M . R., 149, 167 Holt, S . C., 2, 3, 7, 14, 1 7 , 21, 41, 42 Holts, R. B., 67. 121, 127, 150, 171 Hoppe, H . K., 67, 134, 142, 145, 146, 147, 148, 149, 170 Horak, J., 208, 240 Horecker, B. L., 79, 170 Hori, T., 264, 29Y Horn, D., 10, 16, 1 7 , 40, 42 Horne, R. W., 69, 174 H o m e r , T. L., 133, 142, 171 Horton, J . C., 50, 168 Hoshi,M., 80, 81, 90, 170, 171 Hostak, M. B., 116, 170 Hougen, F. W., 142, 170 Howell, S., 262, 263, 299 Hsieh, L. K., 22, 33, 42 Hubbard,S. C., 70, 7 1 , 72, 113, 170 Huberman, J . D., 266,299 Huet, J., 66, 179, 280, 299 Huettel, R. N., 4, 5, 45 Hughes, D. H., 127, 170 Humpers, J., 66, 67, 169 Huneck, S., 154, 168, 170 Hungerer, K. D., 20, 4 2 Hunter, K . , 50, 61, 102, 171, 207, 210, 240 Hurwitz, J., 280, 298 Hutchinson, C. A , , 25, 41 Hutchison, H . T., 106, 170

I Iandola, J . J., 26, 41 Iborra, F., 276, 297 Ichikawa, T., 15, 16, 42, 43 Idriss, J. M., 258, 298 lisima, K . , 15, 34, 42 Ikeda, Y., 25, 43 Iltingworth, R. F., 51, 125, 170 lmae, Y., 21, 36, 39, 42 Iinahori, K . , 25, 44 lngold, C. T., 118, 170 lngraham, J . L., 268,299

Ingram, D. S . , 50, 138, 177 Ingram, J. M . , 235,240 Ingram, M., 197,240 Irwin, W. E., 82, I66 Isobel, M,M., 122, 165 Israelstam, G. F., 234, 240 [to, E., 2 1 , 22, 40

J Jack, C. M., 130, 170 Jack, R. C., 67, 68, 69, 167, 16Y Jack, R. C. M . , 65,80,85,88, 170, 178 Jackson, G. V. H . , 51, 125, 138, 170 Jackson,L. L., 122, 123, 142, 143, 170 Jackson, J. F., 294,299 Jacob, H . , 72, 170 Jacobs, L., 137, 172 Jaenike, L., 294,299 Jager, K., 145, 175 Jakoby, W. B., 233,241 Jakovic, S., 66, 72, 169, 170 James, C. R., 12, 44 Jandric, Z., 95, 168 Jannasch, H . W., 330,331,357,359 Jannsen, S., 294, 295,299 Jarvie, W., 122, 138, 141, 172 Jarvis, F. G., 107, 170 Jayko, M. E., 68, 169 Jeng, Y.-H., 2.5, 42 Jobling, B., 118, 179 Johns, E. W., 251, 252,298 Johnson, B., 72, 170 Johnson, D., 128, 171 Johnson, D. E., 304, 305,357 Johnson, E. J., 291, 240 Johnson, J . M., 7 I , 174 Johnson, M . K., 219,240 Johnson, P. A , , 335,357 Johnson, R. M., 309,310,357 Johnson, T. W., 122, 170 Johnston, I . R., 272,299 Johnston, J . M., 106, I 7 0 Johnson, M. J., 107, 170 Jones, D. F., 76, 7 7 , 171 Jones, H . E., 318, 322,357 .Jones, J . K . , 296, 298 Jones, R. F., 268,300 Jung, P., 82, I71 Jurjitza, G., 159, 171 Just, G., 162, 166

AUTHOR INDEX

K

369

Kish, Z., 67, 171 Kishimoto, Y., 8 0 , 8 1,90, 170, I 7 1 Kaback, D. B., 258,299 Kitazume, Y., 284, 299 Kadota, H., 10, 11, 15, 25, 34, 42, 43, Klein, A., 266, 299 44 Kljaic, K., 80, 178 Klofat, W., 36, 4 1 Kakin, L. E., 75, 177 Klotz, L. C., 254, 255,300 Kakiuchi, Y., 10, 44 Knaysi, G., 28, 42 Kalafer, M. E., 142, 173 Knights, B. A., 66, 69, 150, 162, 171, Kallio, R. E., 333, 335,357 I72 Karnano, T., 25, 44 Knoche, H. W., 123, 129, 133, 142, 143, Kamiya, H., 81, 90, 131, 174 145, 149, 151, 172, 173 Kaneda, T., 6,26, 42,333,335,357 Knowles, C. J., 334, 351,359 Kanetsuma, F., 149, 171 Knox, K. W., 97, 178 Karp, S., 34, 40 Kobayashi, Y., 25, 42 Karst, F., 213, 240 Koch,A., 155, 171 Kates, M., 66, 68, 80, 102, 171, 176 Kodaira, Y., 144, 171 Kato, N., 333,357 Kaufman, B., 112, 165 Koga, S., 33, 42 Kogane, F., 123, 179 Kaufmann, B., 88,91, 171 Kaul, K., 95, 171 Koh, T. Y., 213, 224,240 Kohn, P., 77, 173 Kauss, H., 187, 240 Kok, L. T., 157, 167, 171 Kavenagh, F., 116, 175 Kolenbrander, H. M., 346,359 Kavenagh, V. W., 116, 175 Kondo, M., 10, 12, 13, 14, 15, 16, 42, 43, Kay,Z. D., 338,356 44 Kee, S. G., 268, 269, 270, 299 Keech, D. B., 338,358 KondratPva, E. N., 350,358 Keen, N. T., 138, 144, 178 Kopecka, M., 224,240 Kopp, D. W., 284,298 Keenan, T. W., 60, 174 Korn, E. D., 99,167, 171 Keilin, D., 333,357 Keller, E. B., 280, 299 Kornberg, A,, 6, 1 I , 16, 17, 25, 26, 28, 29, Kelley, W. D., 148, 149, 167 32,40, 41, 42, 43, 44,27 1,299 Kemp, M. B., 338,358 Kornberg, H. L., 342,357 Kennedy, E. P., 103, 105, 113, 174, 175 Kornberg, R. D., 252, 253,299 Korte, I., 314,358 Kennedy, E. R., 105, 166 Koshimizu, K., 80, 85, 172 Kessel, J. F., van, 330,356 Kessly, D. S., 185, 217, 218, 219, 221, Kostiv, L.L., 161, 171 Kotyk, A., 208, 240 235,240 Khuller, G . K., 94, 171 Kovac, L., 65, 175 Kijimoto, S., 99, 169 Kramer, J. K. G., 67, 171 Krebs, H. A., 225, 231,232, 236,240 Kim, W.J., 142, 149, 150, 174 Kreger, D. R., 49, 171 Kimura, A., 80, 85, 171 Kreger-van Rij, N. J. W., 229, 234,241 Kimura, K., 25, 44 Kriss,A. E., 312,358 Kimura, M., 80, 85, 171 Krupa, S., 1 1 7, 153, 171 King, W. L., 13, 14, 1 7 , 23, 41, 42 Kuenen, J. G., 355,358 Kingan, S. L., 23, 42 Kingma Boltjes, T. Y., 304, 306, 308, 31 1, Kuhlwein, H., 159, 171 314, 315, 319, 320, 322, 323, 327, Kuhn, N . J., 105, 171 Kung, H. F., 335,358 329,330,358 Kuo, C. H., 94, 178 Kinoshita, S., 80, 132, 177 KUO,S.-C., 64, 171 Kirby, D. K., 102, 174 Kushwaha, G. C., 67, 171 Kirikova, N. N., 350,358

AUTHOR INDEX

370 Kylin,A., 212, 241 Kyte, J., 75, 171

L Lachmann, E. R., 294, 299 . LaCleve,A.J., 137, 167 Lacroute, F., 213, 240 Ladner, A . , 334,358 Laine, R . A . , 72, 74, 75, 76, 100, 171 Lam, K. B., 255, 256, 260, 271,298 Lambert, E. B., 137, 171 Landsiedel, A , , 90, 171 Lang, D. R., 5, 26, 42 Langcake, P., 151, 171 Langenbach, R. J . , 123, 129, 142, 143, 149, 151, 171, 172 Lanyi, J . K., 195, 213, 241 Large, P.J., 319, 336, 338, 340, 344,357, 358 Lark, K. G., 264,299 Larsh, L. W., 160, 164 Larson, R. J., 329,358 Laseter, J. L., 60, 121, 122, 123, 171, 178 Laskowski, W., 294, 295, 297,298,299 Lasota, 68, 173 Latorella,A. H., 217, 219, 241 Lauer, D. G., 254,255,300 Law, J . H., 27, 44 LPwlor, G. C., 121, 178 Lawrence, P. J., 24, 42, 43 Lazdunski, G., 230,241 Lazzarini, R. A., 25, 42 Leadbetter, E. R., 2, 3, 7 , 14, 42 Leanz, G., 27, 42 Lecadet, M. M., 10, 11, 12, 42 Ledingham, G . A., 51, 116, 141, 142, 143, 148, 168, 170, 175, 177,178 Lee, J. A., 187,241 Lee, J. D., 49, 167 Lee, R. W., 268,300 Lee, T. C., 119, 171 Lee;W. H., 28, 42 Leegwater, D.-C., 67, 127, 130, 171 Lehle, L., 79, 82, 172, 176 Lehmann-Brauns, E., 295,298,299 Lehrian, D. W., 127, I 7 2 Leifson, E., 304, 308, 315, 320, 321, 358 Leloir, L. F., 79, 165 Leman, A., 3 2 , 4 2

Lemieux, R. U., 16, 172 Lenard, J., 59, 60, 177 Lengeler, J., 208, 241 Lennartz, W. J . , 79, 82, 172 Lenton, J. R., 154, 172 Lepidi, A. A., 117, 174 Lessie, P. E., 51, I72 Lester, R. L., 68, 73, 74, 85, 86, 87, 89, 91, 92, 94, 95, 96, 97, 103, 104, 105, 106, 112, 134, 165, 171, 172, 176, 177, 178 Lethbak, A., 251,298 Letnansky, K., 251, 292, 293,302 Letoublon, R., 82, 84, 172 Letters, R., 66, 172 Levitch, M. E., 339,359 Lewin, L. M., 68, I75 Lewis, D., 140, 176 Lewis, D. H., 51, 125, 139, 140, 145, 146, 147, 149, 170, 172, 187, 195, 198, 215,241 Lewis, J . C., 35, 42 Light, R. J., 76, 78, 107, 108, 109, 133, I68 Lin, H. K., 123, 129, 142, 143, 149, 150, I72 Lindeberg, G., 153, 172 Lindeberg, M., 153, 172 Lindegren, C. C., 260,300 Lindegren, G., 260,300 Lingappa, Y., 122, 172 Linnane, A. W., 105, 169 Linnett, P. E., 2 1, 24, 44 Lloyd, G. I., 133, 172 Lochmann, E.-R., 294,300 Loeb, L. A., 273,300 Loeblich, L.A., 217,241 Loginova, N., 338,358,359 Lohr, D., 253,300 London, J., 355,359 Long, P. E., 137, 172 Longley, R. P., 66, 69, 172 Loomis, W. E., 98, I 7 2 Losel, D. M., 48, 51, 66, 67, 85, 95, 102, 116, 125, 131, 139, 140, 142, 145, 146, 147, 149, 150, 166, 172, 174 Lovett, J . S., 51, 172 Lowry, R. J., 51, 172 Lucas, J. J., 79, 172 Luderitz, O., 99, 178 Lumsden, R. D., 148, I 7 2

AUTHOR INDEX

37 1

Marshall, M. O., 102, 171 Martin, A., 318, 352,357 Martin, B., 160, 166 Matches, J. R., 26, 42 Mateles, R. I., 338,356 Mathiesen-Kaarik, A., 156, 157, 173 Matile, P., 55, 56, 57, 96, 173 M Matile, P. L., 247, 251,300 Matsubara, I., 132, 177 McCarty, P. L., 332,359 Matz, L. L., 4 , 5 , 4 3 McClain, R. C., 7 7 , 173 Maurice,A., 78, 165 McClary, D. O., 251,300 Maxwell, D. P., 56, 165 McConnell, W. B., 116, 142, 175 May, R., 292,300 MacElroy, R. D., 219,240 Measures, J. C., 187,241 McGee, E. E. M., 146, 170 Meers, J. L., 187, 241 Machlis, L., 115, 172 Megnet, R., 233,241, 295,299 McIntosh,A. F., 136, 173 Mehta, R. J., 334, 335,358 McKechnie, C., 257, 258,301 McKeen, W. E., 122, 123, 128, 138, 141, Meiberg, J. B. M., 313, 337,358 Melhuish, J . H., 153, 173 172, I73 Mendozza, C. G., 69, 173 MacKenzie, A. P., 35, 43 Mercer, P. C., 51, 127, 173 Mackey, B. M., 3, 6, 42 Merdinger, E., 68, 7 7 , 173 McLachlan, J., 217, 220, 226,240 Mevius, W., Jr, 304, 31 1, 314, 322, 344, McLaughlin, C., 284,300 358 McLean, R. J., 98, 172 Mian, F., 276, 278, 279,301 McMahon, D., 98, 172 Michalenko, G. O., 50, 173 MeMorris, T. C., 115, 172 Michel, G., 24, 42 McMurrough, I., 288,300 Michell, R. H., 72, 169 McNabola, S. S., 138, 144, 178 Michell, R. M., 105, 178 Maeda, Y., 33, 42 Micita, L. K., 25, 40 Magnall, D., 72, 73, 102, 106, 172, 173 Miller, J.J., 136, 164, 173 Maheswari, R., 67, 174 Miller, L. K., 197, 241 Maina, G., 117, 166 Miller, W., 162, 179 Maister, H. G., 131, 172 Miller, W. L., 142, 173 Maitra, U., 280, 298 Mills, G. L., 77, 79, 124, 166, 173 Mandel, M., 309,358 Mims, C., 161, 167 Mandy, W. J., 334,358 Mims, C. W., 123, 173 Manocha, M. S., 128, 151, 173 Minckler, S., 251, 260,300 Mantle, P. G., 144, 173 Minnikin, D. E., 71, 72, 170, 173 Marak, J., 245, 247, 296, 297,300 Mirocha, C. J., 144, 175 Marak, J., 21, 43 Mishra, N. C., 113, 173 March, J. B., 61, 178 Misra, P. C., 232, 241 Marchant, R., 5 1, 173 Mitani, T., 10, 11, 43 Markus, K., 124, 167 Marmur, J., 254, 255, 256, 258, 260, 271, Mitchell, D. T., 144, 167 Mitchell, J. E., 50, 170 297,298,299 Mitchell, N., 122, 138, 141, 172 Marquis, R. E., 33, 38, 41, 43 Mitchell, N . L., 123, 173 Marrk, E., 217, 219, 241 Mitchell, R., 312, 3 5 8 Marschke, C. R., 15, 4 2 Marshall, B., 118, 170 Mitchison, J. M., 261, 300 Marshall, B. J., 32, 33, 42 Miuro Santo, L. Y., 123, 167 Marshall, K. C., 309, 312,356,358,359 Mlodecki, H., 68, 173 Lundgren, D. G., 5, 26, 42 Lutsdorf, U., 233, 241 Lynen, F., 105, 171 Lyon, I., 161, 171

372

AUTHOR INDEX

Moens, P. B., 245, 247, 249, 300 Mogelson, J., 3 1, 43 Molenaar, I., 2 9 3 , 3 0 1 Moll, R., 2 5 3 , 3 0 0 M o o r , H., 247, 2 5 1 , 3 0 0 Moore, D. T., 212, 4 0 Moore, R. L., 306, 309, 318, 320, 321, 322,328,358 M o r , J. R., 288, 300 M o r e n o , R. E., 149, 171 Morita, S., 350,3357 MorrP, P.J., 49, 50, 52, 57, 169 Morris, E. O., 133, 1 7 2 Morris, J. G . , 3, 6, 4 2 Morris, L. J., 101, 144, 173 Mortimer, R. K., 249, 250, 254, 255, 262. 299,300 M o r t o n , J. G . , 282, 297 Mosbach, K., 154, 173 Mosse, B., 139, 141, 173 Mounts, T. L., 26, 41 Muckerjee, K. L., 144,173 M u d d , J. B., 108, 173 Muhlethaler, K . , 57, 169 M u l d o o n , J . J., 2 6 4 , 3 0 0 M u m m a , R. O., 65, 66, 67, 166, 173 Mummery, R. S., 119, 178 Munday, K. A., 187, 240 Murphy, J . A., 7, 43 Murray, R. G. E., 21, 43 Murrell, W. G., 2, 6, 7 , 14, 15, 18, 26, 27, 32, 33, 35, 36, 38, 39, 40, 42, 43, 44 Muscatine, L., 140, 176 Muthukrishnan, S., 2 8 4 , 3 0 0 Myers, A. T., 30, 4 1 Myers, J. S., 2 3 3 , 2 4 1

N Naccareto, W. F., 94, 178 Nachbaur, J., 66, 178 Nakanishi, S., 230, 241 Nakashio, S., 15, 4 2 Nakatani, T., 21, 22, 4 0 Nakayama, T. 0. M., 119, 179 Nakazoto, H., 2 8 4 , 3 0 0 Naliboff, J. A., 262, 263, 299 Napias, C., 78, 165 Naveke, R., 3 0 4 , 3 1 4 , 3 4 0 , 3 4 4 , 3 5 8 Nawlek, J . M., 251, 2 5 2 , 2 9 8 Neihof, R., 33, 43

Nelson, D. L., 25, 28, 29, 32, 40, 42, 43 Nes, W. R., 60, 173 Netrusov, A. I., 3 5 0 , 3 5 8 Newaz, S. S., 341, 3 4 6 , 3 5 8 Newlon, C. S., 254, 266, 270, 27 1 , 3 0 0 N g , A . , 136, 173 Nickerson, K. W., 26, 41 Nickerson, W. J., 49, 50, 162, 166, 175 Nicolson, G. L., 60, 64, 174, 176 Niederpreum, D. J . , 51, 123, 129, 137, 164, 167, 169 Niel, C. B., van, 3 0 4 , 3 5 9 Nishi,A., 129, 174 Nobel, P. S., 182, 241 Noll, M., 2 5 2 , 3 0 0 Norkrans, B., 212, 241 Norris, D. M., 157, 168, 171 N o r m a n , J., 116, 174 Northcote, D. H., 69, 83, 174, 175 Nowak, R., 142, 149, 150, 174 Nurminen, T., 64, 66, 69, 79, 89, 91, 94, ~ 9 8 174,176,177 , Nuti, M . P., 117, 174 Nutting, W. H., 115, 1 7 2 Nyc, J . F., 70, 72, 104, 113, 167, 175 Nygaard, 0. F., 2 6 4 , 3 0 0

0 O’Brien, J. S., 77, 78, 168 O’Connor, M . L., 3 4 6 , 3 5 8 Oda,T., 80, 81, 90, 131, 174 Oeding, V., 3 4 9 , 3 5 8 Ogata, K . , 3 3 , 3 5 7 Ogiso, T., 134, 174 O g u r , M., 2 5 1 , 3 0 0 O g u r , M . S., 2 6 0 , 3 0 0 Ohye, D. F., 6, 7, 14, 15, 18, 38, 43, 44 Onishi, H., 197, 198, 207, 212, 213, 227, 24I Orcival, J., 139, I74 O r d a l , E. J., 3 2 9 , 3 5 8 O r d a l , Z. J., 3, 26, 28, 31, 42, 43 Orndorff, G. R., 35, 43 Ortiz, J. M., 23, 40, 43 Osmond, C. B., 236, 241 O’Sullivan, J., 67, 116, 128, 150, 174 Ou, L.-T., 38, 43 O u r a , E., 2 3 2 , 2 4 1 @yen, R. B., 2 5 8 , 2 9 8

AUTHOR INDEX

@yen, T. B., 258, 259, 2 9 7 , 3 0 0 Ozaki, H., 80, 8 5 , 171

P Paduch, V., 2 7 3 , 2 9 8 Page, R. M . , 119, 174 Paleg, L. G., 187, 2 4 1 Palevitz, B. A . . 139, 168 Paltauf, F., 71, 73, 106, 170, 174 Pankratz, H. S., 3 , 6 , 7 , 12, 4 0 Park,D., 1 1 7 , 176 Parkin, E. A . , 158, 174 Parodi, A. J., 79, I65 Pastan, I . , 230, 2 4 1 Pate, J. L., 3 2 9 , 3 5 8 Patel, R. N., 3 3 4 , 3 3 5 , 3 58 Patton, S . , 6 0 , 127, 174 Paulus, H., 105, 1 7 4 Pearce, S . M., 3 6 , 43 Peat, A , , 4 9 , 5 8 , 174 Peck, R. L., 160, 1 7 4 Pedersen, T. A. 116, 160, 174 Peel, D., 3 1 9 , 3 3 8 , 340, 3 4 4 , 3 5 8 Pelcher, E. A . , 3 1 , 43 Penman, S., 278, 2 9 4 , 3 0 0 Perdue, J. F., 75, 177 Perlman, S., 294, 300 Peterson, J. B., 244, 253, 2 9 9 Peterson, N. S., 2 8 4 , 3 0 0 Peterson, W. H., 80, 166 Petes, T. D., 254, 255, 266, 267, 268, 270, 27 1,300

Petitdemange, H., 2 7 , 4 0 Peveling, E., 139, 154, 174 Peyton, M. P., 15, 43 Pfeifhofer, A . O . , 222, 2 4 1 Pfennig, N., 28, 43, 3 3 8 , 3 5 8 Pheil, C. G., 26, 43 Phillips, S. L., 284, 2 9 9 Pierce, D. A . , 134, 165 Pieringer, R. A , , 9 7 , 174 Pillai, A , , 138, 176 Piovant, M., 230, 2 4 1 Pisano, J . J., 15, 43 Pisano, M . A . , 161, 167 Pitel, D. W., 24, 43 Pitt, J. I., 197, 215, 2 4 1 Planta, R. J., 278, 297 Plattner, J. J., 115, 174 Pockett, H. V., 11, 14, 44

373

Pongratz, E., 3 0 6 , 3 0 9 , 358 Ponta, H., 273, 274, 276, 2 7 8 , 3 0 0 Ponta, U., 273, 274, 276, 2 7 8 , 3 0 0 Posternak, T., 9 6 , 168 Porter, J. S . , 329, 3 5 8 Powell, D. A . , 9 8 , 174 Power, D. M., 9 6 , 174 Pratt, I., 24, 44 Price, R., 2 7 8 , 3 0 0 Priestley, C. A , , 146, 174 Prince, H. N., 1 1 7 , 1 7 4 Pringle, J . R., 248, 265, 294, 2 9 9 , 3 0 0 Pringsheim, E. G., 3 0 9 , 3 5 8 Prostenik, M., 80, 88, 9 1 , 9 5 , 131, 167, 174, 178

Prottey, C., 6 8 , 174 Prouty, C., 3 0 4 , 3 5 8

0 Quayle, J . R., 318, 319, 3 3 0 , 3 3 1 , 3 3 3 , 335, 338, 340, 3 4 1 , 3 4 2 , 3 4 4 , 3 4 5 , 347,354,355,356,357,358,359

R Rabinowitch, S., 222, 2 4 1 Rabinowitz, M., 6 6 , 72, 1 6 9 , 1 7 0 , 271, 2V8

Radin, N. S . , 106, 177 Raetz, C. R. H., 103, 174 Raj, H. G., 123, 174 Raju, K. S., 6 7 , 1 7 4 Ramachandran, S., 123, 129, 135, 143, 166, 1 6 9

Rambo, G. W., 122, 174 Raper, J. R., 115, 174 Raper, K. B., 116, 1 7 0 Rapoport, H., 115, 172, 174 Rapport, G., 245, 2 4 7 , 2 4 9 , 3 0 0 Rast, D., 5 0 , 5 1 , 5 7 , 116, 117, 128, 137, 1 6 5 , 169, 1 7 3 , 174, 176

Ratledge, C., 130, 131, 177 Rattrey, J. B. M., 102, 174 Record, R. R., 22, 43 Rees, D. C., 86, 8 9 , 9 1, 9 2 , 9 5 , 9 6 , 1 7 2 Rees, T. A . , 2 3 6 , 2 4 1 Reid, B. J., 248, 294, 2 9 9 Reindel, F., 76, 1 7 4 Reisner, H., 142, 175 Reisner, H. J., 116, 142, 145, 175

374

AUTHOR INDEX

Reiss, E., 162, 175 Reithel, F. J., 194, 240 Reithetman, R. W., 99, 168 Rheinheimer, G., 312, 314,357 Ribbons, D. W., 338,358 Ribi, E., 3, 4, 41 Richardson, M., 283, 297 Richmond, D. V., 122, I68 Riemersma, J. C., 75, 175, 232,241 Riggs, A. D., 266, 299 Rihova, L., 208,240 Ris, H., 244, 253,299 Rittenberg, S. C., 10, 11, 12, 14, 17, 41, . 44 Rizza, V., 113, 175 Robb, J., 139, 175 Robbins, W. J., 116, 175 Robinow, C. F., 28, 43, 245, 247, 251, 296, 297,300 Robinson, P. M., 117, 169, 175 Rode, L. J., 2, 4 , 5 , 6, 31, 43, 45 Rodriquez, J., 149, 171 Roe, J., 87, 95, 166, 175 Roeder, R. G., 278,301 Rogovin, S. P., 131, 172 Rogolsky, M., 24,42, 43 Rohringer, R., 142, 149, 150, 174 Romanova, A. K., 340,357 Rosbash, M., 283, 294,297,300 Rose, A. H., 50, 51, 61, 66, 69, 102, 105, 125, 170, 172, 178, 207, 210, 240, 288,300 Rose, D., 210, 21 1, 241 Roseman, S., 84, 88, 91, 99, 112, 165, 171, 175 Rosen, S. D., 99, 168 Rosenberg, A., 108, 175 Rosenthal, R., 90, 175 Ross, R. F. A., 32, 43 Rostrepo, A., 176 Roth, R. M., 294,297 Rothman, S. R., 176 Rouser, G., 66, 67, 170 Rownd, R. H., 278,298 Roxburgh, J. M., 333,335,357 Royan, S., 295,301 Rozijn, T. H., 25 1, 292, 293,301 Rubin, G. M., 259, 260,301 Ruinen, J., 132, 176 Rullman, W., 303,359 Rusch, H. P., 118, 168

Russell, A. E., 196, 214,240 Russell, D. W., 122, 166 Rutter, W. J., 274, 275, 276, 217, 278, 279, 280,297,301 Ryter,A., 17, 37, 41

S St. Amant, P. P., 332,359 Sacks,L. E., 18,31,34, 40,43,44 Sadoff, H. L., 25, 41, 43 Sahasrabuddhe, C. G., 252,301 Saito, H., 25, 43 Sajdel-Sulkowska, E. M., 292, 293,301 Sakakibara, Y., 25, 43 Salem, A. R., 355,359 Salkin, L. F., 164, 176 Samsonoff, W. A . , 3, 43 Sanders, F., 139, 141, 152, 167 Sano, K., 15, 16, 42, 43 Santarius, K. A., 234, 240 Sargent, J . A . , 50, 138, 177 Sastry, P. S., 67, 78, 108, 174, 175 Sauer, H . W., 118, 175 Scandella, C. J., 26, 43 Scarborough, G. A , , 64, 104, 175 Schaffner, G., 56,57, 175 SChdtZ, G., 65, 73, 175 Schatzmann, H. J., 78, 175 Scher, M . G., 79, 172 Scherrer, R., 4, 7, 28, 41, 43 Schiebeci, A , , 102, 174 Schindler, D., 293,301 Schipper, A. C., 144, 175 Schisler, L. C., 67, 121, 127, 150, 170, 172, I 7 8 Schlegel, H. G., 349,358 Schmidt, E. W., 138, 175 Schmidt, J. M., 305,359 Schmidt-Nielsen, K., 197, 241 Schneider, E. G., 103, 113, I75 Scholfield, C. R., 81, 176 Scholtz, K., 294,299 Schultz, L. D., 273, 276, 277,297,301 Schwartzenbach, A. M., 56, 175 Schweitzer, E., 251, 257, 258, 268, 298, 301 Scopes,A. W., 260, 273,302 Scott, K. J., 140, 175 Scott, W. J., 182, 197, 215,241 Scribner, H. E., 31, 43

AUTHOR INDEX

Sebastian, J., 260, 273, 276, 278, 279, 291, 292,298,299,301 Seidman, M. M., 68, 174 Seitsma, J. H., 50, 175 Sekura, R. D., 65,67, 175 Selva,A., 117, 166 Semeriva, M., 134, 175 Senior, P. J., 349,359 Sentandreu, R., 83, 175 Sentenac, A., 273, 276, 280, 297, 298, 299 Serafin, F. G., 123, 167 Servettaz, O., 217, 219,241 Setlow, P., 17, 26, 28, 29, 43, 44 Setlow, R. B., 33, 41, 121, 176 Sevaco, C., 27, 40 Seymour, R. C., 152, 175 Shafai, T., 68, 175 Shah, R. G., 310,359 Shankaran, R., 123, 174 Shapiro,A. L., 164, 175 Sharma, C. B., 82, 176 Sharma, R., 124, 138, 176 Shatkin, A. J., 284, 297,300 Shaw, B. R., 252,301 Shaw,M., 139, 140, 144, 173, 175, 176 Shaw, N., 97, 108, 176 Shaw, R., 49, 127, 131, 176 Shepherd, C. J., 123, 129, 176 Sherman, F., 257,298 Sherr, S. I., 73, 104, 176 Shimoda, C., 116, 177 Shishkina, V. N., 305, 310, 314, 317, 318, 319,338,358,359 Short, J., 12, 44 Shortland, F. E., 162, 166 Siddiqui, B., 99, 169 Sillevis-Smitt, W. W., 293, 301 Silva, M. T., 3, 44 Silver, S., 31, 41, 43 Silverstein, Z., 25, 40 Simpson, D. K., 4, 5, 45 Simpson, J. R., 186, 188, 189, 190, 191, 192, 193, 194, 198, 200, 226, 240, 24 I Singer, S. J., 60, 64, 174, 176 Singh, N., 120, 176 Sing, T. N., 187, 241 Sing Klar, A. J., 136, 177 Sison, Y., 96, 168 Skerman, V. B. D., 305,310,359

375

Skucas, G. P., 176 Slater, M. L., 264, 287, 293, 295, 296,301 Slepecky, R.A., 27, 30, 31, 41, 44 Smiganic,A. M., 164, 175 Smith, A. J., 355,359 Smith, D., 140, 176, 251, 292, 293,302 Smith, D. C., 187, 195, 198, 215,241 Smith D. G., 65, 176 Smith, G. G., 56, 178 Smith, J. E., 114, 118, 120, 133, 136, 137, 168,172, 173, 176 Smith,J. S., 197, 241 Smith, R., 122, 138, 141, I72 Smiths., 85, 87,92, 94, 99, 167, 176 Smith, S. W., 86, 87, 89, 91, 92, 94,95,96, 172, 176 Snell, E. E., 68, 158, 169, 175 Snell, N., 3 1, 35, 38, 40 Snell, N. S., 35, 42 Sols, A., 208, 210, 226, 233, 240, 241 Somerville, H. J., 10, 11, 12, 14, 17, 41, 44 Sora, S., 251, 298 Sorokin, H. P., 57, 176 Sousa, J. C. F., 3, 44 Spain, J. C., 341, 342,358 Speer, H. L., 219,240 Spencer, J. F. T., 67, 74, 76, 77, 127, 130, 169, 171, 176, 177, 198, 227,241 Sperl, G. T., 304, 305, 306, 312, 314, 315, 317,331,334,335,359 Spiro, T. G., 34, 44 Sproston, T., 121, I76 Spudich, J. A,, 11, 16, 25, 28, 29, 32, 40, 42, 43, 44 Sripati, C. E., 284,301 Staley, J. T., 329,359 Stanacev, N. Z., 80, 131, 174, 176 Stanier, R. Y., 304, 355,359 Staples, R. C., 123, 142, 143, 176 Starr, M. P., 305, 310,359 Stastna, J., 39, 41 Stauble,E.J., 116, 117, 174, 176 Stavely, J. R., 138, 176 Steenis, P. J., 330,357 Steinberg, W., 25, 44 Steiner, M. R., 68, 87, 103, 104, 106, 172, 176, 178 Steiner, S., 73, 85, 87, 92, 94, 97, 176 Stendrup, A., 251,298 Stephen-Olejniuad, B., 68, 173

AUTHOR INDEX

376

Stevens, R. J., 1 1 7 , 176 Stevenson, I . L., 51, 176 Stewart, G. R., 187, 241 Stewart, J. W., 257, 298 Stewart, P. R., 294,301 Stiers, D. L., 56, 178 Stiller, R. L., 80, 85, 88, 91, 178 Stocks, D. L., 122, 178 Stodola, F. H., 74, 76, 81, 131, 172, 176 Stokes, J . E., 338,359 Stoll, A . , 144, 176 Stone, K. J., 79, 176 Stone, W. B., 165, 175 Storm, E., 255,256, 260, 271,298 Stout, R., 312,358 Strandberg, J. D., 138, 144, 178 Strandberg, J. O., 150, 176 Strange, R. E., 22, 23, 44 Streit, W., 51, 126, 170 Stretton, R. J . , 49, 167 Strohbach, D. R., 92, 95, 167 Strsm, T., 338,359 Strominger, J. L., 6 , 18, 19, 20, 21, 24, 36, 39, 40, 42, 44

Struhl, K . , 285, 286,301 Stubbs, J. M., 13, 14, 17, 4 2 Stumpf, P. K., 100, 176 Stutzer, A., 303, 3 0 8 , 3 5 9 Subcrop, K. F., 51, 176 Subramanian, M. K., 295,301Suggs, F. G., 134, 165 Sugiura, M., 134, 174 Sugiura, Y., 33, 42 Sulston, J . E., 259, 260,301 Suomalainen, H., 64, 66, 69, 79, 89, 91, 94, 96, 98, 174, 176, 177, 232, 241

Surben, R. E., 67, 172 Sussman, A. S., 51, 114, 122, 130, 172, 177

Suzuki, K., 25, 44 Suzuki, T., 69, 132, 177 Svaniszlo, P. J., 161, 167 Svoboda, A., 6 5 , 176, 224,240 Sweeley, G. C., 72, 74, 75, 76, 100, Swift, H., 72, 170

T Takada, N., 305,310,359 Takano, I . , 279,301 Takao, N., 116, 177

Talwalkar, R. T., 73, 74, 105, 177 Tamaki, H., 251,301 Tanaka, K . , 132, 177 Tani, Y., 333,357 Tanner, W., 79, 82, 93, 165, 171, 172, 176, 177

Taskinen, L., 64,98, 174 Tatum, E. L., 113, 124 Taylor, A., 122, 166 Taylor, I. J., 349,359 Tempest, D. W., 187,241 Terano, H., 25, 44 Ter Louw, A . L., 27,45 Tesone, C., 25, 4 0 Thielke, C. N., 128, 177 Thomas, B., 123, 169 Thomas, D. M., 115, 177 Thompson, J. K., 3 3 , 4 3 Thompson, R. O . , 1 2 , 4 4 Thonart, P., 276,301 Thorn, J . A . , 76, 77, 78, 166, 169, 172 Thorpe, R. F., 130, 131, 177 Thornberry, G . D., 123, 125, 178 Tiboni, 0 ., 25 1,298 Tingle, M., 136, 177 Tipper, D. J., 6 , 15, 18, 20, 21, 24, 42, 44 Tochikura, T., 80, 85, 171 Tommerup, I. C., 50, 138, 177 Tonge, G. M., 334,351,359 Tonino, G.J. M., 251,292,301 Tono, H., 2 5 , 4 4 Tonolo, A , , 144, 173 Topiwala, H. H., 330,331,359 Torriani, A . , 25, 4 0 Trevelyan, W. E., 66, 94, 177 Trezzi, F., 219, 241 Trinci, A . P. J., 118, 177 Trione, E. J., 122, 177 Trotsenko,Y. A., 305, 310,314, 317, 318, 319,338,358,359

Trotter, M. D., 163, 165 Truesdell, L. C., 51, 58, 166 Tsai, K.-H., 59, 60, 178 Tsay, Chen, G., 82, 166 Tschierpe, H. J., 137, 177 Tseng, T-C., 134, 177 Tsuji, K., 333, 357 Tsuji, S., 25, 44 Tucker, A . N., 113, 175 Tulloch, A. P., 76, 77, 141, 142, 143, 148, 169, 177

AUTHOR INDEX

Turian, G., 119, 126, 135, 136, 166, 177 Turner, R. S., 98, 177 Tyler, P.A., 309,356,359 Tyorinoja, S., 79, 89, 91, 94, 96, 177 Tyrrell, D., 48, 85, 95, 102, 116, 131, 142, I66

U Uchida, A., 15, 42 Udem, S., 260,301 Uesugi, S., 75, 177 Ujita, S., 25, 44 Ullman, M. D., 106,177 Uribe, L., 165, 175 Utter, M. F., 346,359

V Vadas, R. L., 217,219,241 Valadon, L. R. G., 49, 119, 120, I 7 8 Valenzuela, D., 274, 275, 276, 277, 278, 301

Valenzuela, P., 275, 279, 280,297 Valk, E., 1 1 1, 165 Vanderkelen, B., 66, 67, I69 Vandermolen, G. E., 50, I68 Van Dijken, J. P., 333,359 Van Etten, J. L., 143, 178 Van Holde, K. E., 252, 253,300,301 Vary, J. C., 17, 22,33, 42, 44 Vasil’eva, Z. A., 340,357 Vaughan, M. H., 284,300 Veldkamp, H., 331, 355,358,359 Velez, H., 165, 175 Venkitasubramanian, R. A., 123, 174 Verachtert, H., 236, 240 Verkhoturov, V. N., 350,358 Vichmer, E. E., 161, 171 Vieira, J. R., 164, 178 Vignais, P. M., 66, 178 Vignais, P. V., 66, 178 Villaneuva, V. R., 69, 173 Vincent, W. S., 284,299,302 Vinter,V., 16,23,36,39, 41, 44 Viswanathan, L., 123, 174 Vlak, J. M., 293,301 Vogel, F. S., 51, 128, 178 Von Meyerburg, H . G., 260,297,301 Von Sydow, B., 146, 178 VranP, D., 289,301

377

w Wadzinski,A. M., 338,358 Waechter, C. J., 79, 85, 87, 92, 94, 104, 173, 176, 178 Wagner, C., 335,339,355,358,359 Wagner, H., 80, 87, 89, 90, 92, 178 Wage, R. G., 26, 41 Walker, D. A,, 226, 242 Walker, G. C., 161, I65 Walker, H . W., 26, 42 Walker, P. D., 6, 12, 21, 44 Walker, R. F., 123, 178 Walker, T. A., 236,242 Walkinshaw, C. H., 123, 171 Wallach, D. F. H., 178 Wallis, 0. C., 229, 242 Walsh, H.A., 161, 165 Waltho, J. A , , 200,240 Wardle, K. S., 127, 178 Warner, J. R., 260, 283, 284, 299, 300, 301 Warren, L., 61, 178 Warth, A. D., 6, 14, 15, 18, 19, 20, 21, 22, 23, 24, 27, 33, 35, 36, 38, 39, 43, 44 Watabe, K., 10, 44 Watson, K., 66, 67, 73, 165 Weaver, R. F., 51, 128, 178 Weber, D. J., 51, 116, 122, 123, 128, 129, 139, 142, 165, 169, 170, 171, 178 Weber, D. P., 114, 120, 121, 122, 178 Webster, J., 118, 124, 178 Weete, J. D., 48, 58, 60, 69, 100, 102, 109, 114, 120, 121, 122, 123, 131, 148, 149, 163, 167, 171, 178 Wegener, W. S., 137, 167 Wegman, 0. C., 220,226,242 Weinberg, F., 274, 275, 276, 277,278, 301 Weinberg, R., 294,300 Weiner, R. M., 322,359 Weinert, M., 80, 178 Weinmann, R., 278,301 Weinstein, D. B., 61, 178 Weisrock, W. P., 309, 310,357 Weiss, B., 80, 85, 88, 9 1, I78 Weitkamp,A. W., 164, 175 Wells, G . B., 86, 89, 91, 92, 95,96, 172 Wells, W. W., 94, 178 Westphal, O., 99, 178 Wheeler, B. E. J., 51, 125, 138, 170

378

AUTHOR INDEX

Wheeler, G. E., 105, 178 White, D. C., 113, 175 White, G. L., 66, 104, 106, 178 Whittaker, P. A., 229, 242 Whittenbury, R., 310, 320, 321, 322, 323, 324,359 Wicken,A. J., 97, 179, 188, 194,239 Wickerhan, L. I., 8 I , 131, 172, 176 Wickner, R. B., 295,301 Wickus, C. G., 21, 24, 36, 44 Widdowson, D., 334,351,359 Wiky, W. R., 64, 178 Wilhelms, 0. H . , 99, 178 Wilkinson, T. G., 3 18, 330, 33 1,359 Williams, M. W., 115, 172 Williams, P. H., 51, 138, 144, 178 Williams, W. L., 158, 178 Williamson, D. H., 260, 264, 266, 267, 268, 27 1, 273,300,301,302 Willoughby, L. G., 124, 167 Wills, C., 234, 242 Wilson, A. T., 338,356 Wilson, C. L., 56, 152, 165, 178 Windle, J. J., 34, 43 Wintersberger, E., 253, 271, 272, 273, 274, 276, 278,299,300,302 Wintersberger, N., 27 1 , 3 0 2 Wintersberger, U., 249, 251, 252, 25.5, 271, 272,292, 293,302 Wirth,T. C., 161, 179 Witte, I., 295, 299 Wolf, F. T., 118, 179 Wolf, W., 272, 273,299 W o o d , D . A . , 13, 14, 16, 17, 28, 44 Wood, R. K. S., 51, 58, 127, 139, 169, 173 Wood, W. A., 235,240 Woodhead, S., 236,242 Woodruff, W. H., 34, 44 Woodson, J . , 339,356 Woser, N. A,, 159, 179 Woutern, J . T. M., 50, 175 Wright, P. L., 99, 171

Wu, S., 82, 166 Wyatt, P. J., 33, 44 Wyers, F., 280, 299 Wyckoff, R. W. G., 27, 45 Wynn, W. R., 123, 142, 143, 176

Y Yamamoto, H., 119, 179 Yamamoto, S., 64, 171 Yanagishima, N., 116, 177 Yanagita, T., 123, 179 Ycas, M., 284,299,302 Yen, P. H., 98, 179 Yokiel, R. A., 122, 143, 170 Yokoyama, H., 119, 179 Yolton, D. P., 4, 5, 45 Youatt, J., 50, 179 Youatt, Y., 118, 179 Young, E., 2, 16, 17, 18, 28, 36, 31, 38, 41 Young, I. E., 26, 41 Younger, J., 20, 21, 41 Youngs, C. G.,67, 127, 130, 171

Z Zahler, P. H., 178 Zalokar, M., 135, 179 Zatman, L. J., 308, 317, 333, 334, 335, 336,338,356,357,358 Zavarzin, G. A., 304, 305, 306, 308, 310, 311, 315, 322, 323, 327, 340, 357, 359 Zellner, J., 80, 90, 168, 169, 179 Ziegler, E., 145, 179 Zinker, S., 255, 256, 260, 27 1,298 Zofcsik, W., 87, 89, 90, 92, 178 Zonnenweld, B. J. M., 50, 179 Zweig, G., 190, 240 Zylber, E., 294,300 Zytkovicz, T. H., 39, 45

SUBJECT INDEX

A

Albugo candida, lipids and, 138 Alcohol dehydrogenase in petite mutants, 234 in yeast, glycerol and, 233 Algae halophilism, 183 osmoregulators in, 18 7 physiology, salt relations and, 2 19-223 Allomyces spp. lipids in, 5 0 , 5 1 , 124 sirenin in, 115 Allomyces arbuscula, lipids in, 50 Allomyces javanicum, carotenoid content,

Acanthamoeba castellanii, aggregation, glycosphingolipids and, 99 Acarenoic acid, 154 Acetokinase, 349 N - Acetylglucosaminidase in Bacillus spores, 22 bacterial spore cortex lysed, 23 N-Acetylmuramyl- L-alanine, biosynthe. sis in spore cortex, 24 N-Acetylmuramyl-L-alanineamidase in Bacillus spores, 22 Achlya spp., lipid reserves, 122 Achlya bisexualis,antheridiol in, 115 Acinetobacter sp., methane oxidizers, 330 Acrasiales, aggregation, glycosphingolipids and, 100 Acridine orange in yeast nuclei staining,

119

Alternaria spp. asexual spores, lipid content, 122 morphogenesis, lipid reserves in, 12 1 Alternaris tenuis, spores, surface lipids, 122 Amanita muscaria ceramides in, 80, 85 glycosylceramidesin, 90 sphingolipids in, 88 Amanita rubescens ceramides in, 80 sphingolipids in, 88 a-Amanitin, yeast polymerases and, 27 7-

295

Anyostelium leptosomum, aggregation, hormones and, 116 Aeciospores, lipids in, 5 1 Aeration, polyol production in xerotolerant yeasts and, 227, 228, 236 Agaricus bisporus basidia, lipids in, 128 basidiospores, fatty acid effect on, 1 16 carbon dioxide and, 137 ceramides in, 8 0 , 9 0 glycophosphosphingolipids in, 9 3 , 9 5 lipids, 5 0 , 5 1 phosphoglycerides in, 67 phosphatidylcholine, 68 sphingolipids in, 87,88 sporophores, lipids in, 127 sterols, 150 growth factors, 12 1 sugars in, 7 7 Agaricus clitocybe, sphingolipids in, 88 Agarinw tabescens, sphingolipids in, 88 Aggregation in Hyphomicrobium spp., 327

278

Ambrosial fungi, scolytid beetles and, 156, 157

Amino acids exosporia, appendages and coats, 5 in histones from Saccharomyces cereuisiae, 252

Hyphomicrobium spp., growth inhibition by, 319 in spore coats, 1 1 , 12, 13, 14, 2 6 , 2 8 y-Aminobutyric acid as osmoregulator, 187

Ammonium salts, Hyphomicrobium spp. growth and, 3 18 Amoeba aggregation, glycosphingolipids and, 99

379

SUBJECT INDEX

380

Amphotericin B, arabitol formation by yeasts and, 199 Ancalomicrobium, prosthecae in, 329 Anhydrocerebrin in fungal membranes, 84

Anobiid beetles, fungi and, 155, 159 Antheridial hyphae, lipids in, 50 Antheridiol, 115 Antibiotics, polyene, fungal growth and, 61

Aphanomyces eutiches, lipids in, 50 Apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 Appendages bacterial spores, 2 chemical composition, 4-7 morphology, 3,4 function, 6 Arabinose, apparent inhibitor constants of yeast isocitrate dehydrogenase, 193

Arab i to1 active transport by Saccharomyces rouxii, 209

apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 compatible solute, for Saccharomyces rouxii, 200 for yeasts, 204 distribution in non-tolerant yeasts, 222

in Saccharomyces rouxii, 222 in yeasts, 225 enzyme activity and, 189, 19n extrusion by Saccharomyces rouxii, 232 production by xerotolerant yeasts, aeration and, 227, 228 in yeasts, NADPH and, 236 retention factor in yeast, 202 Saccharomyces rouxii production, 235 in Zygosaccharomyces nectarophilus, 199 Acyria cinerea spores, germination, lipids, 123

Argosterol, fungal, sex hormone for Xyletorus sp., 157 Arsenates in membrane transport of fungi, 73 Arthroderma uncinatum, phosphoglycerides in, 67 Articulospora tetracladia, car0 tenoids, 120

Ascobolus sp., photomorphogenesis, 1 18 Ascoidea hylecoeti, ambrosial fungi and, 156

Ascospores, lipids in, 5 1 Aspartic acid as osmoregulator, 18 7 Aspergzllus spp. mannan in, 98 mannan-protein, biosynthesis, 82 pathogenicity, 164 Aspergillus bisporus, hydroxy acid glycosides in, 74 Aspergillus clavatus, polyols as food reserves, 187 AspergillusJavus conidia, germination, lipids and, 129 spores, germination, lipids, 123 Aspergillusfumigatus lipids in, 5 1 polyprenol-containing glycolipids in, 79

spores, germination, lipids, 123 Aspergillus giganteus, carotenogenesis, 1 18 Aspergdlus nidulans germination, lipids and, 129 lipids, 50 spores, germination, lipids, 123 Aspergillus niger ceramides in, 80,90 conidiation, isocitrate lyase and, 136 diglycosyldiglyceride biosynthesis in, 109

fatty acids in, 1 17 germination, lipids and, 129 glycolipids, biosynthesis, 100, 108 glycophosphosphingolipids in, 95 glycosylceramides in, 90 glycosyldiglyceridesin, 7 7 hydroxy acid glycosides, 74, 75, 76 2 - hydroxyoctadec- trans- 3- enoic acid, biosynthesis, 101 hpases in, 133, 134 phospholipids, 72 polyprenol-containing glycolipids in, 79,82

sphingolipids in, 86, 87, 88, 89 biosynthesis, 1 12, 1 13 spores, germination, lipids, 123 germination, nonanoic acid and, 116

nonanoic acid and, 1 17

SUBJECT INDEX

Aspergillus parasiticus, morphology, manganese and, 162 Aspergillus sydowi, ceramides in, 80 Asticcacaulis biprosthecum, prosthecae in, 329 ATP/ADP ratio, glycerol production and, 232,233,236 Attine ants fungi and, 155 fungus cultivation, 160 Aureobasidium pullulans-See Pullaria pulMans M-Avenasterol in Plasmodiophora brassica spores, 151 Axenic culture, biotrophic fungi, lipids, 143, 144

381

chemical composition, 8, 10 protein, 12 structure, 9 exosporia, chemical composition, 4 phospholipids, 5 water content, 32 sporulation, dehydration mechanism, 37 Bacillus coagulam lytk enzymes, 23 spore coats, amino acids, 13 morphology, 7 structure, 9 Bacillus fastidiosus spores, exosporium, morphology, 3 Bacillus lichmformis spore coats, chemical composition, 15 sulpholactic acid, 28 B Bacillus megaterium Bacillus alesti, spore coats, amino acids, 11 cell wall, 2 1 Bacillus apiarius, spore coats, structure, 9 exosporia, 6 Bacillus brevis, spore cores, sulpholactic lytic enzymes, 22 acid, 28 muramic lactam synthesis, 24 Bacillus brevis 636, spore coats, amino spore coats, amino acids, 13, 14 acids, 15 biosynthesis, 1 7 Bacillus cereus phosphorus, 15 cortex synthesis, 36 cores, amino acids, 28 germ cell wall, 2 1 enzyme protein, 29 lytic enzymes, 22, 23 proteins, 26 spores, carbohydrates, 6 cortex, chemical structure, 18 coats, amino acids, 11, 13, 14 exosporium, morphology, 3 biosynthesis, 16 formation, biochemistry, 23, 24 chemical composition, 15 lipids, 26 morphology, 7 ribosomes, 25 cortex, chemical structure, 18 storage polymers, 27 density, 33 Bacillus megaterium KM exosporium, morphology, 3 spore coats, chemical composition, 8 protein, 12 formation, peptidoglycan synthesis Bacillus polymyxa and, 23 lipids, 26 germ cell wall, 2 1 morphology, 2 spores, lipids, 26 ribosomes, 25 Bacillw sphaericus Bacillus cereus T cortex synthesis, 36 exosporium, protein, 12 peptidoglycan biosynthesis, 24 muramic lactam synthesis, 24 spores, coats, chemical composition, phospholipids, 7 1 15 spores, appendages, chemical comcortex, chemical structure, 18 position, 4 formation, biochemistry, 24 coats, amino acids, 1 1 peptide side chains, 2 1 biosynthesis, 17 walls, peptidoglycan, 20

SUBJECT INDEX

382

Bacillus stearothenophilus spores, coats, structure, 9 cortex, 38 chemical structure, 18 Bacillus subtilis genomes, 26 germ cell wall, 2 1 lytic enzymes, 22, 23 spores, carbohydrates, 6 coats, amino acids, 11, 13, 14 coats, biosynthesis, 16 chemical composition, 10 phosphorus, 15 cores, amino acids, 28 cortex, chemical structure, 18 exosporium, morphology, 3 formation, biochemistry, 24 peptidoglycan from, 19 ribosomes, 25 walls, peptidoglycan, 20 sporulation, low molecular-weight compounds in, 27 Bacillus subtilis 168, spore coats, protein, 12 Bacillus subtilis 36 10, phospholipids, 7 1 Bacillus thiaminolyticus, spore coats, chemical composition, 10 Bacillus thuringiensis lytic enzymes, 23 spores, coats, chemical composition, 10 coats, protein, 12 lipids, 26 protein crystals in, 2 Bacteria halophilism, 183 water activity tolerance, 183 Bacterial spore-See Spores Bacteroides melanogenicus, sphingolipid biosynthesis in, 113 Basidiomycetes phosphoglycerides in, 65, 67 sporophores, carotenoid pigmentation in, 119 Basidiospores germination, lipids and, 123 lipids in, 5 1 Behenic acid in Sphaerotheca humuli spores, 141

-

Beta vulgaris, nicotinamide nucleotide concentrations in, 234 growing in host tissues, lipid metabolism in, 144-155 Biotrophic fungi in axenic culture, lipids, 143, 144 lipids in, 138, 139, 140 spores, lipids and, 141-143 Blakeslea trispora carotene production, trisporic acid and, 117 trisporic acid in, 1 15 Blastocladiales, zoospores, lipids in, 124 Blastocladiella spp. carotene content, 119 glyoxylate-cycle enzymes in, 135 lipids in, 5 1 Blastocladiella emersonii glycosyldiglycerides in, 7 7, 78 zoospores, 58 lipids in, 124 Blastomyces spp., lipids, 161 Blastomyces dermatitidis cell-wall lipids, 149 hyphal walls, lipids, 163 lipids, 161 Blood cells, glycerol and, 197 “Blue stain” fungi, lipids and, 157, 158 Boletus variegatus isobutanol and isobutyric acid in, 153 respiration, fatty acids and, 116 Botryodiplodia theobrome, pathogenicity, 165 Botrytis cinerea, phospholipases in, 134 Botrytis fabae spores, surface lipids, 122 Brassicasterol in dermatophytes, 162 in Plasmodiophora brassica spores, 15 1 Bremia lactucae, lipids in, 50, 138 Brettanomyces truxellensis, phosphoglycerides in, 67 Budding in Hyphomicrobium species, 320 Budding yeast, 247,289 Bullera alba, phosphoglycerides in, 67 Butane-Z,S-diol, effect o n collagen, 196 Butyric acid, hydroxy-, in fungi, 11 7

SUBJECT INDEX

C Calcium in spores, 29,30 cores, 27 dipicolinic acid, 32 heat resistance and, 3 1 Campesterol in Plasrnodiophora brassica spores, 15 1 in rust urediospores, 142 in Uromycesphaseoli, 149, 150 Candida spp. lipid reserves, 130 phosphoglycerides in, 66 phospholipids, fatty acid content, 68 Candida albicans cell-wall lipids, 149 fatty acids, 16 1 hyphal walls, lipids, 163 lipids, 50 Candida bogoriensis glycolipids, biosynthesis in, 107, 108 secondary metabolites in, 133 hydroxyacid glycosides in, 76 sophorosides in, 78, 132 sterol glycoside biosynthesis in, 109 Candida intermedia, free bases in, 80, 85 Candida lipolytica lipases in, 133 phosphoglycerides in, 66 Candida macedoniensis, phosphoglycerides in, 66 Candida parapsilosis,phosphoglycerides in, 66 Candida slooji phosphoglycerides in, 67 respiratory deficient, phospholipids, 73 Candida utilis ceramides in, 80 glycerol permeability and, 208 glycophosphosphingolipids in, 92 glycosylceramides in, 90 lipids in, 50 membranes, phosphoglycerides in, 69 permeability to glycerol, 201 phosphoglycerides in membranes, 72 sphingolipids in, 87,89 Cantherellus infundibulformis, carotenoid content, 119

383

Carbohydrates in exosporia, 6 fungal membranes and, 6 1 in spore appendages, 6 Carbon dioxide fixation, ribulose diphosphate cycle, 338 fungal morphogenesis and, 137 /%Carotene in Blaheslea trispora, 155 fungal, biosynthesis, 119 Carotenoids, fungal morphogenesis and, 1 1 7-120 Catalase in Hyphomicrobium sp., 333 Cell growth, integration with nuclear/cell division, 286-292 Cell membranes fungal, lipids in, 50 permeability, phospholipid unsaturation and, 148 phosphoglycerides in, 75 Piptocephalus uirginiana, 152 Cell walls dimorphic fungi, lipids, 149 fungal glycophosphosphingolipids in, 98 lipids in, 48 phosphoglycerides in, 69 sphingolipids in, 96 Cellular aggregation, glycosphingolipids and, 98 Cephalosporium sp. fatty acids in, 1 17 neutral lipids, 157 phospholipids, 157 Ceramide-(P-inositol), in fungi, 95 Cerarnides acetylated, in fungi, 80, 8 1 in fungal membranes, 85-90 in fungi, 80, 8 1 glycosylation, 112 phosphorylated, in fungi, 80, 8 1 Ceratocystis sp. deliquescence, lysosomes in, 56 insects and, 157 Cerebrosides-See Glycosylceramides Cetraria islandica, protolichesterinic acid in, 154 Chemotaxonomy, fungi, lipids and, 49

SUBJECT INDEX

384

Chlamydomonas sp . gametes, agglutination, mannose and, 98

halofelicity, 2 16 Chlamydosores, lipids in, 5 1 Chlorella spp., NADP+/NADPH ratios in, 234

Chlorella pyrenoidosa, nitrate reductase, glycerol and, 196 Chloroplasts membranes, 59 water activity, 222 Choanophora curcurbitarum,Piptocephalis uirgniana on, lipid metabolism and, 151

Cholesterol in cell membranes, 60 fungal membranes, 6 1 in fungi, 120 in Plasmodiophora brmsica spores, 15 1 in rust urediospores, 142 Chromatin isolation from nuclei, 293 in yeast chromosomes, 252 Chromatographic tltvalue of polyols, enzyme inhibition and, 19 1 sugars, enzyme activity and, 192 Chromosome VII of Saccharomyces cereuisiae, genetic map, 269 Chromosomes yeast, 244,249-260 localization of genes on, 256-260 size, 254-256 Cladonia cristatella, lipid metabolism, 154 Cladosporium carrionii, lipids, 160 Cladosporium herbarum, fatty acids in, 1 1 7 Clauiceps purpurea, axenic culture, lipids, 143, 144

Cleistothecia, lipids in, 5 1 Clitocybe tabescm ceramides in, 80 glycophosphosphingolipids in, 95 glycosylceramidesin, 9 1 Clostridium spp. spores, appendages, morphology, 4 coats, amino acids, 14 exosporium, morphology, 3 morphology, 2 Clostridium bqermentans, spore coats, chemical composition, 10

-

Clostridium butyricum, spores, polysaccharide, 27 Clostridiumpasteurianum, exosporia, 6 Clostridium perfringens, heat resistance, ionic composition and, 3 1 Clostridiumsporogenes spores, coats, amino acids, 13 cortex, chemical structure, 18 Clostridiumtaeniosporum spores, appendages, chemical composition, 4 appendages, morphology, 4 exosporia, chemical composition, 4 Club root, lipids and, 138 Coccidioides immitis, lipids, 16 1 Collagen effect ofglycerol on, 19.6 thermal stability, glycerol and, 214 Colletotrichum coccodes, morphogenesis, lipid reserves and, 127 Colletotrichum lindemuthianum lipids in, 5 1 morphogenesis, lipid reserves and, 127 Compatible solutes, accumulation, regulation, 223-237 in eukaryotic micro-organisms, 18 1242

Conidia germination, lipids and, 123 lipids in, 5 1 Conidiation in Aspergillus niger, isocitrate lyase and, 136 coprinus sp., sterols, is0 Coprinus comatus, triglycerides in, 130 Coprinus lagopus germination, lipids and, 129 lipids in, 5 1 spores, germination, lipids, 123 Cordycepin, Saccharomyces cereuisiae sensitivity to, 294 Cortex bacterial cell, 1 bacterial spores, chemical structure, 18-22

morphology, 18 Cortical membrane, bacterial spores, 18 Cronartiumfusqorme aeciospores, stigmastadienol in, 149 lipids, 148 spores, germination, lipids, 123

SUBJECT INDEX

Cronartium ribicola, lipids and, 139 Cryptococcus spp., lipids, 161 Cunninghamella elegans spores, germination, lipids, 123 nonanoic acid and, 1 16 Cyclic depsipeptides in biotrophic fungi, 144 Cyclohexanetetrol as osmoregulator, 187 Cytochromes in Hyphmicrobium spp., 350, 35 1 Cytoplasm bacterial spores, 2 vegetative cells, fungal, lipids in, 50 Cytosol, water activity, 222

385

22-Dihydro-ergosterol in rust urediospores, 142 Dihydrosphingosine, acetylation, 11 1 Dihydroxyacetone phosphate acylation, 106 photosynthesis and, 226 Di-inositoldiphosphorylceramidein fungi, 86 Dimethyl formamide, enzyme inhibition, 188

Dimethyl sulphoxide, enzyme inhibition, 188 Dimethylamine Hyphomicrobiurn growth and, 3 13 in Hyphomicrobium nutrition, 314 Dimethylamine dehydrogenase, 33 7 D Dimethylamine mono-oxygenase, 336, 33 7 Daedalia unicolor, Xyphidia prolongata and, Dimorphic fungi, cell-wall lipids, 149 158 Daldinia concentrica, Xyphidia prolongata Dimorphism, pathogenic fungi, 162-163 Diphosphatidyglycerol and, 158 in fungi, 63 Debaryomyces hansenii, salt accumulation, biosynthesis, 105 212 membranes, 65 Dehydration, spore cores, mechanism, Dipicolinic acid 35-39 in spores, 34 Density, spores, 32, 33 calcium and, 32 Dermatophytes cells, 2 fatty acids and, 163 cores, 27, 29 lipids, 161 heat resistance and, 35,39 sterols, 162 Deuteromycetes, phosphoglycerides in, Disinfectants, bacterial spores resistance, 65,67 2 Diaminopimelate-ligase, spore pep- Division nuclear/cell, integration of cell growth tidoglycan biosynthesis and, 24 and, 286-292 Dictyostelium discoidium yeast nucleus, 247-249 cellular aggregation, glycosphingoDNA lipids and, 98 bacterial spores, 25 non-aggregating mutant, glycosphinenzymes, in yeast nucleus, 271-282 golipids and, 99 estimation, 294, 295 spores, germination, lipids, 123 Dielectric constant, enzyme activity and, from Saccharomyes cerevisiae, 253 mitochondrial, replication, nuclear con192 trol over, 27 1 Dielectric studies, spores, 33 repetitive, in yeast chromosomes, 244 Differentiation replication in Hyphomicrobium spp., 322 bacteria, 1 in yeast nucleus, 266-27 1 fungal, lipids in, 114 yeast nucleus, molecular weight, 254 peptidoglycan biosynthesis and, 23 DNA polymerases in yeast nucleus, 27 1 Diglycosyldiglycerides DNA synthesis in Aspergillus niger, biosynthesis, 109 inhibition by hydroxyurea, 293 fungal, biosynthesis, 107

386

SUBJECT INDEX

DNA synthesis-cont. initiation in Escherichia coli, cell division and, 260 polymerase activity and, 273 in yeast nucleus, bud emergence and, 248 initiation, 260-265 DNA transcription, glycerol and, 230 Dormancy fungal spores, fatty acids and, 116 spores, ionic composition and, 29 Downy mildew, lipids and, 138 Dunaliella spp. glycerol dehydrogenase in, 234 glycerol, equilibrium status in, 23 I production, 226, 228, 236 regulation in, 225 halofelicity, 2 16 nicotinamide nucleotide ratios, 234 solute concentration in, 22 1 water activity tolerance, 183 Dunaliella parva chloroplast, surface area, 222 glycerol accumulation, 220 halofelicity, 2 16 nitrate reductase, glycerol and, 196 Dunaliella salina chloroplast envelope, 222 halofelicity, 2 16, 2 17 Dunaliella tertiolecta enzyme preparations, salt sensitivity, 220 glucose 6-phosphate dehydrogenase, effect ofglycerol on, 196 glycerol and glucose 6-phosphate dehydrogenase, 185 growth-rate measurements, 2 18 halofelicity, 2 16 salt tolerance, 2 1 7 Dunaliella uiridis enzyme preparations, salt sensitivity, 220 glucose 6-phosphate dehydrogenase, effect of glycerol on, 196 glycerol and glucose 6-phosphate dehydrogenase, 185 growth-rate measurements, 218 halofelicity, 2 16 salt tolerance, 2 1 7

E Ecology, Hyphomicrobium spp., 329-332 Ectotrophic mycorrhiza, lipid physiology in, 153 Elodea densa, nicotinamide nucleotide concentrations in, 234 Encysted zoospores, germination, lipids and, 123 Endogone, lipids and, 139 Endomycopsis, ambrosial fungi and, 156 Endomycopsis jbuligera, surface structure, 224 Endomycopsis selenospora, phosphoglycerides in, 67 Endoplasmic reticulum, lipids as membrane components, 48 Environment, Hyhomicrobium spp. morphology and, 323, 326-329 Enzyme activity dielectric constant and, 19 1 effect ofwater activity, 183 intracellular polyols and, 186 polyolsand 189, 190 Enzymes bacterial spores, 25 resistance, 2 DNA, in yeast nucleus, 27 1-282 glycerol and, 184 methanol oxidation by, 333 polyols and, 188 Enzyme systems, phospholipids and, 7 2 Epidermophyton spp., lipids, 16 1 Epidermophytonjloccosum, sterols, 162 Ergosterol in cell membranes, 60 in dermatophytes, 162 in fungi, 121, 150 Ergot oil in Clauiceps purpurea, 144 Erwinia carotiuora, phospholipases in, 134 Erwinza solani, phospholipases in, 134 Erysiphe cichoracearum conidia, lipids, 14 1 lipids and, 138 spores, surface lipids, 122 Erysiphe graminis lipids, 138 reserves, 128 spores, surface lipids, 122

SUBJECT INDEX

Elysiphe graminis hordei, spores, germination, lipids, 123 Erythritol apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 enzyme activity and, 189, 190 Escherichia coli expression of yeast genes in, 285, 286 gene expression, solute concentration and, 230 initiation of DNA synthesis in, cell division and, 260 reproduction, 320 Ethanol formation by yeasts, 233 in Hyphomicrobium sp. nutrition, 314 Ethanol dehydrogenase, 349 Ethylene glycol apparent inhibitor constants of' yeast isocitrate dehydrogenase, 193 dielectric constant, enzyme activity and, 191, 192 effect on collagen, 196 enzyme activity and, 189, 190 enzyme inactivation.and, 194, 195 enzymological role, 188 Euascomycetes, phosphoglycerides in, 67 Eukaryotes, compatible solutes and extreme water stress in, 18 1-242 E umycotina aggregation, glycosphingo lipids and, 100 Euromyces phaseoli, uredospores, germination, lipids and, 129 Exosporia bacterial spores, 2 chemical composition, 4-7 morphology and, 3 , 4 function, 6 morphology, 2 protein, 12 Expression genes in yeast nucleus, 282-285 of yeast genes in Escherichia coli, 285, 286

F Fat bodies, terminology, 54 Fatty acids

387

determatophytes and, 163 in fungi, 131 biosynthesis, 100 morphogenesis and, 1 16, 1 1 7 phosphoglycerides, 68 in mycoparasites, 152 in niycorrhizal fungi, 153 in pathogenic fungi, 16 1 Ferritin, 58 Fish, glycerol and, 197 Flavobacter spp., methane oxidizers, 330 Flavonoids, photomorphogenesis, 1 19 Fomes annosus fatty acids in, 153 insects and, 157 Food reserves intracellular polyols and, 186 polyols as, 18 7 Formaldehyde fixation, robulose-monophosphate cycle, 338 oxidation by Hyphomicrobium spp., 334 Formaldehyde dehydrogenase, 335 Formates carbon assimilation by Hyphomicrobium spp., 344 Hyphomicrobium spp. growth and, 313, 3 14

Free bases in fungi, 80, 8 1 membranes, 85-90 Fructose apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 enzyme activity and, 189 Fucosyltrigalactosylglucuronosylinositolphosphorylceramide in fungi, 88 Fungal cells, lipids, location, 48-58 Fungal growth, polyene antibiotics and, 61

Fungal membranes general considerations, 59-65 lipids and, 59-100 Fungi glycolipids, 76, 7 7 insect associations, lipids and, 155-1 60 lipids, biosynthesis, 100-1 13 physiology, 47-180

SUBJECT INDEX

388

Fungi-cont. morphogenesis, lipids in, 113-139 water activity tolerance, 183 Fungisterol in rust urediospores, 142 Fungus-host relationship, lipids and, 140-164 Fusarium spp. asexual spores, lipid content, 122 fatty acids in, 1 1 7 Fusarium culmorum, lipids in, 5 1

Fusarium lini ceramides in, 80 sphingolipids in, 88 Fusarium oxysporum lipids in, 5 1 spores, nonanoic acid and, 116 Fusarium solani neutral lipids, 15 7 phospholipids, 157 spores, germination, lipids, 123 sterols in, 156

G P-Galactosidase, induction by sucrose, 230 Galactosylceramide in fungi, 89 n-Galactbsylglycerol as osmoregulator, 187,223 Galactosylmannosylinositolp hosphorylceramide in fungi, 8 7 Gametangia, lipids in, 50 Genes DNA replication and, 268 expression, solute concentration and, 30 yeast, chromosomes, localization, 256260 expression in Escherichia coli, 285, 286 nucleus, expression, 282-285 Genetic map, Saccharomyces cereuisiae, 250 Geotrichum candidum lipases in, 134 lipids, 50 Germ cell wall bacteria, 1 spores, chemical structure, 18-22 morphology, 18 peptidoglycan, 3 1

Germination Clauiceps purpurea, lipids and, 144 fungal, lipids and, 114, 123 lipid reserves and, 129 spores, fatty acids and, 116 lipases and, 133 Gibberella baccata, cyclic depsipeptides in, 144 Giemsa in yeast nuclei staining, 296 Glomerella cingulata, triglycerides in, 130 Glomus mossei lipids and, 139 phosphoglycerides in, 66 Glucitol-See Sorbitol Glucosamine in Bacillus spp. spores, 6 Glucose accumulation, in Saccharomyces cereuisiae, 2 10 in Saccharomyces rouxii, 2 10 apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 consumption by yeasts, 209 dielectric constant, enzyme activity and, 191, 192 in Dunaliella tertiolecta, 220 enzyme activity and, 189 polyol formation by yeasts and, 199 respiratory capacity of yeasts and, 228 Saccharomyces cereuisiae consumption, 205 Saccharomyces rouxii consumption, 205 in yeast growth in salt solutions, 203 Glucose 6 -phosphate dehydrogenase effect of glycerol on, 196 from halophilic algae, 2 19 Glucosylceramide in fungi, 88 Glucosylgalactosylmannosylinositolphosphorylceramide in fungi, 87 Glutarnic acid as osmoregulator, 187 y-Glutamyl diaminopimelate peptidase in Bacillus species sporulation, 24 Glycerol active transport by Saccharomyces rouxii, 209 apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 dielectric sonstant, enzyme activity and, 191, 192 distribution in non-tolerant yeasts, 222

389

SUBJECT INDEX

DNA transcription and, 230 in Dunaliella spp., 22 1 accumulation, 220 production, 236 effect on collagen, 196 on nitrate reductase, 196 enzyme activity and, 189, 190 enzyme inactivation and, 194, 195 enzymological role, 188 extreme water stress in eukaryotes and, 184 as food reserve for yeasts, 204 formation by yeasts, 233 metabolic pathways, 224 as osmoregulator, 187 permeability of yeasts, 208 production, by Dunaliella spp., 226, 227,228 by Saaharomyces cereuisiae, 206, 226 by Saccharomyces rouxii, 200 regulation, 225 by xerotolerant yeasts, aeration and, 227,228 by yeasts, sodium chloride concentration and, 206 as regulatory agent, 229 retention factor in yeast, 202 thermal stability of collagen and, 2 14 Xeromyces bisporus accumulation, 2 15 yeast alcohol dehydrogenase and, 233 yeast isocitrate dehydrogenase inhibition by, 194 yeast surface structure and, 224 in Zygosaccharomyces nectarophilus, 199 Glycerol dehydrogenase in Dunaliella spp., 234 from halphilic algae, 2 19 a-Glycerophosphate, acylation by yeast extract, 105 Glycolipids fungal, 48, 74-84 biosynthesis, 107-109 membranes, 59 polyprenol-containing, in fungal membranes, 79-84 Glycosphosphosphingolipids in fungi, 92-95,98 biosynthesis, 1 12 cell surfaces, 60 Glycosylceramides in fungi, 90, 9 1

Glycosyldiglyceridesin fungi, 7 7 membranes, 7 8 in Gram-positive bacteria biosynthesis, 108 Glyoxylate-cycle enzymes in fungal morphogenesis, 135-139 Glyoxysomes, terminology, 5 7 Golgi apparatus from Schizosaccharomyces pombe, isolation, 65 Gram-positive bacteria glycosyldiglyceridebiosynthesis in, 108 membranes, 59 phosphoglycolipids in, 97 Graphium sp. neutral lipids, 157 phospholipids, 157 Great Salt Lake, Dunaliella spp. in, 216 Growth rate Hyphornicrobiurn X , hyphal length and, 326,327 Saccharomyces rouxii, polyol production, aeration and, 227, 228 yeast, DNA-dependent RNA polymerases and, 2 7 9 Growth regulating factors, fungal morphogenesis and, 114

-

H Halobacterium salinarium isocitrate dehydrogenase inhibition by salts, I88 NADP-specific isocitrate dehydrogenase, 194 Halophilic algae, 2 15-223 Halophilic bacteria, osmoregulators, 187 Hanseniaspora ualbyensis phospholipids, fatty-acid content, 68 sphingolipid biosynthesis in, 109 Hansenula spp. lipid reserves, 130 sphingolipids in, 131 Hansenula anomala, phosphoglycerides in, 61 Hansenula cqerri free bases in membranes, 85 glycosylceramides in, 9 1 sphingolipids in, 88, 131, 132 biosynthesis in, 109, 1 1 1

390

SUBJECT INDEX

Hansenula cc&erri--cont. sphingosine in, 131 tetra-acetylphytosphingosine in, 8 1 triacetylphosphingosine in, 8 1 Hansenula holstii, polyprenol-containing glycolipids in, 82 Hansenula wingei, agglutination factor, mannan in, 98 Heat resistance bacterial spores, 2 spores, calcium and, 3 1 coats and, 1 7 , 37 cortex, 36 dipicolinic acid and, 39 ionic composition and, 29,3 1 ionic content and, 38 water content and, 34,35 Hebeloma sarcophyllum, lipid metabolism in, 153 Helixpomatia, lytic enzymes in, 64 Hemiascomycetes, phosphoglycerides in, 66 Histones from Saccharomyces cereuisiae, aminoacid composition, 252 in yeast chromosomes, 244,25 1-253 Histoplasma spp., lipids, 161 Histoplasma capsulatum cell-wall lipids, 149 hyphal walls, lipids, 163 lipids, 160 Homobasidiomycetes, basidiospores, germination, lipids and, 130 Honey, fermentation, 197 Hormones, fungal morphogenesis and, 114 Host-parasite interactions, spherosomes in, 56 Humicola grisea var. thermoidea, phosphoglycerides, 65, 67 Hydroxy acid glycosides fungal, 74-78 biosynthesis, 107 3-Hydroxybutyrate in Hyphomicrobium sp. nutrition, 314 8-Hydroxyquinoline RNA synthesis inhibition by, 294 rRNA synthesis and, 278 Hydroxyurea, ribonucleotide reductase inhibition by, 293

Hylocoetus denestoides, ambrosial fungi and, 156 Hyphal filaments in Hyphomicrobium spp., function, 327 Hyphal length, Hyphomicrobium X, growth rate and, 326,327 Hypholomafasiculare, ceramides in, 80 Hyphomicrobium spp. biochemistry, 332-355 classification, 305-3 10 enrichment and isolation, 3 1 1-3 14 growth requirements, 308 new spp., 310 physiology and biochemistry, 303-332 Hyphomicrobium 10099, classification, 309 Hyphomicrobium B 5 2 2 growth rate, 3 18 nutrition, 3 1 7 synchronous culture, 32 1 Hyphomicrobium C, environment and growth, 324 Hyphomicrobium G, nutrition, 3 1 7 Hyphomicrobium X growth rate, 3 18 nutrition, 3 17 Hyphomicrobium indicum, classification, 309 Hyphomicrobium neptunium, classification, 308 Hyphomonas polymorpha, classification, 309 Hyphomicrobium uulgare, 303 isolation, 304 life cycle, 324 Hyphomicrobium vulgare N Q , nutrition, 314 Hyphomonas, classification, 305

I Influenza virions, membranes, 59 Inhibition of yeast nuclear functions, 293,294 Initiation DNA synthesis, in Eschrichia coli, cell division and, 260 in yeast nucleus, 260 Inositolphosphorylceramide biosynthesis in Aspergillus niger, 1 13 in fungi, 86, 94

39 1

SUBJECT INDEX

Insects fungi associated with, lipids and, 155160

glycerol and, 19 7 Intranuclear spindle, yeast, 248 Ion exchange, phospholipids and, 75 Ionic composition, spores, 29-32 Ionic content, spores, heat resistance and, 38 Isobutanol in rnycorrhizal fungi, 153 Isobutyric acid in mycorrhizal fungi, 153 Isocitrate dehydrogenase apparent inhibitor constants with polyols and sugars, 193 inhibition by sugars, 192 polyols and, 188 Isocitrate lyase in fungal rnorphogenesis, 135-139 in Hyphomicrobium sp., 34 2 28 -1sofucosterol in rust urediospores, 142 in Uromycesphaseoli, 149

J Jack-bean urease, effect of polyols on, 194

K Keratins, spore coats and, 15 Kloeckera apiculata, phospholipases in, 134

L Labelling in DNA estimation, 294, 295 Lactose, Saccharomyces rouxii permeability, 209

Leaf-cutting ants, fungus cultivation, 160 Lecanora rupicola, rocellic acid in, 154 Leccinum aurantiacum, phosphatidylcholine in, 68 Leccinum scabrum, phosphatidylcholine in, 68

Lemmonnaiera aquatica, cartotenoids, 120 Leptographium, insects and, 157

Leucosporidium frigtdum, phosphoglycerides in, 67 Lichen lipids and, 139 metabolism, 154 Life cycles in Hyphomicrobium sp., 319-326 Light, Hyphomicrobium sp. growth and, 3 19 Linoleic acid in rnycorrhizal fungi, 154 mushroom production and, 127 y-Linoleic acid, synthesis by mycoparasites, 152 Lipases in lipid metabolism in fungal rnorphogenesis, 133 Lipid bodies in Phycomyces spp., 58 terminology, 54, 56 Lipid inclusions, fungal, 49 Lipids bacterial spores, 26 in biotrophic fungi in axenic culture, 143. 144

fungal, biosynthesis, lo&] I3 location, 48-58 membranes and, 59-100 metabolism in morphogenesis, 133139

morphogenesis, 113-139 physiology, 47-1 80 reserves, in morphogenesis, 121-130 secondary metabolites, 130-1 33 fungus-host relationships and, 140165

yeast production, salt solutions, 207 Lipomyces lipoferus phospholipids, fatty-acid content, 68 triglycerides in, 130 Lipomyces starkeyi, phospholipids, fattyacid content, 69 Liposornes, terminology, 124 Lipoteichoic acid in cell walls, 98 in Gram-positive bacteria, 97 Lornofungin, RNA synthesis inhibition by, 294 Lycopene in Phagomyces blakesleeanus, 1 19 Lycoperdon bouista, glycosylcerarnides in, 90

Lysine uptake by Neurospora phosphatidylcholine and, 73

crassa,

392

SUBJECT INDEX

Lysosomes, spherosomes as, 56 Lytic enzymes, spore cortex and germ cell wall and, 22,23

.

Magnesium in spores, transport systems, 31 Malate synthetase in fungal morphogenesis, 135-139 Manganese Aspergillus parasiticw morphology and, 162 in spores, 29 transport systems, 3 1 Mannan in cell membranes, 98 synthesis, polyprenols in, 48 Mannan-protein, biosynthesis, 82 Mannitol enzyme activity and, 190 storage in Aspergillus clauatus, 187 yeast surface structures and, 224 4- 0 - p - D-Mannopyranosyl-D-erythritol in smut fungi, 132 Mannose, transport across plasma membranes, 84 Mannosyldi-inositoldiphosphorylceramide in fungi, 87,92, 93 Mannosylinositolphosphorylceramide in fungi, 87 Mannosylmono-inositolmonophosphorylceramide in fungi, 92 Marasmius scorodonius ceramides in, 80 glycosylceramides in, 90 Melampsora lini lipids and, 139 spores, germination, lipids, 123 sterols, 142 Membrane systems, fungal cells, 58 Membrane transport in fungi, phospholipids and, 73 Methane oxidizers, Hyphomicrobium sp. growth and, 330 Methanol apparent inhibitor constants of yeast isocitrate dehydrogenase, 193

dielectric constant, enzyme activity and, 191, 192 enzyme activity and, 189, 195 Hyphomicrobium sp. growth on, 3 12,314, 316 Methanol dehydrogenase formaldehyde oxidation by, 335 in HjFhomicrobium sp., 333,334 Methylamine, Hyphomicrobium sp. growth and, 313,314,318 Methylotrophs, electron transport chains, 350 Methylotrophy, restricted in Hyphomicrobium sp., 352-355 Microbodies, terminology, 57 Microciona prolifera, re-aggregation, glucuronic acid in, 98 Microspora adouinii, squalene and, 164 Microsporon, lipids, 161 Microsporum spp., sterols, 16 1 Milk-fat globule membranes, 59 Mitochondria DNA replication, nuclear control over, 27 1 water activity, 222 Mitochondria1 electron-transport chain, phospholipids and, 72 Mitosis in yeast, 244 nuclear morphology and, 247 Molecular structure, bacterial spore, 145 Molecular weight polyols, enzyme activity and, 192 sugars, enzyme activity and, 192 Moniliaferruginea, insects and, 157 Moniliella tomentoja, phosphofructokinase activity, 236 Monogalactosyldiacylglyceroldigalactosyldiacylglycerol, fungus host relationships and, 148 Monoglucosyloxyhexadecenoic acid in Aspergillus bisporus, 74 Monoglucosyloxyoctadecenoic acid in Aspergillus niger, 74, 75 in fungi, 78 Monoglucosyltrigalactosyfceramide in fungi, 89 Monotypha microspora, Piptocephalis virginiana, lipids and, 56

SUBJECT INDEX

Morp hogenesis fungal, lipids, 48, 113-139 lipid metabolism in, 133-139 lipid reserves in, 121-130 Morphology Hyphomicrobium sp., environment and, 326-329

yeast nucleus, 245-247 Moulds, water activity tolerance, 183 Mucorjavanicus, lipases in, 134 Mucor rouxii cell-wall lipids, 149 glycophosphosphingolipids in, 98 hyphal walls, lipids, 163 lipids in, 49, 50 spores, surface lipids, 122 Multiplication in Hyphomicrobium sp., 208 Muramic lactam in Bacillus cereus T biosynthesis, 24 in bacterial spore peptidoglycan, 20, 21

Muramyl- L-alanine amidase in Bacillus subtilis biochemistry, 24 Mushroom spores, lipid bodies in, 57 Mycoparasites, lipids and, 15 1 Mycoplasma, membranes, 59 Mycorrhiza, lipids and, 139 Mycorrhizal fungi, lipid metabolism and, 152, 153, 154

Mycotypha microspora, Piptocephalus virginiana, on, 152 Myxomycete spores, germination, lipids and, 123

393

Neurospora crassa ascospores, germination, lipids and, 130

carotenogenesis, 1 18 conidiation, isocitrate lyase and, 136 glycophosphosphingolipids in, 95 lysine uptake, phosphatidylcholine and, 73 mutants, glycolipids, 72 phospholipid synthesis and, 70 phosphoglycerides in, 67 phospholipases in, I34 phospholipid biosynthesis in, 104 plasma membranes, isolation, 65 polyglycosylceramidesin, 9 1 polyprenol-containing glycolipids, 82 sphingolipids in, 8 6 , 8 9 functional features, 96 spores, surface lipids, 122 Neurospora lanceolata lipids in, 5 1 morphogenesis, lipids and, 126 Neurospora tetrasperma ascospores, lipid reserves, 122 lipids in, 5 1 Neurosporene in fungi, 119 Nicotinamide nucleotide coenzymes, glycerol and, 23 1 Nicotinamide nucleotide ratios in Dunaliella spp., 234 Nitrate reductase, effect of glycerol on, 196

Nitrates, Hyphomicrobium sp. growth on, 313,318,331

N

Nitrosobacteriumformat- novae, 303 Nitrosoguanidine, DNA replication and, 268

NADP'-specific isocitrate dehydrogenase Halobacterium salinarium, 194 inhibition by glycerol, 188 polyhydric alcohols and, 189 Nectria galligena, spores, surface lipids, 122

Neurospora spp. conidia, glyoxylate-cycle enzymes in, 135

membrane systems, 58 morphogenesis, lipids and, 126 mutants with phospholipid synthesis defects, 113

Nonanal, Puccinia graminis urediospores and, 116 Nonanoic acid, Aspergillus niger spore germination and, 116 Nucleic acids in spore cores, 29 Nucleohistone fibres from Saccharmnyces cerevisiae sphaeroplasts, 253 Nucleolar fractions in yeast nuclei, isolation, 293 Nucleotide sugars in yeast mannan biosynthesis, 83 Nucleus water activity, 222

SUBJECT INDEX

394 N ucleus-cont

.

yeast, 243-302 isolation, 292,293 morphology, 245-247 Nutrition of Hyphomicrobium spp., 3 14319

0 Octadecenoic acid, epoxy-, in Uromyces phaseoli urediospores, 148 -, 2-hydroxyin Aspergillus niger, biosynthesis, 10 1 monoglucoside, in Aspergillus niger, 100 Oil droplets, spherosomes as precursors of, 57 Oleic acid in fungal phosphoglycerides, 69 mushroom production and, 127 in mycorrhizal fungi, 154 One-carbon compounds assimilation by Hyphomicrobium sp., 338-344 Hyphomicrobiurn sp. growth on, biochemistry, 332 oxidation, Hyphomicrobium sp. growth and, 333-338 Oogonium, fungal, lipids in, 50 Ooidia, germination, lipids and, 123 Oomycetes, phosphoglycerides in, 66 Oospora destructor, cyclic depsipeptides in, 144 Ophiostomapini, biotin in, 158 Ophiostoma ulnii, Ceratocystis sp. and, 158 Orcein in yeast nuclei staining, 296 Organelles lipids as membrane components, 48 water activity, 222 Osmiophilic globulues, terminology, 54 Osmoregulation, definition, 186 Osmoregulators, intracellular polyols and, 186 Osmotic pressure, terminology, 182 Oxidation of one-carbon compounds, Hyphomicrobium sp. growth and, 333339

P Palmitic acid in mycorrhizal fungi, 154 Palmitoleic acid in fungal phosphoglvcerides. 69 L

I

v

,

Palstomyces dermatitidis, lipids, 160 Paracoccidioides, hyphal walls, lipids, 163 Paracoccidioides brasiliensis cell-wall lipids, 149 hyphal walls, lipids, 163 Pathogenic fungi dimorphism, 162 lipid content, 160-162 Phaseolus graminis on wheat tissue, lipids and, 145 Phaseolus uulgaris infected with Thielauiopsis basicola, phosphatidase activity in, 148 Uromyces phaseoli on, fatty-acid composition and, 144 membrane lipids and, 146 Phialophora rechardsiae, dimorphism, 162 Phialopholra uerrucosa dimorphism, 162 lipids, 160 Phosphatidase in Phaseolus uulgaris infected with Thielauiopsis basicola, 148 Phosphatidic acid in fungi, 62 in rust infections of Tussilago sp., 146 Phosp hatidylcholine in cell membranes, 60 in fungi, 63,68 membranes, 65 fungus-host relationships and, 148 in leaf tissue infected with Puccinia poarum, 146 lysine uptake by Neurospora crassa and, 73 Phosphatidylethanolamine in fungi, 62,68 membranes, 65 fungus-host relationships and, 148 in leaf tissue infected with Puccinia poarum, 146 Phosphatidyglycerol in fungi, 62 membranes, 65 fungus-host relationships and, 148 Phosphatidylinositol in fungi, 68 biosynthesis, 104, 105 membranes, 65 in Neurospora crassa, 96

SUBJECT INDEX

Phosp hatidylmonomethylethanolamine in fungi, 62 Phosphatidylmyo-inositol in fungi, 63 Phosphatidylserine in fungi, 62 membranes, 65 in Pulhlaria pullulans, 68 in rust infections of Tussilago sp., 146 Phosphoenolpyruvate carboxylase activity in Pseudomonas MA, 346 Phosphoglycerate inutase in Hyphomicrobium sp., 342,343,344 Phosphoglycerides in fungi, 66, 67 membranes, 48, 65 physiological aspects, 69-74 in Gram-positive bacteria, 97 Phospholipases in fungal morphogenesis, 134, 135 Phospholipids in Bacillus cereus T exosporium, 5 bacterial spores, 26 fungal, 62, 63 biosynthesis, 102-1 0 7 membranes, 59,65-69 unsaturation, host-fungus relationships and, 148 Phosphorus, spore coats, 14 Phosphosphingolipids in Fungi, 92-95 Phosphotransacetylase, 349 Pho tomorphogenesis fungal, 12 1 carotenoids and, 11 7 Photosynthesis, dihydroxyactone phosphate and, 226 Phycomyces spp. lipid bodies in, 58 sporangiophore, 49 photomorphogenesis, 1 18 spores, dormancy-breaking and germination, fatty acids and, 116 Phycomyces blakesleeanus carotene biosynthesis, 119 free bases in, 80 lipid bodies in, 58 lipids in, 5 1 phosphoglycerides in, 66 phospholipids, Fatty-acid content, 68 sphingolipids in, 88 triglycerides in, 130

395

Phycomycetes, phosphoglycerides in, 65 Phymatotrichum omniuorum +sexual spores, lipid content, 122 sterol growth factors, 12 1 Physarum polycephalum, pho tomorpho genesis, 1 18 Phytophthora spp., sterol growth factors, 120,121 Phytophthora capsici, lipids in, 50 Phytophthora cinnamomi fatty acids in, 153 lipids, 50 Phytophthora erythroseptica sporangia, lipids in, 124 zoospores, glyoxylate-cycle enzymes in, 135 Phytophthora infestans lipids in, 50 sporulation, sterols and, 151 Phytophthora palmiuora lipids in, 5 1 spores, germination, lipids, 123 zoospores, lipids in, 124 Phytophthora parasitica lipids in, 5 1 phosphoglycerides in, 66 zoospores, lipid reserve, 122, 124 Phytosphingolipids-See Glycophosphosphingolipids Phytosphingosine acetylation, 1 1 1 C18, in Candida intermedia, 85 Pedomicrobium, classification, 305, 309 Penicillium altrouenetum spores, germination, lipids, 123, 129 lipids, 143 penicillium baanense, polyketides in, 154 Penicillium expansum pentose-phosphate cycle, 236 spores, surface lipids, 122 Penicilliumfrequentens, fatty acids in, 1 1 7 Penicillium notatum, ceramides in, 80 Penicillium oxalicum, conidia, glyoxylatecycle enzymes in, 135 Pep tidoglycan Bacillus subtilis spores, 19 . in bacterial spores, 20 biosynthesis, 23-25

SUBJECT INDEX

396

Pep tidoglycan-cont. spore cortex, 39 spore walls, structure, 20 Perithecia, lipids in, 5 1 Permeability appendages and, 6 cell membranes, phospholipid unsaturation and, 148 exosporia and, 6 . glycerol, in yeasts, 208 spore coats and, 1 7 of yeasts, 207, 21 1, 225 Pestalotia rhododendri, fatty acids and, 116

Petite mutant alcohol dehydrogenase activity, 234 Saccharomyces rouxii, 229 Pezizella ericae, photomorphogenesis, 1 18 Pigmentation, fungal, 12 1 Pilobus sp. sporangiophores, flavonoids and, 1 19 photomorphogenesis, 118 Piptocephalus uirgtniana cell membrane, 15 1 lipid metabolism, 15 1 on Monotypha microspora, lipids and, 56 Pithomyces chartarum, spores, surface lipids, 122 Pityrosporium orbiculare, lipids, 160 Plant pathogens, lipid metabolism of biotrophic fungi and, 144 Plant tissues, fungal associations, 140155

Plaque duplication in yeast nucleus, 247 Plasmalemma, lipids as membrane components, 48 Plasma membranes, sphingolipids in, 96 Plasmodiophora sp. release from host tissue, 57 sterols, 162 Plasmodiophora brassicae lipids and, 138 in cabbage hypocotyls infected with, 144

spores, sterols in, 15 1 Platypus cylindrus, ambrosial fungi and, 156

Pleomorphism in Hyphomicrobium sp., 3 19-326

Poa spp., rust-infected, sterols, 149

Poa pratensis, Puccinia poalum on, lipids and, 140 Poly(A)polymerases in yeast nucleus, 280-282

Polyene, antibiotics, fungal growth and, 61

Polyethylene glycol effect on Saccharomyces rouxiz, 224 polyol formation by yeasts and, 199 yeast isocitrate dehydrogenase inhibition by, 188 Polyglycosylceramide in fungi, 9 1, 92 Polyols enzymologicalrole, 188-197 extreme water stress in eukarotes and, 184- 19 7 as osmoregulators, 223 production by xerotolerant yeasts, 198 transport mechanisms in yeast, 208 xerotolerant yeast production, aerationand, 227,228 in yeast growth in salt solutions, 203 Polyphospho-inositides proton transport across yeast membranes and, 73 in yeasts, biosynthesis, 105 Polypoms pinocola, glycosylceramides in, 90

Polyprenols in fungal membrane glycolipids, 79 in mannan synthesis, 48 in yeast mannan biosynthesis, 83 Polyprenolphosphate-mannose, fungal, biosynthesis, 107 Polyporus uesicolor, phosphoglycerides in, 67

Potassium as osmoregulators, 187 in spores, transport systems, 3 1 Potassium chloride NADP'-specific isocitrate dehydrogenase inhibition by, 194 as osmoregulators, 18 7 yeast surface structures and, 224 Powdery mildew, lipids and, 138 Proline as osmoregulator, 187, 223 Propanediol, enzyme inactivation and, 194

Propane- 1,2-diol, effect on collagen, 196 Propane- 1,3-diol, effect on collagen, 196

397

SUBJECT INDEX

Prosthecomicrobium sp., prosthecae in, 329 Proteins in bacterial spore cores, 26 in cell membranes, 64 crystals, in bacterial spores, 2 in spores, coats, 10, 29 heat resistance and, 34 Protein synthesis, DNA synthesis initiation in yeast nucleus and, 264 Protolichesterinic acid in Cetraria islandica, 154 Protons, efflux, 232 Proton transport across yeast membranes, polyphospho-inositides and, 13

Pseudomonas aeruginosa, glycolipid biosynthesis in, 107 Pseudomonas AM I formate dehydrogenase in, 335 methanol dehydrogenase in, 334 Pseudomom diminuta, glycolipids in membranes of, 7 1 Pseudmonm extorqwm, & value, 33 1 Pseudomonas fluorescens, membranes, glycolipids in, 7 1 Pseudomonus MA, phosphoenolpyruvate carboxylase in, 346 Pseudomom RJ I, formaldehyde dehydrogenase in, 335 Puccinia spp., spores, lipids, 142 Puccinia graminis lipases in, 133 lipids in, 5 1 spores, germination, lipids, 123 sterols, 142 urediospores, lipase preparations from, 142 nonanal and, 116 wheat leaves infected with, lipids and, 144

Puccinia helianthi, lipids and, 139 Puccinia poarum lipids in, 51, 139 metabolism, 147 morphogenesis, lipids and, 125 on Tussulago fafara, lipids and, 140, 145

on Tussilago sp., infected by, sterols and, 149

membrane lipids and, 146

Pucciniapratenris, lipids and, 145 Puccinia strii)om's spores, surface lipids, 122 urediospores, lipids, 143 Pullularia pulldam pathogenicity, 164 phospholipids in, 68 sugars in, 7 7 Purity of Hyphomicrobium spp. cultures, 313

Pyronema omphalodes, photomorphogenesis, 1 1 8 Pyruvate dehydrogenase in Hyphomicrobium sp., 354 Pythium spp. hyphal tip region, 49,52, 57 sterol growth factors, 120 Pythium ultimum lipids in, 50 phosphoglycerides in, 66

R Radiation, bacterial spores resistance, 2 Radiation resistance, spore coats and, 1 7 Ramalina muciyormis, Trebouxia sp. in, lipid metabolism, 154 Ramlina munyomis, lipids and, 139 Rangiferic acid, 154 Red blood cells, membranes, 59 Refractive index, spores, 32 Refractivity, spore coats and, 37 Renaturation of DNA from Sacchromyces carlsbergenris nuclei, 25 1 Replication of DNA in yeast nucleus, 266-27 1 mitochondria1 DNA, nuclear control over, 27 1 Reduction in Hyphomimobium species, 320 Reproductive structures, fungal, lipids in, 49

Respiratory capacity of yeasts, polyols and, 228 Rhamnose, apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 L- RhamnosyU 1+ 3)-L-rhamnosylhydroxy-

decanoylhydroxydecanoate, fungal biosynthesis, 107

398

SUBJECT INDEX

Rhizina undulata, fatty acids in, 153 Rhizoctonia solani, phospholipases in, 134 Rhizophydiurn rnacrosporum, Septosperma sp. on, lipid metabolism and, 152 Rhizophlyetis rosea, carotene production, 119 Rhizopus arrhizus lipases in, 134 lipids in, 5 1 spores, germination, lipids, 123, 129 sporulation, sterol growth factors and, 121 Rhizopus sexualis, lipids in, 5 1 Rhizopus stolonger, spores, surface lipids, 122 Rhodomicrobium Rm5, DNA in, 323 Rhodotorula spp. lipid reserves, 130 ustilagic acids in, 132 Rhodotorula graminis sugar alcohols in, 7 7 ustilagic acids in, 132 Ribitol apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 enzyme activity and, 189, 190 storage in Aspergillus clavatus, 187 Ribonucleotide reductase, inhibition by hydroxyurea, 293 Ribose, enzyme activity and, 189 Ribosomal cistrons Saccharomyces cerevisiae, replication, 268 in yeast nucleus, localization, 258 Ribosomes in bacterial spores, 25 Ribulose diphosphate cycle, carbon dioxide fixation, 338 Ribulose monophosphate cycle, formaldehyde fixation, 338 Ricinoleic acid in Clavicepspurpurea, 144 Rzcinus sp., spherosomes, lipids in, 56 RNA polymerases yeast, 245 nucleus, DNA-dependent, 273-280 RNA synthesis Hyphomicrobium spp., 323 inhibitors, 294 mRNA from yeast, 284 rRNA, gene coding for, 25 7 rRNA synthesis, RNA polymerase and, 278

tRNA bacterial spores, 25 gene coding for, 257 Rocellic acid, 154 Rust fungi, sterols in, 151 Rust urediospores fatty-acid effect of, 1 16 sterols, 142 Rusts, lipids and, 138

S

Saccharomyces carlsbergensis DNA from nuclei, renaturation, 25 1 inositol-deficient cells, phosphatidylserine in, 7 1 isolation of nucleolar fraction, 293 phospholipases in, 134 phospholipids, fatty-acid content, 68 Saccharomyces cerevisiae anhydroceramide in, 80 arabitol production and, 204 ascospores, germination, lipids and, 130 cell-division cycle, model, 262, 288 cell volume and, 2 10 ceramide phosphate in, 8 1 dolichols in, 82 fatty acids in, 1 17 free bases in, 802 genetic map, 250 glucose, accumulation in, 2 10 consumption, 205, 209 glycerol, distribution in, 222 production, 206, 223, 226 sodium chloride concentration and, 206 regulation in, 225 retention factor in, 202 glycophosphosphingolipids in, 92, 93, 94,95 glycosylceramidesin, 90 glyoxylate-cycle enzymes in, 136 growth in salt solutions, polyol concentrations, 203 lipid droplets in, 57 lipids in, 50, 5 1 production, salt solutions and, 207 morphogenesis, lipids and, 125

SUBJECT INDEX

399

NADP-specific isocitrate dehydrogrowth and, 205 genase, 185 production,. sodium chloride connucleus, 243-303 centration and, 206 pentose-phosphatepathway, 235 regulation in, 225 phosphoglycerides in, 66 NADP’-specific isocitrate dehydromembranes of, 69 genase, 185 phospholipases in, 134 inhibition by glycerol, 188 phospholipids in, 68 polycyclic alcohols and, 189 phospholipids, biosynthesis in, 103, non-metabolic regulatory mechanism, 104 224 fatty acid contents, 68 polyols, active transport in, 209 polyols, permeability in, 208 dehydrogenases in, 226 production, 201 distribution in, 222 promitochondria, 65 production, 201 respiratory capacity, 228 aeration and, 227,228 glucose and, 229 transport mechanism in, 208 . . respiratory-deficient, phospholipids respiratory capacity, 228 and, 73 glucose and, 229 salt accumulation, 2 12 mitochondria, phospholipids and, 73 in salt solutions, 200 in salt solution, 200 polyol concentrations, 203 sexual hormones and, 1 16 trehalose production, 207 spherosomes, 57 water activity tolerance, 183 lipids in, 56 xerofekity, 2 13 sphingolipids in, 86, 87, 89 as xerotolerant yeast. , , 198 biosynthesis in, 112 Saccharomyces telluris, phosphoglycerides functional features, 96 in, 66 sugars in, 7 7 Salt relations, algae, physiology and, 219 surface structure, 224 Salt solutions trehalose production, water stress and, lipid production by yeast and, 207 20 7 polyols, concentration in yeast growth xerofelicity, 2 13 in, 203 Saccharomyces cerevisiae var. ellipsoideus, production by yeasts in, 20 1 Saccharomyces rouxii growth in, 200 “sodium yeast” colonies, 2 12 Salts, accumulation by yeasts, 212 Saccharomyces fragilis, phosphoglycerides Saprolegniaferax in, 66 glycophosphosphingolipids in, 98 Saccharmyces marrianus, phosphoglycerlipids in, 50 ides in, 66 Saccharomyces pastorianus, phosphoglycer- Schizophyllum commune fruiting, isocitrate lyase and, 137 ides in, 66 germination, lipids and, 129 Saccharomyces rouxii pathogenicity, 164 arabitol, and glycerol retention factors phosphoglycerides in, 67 in, 202 spores, germination, lipids, 123 as compatible solute, 200 Schizosaccharomyces pombe distribution in, 222 cell volume and, 2 10 production, 235 membranes, isolation, 65 cell volume and, 210 phosphoglycerides in, 66 glucose accumulation in, 2 10 phospholipid biosynthesis in, 104 glucose consumption, 205, 209 glycerol, consumption, 204 surface structure, 224

SUBJECT INDEX

400

Sclerotia, lipids in, 5 1 Sclerotinea sclerotiorum, phosphatidase production, 148 Sclerotium rolfsii, phospholipases in, 134 Scolytid beetles ambrosial fungi and, 155, 156 fungi and, 155 Septosperma sp. on Rhizophydium macrosporum, lipid metabolism and, 152 Serine pathway in Hyphomicrobium sp., regulation sites, 344-349 Serine transhydroxymethylase, 346 Sex hormones, fungal, 115, 116 Sheathing, lipid physiology in, 153 Sinapis alba, sterols in, 150 Sirenin, 115 Sirex sp., wood-rotting fungi and, 158 Sirex cyaneus, wood-rotting fungi and, 158 Sitodrepapanicea, fungal symbionts, 159 p-Sitosterol in fungi, 120 in Plasmodiophora brassica spores, 151 in rust urediospores, 142 in Uromycesphaseoli, 149 Slime moulds, photomorphogenesis, 118 Smut spores, germination, lipids and, 123 Smuts, lipids and, 139 Sodium acetate in Hyphomicrobium sp. nutrition, 3 14 Sodium chloride concentration, yeast growth, glycerol production and, 206,207 isocitrate dehydrogenase and, 188 NAD P - specific isocitrate dehydrogenase inhibition by, 194 Saccharomyces spp. morphology and, 224 Solute concentration in Dunaliella sp., 221 Sophorosides fungal, 78, 132 biosynthesis, 107 Sorbitol apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 enzyme activity and, 189, 190 Spermatozoa, glycerol and, 197 Sphaeroplasts, isolation, 64 Sphaerotheca humuli, spores, behenic acid in, 141 +

Sphaerotheca macularis spores, germination, lipids, 123 surface lipids, 122 Sphaerotheca mors - uvae lipids and, 51, 138 morphogenesis, lipids in, 125 Spherosomes developmental sequence, 57 physiology, 48 terminology, 55, 56 Sphingolipidoses, 96 Sphingolipids in fungi, 86-89, 131 fungal, biosynthesis, 109-1 13 membranes, 84-100 functional features, 96-100 Sphingomyelin biosynthesis, 107 in cell membranes, 60 in fungi, 63 Sphingosine acetylation, 111 in fungi, 131 Spinacea oleracea, nicotinamide nucleotide concentrations in, 234 Spindle plaques, haploid strain of Saccharomyces cerevisiae, electron micrographs, 245-241 Spirillum sp., growth on methanol, 33 1 Sporangiospores germination, lipids and, 123 lipids in, 5 1 Spore coats, 1 biosynthesis, 16, 17 chemical composition, 8-16 function, 17, 18 heat resistance and, 37 morphology, 2, 7, 8 phosphorus, 3 1 structure, 8-16 Spore cores dehydration, mechanism, 35-39 low molecular-weight compounds, 2729 macromolecular composition, 25-2 7 water content and, 32-35 Spore cortex heat resistance and, 36 peptidoglycan, 3 1,39

SUBJECT INDEX

Spores fungal, lipids in, 48 ionic composition, 29-32 molecular structure, 1 4 5 morphology, 2 Sporulation fungal, lipids in, 114, 12 1 glyoxylate-cycleenzymes and, 136 Squalene, determatophytes and, 163 Staining, yeast nuclei, 295, 296 Stearic acid in mycorrhizal fungi, 154 Stemphylium solani, sterol growth factors, 121

Stereochemistry, sugars, enzyme activity and, 192 Stereum spp., Xyphidia prolongata and, 158 Sterol glucosides, fungal, biosynthesis, 107 Sterol glycosides, fungal, biosynthesis,

40 1

inhibition by, 194 yeast surface structure and, 224 Sugar alcohols in fungi, 7 7 membranes, 78, 79 Sugar-beet rust, lipids and, 138 Sugars acetylated, fungal, 78, 79 in fungi, 7 7 molecular weight, enzyme activity and, 192

Sulpholactic acid in spore cores, 28 Synchytrium sp., zoospores, lipids in, 124 Synchytrium endobioticum lipids and, 138 spores, lipids in, 124

T

109

Sterol growth factors, fungal, 120, 12 1 Sterols in ambrosial fungi, scolytid beetles and, 157 anobiid beetle larvae and, 155 in cell membranes, 60 dermatophytes, 162 fungal, biosynthesis, 100 host-fungus relationships and, 149 in pathogenic fungi, 16 1 in rust fungi, 151 in Xanthoriaparietina, 155 S tigmastadienol in Cronartiumfusgorme aeciospores, 149 in rust urediospores, 142 in Uromycesphaseoli, 149 S tigmasten-3p-01 in rust urediospores, 142 in Uromycesphaseoli, 149 Stigmasterol in Plasmodiophora brassica spores, 15 1 in rust urediospores, 142 in Uromycesphaseoli, 149 Sucrose dielectric constant, enzyme activity and, 191,192 enzyme activity and, 189, 195 enzymologicalrole, 18 8 yeast isocitrate dehydrogenase and, 188

Taphrina sp., fungal symbionts, 159 Teliospores, lipids in, 5 1 Temperature Dunaliella tertiolecta salt relations and, 218

high solute concentration tolerance and, 213 water activity and, 2 14 Terpenoids, growth regulating, fungal morphogenesis and, 117-12 1 Tetra-ace!$ phytosphingosine in fungi, 131, 132

Tetrahydrofolate, one-carbon complex, 347

Thielaviopsis basicola Phaseolus vulgaris infected with, phosphatidase activity, 148 phospholipases in, 134 Thuringtensis berliner, spore coats, amino acids, 11 Tilletia spp., spores, surface lipids, 122 Tilletia caries, lipids, 5 1, 139 Tilletia controversa lipids and, 139 teliospores, lipid reserves, 122 Torulopsis spp., sophorosides in, 132 Torulopsis apicola cell volume and, 2 10 hydroxyacid glycosides in, 7 6 Torulopsis bovinu, phosphoglycerides in, 6 7

402

SUBJECT INDEX

Torulopsis globosa, cell volume and, 2 10 Torulopsis gropengensis, hydroxyacid glycosides in, 76 Torulopsis halonitratophila, halophilic, 2 12 Training for Dunaliella sp. salt tolerance, 2 17 for salt tolerance, 2 18 Transport kinetics in yeasts, 225 Transport mechanisms of polyols, yeast, 208 of yeasts, 207 Trasiquone, DNA synthesis inhibition by, 294 Trebouxia sp. in Ramalina macformis, lipid metabolism, 153 sterols, 154 Tehalose, production by Saccharomyces cerevisiae, 207 Tremax sp., wood-rotting fungi and, 158 Tricholoma sp., sporophores, lipids in, 127 Tricholoma nudum lipid reserves, 130 morphogenesis, lipids and, 127 phosphoglycerides in, 67 Trichophyton discoides, sterols, 16 1 Trichophytonmegnini, sterols, 16 1 Trichophyton mentagrophytes, fatty acids, 161 Trichophyton rubrum fatty acids, 16 1 phospholipids, 161 squalene and, 163 sterols, 161 Trichophyton violaceum, sterols, 16 1 Trichosporium tingens, scolytid beetles and, 156 Trigalactosyldimannosylinositolphosphorylceramide in fungi, 88 Triglycerides, fungal, 130 Trigonopsis uariabilis, phosphoglycerides in, 67 Trimethylamine aerobic degradation to dimethylamine, 336 Hyphomicrobium sp. growth and, 313,s 14 Trimethylamine dehydrogenase, 33 7 Trisporic acid, 115, 1 1 7 Trypodendron lineatum, blue-stain fungus, 157

Tryptophanase, induction by sucrose, 230 Tussilago sp. Puccinia poarum on, membrane lipids and, 146 rust-infected, sterols, 149 Tussilagof a f a r a , Puccinia poarum on, lipids and, 140, 145 Two-carbon compounds, Hyphomicrobium sp. growth on, biochemistry, 349350

U Undecylenic acid, fungistatic activity, 117 Urea, yeast surface structure and, 224 Urediospores germination, lipids and, 123 lipids in, 51 Urocerus sp., wood-rotting fungi and, 158 Urocerus gzgas, wood-rotting fungi and, 159 Uromyces appendiculatus, membrane systems, 58 Uromycesphaseoli lipids and, 139 on cowpea leaves, lipids and, 141 on Phaseolus vulgaris, fatty acids and, 145 membrane lipids and, 146 phosphoglycerides in, 67 spores, germination, lipids, 123 sterols, 142, 149 synthesis in rust-infected bean leaves and, 149 urediospores, lipids in, 148 Ustilagic acid in fungi, 76, 7 7 , 132 Ustilago hordei, lipids and, 139 Ustilago may& acylated disaccharides in, 78 axenic culture, lipids, 143 germination, lipids and, 130 spores, germination, lipids, 123 sugars in, 7 7 ustilagic acid in, 76, 132 Ustilago nuda axenic culture, lipids, 143 ustilagic acids in, 77, 132

SUBJECT INDEX

V Vegetative cell walls, fungal, lipids in, 50 Vegetative hyphae fine structure, 49 lipid inclusions, 49 Vegetative mycelium, lipid inclusions, 49 Verticillium alboatrum spores, germination, lipids, 123 surface lipids, 122 Vitamins, anobiid beetle larvae and, 155

w Walled structures, fungal membrane systems and, 58 Wart disease of potatoes, lipids and, 138 Water activity temperature and, 2 14 terminology, 182 Water content, spore cores, 32 Water potential, terminology, 182 Water stress in eukaryotic micro-organisms, 18 1242

Saccharomyces cereuisiae, trehalose production, 207 White rust, lipids and, 138 Wood-rotting fungi, lipids in, 158 Wood wasps, wood-rotting fungi and, 158

Xyletorus sp., fungal ergosterol as sex hormone for, 157 Xyletorusferrugineus Fusarium soluni from mycangia of, sterols in, 156 neutral lipids, 157 phospholipids, 157 Xylose apparent inhibitor constants of yeast isocitrate dehydrogenase, 193 enzyme activity and, 189 Xyphidia prolongata, wood-rotting fungi and. 158

Y Yeasts lipid production, salt solutions, 207 mannan in, 98 mannan-protein, biosynthesis, 82 nucleus, 243-302 osmoregulators, 187 polyphospho-inositides, biosynthesis, 105

salt accumulation, 2 12 transport mechanisms of polyols in, 208

water activity tolerance, 183 Yeast isocitrate dehydrogenase, forms, 194

Yeast mannan, biosynthesis, 83, 84 Yeast mitochondria, phospholipids and,

X Xantheoria parietina, sterols, 154 Xeromyces bisporus glycerol accumulation, 2 15 water activity tolerance, 183 Xerophilic yeasts, 212-215 Xerotolerance intermediate, 223 physiology, 197-223 Xerotolerant fungi, 2 15 Xerotolerant moulds, glycerol in, 187 Xerotolerant yeasts, 198-2 12 natural habitats, 197 polyol production, aeration and, 227, 228

403

72

Yeast plasma membrane, mannan in, 98 Yeast protoplasts, preparation, 64

Z Zoosporangia, lipids in, 50 Zoospores, lipids in, 5 1 Zygomycetes asexual spores, lipid content, 122 phosphoglycerides in, 66 Zygosaccharomyces nectarophilus polyol formation, 199 xerofehcity, 2 13 Zygospores, lipids in, 5 1

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